Bailey & Scott's Diagnostic Microbiology, 15th Edition (Complete PDF) [15 ed.] 0323681050, 9780323681056, 9780323681063

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Bailey & Scott’s

DIAGNOSTIC MICROBIOLOGY F i f te e n t h E d i t i o n

PATRICIA M. TILLE, PhD, MLS(ASCP), AHI(AMT), FACSc C h a i r o f M i c ro b i o l o g y A d v i s o r y Co m m i t t e e ; Ed i to r i n C h i e f I J B L S ; I n te r n a t i o n a l Fe d e r a t i o n o f ­B i o m e d i c a l L a b o r a to r y S c i e n ce G r a du a te P ro g r a m D i re c to r / Fa c u l t y M e d i c a l L a b o r a to r y S c i e n ce University of Cincinnati Cincinnati, Ohio

Elsevier 3251 Riverport Lane St. Louis, Missouri 63043

BAILEY & SCOTT’S DIAGNOSTIC MICROBIOLOGY, FIFTEENTH EDITION Copyright © 2022 by Elsevier, Inc. All rights reserved.

ISBN: 978-0-323-68105-6

No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein).

Notice Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds or experiments described herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made. To the fullest extent of the law, no responsibility is assumed by Elsevier, authors, e­ ditors or ­contributors for any injury and/or damage to persons or property as a matter of products liability, ­negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Previous editions copyrighted 2017, 2014, 2007, 2002, 1998, 1994, 1990, 1986, 1982, 1978, 1974, 1970, 1966, 1962. International Standard Book Number: 978-0-323-68105-6

Content Strategist: Heather Bays-Petrovic Director, Content Development: Laurie Gower Content Development Specialist: Betsy McCormac Publishing Services Manager: Catherine Jackson Health Content Management Specialist: Kristine Feeherty Design Direction: Bridget Hoette Printed in Canada Last digit is the print number: 9 8 7 6 5 4 3 2 1

Life is fleeting, but you must always remember friends, colleagues, and family. I will always thank my husband, David, and our children, Chrissy, Malissa, D.J., and Katie, along with their significant others. I would be remiss if I did not mention the seven little “smiles” that bring daily joy to our lives: Aedan, Milan Jr., Julia, Maja, Jayce, Riley, and Mila! Lastly, no endeavor such as this would continue to evolve from one edition to the next without the insightful comments and input from numerous professional users and students. Thank you for your dedication, hard work, and humor. This edition was created during a time of great unrest and challenges in the medical field, with the world facing the COVID-19 pandemic. I myself was struck with the virus and hospitalized for a period. I can say firsthand that the care and compassion of the health care workers was amazing, from physical, occupational, and respiratory therapy; nursing staff; phlebotomists; and laboratory professionals. They all came together to ensure the best care! This edition is dedicated to all the essential workers who keep the country going and the health care professionals who continue to save lives as we battle this formidable viral adversary! To all the lives lost, may you rest in peace and know that the knowledge developed during the pandemic continues to save more lives every day!

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Reviewers

Shari Batson, BSc, ART(Microbiology) Professor Health Sciences St. Lawrence College Kingston, Ontario, Canada

Jimmy L. Boyd, MS/MHS, MLS(ASCP)CM

Program Director/Department Chair (Tenured) Medical Laboratory Sciences Arkansas State University-Beebe Beebe, Arkansas

Lisa K. Cremeans, MMDS, MLS(ASCP)CM, SMCM, MBCM Assistant Professor Department of Allied Health Sciences Division of Clinical Laboratory Science The University of North Carolina at Chapel Hill Chapel Hill, North Carolina

Guyla Corbett Evans, PhD, MLS(ASCP)CMSCCM Clinical Assistant Professor Clinical Laboratory Science East Carolina University Greenville, North Carolina

Kathleen J. Fennema, BS, MT(ASCP)

Clinical Laboratory Scientist Infectious Diseases Diagnostic Laboratory (Mycology and Parasitology Section) University of Minnesota Medical Center (M Health) Minneapolis, Minnesota

Michele G. Harms, MS, MLS(ASCP) Program Director Medical Laboratory Science Program UPMC Chautauqua Hospital Jamestown, New York

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Janet Hudzicki, PhD, MLS(ASCP)CM, SM(ASCP)CM Associate Professor Department of Clinical Laboratory Sciences KU Medical Center The University of Kansas Kansas City, Kansas

Cynthia Kaufman, MS, MT(ASCP)SM

Assistant Clinical Professor Department of Pathology and Laboratory Medicine Indiana University School of Medicine Indianapolis, Indiana

Louise Millis, MS, MLS(ASCP)CM

MLS Certification Associate Professor of Biology and MLS Program Director Department of Biology St. Cloud State University St. Cloud, Minnesota

Mathumathi Rajavel, PhD

Associate Professor Medical Technology Program School of Computer, Mathematical and Natural Sciences Morgan State University Baltimore, Maryland

Katherine M. Steele, MPH, MLS(ASCP)CM Assistant Clinical Professor Pathology and Laboratory Medicine Indiana University School of Medicine Indianapolis, Indiana

Contributors

Hassan A. Aziz, PhD, MSc

Associate Dean for Academic, Faculty, and Student Affairs College of Health Professions Professor Clinical Laboratory Science The University of Tennessee Health Science Center Memphis, Tennessee Chapter 34: Legionella Chapter 77: Quality in the Clinical Microbiology ­Laboratory Chapter 78: Infection Control Chapter 79: Sentinel Laboratory Response to Bioterrorism

Erin Barger, BS MLS, MA Education

Medical Technologist Microbiology UC Health Cincinnati, Ohio Chapter 21: Pseudomonas, Burkholderia, and Similar Organisms Chapter 25: Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum

Janice Conway-Klaassen, PhD, MLS(ASCP)CM, SMCM, FACSc

Stephanie Jacobson, MS, MLS(ASCP)CM

MLS Upward Mobility Online Instructor Medical Laboratory Science South Dakota State University Brookings, South Dakota; Microbiologist Laboratory Monument Health Rapid City, South Dakota Chapter 15: Bacillus and Similar Organisms Chapter 16: Listeria, Corynebacterium, and Similar Organisms Chapter 17: Erysipelothrix, Lactobacillus, and Similar Organisms Chapter 47: Intestinal Protozoa Chapter 49: Protozoa From Other Body Sites Chapter 50: Intestinal Nematodes Chapter 51: Tissue Nematodes Chapter 52: Blood and Tissue Filarial Nematodes Chapter 53: Intestinal Cestodes Chapter 54: Tissue Cestodes Chapter 55: Intestinal Trematodes Chapter 56: Liver and Lung Trematodes Chapter 57: Blood Trematodes

Director Medical Laboratory Sciences University of Minnesota Minneapolis, Minnesota Chapter 46: Overview of the Methods and Strategies in Parasitology Chapter 48: Blood and Tissue Protozoa

James March Mistler, MS

April Harkins, PhD, MT(ASCP)

Associate Professor Biomedical Sciences University of New England Biddeford, Maine Chapter 44: Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma

Associate Professor and Department Chair Clinical Laboratory Science Marquette University Milwaukee, Wisconsin Chapter 60: Dematiaceous (Melanized) Molds Chapter 61: Atypical and Parafungal Agents

Program Director/Lecturer Department of Medical Laboratory Science University of Massachusetts Dartmouth North Dartmouth, Massachusetts Chapter 39: Neisseria and Moraxella catarrhalis

Meghan May, PhD, MS

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Contributors

Caterina Miraglia, DC, MLS(ASCP)CM Assistant Professor Medical Laboratory Science University of Massachusetts Dartmouth North Dartmouth, Massachusetts Chapter 45: The Spirochetes

Nicholas M. Moore, PhD

Assistant Professor Department of Medical Laboratory Science and Pathology Assistant Director Division of Clinical Microbiology Rush University Medical Center Chicago, Illinois Chapter 10: Principles of Antimicrobial Action and ­Resistance Chapter 11: Laboratory Methods and Strategies for ­Antimicrobial Susceptibility Testing Chapter 20: Acinetobacter, Stenotrophomonas, and Other Organisms Chapter 62: The Yeasts and Yeastlike Organisms Chapter 63: Antifungal Susceptibility Testing, Therapy, and Prevention

Rodney E. Rohde, PhD, MS, BS

Chair and Professor Clinical Laboratory Science Associate Dean for Research College of Health Professions Associate Director for Translational Health Research Initiative Texas State University San Marcos, Texas; Associate Adjunct Professor of Biology Department of Biology Austin Community College Austin, Texas Chapter 8: Nucleic Acid–Based Analytic Methods for ­Microbial Identification and Characterization Chapter 13: Staphylococcus, Micrococcus, and Similar Organisms Chapter 35: Brucella Chapter 37: Francisella Chapter 44: Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma Chapter 64: Overview of the Methods and Strategies in Virology Chapter 65: Viruses in Human Disease

Frank Scarano, PhD, MS, BA, AAS

Professor Medical Laboratory Science University of Massachusetts Dartmouth Dartmouth, Massachusetts Chapter 28: Eikenella corrodens and Similar Organisms Chapter 29: Pasteurella and Similar Organisms

Tim Southern, MS, PhD, D(ABMM)

Laboratory Director South Dakota Public Health Laboratory South Dakota Department of Health Pierre, South Dakota Chapter 12: Overview of Bacterial Identification Methods and Strategies Chapter 19: Enterobacterales Chapter 43: Obligate Intracellular and Nonculturable Bacterial Agents

Shannon Weigum, BA, MS, PhD

Associate Professor Department of Biology Materials Science, Engineering, and Commercialization Program Texas State University San Marcos, Texas Chapter 8: Nucleic Acid–Based Analytic Methods for ­Microbial Identification and Characterization Chapter 64: Overview of the Methods and Strategies in Virology Chapter 65: Viruses in Human Disease

Preface

This, the fifteenth edition of Bailey & Scott’s Diagnostic Microbiology, is the third edition that I have had the great pleasure to edit and author with some amazing colleagues. The dynamics of infectious disease trends, along with the technical developments available for diagnosing, treating, and controlling these diseases, continues to present major challenges in the laboratory and medical care. In meeting these challenges, the primary goal for the fifteenth edition is to provide an updated and reliable reference text for practicing clinical microbiologists and technologists, while also presenting this information in a format that supports the educational efforts of all those responsible for preparing others for a career in diagnostic microbiology. The text retains the traditional information needed to develop a solid, basic understanding of diagnostic microbiology while integrating the dynamic expansion of molecular diagnostics and advanced techniques such as matrix-assisted laser desorption time-of-flight mass spectrometry. We have kept the favorite features and made adjustments in response to important critical input from users of the text. The succinct presentation of each organism group’s key laboratory, clinical, epidemiologic, and therapeutic features in tables and figures has been kept and updated. Regarding content, the major changes reflect the changes that the discipline of diagnostic microbiology continues to experience. Also, although the grouping of organisms into sections according to key features (e.g., Gram reaction, catalase or oxidase reaction, growth on MacConkey agar) has remained, changes regarding the genera and species discussed in these sections have been made. These changes, along with changes in organism nomenclature, were made to accurately reflect the changes that have occurred, and continue to occur, in taxonomy. Also, throughout the text, the content has been enhanced with new photographs and artistic drawings. Finally, although some classic methods

for bacterial identification and characterization developed over the years (e.g., catalase, oxidase, Gram stain) still play a critical role in today’s laboratory, others have given way to commercial identification systems. We realize that in a textbook such as this, a balance is needed for practicing and teaching diagnostic microbiology; our selection of identification methods that received the most detailed attention may not always meet the needs of both groups. However, we have tried to be consistent in selecting those methods that reflect the most current and common practices of today’s clinical microbiology laboratories, along with those that p ­resent historical information required within an ­educational program. Finally, in terms of organization, the fifteenth edition is similar in many aspects to the fourteenth edition, but some changes have been made. Various instructor ancillaries, specifically geared for the fifteenth edition, are available on the Evolve website, including an expanded test bank, updated PowerPoints, a laboratory manual with answers, review questions with answer key, and an electronic image collection. Student resources include a laboratory manual, review questions, online case studies, and online procedures. We sincerely hope that clinical microbiology practitioners and educators find Bailey & Scott’s Diagnostic Microbiology, fifteenth edition, to be a worthy and useful tool to support their professional activities.

Acknowledgments I would like to acknowledge the help of my colleagues at Elsevier who guided me through this project: Kristine Feeherty, Health Content Management Specialist, and Betsy McCormac, Content Development Specialist. Patricia M. Tille

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Contents

Part I  Basic Medical Microbiology, 1 1 Microbial Taxonomy, 1 Classification, 2 Nomenclature, 3 Identification, 4 2 Bacterial Genetics, Metabolism, and Structure, 6 Bacterial Genetics, 6 Bacterial Metabolism, 17 Structure and Function of the Bacterial Cell, 21 3 Host-Microorganism Interactions, 26 The Encounter Between Host and Microorganism, 27 Microorganism Colonization of Host Surfaces, 29 Microorganism Entry, Invasion, and Dissemination, 32 Outcome and Prevention of Infectious Diseases, 41 

Part II  General Principles in Clinical Microbiology, 44 Section 1  Safety and Specimen Management, 44 4 Laboratory Safety, 44 Sterilization, Disinfection, and Decontamination, 45 Chemical Safety, 48 Fire Safety, 49 Electrical Safety, 50 Handling of Compressed Gases, 50 Biosafety, 50 Exposure Control Plan, 50 Employee Education and Orientation, 50 Disposal of Hazardous Waste, 51 Standard Precautions, 51 Laboratory Design and Engineering Controls, 52 Biological Safety Levels, 52 Mailing Biohazardous Materials, 56 5 Specimen Management, 58 General Concepts for Specimen Collection and Handling, 58 Specimen Preservation, 59 Specimen Work-Up, 74 Expediting Results Reporting: Computerization, 76  viii

Section 2  Approaches to Diagnosis of Infectious Diseases, 77 6 Role of Microscopy, 77 Bright-Field (Light) Microscopy, 78 Phase-Contrast Microscopy, 85 Fluorescent Microscopy, 86 Dark-Field Microscopy, 89 Digital Automated Microscopy, 90 Digital Holographic Microscopy, 90 7 Overview of Conventional Cultivation and Systems for Identification, 91 Organism Identification, 91 Principles of Bacterial Cultivation, 91 Bacterial Cultivation, 101 Principles of Identification, 105 Principles of Phenotypic Identification Schemes, 111 Commercial Identification Systems and Automation, 116 8 Nucleic Acid–Based Analytic Methods for Microbial Identification and Characterization, 119 Overview of Nucleic Acid–Based Methods, 120 Postamplification End-Point Analysis, 139 9 Overview of Immunochemical Methods Used for Organism Detection, 149 Features of the Immune Response, 149 Features of the Humoral Immune Response Useful in Diagnostic Testing, 151 Interpretation of Serologic Tests, 152 Production of Antibodies for Use in Laboratory Testing, 152 Immunoglobulin M Clinical Significance, 154 Separating Immunoglobulin M From Immunoglobulin G for Serologic Testing, 154 Principles of Immunochemical Methods Used for Organism Detection, 154 Summary, 163 

Contents

Section 3  Evaluation of Antimicrobial Activity, 165 10 Principles of Antimicrobial Action and Resistance, 165 Antimicrobial Action, 165 Mechanisms of Antibiotic Resistance, 174 11 Laboratory Methods and Strategies for Antimicrobial Susceptibility Testing, 182 Goal and Limitations, 182 Testing Methods, 183 Laboratory Strategies for Antimicrobial Susceptibility Testing, 200 Accuracy, 203 Communication, 206 

Part III  Bacteriology, 208 Section 1  Principles of Identification, 208

Antimicrobial Susceptibility Testing and Therapy, 296 Prevention, 297

16 Listeria, Corynebacterium, and Similar Organisms, 298 General Characteristics, 298 Epidemiology, 298 Pathogenesis and Spectrum of Disease, 300 Laboratory Diagnosis, 302 Antimicrobial Susceptibility Testing and Therapy, 309 Prevention, 309 Treatment, 309 

Section 5  Non-Branching, Catalase-Negative, Gram-Positive Bacilli, 314

12 Overview of Bacterial Identification Methods and Strategies, 208 Rationale for Approaching Organism Identification, 208 Future Trends of Organism Identification, 209 

17 Erysipelothrix, Lactobacillus, and Similar Organisms, 314 General Characteristics, 314 Epidemiology, 314 Pathogenesis and Spectrum of Disease, 314 Laboratory Diagnosis, 316 Prevention, 320 

Section 2  Catalase-Positive, Gram-Positive Cocci, 251

Section 6  Branching or Partially Acid-Fast, Gram-Positive Bacilli, 322

13 Staphylococcus, Micrococcus, and Similar Organisms, 251 General Characteristics, 252 Epidemiology, 252 Pathogenesis and Spectrum of Disease, 253 Laboratory Diagnosis, 255 Antimicrobial Susceptibility Testing and Therapy, 262 Prevention, 267 

18 Nocardia, Streptomyces, Rhodococcus, and Similar Organisms, 322 General Characteristics, 323 Epidemiology and Pathogenesis, 326 Laboratory Diagnosis, 327 Antimicrobial Susceptibility Testing and Therapy, 333 Prevention, 333 

Section 3  Catalase-Negative, Gram-Positive Cocci, 269

Section 7  Gram-Negative Bacilli and Coccobacilli (MacConkey-Positive, Oxidase-Negative), 335

14 Streptococcus, Enterococcus, and Similar Organisms, 269 General Characteristics, 270 Epidemiology, 270 Pathogenesis and Spectrum of Disease, 274 Laboratory Diagnosis, 276 Antimicrobial Susceptibility Testing and Therapy, 284 Prevention, 285 

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Section 4  Non-Branching, Catalase-Positive, Gram-Positive Bacilli, 287 15 Bacillus and Similar Organisms, 287 General Characteristics, 287 Laboratory Diagnosis, 291

 nterobacterales, 335 E General Characteristics, 336 Epidemiology, 336 Pathogenesis and Spectrum of Diseases, 336 Specific Organisms, 339 Laboratory Diagnosis, 345 Prevention, 361

20 Acinetobacter, Stenotrophomonas, and Other Organisms, 362 General Characteristics, 362 Epidemiology, 362 Pathogenesis and Spectrum of Disease, 363 Laboratory Diagnosis, 364 Cultivation, 364

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Contents

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Antimicrobial Resistance and Antimicrobial Susceptibility Testing, 367 Antimicrobial Therapy, 368 Prevention, 368

Section 8  Gram-Negative Bacilli and Coccobacilli (MacConkey-Positive, Oxidase-Positive), 370 21 Pseudomonas, Burkholderia, and Similar Organisms, 370 General Characteristics, 371 Epidemiology, 371 Pathogenesis and Spectrum of Disease, 372 Laboratory Diagnosis, 373 Antimicrobial Susceptibility Testing and Therapy, 381 Prevention, 381 22 Achromobacter, Rhizobium, Ochrobactrum, and Similar Organisms, 383 General Characteristics, 384 Epidemiology, 384 Pathogenesis and Spectrum of Disease, 384 Laboratory Diagnosis, 385 Antimicrobial Susceptibility Testing and Therapy, 389 Prevention, 389 23 Chryseobacterium, Sphingobacterium, and Similar Organisms, 392 General Characteristics, 392 Epidemiology, 392 Pathogenesis and Spectrum of Disease, 393 Laboratory Diagnosis, 394 Antimicrobial Susceptibility Testing and Therapy, 395 Prevention, 395 24 Alcaligenes, Comamonas, and Similar Organisms, 398 General Characteristics, 398 Epidemiology, 398 Pathogenesis and Spectrum of Disease, 398 Laboratory Diagnosis, 400 Antimicrobial Susceptibility Testing and Therapy, 402 Prevention, 403 25 Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum, 404 General Characteristics, 405 Epidemiology, 406 Pathogenesis and Spectrum of Disease, 406 Laboratory Diagnosis, 407 Antimicrobial Susceptibility Testing and Therapy, 411 Prevention, 411

Section 9  Gram-Negative Bacilli and Coccobacilli (MacConkey-Negative, Oxidase-Positive), 414 26 Sphingomonas and Similar Organisms, 414 General Considerations, 414 Epidemiology, Spectrum of Disease, and Antimicrobial Therapy, 414 Laboratory Diagnosis, 414 Prevention, 417 27 Moraxella and Neisseria spp., 419 General Characteristics, 419 Epidemiology, Spectrum of Disease, and Antimicrobial Therapy, 419 Laboratory Diagnosis, 420 Antimicrobial Susceptibility, 423 Prevention, 423 28 Eikenella corrodens and Similar Organisms, 425 General Characteristics, 425 Epidemiology, Spectrum of Disease, and Antimicrobial Therapy, 425 Laboratory Diagnosis, 426 Serodiagnosis, 428 Prevention, 428 29 Pasteurella and Similar Organisms, 430 General Characteristics and Taxonomy, 430 Epidemiology, Spectrum of Disease, and Antimicrobial Therapy, 430 Laboratory Diagnosis, 432 Serodiagnosis, 432 Serodiagnosis, 434 Prevention, 434 30 Actinobacillus, Kingella, Cardiobacterium, Capnocytophaga, and Similar Organisms, 435 General Characteristics, 436 Epidemiology, Pathogenesis, and Spectrum of ­Disease and Antimicrobial Therapy, 436 Laboratory Diagnosis, 437 Nucleic Acid Detection, 439 Serodiagnosis, 439 Prevention, 442

Section 10  Gram-Negative Bacilli and Coccobacilli (MacConkey-Negative, Oxidase-Variable), 443 31 Haemophilus, 443 General Characteristics, 443 Epidemiology, 443 Pathogenesis and Spectrum of Disease, 445 Laboratory Diagnosis, 445 Antimicrobial Susceptibility Testing and Therapy, 450 Prevention, 450

Contents

Section 11  Gram-Negative Bacilli That Are Optimally Recovered on Special Media, 452 32 Bartonella, 452 Bartonella, 452 Direct Detection Methods, 454 33 Campylobacter, Arcobacter, and Helicobacter, 457 Campylobacter and Arcobacter, 457 Helicobacter spp., 464 34 Legionella, 468 General Characteristics, 468 Pathogenesis and Spectrum of Disease, 469 Laboratory Diagnosis, 471 Antimicrobial Susceptibility Testing and Therapy, 474 Prevention, 474 35 Brucella, 476 General Characteristics, 476 Epidemiology and Pathogenesis, 476 Spectrum of Disease, 477 Laboratory Diagnosis, 477 Antimicrobial Susceptibility Testing and Therapy, 481 Prevention, 481 36 Bordetella pertussis, Bordetella parapertussis, and Related Species, 482 General Characteristics, 482 Spectrum of Disease, 483 Laboratory Diagnosis, 484 Antimicrobial Susceptibility Testing and Therapy, 485 Prevention, 486 37 Francisella, 487 General Characteristics, 487 Epidemiology and Pathogenesis, 487 Spectrum of Disease, 488 Laboratory Diagnosis, 489 Antimicrobial Susceptibility Testing and Therapy, 491 Prevention, 491 38 Streptobacillus spp. and Spirillum minus, 492 Streptobacillus spp., 492 Spirillum minus, 494 

Section 12  Gram-Negative Cocci, 495 39 Neisseria and Moraxella catarrhalis, 495 General Characteristics, 495 Epidemiology, 495 Pathogenesis and Spectrum of Disease, 496 Laboratory Diagnosis, 498 Antimicrobial Susceptibility Testing and Therapy, 503 Prevention, 503 

Section 13  Anaerobic Bacteriology, 505 40 Overview and General Laboratory Considerations, 505 General Characteristics, 505 Specimen Collection and Transport, 505 Macroscopic Examination of Specimens, 506 Direct Detection Methods, 506 Specimen Processing, 509 Anaerobic Media, 510 Approach to Identification, 512 Antimicrobial Susceptibility Testing and Therapy, 513 41 Overview of Anaerobic Organisms, 515 Epidemiology, 516 Pathogenesis and Spectrum of Disease, 516 Nucleic Acid Detection and MALDI-TOF MS (Gram-Positive), 524 Nucleic Acid Detection and MALDI-TOF MS (Gram-Negative), 525 Nucleic Acid Detection and MALDI-TOF MS, 528 Prevention, 528 

Section 14  Mycobacteria and Other Bacteria With Unusual Growth Requirements, 530 42 Mycobacteria, 530 Mycobacterium tuberculosis Complex, 531 Nontuberculous Mycobacteria, 534 Laboratory Diagnosis of Mycobacterial Infections, 541 Antimicrobial Susceptibility Testing and Therapy, 555 Prevention, 559 43 Obligate Intracellular and Nonculturable Bacterial Agents, 562 Chlamydia, 563 Rickettsia, Orientia, Anaplasma, and Ehrlichia, 570 Coxiella, 573 Tropheryma whipplei, 573 Klebsiella granulomatis, 574 44 Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma, 576 General Characteristics, 576 Epidemiology and Pathogenesis, 577 Spectrum of Disease, 580 Laboratory Diagnosis, 580 Cultivation, 581 Susceptibility Testing and Therapy, 584 Prevention, 584 45 The Spirochetes, 586 Treponema, 587 Borrelia, 593 Brachyspira, 597 Leptospira, 598 Prevention, 600 

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Part IV  Parasitology, 601 46 Overview of the Methods and Strategies in Parasitology, 601 Epidemiology, 601 Pathogenesis and Spectrum of Disease, 609 Laboratory Diagnosis, 609 Approach to Identification, 617 Prevention, 639 Ectoparasites, 640 47 Intestinal Protozoa, 642 Amoebae, 642 Flagellates, 662 Ciliates, 669 Sporozoa (Apicomplexa), 670 Microsporidia, 678 48 Blood and Tissue Protozoa, 683 Plasmodium spp., 683 Babesia spp., 696 Trypanosoma spp., 697 Leishmania spp., 702 49 Protozoa From Other Body Sites, 706 Free-Living Amoebae, 706 Naegleria fowleri, 706 Acanthamoeba spp., 710 Acanthamoeba Keratitis, 712 Balamuthia mandrillaris, 712 Trichomonas vaginalis, 713 Toxoplasma gondii, 715 50 Intestinal Nematodes, 720 Ascaris lumbricoides, 720 Enterobius vermicularis, 723 Strongyloides stercoralis, 725 Trichostrongylus spp., 727 Trichuris trichiura, 728 Capillaria philippinensis, 729 Hookworms, 729 Ancylostoma duodenale, 730 Necator americanus, 730 Ancylostoma ceylonicum, Ancylostoma braziliense, and Ancylostoma caninum, 731 Results and Reporting, 732 51 Tissue Nematodes, 733 Trichinella spp., 733 Toxocara canis (Visceral Larva Migrans) and Toxocara cati (Ocular Larva Migrans), 734 Baylisascaris procyonis (Neural Larva Migrans), 737 Ancylostoma braziliense and Ancylostoma caninum (Cutaneous Larva Migrans), 738 Dracunculus medinensis, 738 Parastrongylus cantonensis (Cerebral Angiostrongyliasis), 740

Parastrongylus costaricensis (Abdominal Angiostrongyliasis), 741 Gnathostoma spinigerum, 741 Capillaria hepatica, 741 Dirofilaria immitis and Other Species, 742

52 Blood and Tissue Filarial Nematodes, 744 Wuchereria bancrofti, 744 Brugia malayi and Brugia timori, 748 Loa loa, 748 Onchocerca volvulus, 749 Mansonella spp. (M. ozzardi, M. streptocerca, M. perstans), 750 Dirofilaria spp. (D. immitis, D. repens, D. tenuis), 751 53 Intestinal Cestodes, 753 Diphyllobothrium latum, 753 Dipylidium caninum, 755 Hymenolepis nana, 757 Hymenolepis diminuta, 759 Taenia solium, 759 Taenia saginata, 762 Taenia asiatica, 762 Taenia crassiceps, 763 54 Tissue Cestodes, 765 Taenia solium, 765 Echinococcus granulosus Complex, 767 Echinococcus multilocularis, 769 Echinococcus oligarthrus and Echinococcus vogeli, 770 Taenia multiceps and Other Species, 771 Taenia serialis, 771 Spirometra mansonoides, 772 55 Intestinal Trematodes, 774 Echinostoma spp., 774 Fasciolopsis buski General Characteristics, 775 Gastrodiscoides hominis, 776 Heterophyes: Metagonimus yokogawai, Centrocestus spp., Haplorchis spp., Stellantchamus spp., and Pygidiopsis spp. General Characteristics, 776 56 Liver and Lung Trematodes, 780 The Liver Flukes, 780 The Lung Flukes, 783 57 Blood Trematodes, 786 General Characteristics, 786 Epidemiology, 787 Pathology and Spectrum of Disease, 787 Laboratory Diagnosis, 789 Therapy, 790 Prevention, 790

Contents

Part V  Mycology, 791

Part VI  Virology, 884

58 Overview of Fungal Identification Methods and Strategies, 791 Epidemiology, 791 General Features of the Fungi, 792 Taxonomy of the Fungi, 792 Clinical Classification of the Fungi, 794 Pathogenesis and Spectrum of Disease, 795 Laboratory Diagnosis, 795 General Considerations for the Identification of Yeasts, 803 General Considerations for the Identification of Molds, 806 General Morphologic Features of the Molds, 808 Clinical Relevance for Fungal Identification, 811 Laboratory Safety, 812 Prevention, 812

64 Overview of the Methods and Strategies in Virology, 884 General Characteristics, 885 Epidemiology, 888 Pathogenesis and Spectrum of Disease, 888 Prevention and Therapy, 888 Viruses That Cause Human Diseases, 889 Laboratory Diagnosis, 889

59 Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses, 814 The Mucorales, 815 The Entomophthorales and Basidiobolales, 819 The Dermatophytes, 819 The Opportunistic Mycoses, 827 Systemic Mycoses, 833 60 Dematiaceous (Melanized) Molds, 845 General Characteristics, 846 Epidemiology and Pathogenesis Superficial Infections, 846 Pathogenesis and Spectrum of Disease, 849 Laboratory Diagnosis, 850 61 Atypical and Parafungal Agents, 858 PNEUMOCYSTIS, 858

General Characteristics, 858 Epidemiology, 858 Pathogenesis and Spectrum of Disease, 859 Laboratory Diagnosis, 859 Rare Atypical and Parafungal Agents, 860

62 The Yeasts and Yeastlike Organisms, 863 General Characteristics, 863 Epidemiology, 865 Pathogenesis and Spectrum of Disease, 867 Laboratory Diagnosis, 870 Commercial Yeast Identification Systems, 875 63 Antifungal Susceptibility Testing, Therapy, and Prevention, 880 Antifungal Susceptibility Testing, 880 Antifungal Therapy and Prevention, 881

65 Viruses in Human Disease, 918 Viruses in Human Disease, 918 Adenoviridae, 918 Arenaviridae, 921 Astroviridae, 922 Caliciviridae, 922 Coronaviridae, 922 Filoviridae, 924 Flaviviridae, 925 Hantaviridae, 927 Hepadnaviridae, 928 Hepeviridae, 929 Herpesviridae, 929 Herpesviruses, 930 Varicella-Zoster Virus, 931 Epstein-Barr Virus, 932 Cytomegalovirus, 932 Orthomyxoviridae, 933 Papillomaviruses, 934 Paramyxoviridae, 935 Measles Virus, 935 Parvoviridae, 936 Picornaviridae, 936 Enteroviruses, Parechoviruses, and Polioviruses, 936 Rhinovirus, 937 Hepatitis A Virus, 938 Pneumoviridae, 939 Polyomaviridae, 939 Poxviridae, 940 Reoviridae, 940 Retroviridae, 941 Rhabdoviridae, 943 Togaviridae, 943 Prions in Human Disease, 943 66 Antiviral Therapy, Susceptibility Testing, and Prevention, 946 Antiviral Therapy, 946 Antiviral Resistance, 946 Methods of Antiviral Susceptibility Testing, 948 Prevention of Other Viral Infections, 951

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Part VII  Diagnosis by Organ System, 953 67 Bloodstream Infections, 953 General Considerations, 954 Detection of Bacteremia, 960 Special Considerations for Other Relevant ­Organisms Isolated From Blood, 966 68 Infections of the Lower Respiratory Tract, 970 General Considerations, 970 Diseases of the Lower Respiratory Tract, 973 Laboratory Diagnosis of Lower Respiratory Tract Infections, 978 69 Upper Respiratory Tract Infections and Other Infections of the Oral Cavity and Neck, 985 General Considerations, 985 Diseases of the Upper Respiratory Tract, Oral Cavity, and Neck, 985 Diagnosis of Upper Respiratory Tract Infections, 989 Diagnosis of Infections in the Oral Cavity and Neck, 991 70 Meningitis and Other Infections of the Central Nervous System, 993 General Considerations, 993 Shunt Infections, 999 Laboratory Diagnosis of Central Nervous System Infections, 999 71 Infections of the Eyes, Ears, and Sinuses, 1005 Eyes, 1005 Ears, 1012 Sinuses, 1014 72 Infections of the Urinary Tract, 1017 General Considerations, 1017 Infections of the Urinary Tract, 1018 Laboratory Diagnosis of Urinary Tract Infections, 1022 73 Genital Tract Infections, 1029 General Considerations, 1029 Genital Tract Infections, 1030 Laboratory Diagnosis of Genital Tract Infections, 1037 74 Gastrointestinal Tract Infections, 1045 Anatomy, 1045 Resident Gastrointestinal Microbiome, 1046 Gastroenteritis, 1047 Other Infections of the Gastrointestinal Tract, 1055 Laboratory Diagnosis of Gastrointestinal Tract Infections, 1058

75 Skin, Soft Tissue, and Wound Infections, 1064 General Considerations, 1064 Skin and Soft Tissue Infections, 1065 Laboratory Diagnostic Procedures, 1072 76 Normally Sterile Body Fluids, Bone and Bone Marrow, and Solid Tissues, 1075 Specimens From Sterile Body Sites, 1075 Laboratory Diagnostic Procedures, 1081 

Part VIII  Clinical Laboratory Management, 1085 77 Quality in the Clinical Microbiology Laboratory, 1085 Quality Program, 1086 Specimen Collection and Transport, 1086 Standard Operating Procedure Manual, 1087 Personnel, 1087 Reference Laboratories, 1087 Patient Reports, 1087 Proficiency Testing, 1088 Performance Checks, 1088 Antimicrobial Susceptibility Tests, 1089 Maintenance of Quality Control Records, 1089 Maintenance of Reference Quality Control Stocks, 1090 Quality Assurance Program, 1090 Types of Quality Assurance Audits, 1090 Conducting a Quality Assurance Audit, 1091 Continuous Daily Monitoring, 1091 78 Infection Control, 1094 Incidence of Health Care–Associated Infections, 1095 Types of Health Care–Associated Infections, 1095 Emergence of Antibiotic-Resistant Microorganisms, 1096 Hospital Infection Control Programs, 1097 Role of the Microbiology Laboratory, 1097 Characterizing Strains Involved in an Outbreak, 1097 Preventing Health Care–Associated Infections, 1098 Surveillance Methods, 1099 79 Sentinel Laboratory Response to Bioterrorism, 1103 General Considerations, 1103 Government Laws and Regulations, 1103 Laboratory Response Network, 1105 Glossary, 1108 Index, 1126

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Microbial Taxonomy OBJECTIVES 1. Define classification, identification, species, genus, type genus, and binomial nomenclature. 2. Properly use binomial nomenclature in the identification of microorganisms, including syntax, capitalization, and punctuation. 3. Identify a microorganism’s characteristics as either phenotypic or genotypic. 4. Define polyphasic taxonomy and chemotaxonomic methods and how they are being applied to the classification of microorganisms. 5. Describe how the classification, naming, and identification of organisms play a role in diagnostic microbiology in the clinical setting.

T

he science of taxonomy is a systematic process applied to all living entities, providing a consistent means to classify, name (nomenclature), and identify organisms. This consistency allows biologists worldwide to use a common label for every organism studied within the multitude of biologic disciplines. The common language of taxonomy minimizes confusion about organisms’ names, physiology, and biologic relatedness. Taxonomy is important in the phylogeny (the evolutionary history of organisms) and scientific study of all living things in virtually every biologic discipline, including microbiology. As a result of the advances in molecular biology, traditional taxonomy based on genotypic, phenotypic, and phylogenetic or evolutionary relationships currently encompasses a multifaceted analysis of epigenetic (variations in gene expression not caused by nucleic acid sequence similarities or differences) and chemotaxonomic methods. This method of classification or polyphasic taxonomy provides a more detailed but very complex analysis of the current classification system using ribosomal ribonucleic acid (rRNA) sequences, whole genome sequences, epigenetics, and mass spectrometry (MS). The “gold standard” for classification of bacterial species has historically been based on deoxyribonucleic acid (DNA) including DNA hybridization (DDH) patterns and 16S rRNA gene (16S rDNA) sequence homology. With the implementation of next generation sequencing, a more detailed analysis of organism

genomes including the average nucleotide identity (ANI), multilocus phylogenetic, and genome-to-genome distance (GGD) analysis permit the resolution of microorganisms from closely related subspecies to specific species. Not all parameters clearly delineate each organism to the species level. In other words, some characteristics may strengthen the organization of the genus, and some may be useful at the species level. Species identification techniques have distinct variations in cutoff values or thresholds for the differentiation of organisms at the genus and species levels. The comparative thresholds indicate the likelihood that two genomes are from the same organism (Table 1.1). When using a single sequence such as the 16S rRNA, the possibility of gene transfer may also affect genotypic classification. Although 16S rRNA sequences are evolutionarily highly conserved, ANI evaluates multiple coding regions across an entire genome, making the genomic analysis more detailed and accurate. Finally, lateral gene transfer among organisms, particularly bacteria, creates difficulty in the classification of organisms according to phenotypic traits or biochemical traits and genotypic criteria such as DNA G + C content, which has historically been the hallmark of diagnostic microbiology. Molecular methods have provided a means for identifying the historical core genomes used in classification and species identification. However, it is important to recognize that phenotypic expression and classification of organisms will continue to be compounded by the variation in genomes as a result of gene transfer among organisms. In addition to more advanced genomic analysis, chemotaxonomic methods are more frequently being applied to the identification and classification of microorganisms. These methods include protein studies, fatty acid analysis, and cell wall composition. MS and matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) use the separation and analysis of high-abundance proteins and peptides for the classification and identification of bacterial isolates. Techniques such as rapid evaporative ionization mass spectrometry (REIMS) are able to identify molecules and create images of tissues and microorganisms from laboratory growth medium. This polyphasic analysis beyond genomics provides a mechanism to use the MS data in conjunction with the genomic analysis and phenotypic characteristics to identify and classify organisms, as well as monitor biochemical therapies in complex disease states. 1

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TABLE   Identification Criteria and Characteristics for Microbial Classification 1.1 

Criteria

Characteristics

Phenotypic Macroscopic morphology

The microbial growth patterns on artificial media as observed when inspected with the unaided eye. Examples include the size, texture, and pigmentation of bacterial colonies.

Microscopic morphology

The size, shape, intracellular inclusions, cellular appendages, and arrangement of cells when observed with the aid of microscopic magnification.

Staining characteristics

The ability of an organism to reproducibly stain a particular color with the application of specific dyes and reagents. Staining is used in conjunction with microscopic morphology for bacterial identification. For example, the Gram stain for bacteria is a critical criterion for differential identification.

Environmental requirements

The ability of an organism to grow at various temperatures, in the presence of oxygen and other gases, at various pH levels, or in the presence of other ions and salts, such as NaCl.

Nutritional requirements

The ability of an organism to use various carbon and nitrogen sources as nutritional substrates when grown under specific environmental conditions.

Resistance profiles

The exhibition of a characteristic inherent resistance to specific antibiotics, heavy metals, or toxins.

Antigenic properties

The profiles of microorganisms established by various serologic and immunologic methods to determine relatedness among various microbial groups.

Subcellular properties

Molecular constituents of the cell that are typical of a particular taxon, or organism group, as established by various analytic methods. Some examples include cell wall components, components of the cell membrane, and enzymatic content of the microbial cell.

Chemotaxonomic properties

The chemical constituents of the cell, such as the structure of teichoic acids, fatty acid analysis, and protein profiles, as determined by analytical methods.

Genotypic DNA base composition ratio

DNA comprises four bases (guanine, cytosine, adenine, and thymine). The extent to which the DNA from two organisms is made up of cytosine and guanine (i.e., G + C content) relative to their total base content can be used as an indicator of relatedness or lack thereof. For example, an organism with a G + C content of 50% is not closely related to an organism with a G + C content of 25%.

Nucleic acid (DNA and RNA) base sequence characteristics, including those determined by hybridization assays

The order of bases along a strand of DNA or RNA is known as the base sequence. The extent to which sequences are homologous (similar) between two microorganisms can be determined directly or indirectly by various molecular methods. The degree of similarity in the sequences may be a measure of the degree of organism relatedness, specifically, the rRNA sequences that remain stable in comparison to the genome as a whole.

Average nucleotide identity (ANI)

This method analyses multiple coding sequences in a microorganism’s genome to determine the average nucleotide identity using genome sequencing and computer algorithms. The relatedness of microorganisms is accurate at 95%–96% threshold for organism identification.

Genome-to-Genome Distance (GGD)

This is a computerized calculation that uses inference by in-silico genome comparisons eliminating the limitations and errors associated with wet-lab techniques. Organisms are related with a GGD threshold score of 70% or greater.

DNA, Deoxyribonucleic acid; RNA, ribonucleic acid; rRNA, ribosomal RNA.

As technology improves, the classification and identification of organisms will undoubtedly continue to evolve along with the changes in the populations of organisms. In diagnostic microbiology, classification, nomenclature, and identification of microorganisms play a central role in providing an accurate, timely diagnosis and monitoring the management of infectious disease. A brief, detailed discussion of the major components of taxonomy is important for a basic understanding of bacterial identification and application to diagnostic microbiology.

Classification Classification is a method for organizing microorganisms into groups or taxa based on similar morphologic, physiologic, and genetic traits. The hierarchical classification system consists of the following taxa: • Domains (Bacteria, Archaea, and Eukarya) • Kingdom (contains similar divisions or phyla; most inclusive taxa) • Phylum (contains similar classes; equivalent to the Division taxa in botany)

CHAPTER 1  Microbial Taxonomy

• • • • •

 lass (contains similar orders) C Order (contains similar families) Family (contains similar genera) Genus (contains similar species) Species (specific epithet; lowercase Latin adjective or noun; most exclusive taxa) Bacteria or prokaryotes (prenucleus) are separated into two domains, the Bacteria and the Archaea (ancient bacteria). The Bacteria contain the environmental prokaryotes (blue green or cyanobacteria) and the heterotrophic medically relevant bacteria. The Archaea are environmental isolates that live in extreme habitats such as high salt concentrations, jet fuel, or high temperatures. The third domain, Eukarya, eukaryotes (true nucleus), also contains medically relevant organisms, including fungi and parasites. There are several other taxonomic sublevels below the domains, as noted previously; however, the typical application of organism classification in the diagnostic microbiology laboratory primarily uses the taxa beginning at the family designation.

Family A family encompasses a group of organisms that may contain multiple genera and consists of organisms with a common attribute. The name of a family is formed by adding the suffix -aceae to the root name of one of the group’s genera, called the type genus; for example, the Streptococcaceae family type genus is Streptococcus. One exception to the rule in microbiology is Enterobacterales; it is named after the “enteric” group of bacteria rather than the type species Escherichia coli. Bacterial (prokaryotic)-type species or strains are determined according to guidelines published by the International Committee for the Systematics of Prokaryotes (ICSP) in The International Code of Nomenclature of Prokaryotes (ICNP). This code provides the guidelines for linking nomenclature, classification, and characterization of organisms using the physiologic, biochemical, genetic, and phenotypic traits of organisms. Microorganism type species should be described in detail using diagnostic and comparable methods that are reproducible, and all authentic strains must be available for further analysis. 

Genus Genus (plural, genera), the next taxon, contains different species that have several important features in common. Each species within a genus differs sufficiently to maintain its status as an individual species. Placement of a species within a particular genus is based on various genetic and phenotypic characteristics shared among the species. Microorganisms do not possess the multitude of physical features exhibited by higher organisms such as plants and animals. For instance, they rarely leave any fossil record, and they exhibit a tremendous capacity to intermix genetic material among seemingly unrelated species and genera. For these reasons, confidently establishing a microorganism’s relatedness in higher taxa beyond the genus level is difficult. Although

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grouping similar genera into common families and similar families into common orders is used for classification of plants and animals, these higher taxa designations (i.e., division, class, and order) are not useful for classifying bacteria. 

Species Species (abbreviated as sp., singular, or spp., plural) is the most basic of the taxonomic groups and can be defined as a collection of bacterial strains that share common physiologic and genetic features and differ notably from other microbial species. Occasionally, taxonomic subgroups within a species, called subspecies, are recognized. Furthermore, designations such as biotype, serotype, or genotype may be given to groups below the subspecies level that share specific but relatively minor characteristics. For example, Klebsiella pneumoniae and Klebsiella oxytoca are two distinct species within the genus Klebsiella. Serratia odorifera biotype 2 and Treponema pallidum subsp. pallidum are examples of a biotype and a subspecies designation. A biotype is considered the same species with the same genetic makeup but displays differential physiologic characteristics. Subspecies do not display significant enough divergence to be classified as a biotype or a new species. Although these subgroups may have some taxonomic importance, their usefulness in diagnostic microbiology is limited. 

Nomenclature Nomenclature is the naming of microorganisms according to established rules and guidelines set forth in the ICNP. It provides the accepted labels by which organisms are universally recognized. Because genus and species are the groups commonly used by microbiologists, the discussion of rules governing microbial nomenclature is limited to these two taxa. In this binomial (two name) system of nomenclature, every organism is assigned a genus and a species of Latin or Greek derivation. Each organism has a scientific “label” consisting of two parts: the genus designation, in which the first letter is always capitalized, and the species designation, in which the first letter is always lowercase. The two components are used simultaneously and are printed in italics or underlined in script. For example, the streptococci include Streptococcus pneumoniae, Streptococcus pyogenes, Streptococcus agalactiae, and Streptococcus bovis, among others. Alternatively, the name may be abbreviated by using the uppercase form of the first letter of the genus designation followed by a period (.) and the full species name (e.g., S. pneumoniae, S. pyogenes, S. agalactiae, and S. bovis). Finally, when discussing a single specific organism, the species may be designated using sp., and a group of species within the genus using spp. (e.g., Staphylococcus sp. and Staphylococcus spp.). Frequently an informal designation (e.g., staphylococci, streptococci, enterococci) may be used to label a particular group of organisms. These designations are not capitalized or italicized.

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As more information is gained regarding organism classification and identification, a particular species may be moved to a different genus or assigned a new genus name. The rules and criteria for these changes are beyond the scope of this chapter, but such changes are documented in the International Journal of Systemic and Evolutionary Microbiology. Published nomenclature may be found at http:// www.bacterio.net for bacteria, http://www.ictvonline. org for viruses, http://www.iapt-taxon.org/nomen/main. php for fungi, and http://www.iczn.org for parasites. It is important to note that the fungi and parasite lists are difficult to maintain and may not reflect the current validity at the time of review. In the diagnostic laboratory, changes in nomenclature are phased in gradually so that physicians and laboratorians have ample opportunity to recognize that a familiar pathogen has been given a new name. This is usually accomplished by using the new genus designation while continuing to provide the previous designation in parentheses; for example, Stenotrophomonas (Xanthomonas) maltophilia or Burkholderia (Pseudomonas) cepacia. 

Identification Microbial identification is the process by which a microorganism’s key features are delineated. Once those features have been established, the profile is compared with those of other previously characterized microorganisms. The organism can then be assigned to the most appropriate taxa and can be given appropriate genus and species names; both are essential aspects of taxonomy in diagnostic microbiology and the management of infectious disease (Box 1.1).

Identification Methods A wide variety of methods and criteria are used to establish a microorganism’s identity. These methods can be separated into either of two general categories: genotypic or phenotypic characteristics. Genotypic characteristics relate to an organism’s genetic makeup, including the nature of the organism’s genes and constituent nucleic acids (see Chapter 2 for more information about microbial genetics). Phenotypic characteristics are based on features beyond the genetic level, including both readily observable characteristics and features that may require extensive analytic procedures to be detected. Examples of characteristics used as criteria for bacterial identification and classification are provided in Table 1.1. Modern microbial taxonomy uses a combination of several methods to characterize microorganisms thoroughly to classify and name each organism. Although the criteria and examples in Table 1.1 are given in the context of microbial identification for classification purposes, the principles and practices of classification parallel the approaches used in diagnostic microbiology for the identification and characterization of microorganisms encountered in the clinical setting. Fortunately, because of the previous efforts and accomplishments of microbial taxonomists, microbiologists do not have to use several burdensome classification

• BOX 1.1 Role of Taxonomy in Diagnostic

Microbiology

• E  stablishes and maintains records of key characteristics of clinically relevant microorganisms • Facilitates communication among technologists, microbiologists, physicians, and scientists by assigning universal names to clinically relevant microorganisms. This is essential for: • Establishing an association of particular diseases or syndromes with specific microorganisms • Epidemiology and tracking outbreaks • Accumulating knowledge regarding the management and outcome of diseases associated with specific microorganisms • Establishing patterns of resistance to antimicrobial agents and recognition of changing microbial resistance patterns • Understanding the mechanisms of antimicrobial resistance and detecting new resistance mechanisms exhibited by microorganisms • Recognizing new and emerging pathogenic microorganisms • Recognizing changes in the types of infections or diseases caused by characteristic microorganisms • Revising and updating available technologies for the development of new methods to optimize the detection and identification of infectious agents and the detection of resistance to antiinfective agents (microbial, viral, fungal, and parasitic) • Developing new antiinfective therapies (microbial, viral, fungal, and parasitic)

and identification schemes to identify infectious agents. Instead, microbiologists use key phenotypic and genotypic features on which to base their identification to provide clinically relevant information in a timely manner (Chapter 12). This should not be taken to mean that the identification of all clinically relevant organisms is easy and straightforward. This is also not meant to imply that microbiologists can identify or recognize only organisms that have already been characterized and named by taxonomists. Indeed, the clinical microbiology laboratory is well recognized as the place where previously unknown or uncharacterized infectious agents are initially encountered, and as such it has an ever-increasing responsibility to be the source of information and reporting for emerging etiologies of infectious disease.

Visit the Evolve site for a complete list of procedures, review questions and answers, and case studies.

Bibliography Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2015, Elsevier-Saunders. Bhandari V, Naushad HS, Gupta RS: Protein based molecular markers provide reliable means to understand prokaryotic phylogeny and support darwinian mode of evolution, Front Cell Infect Microbiol 2:98, 2012. Brock TD, Madigan M, Martinko J, et al.: Biology of microorganisms, Englewood Cliffs, NJ, 2009, Prentice Hall.

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Clark AE, Kaleta EJ, Arora A, Wolk DM: Matrix-assisted laser desorption ionization time-of-flight mass spectrometry: a fundamental shift in the routine practice of clinical microbiology, Clin Microbiol Rev 26:547–603, 2013. Cleary J, Sanchez L: Large scale MALDI-T of imaging of metabolites from filamentous fungi, Billerica MA, 2018, Ebook Bruker Daltonics Inc. Dworkin M, Falkow S, Rosenberg E, et al.: The prokaryotes: a handbook on the biology of bacteria: ecophysiology, isolation, identification, applications (Vol. 1–4). New York, 2006, Springer. Figueras M, Beaz-Hidalgo R, Hossain MJ, Liles MR: Taxonomic affiliation of new genomes should be verified using average nucleotide identity and multilocus phylogenetic analysis, Genome Announc 2(6):e00927–e01014, 2014, https://doi.org/10.1128/ genomeA.00927-14. Golf O, Strittmatter N, Karancsi T, et al.: Rapid evaporative ionization mass spectrometry imaging platform for direct mapping from bulk tissue and bacterial growth media, Anal Chem 87:2527–2534, 2015. Jorgensen J, Pfaller M, Carroll K, et al.: Manual of clinical microbiology, ed 11, Washington, DC, 2015, ASM Press.

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Kook JK, Park SN, Lim YK, et  al.: Genome-based reclassification of fusobacterium nucleatum subspecies at the species level, Curr Microbiol 74:1137–1147, 2017. Martins MD, Machado-de-Lima NM, Branco LHZ: Polyphasic approach using multilocus analyses supports the establishment of the new aerophytic cyanobacterial genus Pycnacronema (Coleofas ciculaceae, Oscillatoriales), J Phycol 55:146–159, 2018, https://doi. org/10.1111/jpy.12805. Mohr KI, Wolf C, Nübel U, et al.: A polyphasic approach leads to seven new species of cellulose decomposing genus sorangium, Sorangium ambruticinum sp. nov., sorangium arenae sp. nov., sorangium bulgaricum sp. nov., sorangium dawidii sp. nov, sorangium kjenyense sp. nov., sorangium orientale sp. nov. and sorangium richenbachii sp. nov, Int J Syst Evol Microbiol 68:3576–3586, 2018. Parker CT, Tindall BJ, Garrity GM: International code of nomenclature for prokaryotes, Int J Syst Evol Microbiol 69:S1–S111, 2019, Available at http://ijs.microbiologyresearch.org/content/journal/ij sem/10.1099/ijsem.0.000778#tab9.

Chapter Review 1. The most specific and exclusive taxon used in the classification of microorganisms is: a. Family b. Order c. Species d. Genus 2. The process consisting of a series of methods designed to provide the microbiologist with relevant and useful clinical information about a microorganism is: a. Classification b. Identification c. Organization d. Taxonomy 3. Classification and naming of organisms are useful in diagnostic microbiology for all of the following except: a. Providing standardized groupings for identification b. Standardized groupings are always genotypically similar at 0.98% c. Standardized groupings share similar phenotypic traits d. The ability of organisms within a standard group may be identified using similar methods 4. Which of the following is not a correct use of the binomial nomenclature system? (Select all that apply.) a. Staphylococcus Aureus b. S. aureus c. Staphylococcus aureus d. Staphylococcus aureus

5. Labeling: Label each of the following characteristics as either a phenotypic (P) or a genotypic (G) characteristic. _____ Color of growth on artificial media _____ The presence of an antibiotic-resistance DNA sequence _____ The shape of the bacterial cell _____ The arrangement of the bacterial cells on a microscope slide _____ The ability of the organism to ferment lactose 6. Mass spectrometry is a technique used to separate and identify the spectrum of proteins and peptides that are expressed by microorganisms. This method is considered a ________method for the characterization and classification of organisms. a. phenotypic b. chemotaxonomic c. genotypic d. polyphasic 7. Which of the following methods would be considered chemotaxonmic? a. Fatty acid analysis b. Protein mass spectrometry c. Cell wall composition d. All of the answers are correct

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Bacterial Genetics, Metabolism, and Structure OBJECTIVES 1. Describe the basic structure and organization of prokaryotic (bacterial) chromosomes, including number, relative size, and cellular location. 2. Outline the basic processes and essential components required for information transfer in replication, transcription, translation, and regulatory mechanisms. 3. Define mutation, recombination, transduction, transformation, and conjugation. 4. Describe how genetic alterations and diversity provide a mechanism for the evolution and survival of microorganisms. 5. Differentiate environmental oxygenation and final electron acceptors (aerobes, facultative anaerobes, and strict anaerobes) in the formation of energy. 6. Compare and contrast the key structural elements, cellular organization, and types of organisms classified as prokaryotic and eukaryotic. 7. State the functions and biologic significance of the following cellular structures: the outer membrane, cell wall, periplasmic space, cytoplasmic membrane, capsule, fimbriae, pili, flagella, nucleoid, and cytoplasm. 8. Differentiate the organization and chemical composition of the cell envelope for a gram-positive and a gram-negative bacterium.

M

icrobial genetics, metabolism, and structure are the keys to microbial viability and survival. These processes involve numerous pathways that are widely varied, often complicated, and frequently interactive. Essentially, survival requires nutrients and energy to fuel the synthesis of materials necessary to grow, propagate, and carry out other metabolic processes (Fig. 2.1). Although the goal of survival is the same for all organisms, the strategies microorganisms use to accomplish this vary substantially. Knowledge regarding genetic, metabolic, and structural characteristics of microorganisms provides the basis for understanding almost every aspect of diagnostic microbiology, including: • The mechanisms by which microorganisms cause disease • The development and implementation of techniques for microbial detection, cultivation, identification, and characterization 6

• A  ntimicrobial action and resistance • The development and implementation of tests for the detection of antimicrobial resistance • Potential strategies for disease therapy and control of microorganisms Microorganisms vary significantly in their genomic and metabolic pathways and therefore structure. A detailed consideration of these differences is beyond the scope of this textbook. Therefore, a generalized description of bacterial systems is used as a model to discuss microbial physiology and structure. Information regarding characteristics of fungi, parasites, and viruses can be found in subsequent chapters for each specific taxonomic group.

Bacterial Genetics Genetics, the process of heredity and variation, is the starting point from which all other cellular pathways, functions, and structures originate. The ability of a microorganism to maintain viability, adapt, multiply, and cause disease is determined by the organism’s genetic composition. The three major aspects of microbial genetics that require discussion include: • The structure and organization of genetic material • Replication and expression of genetic information • The mechanisms by which genetic information is altered and exchanged among bacteria

Energy and nutrients

Motion and other responses to environment

Genetic processes Biosynthesis

Assembly of cell structure

Waste removal

Bacterial cell

• Fig. 2.1  General overview of bacterial cellular processes.

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

or prenuclear, organisms do not have membrane-bound organelles, and the cells’ genetic material is therefore not enclosed in a nucleus. Eukaryotic, or “true nucleus,” organisms have the genetic material enclosed in a nuclear envelope.

Nucleic Acid Structure and Organization For all living entities, hereditary information resides or is encoded in nucleic acids. The two major classes of nucleic acids are deoxyribonucleic acid (DNA), which is the most common macromolecule that encodes genetic information, and ribonucleic acid (RNA). In some forms, RNA encodes genetic information for various viruses; in other forms, RNA plays an essential role in several of the genetic processes in prokaryotic and eukaryotic cells, including the regulation and transfer of information. Prokaryotic, Nucleotide HO

H H

H

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Phosphodiester bond

5’

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-O P= O

H

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Base

CH2

DNA consists of deoxyribose sugars connected by phosphodiester bonds (Fig. 2.2A). The bases that are covalently linked to each deoxyribose sugar are the key to the genetic code within the DNA molecule. The four nitrogenous bases

Deoxyribose 3’ sugar

O O - P O - Base-pair O

Nucleotide Structure and Sequence

3’ hydroxyl

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H

O O

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Fig. 2.2 (A) Molecular structure of deoxyribonucleic acid (DNA) depicting nucleotide structure, phosphodiester bonds connecting nucleotides, and complementary base pairing (A, adenine; T, thymine; G, guanine; C, cytosine) between antiparallel nucleic acid strands. (B) 5′ and 3′ antiparallel polarity and double-helix configuration of DNA.

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include two purines, adenine (A) and guanine (G), and the two pyrimidines, cytosine (C) and thymine (T) (Fig. 2.3). In RNA, uracil replaces thymine. The combined sugar, phosphate, and a base form a single unit referred to as a nucleotide (adenosine triphosphate [ATP], guanine triphosphate [GTP], cytosine triphosphate [CTP], and thymine triphosphate [TTP] or uridine triphosphate [UTP]). DNA and RNA are nucleotide polymers (i.e., chains or strands), and the order of bases along a DNA or RNA strand is known as the base sequence. This sequence provides the information that codes for the proteins that will be synthesized by microbial cells; that is, the sequence is the genetic code. 

an entire genome are usually expressed in the number of bp present (e.g., kilobases [103 bases], megabases [106 bases]). Certain genes are widely distributed among various organisms, whereas others are limited to a particular species. In addition, the base pair sequence for individual genes may be highly conserved (i.e., show limited sequence differences among different organisms) or be widely variable. As discussed in Chapter 8, these similarities and differences in genetic content and sequences are the basis for the development of molecular methods used to detect, identify, and characterize microorganisms. 

Deoxyribonucleic Acid Molecular Structure

The genome is organized into discrete elements known as chromosomes. The set of genes within a given chromosome is arranged in a linear fashion, but the number of genes per chromosome is variable. Similarly, although the number of chromosomes per cell is consistent for a given species, this number varies considerably among species. For example, human cells contain 23 pairs (i.e., diploid) of chromosomes, whereas bacteria contain a single, unpaired (i.e., haploid) chromosome. Bacteria are classified as prokaryotes; therefore the chromosome is not located in a membrane-bound organelle (i.e., nucleus). The bacterial chromosome contains the genes essential for viability and exists as a double-stranded, closed, circular macromolecule. The molecule is extensively folded and twisted (i.e., supercoiled) to fit within the confined space of the bacterial cell. The linearized, unsupercoiled chromosome of the bacterium Escherichia coli is approximately 130 μm long, but it fits within a cell 1 × 3 μm; this attests to the extreme compact structure of the supercoiled bacterial chromosome. For genes in the compacted chromosome to be expressed and replicated, unwinding or relaxation of the molecule is required. In contrast to the bacterial chromosome, the chromosomes of parasites and fungi number more than one per cell, are linear, and are housed within a membrane-bound organelle (the nucleus) of the cell. This difference is a major criterion for classifying bacteria as prokaryotes and fungi and parasites as eukaryotes. The genetic makeup of a virus may consist of DNA or RNA contained within a protein coat rather than a cell. 

The intact DNA molecule is composed of two nucleotide polymers. Each strand has a 5′ (prime) phosphate and a 3′ (prime) hydroxyl terminus (Fig. 2.2A). The two strands run antiparallel, with the 5′ of one strand opposed to the 3′ terminal of the other. The strands are also complementary. This adherence to A-T and G-C base pairing results in a doublestranded DNA (dsDNA) molecule (double helix). The two antiparallel single strands of DNA form a “twisted ladder” structure (Fig. 2.2B). In addition, the dedicated base pairs (bp) provide the format for consistent replication and expression of the genetic code. In contrast to DNA, which carries the genetic code, RNA rarely exists as a double-stranded molecule. There are four major types of RNA (messenger RNA [mRNA], transfer RNA [tRNA], and ribosomal RNA [rRNA]) along with a variety of noncoding RNA (ncRNA) molecules such as microRNA (miRNA) that play key roles in posttranscriptional regulation of gene expression. 

Genes and the Genetic Code A DNA sequence that encodes a specific product (RNA or protein) is defined as a gene. Thousands of genes in an organism encode messages or blueprints for the production of one or more proteins and RNA products that play essential metabolic roles in the cell. All the genes in an organism comprise the organism’s genome. The genome of a microorganism includes the chromosomes and the mobilome (extrachromosomal mobile genetic elements). The size of a gene and

NH2

O

N H

N H

O

N H

Cytosine C

Thymine T

Uracil U

DNA and RNA

DNA only

RNA only

•

N

N

N O

NH2

O

H3C N

Chromosomes

O

N H

O N

N Adenine A

N

N

N H

N

NH2

Guanine G DNA and RNA

Fig. 2.3 Molecular structure of nucleic acid bases. DNA, Deoxyribonucleic acid; RNA, ribonucleic acid.  Pyrimidines: cytosine, thymine, and uracil. Purines: adenine and guanine.

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

Nonchromosomal Elements (Mobilome) Although the bacterial chromosome represents the majority of a cell’s genome, not all genes are confined to the chromosome. Many genes may also be located on plasmids and transposable elements. Both of these extrachromosomal elements are able to replicate and encode information for the production of various cellular products. Many of these elements replicate by integration into the host chromosome, whereas others, referred to as episomes, are capable of replication independently of the host chromosome. Although considered part of the bacterial genome, they are not as stable as the chromosome and may be lost during cellular replication, often without any detrimental effects on the viability of the cell. Plasmids exist as double-stranded, closed, circular, autonomously replicating extrachromosomal genetic elements ranging in size from 1 to 2 kilobases up to 1 megabase or more. The number of plasmids per bacterial cell varies extensively, and each plasmid is composed of several genes. Some genes encode products that mediate plasmid replication and transfer between bacterial cells, whereas others encode products that provide a specialized function, such as a determinant of antimicrobial resistance or a unique metabolic process. Unlike most chromosomal genes, plasmid genes do not usually encode for products essential for viability. Plasmids, in whole or in part, may also become incorporated into the chromosome. Transposable elements are pieces of DNA that move from one genetic element to another, from plasmid to chromosome or vice versa. Unlike plasmids, many are unable to replicate independently and do not exist as separate entities in the bacterial cell. The two types of transposable elements are the simple transposon or insertion sequence (IS) and the composite or complex transposon. Insertion sequences are limited to containing the genes that encode information required for movement from one site in the genome to another. Composite transposons are cassettes (grouping of genes) flanked by insertion sequences. The internal gene embedded in the IS encodes for an accessory function, such as antimicrobial resistance. Plasmids and transposable elements coexist with chromosomes in the cells of many bacterial species. These extrachromosomal elements play a key role in the exchange of genetic material throughout the bacterial microbiosphere, including genetic exchange among clinically relevant bacteria. 

DNA Replication Replication Bacteria multiply by binary fission (a form of cell division), resulting in the production of two daughter cells from one parent cell. As part of this process, the genome must be replicated and each daughter cell receives an identical copy of functional DNA. Replication is a complex process mediated by various enzymes, such as DNA polymerase and

9

cofactors; replication must occur quickly and accurately. For descriptive purposes, replication may be considered in four stages (Fig. 2.4): 1. Unwinding or relaxation of the chromosome’s supercoiled DNA 2. Separation of the complementary strands of the parental DNA. Each strand may serve as a template (i.e., pattern) for synthesis of new DNA strands, referred to as semiconservative replication 3. Synthesis of the new (i.e., daughter) DNA strands 4. Termination of replication, releasing two identical chromosomes, one for each daughter cell Relaxation of supercoiled chromosomal DNA is required, which permits the enzymes and cofactors involved in replication access to the DNA molecule at the site where the replication process will originate (i.e., origin of replication). The origin of replication (a specific sequence of approximately 300 bp) is recognized by several initiation proteins, followed by the separation of the complementary strands of parental DNA. Each parental strand serves as a template for the synthesis of a new complementary daughter strand. The site of active replication is referred to as the replication fork; two bidirectional forks are involved in the replication process. Each replication fork moves through the parent DNA molecule in opposite directions as a bidirectional process. Activity at each replication fork involves different cofactors and enzymes, with DNA polymerase playing a central role. Using each parental strand as a template, DNA polymerase adds nucleotide bases to each growing daughter strand in a sequence that is complementary to the base sequence of the template (parent) strand. The complementary bases of each strand are then held together by hydrogen bonding between nucleotides and the hydrophobic nature of the nitrogenous bases. The new nucleotides can be added only to the 3′ hydroxyl end of the growing strand. The synthesis for each daughter strand occurs in the 5′ to 3′ direction. Termination of replication occurs when the replication forks meet. The result is two complete chromosomes, each containing two complementary strands, one of parental origin and one newly synthesized daughter strand. Although the time required for replication can vary among bacteria, the process generally takes approximately 20 to 40 minutes in rapidly growing bacteria such as E. coli. The replication time for a particular bacterial strain can vary depending on environmental conditions, such as the availability of nutrients or the presence of toxic substances (e.g., antimicrobial agents). 

Expression of Genetic Information Gene expression is the processing of information encoded in genetic elements (i.e., chromosomes, plasmids, and transposons) that results in the production of biochemically functional molecules, including RNA and proteins. The overall process of gene expression is composed of two steps, transcription and translation. Gene expression

10 PA RT I     Basic Medical Microbiology

5’

C A G G T C A

C G A G T G C C

G

G 5’ 3’

Daughter strand

3’

3’

T T A C A T G

5’ 3’

5’

G C G T A

3’ 5’

C

5’

3’ 5’ 3’ 5’ 5’ 3’

C G A

T

3’

T

C

Origin of replication

Replication

DNA polymerase

G C T A C G C G

Replication fork

Replication

A

5’

A A T T T A C

3’

Daughter strand

T A C G G C

Bidirectional replication

A T A T T A G C C G G C A T 3’ 5’ Parent strands

Terminus

• Fig. 2.4  Bacterial deoxyribonucleic acid (DNA) replication with bidirectional movement of two replication forks from the origin of replication. Each parent strand serves as a template for production of a complementary daughter strand and, eventually, two identical chromosomes.

requires various components, including a DNA template representing a single gene or cluster of genes, various enzymes and cofactors, and RNA molecules of specific structure and function. Transcription

Gene expression begins with transcription. During transcription the DNA base sequence of the gene (i.e., the genetic code) is converted into an mRNA molecule that is complementary to the gene’s DNA sequence (Fig. 2.5). Usually only one of the two DNA strands (sense strand) encodes for a functional gene product. This same strand is the template for mRNA synthesis. RNA polymerase is the enzyme central to the transcription process. The enzyme is composed of four protein subunits and a sigma (σ) factor. Sigma factors are required for the RNA polymerase to identify the appropriate site on the DNA template where transcription of mRNA is initiated. This initiation site is also known as the promoter sequence. The remainder of the enzyme functions to unwind the dsDNA at the promoter sequence and use the DNA strand as a template to sequentially add ribonucleotides (ATP, GTP, UTP, and CTP) to form the growing mRNA strand. Transcription proceeds in a 5′ to 3′ direction. However, in mRNA, the TTP of DNA is replaced with UTP. TTP

DNA coding strand 5’• • • C T T

T T T G T T A T T C A G C A T • • • 3’

3’ • • • G A A A A A C A A T A A G T C G T A • • • 5’ RNA polymerase

Transcription

DNA template

5’ C U U U U U G U U A U U C A G C A U 3’ mRNA

Ribosomes, tRNA, amino acids, cofactors

Translation H2N

leu

phe

val

iso

glu

his

COOH

Polypeptide

• Fig. 2.5  Overview of gene expression components: transcription for

production of messenger ribonucleic acid (mRNA) and translation for production of a polypeptide (protein). DNA, Deoxyribonucleic acid; RNA, ribonucleic acid; tRNA, transfer RNA.

contains thymine, and UTP contains uracil. Both molecules contain a heterocyclic ring and are classified as pyrimidines. During synthesis and modification of these molecules, a portion of the molecules are dehydroxylated, forming a

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

2′-deoxynucleotide monophosphate. The dehydroxylated uracil monophosphate (dUMP) is then methylated, forming dehydroxylated thymine monophosphate (dTMP). After phosphorylation, thymine is found only in the final state as deoxythymidine and therefore cannot be incorporated into an RNA molecule. Synthesis of the single-stranded mRNA product ends when specific nucleotide base sequences on the DNA template are encountered. Termination of transcription may be facilitated by a rho (a prokaryotic protein) cofactor or an intrinsic termination sequence. Both of these mechanisms disrupt the mRNA-RNA polymerase template DNA complex. In bacteria, the mRNA molecules that result from the transcription process are polycistronic; that is, they encode for several gene products. Polycistronic mRNA may encode several genes whose products (proteins) are involved in a single or closely related cellular function. When a cluster of genes is under the control of a single promoter sequence, the gene group is referred to as an operon. The transcription process not only produces mRNA but also tRNA, rRNA, and regulatory non coding (ncRNA) molecules. All types of RNA molecules have key roles in protein synthesis. To initiate transcription, accessory factors are needed to localize the RNA polymerase to the promoter upstream of the coding sequence. In bacteria, the σ factor binds to the RNA polymerase and recognizes the gene-specific promotor. In some bacteria a small regulatory RNA (sRNA), 6S RNA, binds the sigma factor to repress transcription in the late stationary phase of bacterial growth. The 6S RNA binds and forms a bulge or loop. The loop serves as an RNA-dependent site for RNA synthesis. The RNA synthesized from the loop is referred to as pRNA. When sufficient pRNA is produced, it causes the 6S RNA to detach from the promotor, permitting transcription to continue. Transfer RNA (tRNA) binds to the A site in the ribosome and delivers the appropriate amino acid during elongation. However, tRNAs exist in many more diverse forms than once believed. In bacteria, the initiation codon codes for an N-formylmethionine. This modified amino acid is never placed inside the coding sequence of a bacterial protein. In other words, there are two forms of tRNA that are produced in bacteria that are cable of carrying methionine. One is the initiator tRNAMet and the other is the elongation tRNAMet. The elongation tRNAMet binds to the A site of the ribosome, whereas the initiation tRNAMet is capable of binding only to the P site within the ribosome. The binding of the elongation-specific tRNA is controlled by transcription elongation factor 1. Ribosomal RNA, specifically the 16S rRNA, has historically been associated with classification of organisms based on evolutionary relatedness. The 16S rRNA is present in all organisms and is responsible for catalyzing the peptidyl transferase reaction during protein synthesis. A very small portion of the molecule is capable of undergoing genetic changes without deleterious effects to the transcription process, providing a means to monitor the evolutionary development of bacterial species.

11

In addition to the differences in tRNA specificity, bacteria have developed numerous mechanisms to regulate gene transcription and respond to the environment, including transcriptional and posttranscriptional regulation. Many sensory and regulatory RNA molecules have now been identified that serve as RNA thermosensors and riboswitches. These molecules may undergo structural alterations during temperature changes or serve as antisense RNAs and sRNAs that bind to either nucleic acid–binding proteins modulating their activity or directly to mRNA sequences to suppress and alter gene expression. This reversible regulation is clearly evident in the expression of virulence genes in many known pathogens including E. coli, Shigella spp., and Yersinia spp. The global changes of RNA expression within the transcriptome of a pathogenic bacteria allows the organism to rapidly adjust to changes in the environment associated with temperature, ionic conditions, oxygen conditions, pH, calcium, iron, and other metals to maintain growth and survival.  Translation

The next phase in gene expression, translation, involves protein synthesis. Through this process the genetic code in mRNA molecules is translated into specific amino acid sequences that are responsible for protein structure and function (Fig. 2.5). The process of protein translation requires the use of a genetic alphabet or code. The code consists of triplets of nucleotide bases, referred to as codons; each codon encodes for a specific amino acid. Because there are 64 different codons for 20 amino acids, an amino acid can be encoded by more than one codon (Table 2.1). Each codon is specific for a single amino acid. The codon sequences in mRNA direct which amino acids are added and in what order. Translation ensures that proteins with proper structure and function are produced. Errors in the process can result in aberrant proteins that are nonfunctional, emphasizing the need for translation to be well controlled and accurate. To accomplish the task of translation, intricate interactions between mRNA, tRNA, and rRNA are required. Sixty different standard types of tRNA molecules are responsible for transferring different amino acids from intracellular locations to the site of protein synthesis. These molecules, which have a structure that resembles an inverted t, contain one anticodon (sequence recognition site) for binding to specific codons (3-base sequences) on the mRNA molecule (Fig. 2.6). A second site binds specific amino acids, the building blocks of proteins. Each amino acid is joined to a specific tRNA molecule through the enzymatic activity of aminoacyl-tRNA synthetases. Transfer RNA molecules use the codons of the mRNA molecule as the template for precisely delivering a specific amino acid for polymerization. This process occurs in ribosomes, which are compact nucleoproteins, composed of rRNA and proteins. They are central to translation, assisting with the coupling of all required components and controlling the translational process.

12 PA RT I     Basic Medical Microbiology

TABLE a 2.1    The Genetic Code as Expressed by Triplet-Base Sequences of Messenger Ribonucleic Acid

Codon

Amino Acid

Codon

Amino Acid

Codon

Amino Acid

Codon

Amino Acid

UUU

Phenylalanine

CUU

Leucine

GUU

Valine

AUU

Isoleucine

UUC

Phenylalanine

CUC

Leucine

GUC

Valine

AUC

Isoleucine (start)b

UUG

Leucine

CUG

Leucine

GUG

Valine

AUG

Methionine

UUA

Leucine

CUA

Leucine

GUA

Valine

AUA

Isoleucine

UCU

Serine

CCU

Proline

GCU

Alanine

ACU

Threonine

UCC

Serine

CCC

Proline

GCC

Alanine

ACC

Threonine

UCG

Serine

CCG

Proline

GCG

Alanine

ACG

Threonine

UCA

Serine

CCA

Proline

GCA

Alanine

ACA

Threonine

UGU

Cysteine

CGU

Arginine

GGU

Glycine

AGU

Serine

UGC

Cysteine

CGC

Arginine

GGC

Glycine

AGC

Serine

UGG

Tryptophan

CGG

Arginine

GGG

Glycine

AGG

Arginine

UGA

None (stop signal)

CGA

Arginine

GGA

Glycine

AGA

Arginine

UAU

Tyrosine

CAU

Histidine

GAU

Aspartic

AAU

Asparagine

UAC

Tyrosine

CAC

Histidine

GAC

Aspartic

AAC

Asparagine

UAG

None (stop signal)

CAG

Glutamine

GAG

Glutamic

AAG

Lysine

UAA

None (stop signal)

CAA

Glutamine

GAA

Glutamic

AAA

Lysine

aThe

codons in deoxyribonucleic acid (DNA) are complementary to those given here. Thus U is complementary to the A in DNA, C is complementary to G, G to C, and A to T. The nucleotide on the left is at the 5′ end of the triplet. bAUG codes for N-formylmethionine at the beginning of messenger ribonucleic acid (mRNA) in bacteria. Modified from Brock TD, Madigan M, Martinko J, et al., eds. Biology of Microorganisms. Upper Saddle River, NJ: Prentice Hall; 2009.

Translation, diagrammatically shown in Fig. 2.6, involves three steps: initiation, elongation, and termination. After termination, bacterial proteins often undergo posttranslational modifications as a final step in protein synthesis. Initiation begins with the association of ribosomal subunits, mRNA, formylmethionine (f-met) tRNA (carrying the initial amino acid of the protein to be synthesized), and various initiation factors (Fig. 2.6A). Assembly of the complex begins at a specific 3- to 9-base sequence (ShineDalgarno sequence) on the mRNA approximately 10 bp upstream of the AUG start codon. After the initial complex has been formed, addition of individual amino acids begins. Elongation involves tRNAs and a host of elongation factors that mediate the addition of amino acids in a specific sequence dictated by the codon on the mRNA molecule (Fig. 2.6B and C and Table 2.1). As the mRNA molecule threads through the ribosome in a 5′ to 3′ direction, peptide bonds are formed between adjacent amino acids, still bound by their respective tRNA molecules in the peptide (P) and acceptor (A) sites of the ribosome. During the process, the forming peptide is moved to the P site, and the 5′ tRNA is released from the exit (E) site. This movement vacates the A

site, which contains the codon specific for the next amino acid, so that the incoming tRNA–amino acid can join the complex (Fig. 2.6C). Because multiple proteins encoded on an mRNA strand can be translated at the same time, multiple ribosomes may be simultaneously associated with one mRNA molecule. Such an arrangement is referred to as a polysome; its appearance resembles a string of pearls. Termination, the final step in translation, occurs when the ribosomal A site encounters a stop or non sense codon that does not specify an amino acid (i.e., a “stop signal”; Table 2.1). At this point, the protein synthesis complex disassociates and the ribosomes are available for another round of translation. After termination, most proteins must undergo modification, such as folding or enzymatic trimming, so that protein function, transportation, or incorporation into various cellular structures can be accomplished. This process is referred to as posttranslational modification. 

Regulation and Control of Gene Expression The vital role that gene expression and protein synthesis play in the survival of cells dictates that bacteria judiciously

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

E site Ribosome

Amino acid

arg

A site

P site

tRNA

f-met

tRNA

Initiation

U C U

A

13

U

A

C

A

U

G

A

A

G

A

C

C

G

C

Messenger RNA codons

G

Start codon

Peptide bond A site P site

E site

f-met

thr

arg

U U

A

C

U

C

U

A

U

G

A G

A

A

C

C

G

G G

C

G

Elongation A

G

G

G

A

U

B

thr

arg

f-met

A site

P site

ala

asp

arg

E site

U

5’

A

U

G

U

Release of discharged tRNA

U

G

G

C

A G

A

A

C

C G G C

C

U

C

C

G

A

G

G

G

A

U

A

A

3’

C • Fig. 2.6  Overview of translation in which messenger ribonucleic acid (mRNA) serves as the template for

the assembly of amino acids into polypeptides. The three steps include initiation (A), elongation (B and C), and termination (not shown). tRNA, transfer RNA.

control these processes. The cell must regulate gene expression and control the activities of gene products so that a physiologic balance is maintained. Regulation and control are also key factors. These are highly complex mechanisms by which single-cell organisms are able to respond and adapt to environmental challenges, regardless of whether

the challenges occur naturally or result from medical intervention (e.g., antibiotics). Regulation occurs at one of three levels of information transfer from the gene expression and protein synthesis pathway: transcriptional, translational, or posttranslational. The most common is transcriptional regulation. Because

14 PA RT I     Basic Medical Microbiology

direct interactions with genes and their ability to be transcribed to mRNA are involved, transcriptional regulation is also referred to as genetic control. Genes that encode enzymes involved in anabolic processes (biosynthesis) and genes that encode enzymes for catabolic processes (biodegradation) are examples of genetic control. In general, genes that encode anabolic enzymes for the synthesis of particular products are repressed (i.e., are not transcribed and therefore are not expressed) in the presence of the gene end product. This strategy prevents waste and overproduction of products that are already present in sufficient supply. In this system, the product acts as a corepressor that forms a complex with a repressor molecule. In the absence of corepressor product (i.e., gene product), transcription occurs (Fig. 2.7A). When present in sufficient quantity, the product forms a complex with the repressor. The complex then binds to a specific base region of the gene sequence known as the operator region (Fig. 2.7B). This binding blocks RNA polymerase progression from the promoter sequence and inhibits transcription. As the supply of product (corepressor) dwindles, an insufficient amount remains to form a complex with the repressor. The operator region is no longer bound to the repressor molecule. Transcription of the genes for the anabolic enzymes commences and continues until a sufficient supply of end product is again available. In contrast to repression, genes that encode catabolic enzymes are usually induced; that is, transcription occurs when the substrate to be degraded by enzymatic action is present. Production of degradative enzymes in the absence of substrates would be a waste of cellular energy and resources. When the substrate is absent in an inducible system, a repressor binds to the operator sequence of the DNA and blocks transcription of the gene for the degradative enzyme (Fig. 2.7C). In the presence of an inducer, which often is the target substrate for degradation, a complex is formed between the inducer and the repressor that results in the release of the repressor from the operator site, allowing transcription of the genes encoding the specific catabolic enzyme (Fig. 2.7D). Certain genes are not regulated; that is, they are not under the control of inducers or repressors. These genes are referred to as constitutive. Because they usually encode for products that are essential for viability under almost all growth and environmental conditions, these genes are continuously expressed. In addition, not all regulation occurs at the genetic level (i.e., transcriptional regulation). For example, the production of some enzymes may be controlled at the protein synthesis (i.e., translational) level. The activities of other enzymes that have already been synthesized may be regulated at a posttranslational level; that is, certain catabolic or anabolic metabolites may directly interact with enzymes either to increase or to decrease their enzymatic activity. Among different bacteria and even different genes in the same bacterium, the mechanisms by which inducers and corepressors are involved in gene regulation vary widely. Furthermore, bacterial cells have mechanisms to detect

Repression Promotor

Operator

RNA polymerase

Gene 1

Gene 2

Gene 3

Transcription occurs Absence of corepressor (gene product)

Repressor

A Promotor

Operator

RNA Repressor polymerase

Gene 1

Gene 2

Gene 3

Transcription blocked Corepressor (gene product)

B Induction Promotor

Operator

RNA Repressor polymerase

C

Gene 1

Gene 2

Gene 3

Transcription blocked by complex

Absence of substrate (inducer)

Promotor

Operator

RNA polymerase

Gene 1

Gene 2

Gene 3

Transcription occurs

Inducer Repressor

D

Substrate (inducer) present

• Fig. 2.7  Transcriptional control of gene expression. (A and B) Gene repression. (C and D) Induction. RNA, Ribonucleic acid.

environmental changes. These changes can generate signals that interact with the gene expression mechanism, ensuring that appropriate products are made in response to the environmental change. In addition, several complex interactions between different regulatory systems are found within a single cell. Such diversity and interdependence are necessary components of metabolism that allow an organism to respond to environmental changes in a rapid, well-coordinated, and appropriate way. 

Genetic Exchange and Diversity In eukaryotic organisms, genetic diversity is achieved by sexual reproduction, which allows for the mixing of genomes through genetic exchange. Bacteria multiply by simple binary cell division in which two identical daughter

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

cells result by division of one parent cell. Each daughter cell receives the full genetic complement contained in the original parent cell. This process does not allow for the mixing of genes from other cells and leaves no means of achieving genetic diversity among bacterial progeny. Without genetic diversity and change, the essential ingredients for evolution are lost. However, microorganisms have been on earth for billions of years, and microbiologists have witnessed their ability to change as a result of exposure to chemicals (i.e., antibiotics) and environmental conditions (i.e., temperature or oxygenation). It is evident that these organisms are fully capable of evolving and altering their genetic composition. Genetic alterations and diversity in bacteria are accomplished by three basic mechanisms: mutation, genetic recombination, and genetic exchange, with or without recombination. Throughout diagnostic microbiology and infectious diseases, there are numerous examples of the effect these genetic alteration and exchange mechanisms have on clinically relevant bacteria and the management of the infections they cause.

Mutation Mutation is defined as an alteration in the original nucleotide sequence of a gene or genes within an organism’s genome (i.e., a change in the organism’s genotype). This alteration may involve a single DNA base in a gene, an entire gene, or several genes. Mutational changes in the sequence may arise spontaneously, perhaps by an error made during DNA replication. Alternatively, mutations may be induced by mutagens (i.e., chemical or physical factors) in the environment or by biologic factors, such as the introduction of foreign DNA into the cell. Alterations in the DNA base sequence can result in changes in the base sequence of mRNA during transcription. This, in turn, can affect the types and sequences of amino acids that will be incorporated into the protein during translation. Depending on the site and extent of the mutation, various outcomes may affect the physiologic functions of the organism. For example, a mutation may be so devastating that it is lethal to the organism; therefore the mutation “dies” along with the organism. In another instance, the mutation may be silent so that no changes are detected in the organism’s phenotype (i.e., observable properties). Alternatively, the mutation may result in a noticeable alteration in the organism’s phenotype, and the change may provide the organism with a survival advantage. This outcome, in Darwinian terms, is the basis for prolonged survival and evolution. Nonlethal mutations are considered stable if they are passed on from one generation to another as an integral part of the cell’s genotype (i.e., genetic composition). In addition, genes that have undergone stable mutations may also be transferred to other bacteria by one of the mechanisms of genetic exchange. In other instances, the mutation may be lost as a result of cellular repair mechanisms capable of restoring the original genotype and phenotype, or it may be lost spontaneously during subsequent cycles of DNA replication. 

Genetic Recombination Besides mutations, bacterial genotypes can be altered through recombination. In this process, a segment of

15

DNA originating from one bacterial cell (i.e., the donor) enters a second bacterial cell (i.e., the recipient) and is exchanged with a DNA segment of the recipient’s genome. This is also referred to as homologous recombination, because the pieces of DNA that are exchanged usually have extensive homology or similarities in their nucleotide sequences. Recombination involves a number of binding proteins, with the bacterial recombinase protein (RecA) playing a central role (Fig. 2.8A). RecA is capable of binding single-stranded DNA (ssDNA) to the complementary dsDNA, providing a mechanism for DNA repair and recombination to occur. After recombination, the recipient DNA consists of one original, unchanged strand and a second strand from the donor DNA fragment that has been recombined. Recombination is a molecular event that occurs frequently in many varieties of bacteria, including most of the clinically relevant species, and it may involve any portion of the organism’s genome. However, the recombination event may go unnoticed unless the exchange of DNA results in a distinct alteration in the phenotype. Nonetheless, recombination is a major means by which bacteria may achieve genetic diversity. 

Genetic Exchange An organism’s ability to undergo recombination depends on the acquisition of “foreign” DNA from a donor cell. The three mechanisms by which bacteria physically exchange DNA are transformation, transduction, and conjugation. Transformation Transformation involves recipient cell uptake of naked (free) DNA released into the environment when another bacterial cell (i.e., the donor) dies and undergoes lysis (Fig. 2.8B). This genomic DNA exists as fragments in the environment. Certain bacteria are able to take up naked DNA from their surroundings; that is, they are able to undergo transformation. Such bacteria are said to be competent. Among the bacteria that cause human infections, competence is a characteristic commonly associated with members of the genera Haemophilus, Streptococcus, and Neisseria. Once the donor DNA, usually as a single strand, gains access to the interior of the recipient cell, recombination with the recipient’s homologous DNA can occur. The mixing of DNA between bacteria via transformation and recombination plays a major role in the development of antibiotic resistance and in the dissemination of genes that encode factors essential to an organism’s ability to cause disease. In addition, genetic exchange by transformation is not limited to organisms of the same species, thus allowing important characteristics to be disseminated to a greater variety of medically important bacteria.  Transduction Transduction is a second mechanism by which DNA from two bacteria may come together in one cell, thus allowing for recombination (Fig. 2.8C). This process is mediated

16 PA RT I     Basic Medical Microbiology

A Recombination Rec A protein

Recipient DNA

Uptake of donor ("foreign") DNA

Alignment of donor DNA with homologous recipient DNA

Recombined DNA fragment (blue)

B Transformation Recipient

Donor Free DNA

Cell lysis and release of free DNA

Uptake and recombination

C Transduction

Donor cell DNA packaged in bacteriophage

Release of bacteriophage from donor cell

Bacteriophage infects and releases donor DNA

D Conjugation: Chromosome transfer Recipient (Final)

Recipient

Donor

Transfer of newly synthesized chromosomal DNA mobilized through intercellular bridge

E Conjugation: Plasmid transfer Recipient

Donor

Chromosome

Recipient (Final)

Plasmid

Transfer of newly synthesized plasmid DNA through intercellular bridge

• Fig. 2.8  (A) Genetic recombination. The mechanisms of genetic exchange between bacteria are transformation (B), transduction (C), and conjugational transfer of chromosomal (D) and plasmid (E) deoxyribonucleic acid (DNA).

through viruses capable of infecting bacteria (i.e., bacteriophages). In their “life cycle,” these viruses integrate their DNA into the bacterial cell’s chromosome, where viral DNA replication and expression occur. When the production of viral products is complete, viral DNA is excised (cut) from the bacterial chromosome and packaged within a protein coat. The excision process is not always accurate, resulting in the removal of genetic material that contains both the bacterial and viral DNA. The newly formed recombinant virion (virus particle), along with the additional multiple virions, is released when the infected bacterial cell lyses. The bacterial DNA may be randomly incorporated with viral DNA (generalized transduction), or it may be

incorporated along with specific adjacent viral DNA (specialized transduction). In generalized transduction, the viral DNA is inserted randomly into any area of the bacterial genome. However, in specialized transduction, the virus inserts into particular genes in an organism based on sequence specificity and resulting in a higher frequency of genetic material in those regions being transferred through recombination. In either case, when the virus infects another bacterial cell, it releases its DNA, which includes the previously incorporated bacterial donor DNA. The newly infected cell is then the recipient of donor DNA introduced by the bacteriophage, and recombination between DNA from two different cells occurs. 

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

•

Fig. 2.9  Photomicrograph of an Escherichia coli sex pilus between a donor and a recipient cell. (From Brock TD, Madigan M, Martinko J, et al, eds. Biology of Microorganisms. Upper Saddle River, NJ: Prentice Hall; 2009.)

Conjugation

The third mechanism of DNA exchange between bacterial cells is conjugation. This process involves cell-to-cell contact and requires mobilization of the donor bacterium’s chromosome or other mobile genetic element. The nature of intercellular contact is not well characterized in all bacterial species capable of conjugation. However, in E. coli, contact is mediated by a sex pilus (Fig. 2.9). The sex pilus originates from the donor and establishes a conjugative bridge that serves as the conduit for DNA transfer from donor to recipient cell. With intercellular contact established, mobilization of the genetic element is undertaken and involves DNA synthesis. One new DNA strand is produced by the donor and is passed to the recipient (Fig. 2.8D). The amount of DNA transferred depends on how long the cells are able to maintain contact, but usually only portions of the donor molecule are transferred. In any case the newly introduced DNA is then available to recombine with the recipient’s genome. In addition to chromosomal DNA, genes encoded in extrachromosomal genetic elements, such as plasmids and transposons, may be transferred by conjugation (Fig. 2.8E). Not all plasmids are capable of conjugative transfer, but for those that are, the donor plasmid usually is replicated so that the donor retains a copy of the plasmid transferred to the recipient. (See the discussion of the F plasmid in the section Cellular Appendages, later in the chapter.) Plasmid DNA may also become incorporated into the host cell’s chromosome. In contrast to plasmids, most transposons do not exist independently in the cell. Except when they are moving from one location to another, many transposons must be incorporated into the chromosome, plasmids, or both. These elements are often referred to as “jumping genes” because of their ability to change location within and even between the genomes of bacterial cells. Transposition is the process by which these genetic elements excise from one genomic location and insert into another. Transposons carry genes that have products that help to mediate the transposition process, in addition to genes that encode for other accessory characteristics, such as antimicrobial resistance.

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Homologous recombination between the genes of plasmids or transposons and the host bacterium’s chromosomal DNA may occur. Plasmids and transposons play a key role in genetic diversity and the dissemination of genetic information among bacteria. Many characteristics that significantly alter the activities of clinically relevant bacteria are encoded and disseminated on these elements. Furthermore, as shown in Fig. 2.10, the variety of strategies that bacteria can use to mix and match genetic elements provides them with a tremendous capacity to genetically adapt to environmental changes, including those imposed by human medical practices. A good example of this is the emergence and widespread dissemination of resistance to antimicrobial agents among clinically important bacteria. Bacteria have used their capacity for disseminating genetic information to establish resistance to many of the commonly prescribed antibiotics. (See Chapter 10 for more information about antimicrobial resistance mechanisms.) 

Bacterial Metabolism Fundamentally, bacterial metabolism involves all the cellular processes required for the organism’s survival and replication. Familiarity with bacterial metabolism is essential to understand bacterial interactions with human host cells, the mechanisms bacteria use to cause disease, and the basis of diagnostic microbiology (i.e., the tests and strategies used for laboratory identification of infectious organisms). Because metabolism is an extensive and complicated topic, this section focuses on processes typical of medically relevant bacteria. For the sake of clarity, metabolism is discussed in terms of four primary, but interdependent, processes: fueling, biosynthesis, polymerization, and assembly (Fig. 2.11).

Fueling Fueling is considered the utilization of metabolic pathways involved in the acquisition of nutrients from the environment, production of precursor metabolites, and energy production.

Acquisition of Nutrients Bacteria use various strategies for obtaining essential nutrients from the external environment and transporting these substances into the cell’s interior. For nutrients to be internalized, they must cross the bacterial cell wall and membrane. These complex structures help to protect the cell from environmental insults, maintain intracellular equilibrium, and transport substances into and out of the cell. Although some key nutrients (e.g., water, oxygen, and carbon dioxide) enter the cell by simple diffusion across the cell membrane, the uptake of other substances is controlled by membrane-selective permeability; still other substances use specific transport mechanisms. Active transport is among the most common methods used for the uptake of nutrients such as certain sugars, most

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Donor organism Chromosome Plasmids Transposon

Potential for subsequent dissemination of plasmids and transposons to a variety of other recipients

Donor

Recipient

• Fig. 2.10  Pathways for bacterial dissemination of plasmids and transposons, together and independently.

amino acids, organic acids, and many inorganic ions. The mechanism, driven by an energy-dependent pump, involves carrier molecules embedded in the membrane portion of the cell structure. These carriers combine with the nutrients, transport them across the membrane, and release them inside the cell. Group translocation is another transport mechanism that requires energy but differs from active transport in that the nutrients being transported undergo chemical modification. Many sugars, purines, pyrimidines, and fatty acids are transported by this mechanism. 

Production of Precursor Metabolites Once inside the cell, many nutrients serve as the raw materials from which precursor metabolites for subsequent biosynthetic processes are produced. These metabolites, listed in Fig. 2.11, are produced through two central pathways: the Embden-Meyerhof-Parnas (EMP) pathway (glycolysis) and the tricarboxylic acid (TCA) cycle. The two major pathways and their relationship to one another are shown in Fig. 2.12; not shown are the alternative pathways (e.g., the Entner-Doudoroff and the pentose phosphate pathway) that play key roles in redirecting and replenishing the precursors as they are used in subsequent processes. The EntnerDoudoroff pathway catalyzes the degradation of gluconate and glucose. The gluconate is phosphorylated, dehydrated, and converted into pyruvate and glyceraldehyde, leading to ethanol production. Alternatively, the pentose phosphate pathway uses glucose to produce reduced nicotinamide adenine dinucleotide phosphate (NADPH), pentoses, and tetroses for biosynthetic reactions such as nucleoside and amino acid synthesis.

The production efficiency of a bacterial cell resulting from these precursor-producing pathways can vary substantially, depending on the growth conditions and availability of nutrients. This is an important consideration because the accurate identification of medically important bacteria has traditionally depended on methods that measure the presence of products and byproducts of these metabolic pathways. 

Energy Production The third type of fueling pathway is one that produces the energy required for nearly all cellular processes, including nutrient uptake and precursor production. Energy production is accomplished by the breakdown of chemical substrates (i.e., chemical energy) through the degradative process of catabolism coupled with oxidation-reduction reactions. In this process, the energy source molecule (i.e., substrate) is oxidized as it donates electrons to an electron-acceptor molecule, which is then reduced. The transfer of electrons is mediated through carrier molecules, such as nicotinamide-adenine-dinucleotide (NAD+) and nicotinamide-adenine-dinucleotide-phosphate (NADP+). The energy released by the oxidation-reduction reaction is transferred to phosphate-containing compounds, where high-energy phosphate bonds are formed. ATP is the most common of such molecules. The energy contained in this compound is eventually released by the hydrolysis of ATP under controlled conditions. The release of this chemical energy, coupled with enzymatic activities, specifically catalyzes each biochemical reaction in the cell and drives cellular reactions.

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

Precursor metabolites • Glucose 6-phosphate • Fructose 6-phosphate • Pentose 5-phosphate • Erythrose 4-phosphate • 3-Phosphoglycerate • Phosphoenolpyruvate • Pyruvate • Acetyl CoA • α-Ketoglutarate • Succinyl CoA • Oxaloacetate Biosynthetic reactions

Metabolic reactions Precursor metabolites

Assembly reactions

Polymerizations Lipid

Inclusions

Fatty acids Lipopolysaccharide

Metabolic energy

Glucose

Sugars

Glycogen

Envelope Flagella

Murein Pili

Amino acids

Nutrients

Protein Cytosol RNA Polyribosomes

Nucleotides DNA Metabolic products

Building blocks

Macromolecules

Nucleoid Structures

Nutrients • Gases Carbon dioxide (CO2) Oxygen (O2) Ammonia (NH3) • Organic compounds, including amino acids • Water (H2O) • Nitrate (NO3-) • Phosphate (PO43-) • Hydrogen sulfide (H2S) • Sulfate (SO42-) • Potassium (K+) • Magnesium (Mg2+) • Calcium (Ca2+) • Sodium (Na+) • Iron (Fe3+) Organic iron complexes

• Fig. 2.11  Overview of bacterial metabolism, which includes the processes of fueling, biosynthesis, polym-

erization, and assembly. CoA, Coenzyme A; DNA, deoxyribonucleic acid; RNA, ribonucleic acid. (Modified from Niedhardt FC, Ingraham JL, Schaechter M, eds. Physiology of the Bacterial Cell: A Molecular Approach. Sunderland, MA: Sinauer Associates; 1990.)

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Glucose

NADPH2

P Glucose 6-phosphate

6-Phosphogluconolactone

6-Phosphogluconate NADPH2 Pentose 5-phosphate*

Fructose 6-phosphate P

Erythrose 4-phosphate

Fructose 1,6-diphosphate Pentose phosphate cycle Triose 3-phosphate FADH2 NADH2 1,3-Diphosphoglycerate

Succinate

Fumarate

P

P

3-Phosphoglycerate P Malate

2-Phosphoglycerate

NADH2 Phosphoenolpyruvate P

Succinyl CoA

TCA cycle

NADH2

P

α-Ketoglutarate

Oxaloacetate

NADH2

PYRUVATE

NADPH2 Acetyl CoA

Citrate

Isocitrate

EMP Pathway

• Fig. 2.12  Overview of the central metabolic pathways (Embden-Meyerhof-Parnas [EMP], the tricarboxylic

acid [TCA] cycle, and the pentose phosphate shunt). Precursor metabolites (Fig. 2.11) that are produced are highlighted in red; production of energy in the form of adenosine triphosphate (〜P) by substrate-level phosphorylation is highlighted in yellow; and reduced carrier molecules for transport of electrons used in oxidative phosphorylation are highlighted in green. (Modified from Niedhardt FC, Ingraham JL, Schaechter M, eds. Physiology of the Bacterial Cell: A Molecular Approach. Sunderland, MA: Sinauer Associates; 1990.)

The two general mechanisms for ATP production in bacterial cells are substrate-level phosphorylation and electron transport, also referred to as oxidative phosphorylation. In substrate-level phosphorylation, high-energy phosphate bonds produced by the central pathways are donated to adenosine diphosphate (ADP) to form ATP directly from the substrate as opposed to generation via the electron transport chain (Fig. 2.12). In addition, pyruvate, a primary intermediate in the central pathways, serves as the initial substrate for several other pathways to generate ATP by substrate-level phosphorylation. These other pathways constitute fermentative metabolism, which does not require oxygen and produces various end products, including alcohols, acids, carbon dioxide, and hydrogen. The specific fermentative pathways and the end products produced vary with different bacterial species. Detection of these

products is an important basis for laboratory identification of bacteria. (See Chapter 7 for more information on the biochemical basis for bacterial identification.) Oxidative Phosphorylation

Oxidative phosphorylation involves an electron transport system that conducts a series of electron transfers from reduced carrier molecules such as NADH2, NADPH2, and FADH2 (flavin adenine dinucleotide), produced in the central pathways (Fig. 2.12), to a terminal electron acceptor. The energy produced by the series of oxidation-reduction reactions is used to generate ATP from ADP. When oxidative phosphorylation uses oxygen as the terminal electron acceptor, the process is known as aerobic respiration. Anaerobic respiration refers to processes that use final electron acceptors other than oxygen.

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

A knowledge of which mechanisms bacteria use to generate ATP is important for designing laboratory protocols for cultivating and identifying these organisms. For example, some bacteria depend solely on aerobic respiration and are unable to grow in the absence of oxygen (strictly aerobic bacteria). Others can use either aerobic respiration or fermentation, depending on the availability of oxygen (facultative anaerobic bacteria). For still others, oxygen is absolutely toxic (strictly anaerobic bacteria). 

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cell types share many common features, they have important differences in terms of structure, metabolism, and genetics.

Eukaryotic and Prokaryotic Cells

The fueling reactions essentially bring together all the raw materials needed to initiate and maintain all other cellular processes. The production of precursors and energy is accomplished through catabolic processes and the degradation of substrate molecules. The three remaining pathways for biosynthesis, polymerization, and assembly depend on anabolic metabolism. In anabolic metabolism, precursor compounds are joined for the creation of larger molecules (polymers) required for assembly of cellular structures (Fig. 2.11). Biosynthetic processes use the precursor products in dozens of pathways to produce a variety of building blocks, such as amino acids, fatty acids, sugars, and nucleotides (Fig. 2.11). Many of these pathways are highly complex and interdependent, whereas other pathways are completely independent. In many cases the enzymes that drive the individual pathways are encoded on a single mRNA molecule that has been transcribed from contiguous genes in the bacterial chromosome (i.e., an operon). As previously mentioned, bacterial genera and species vary extensively in their biosynthetic capabilities. Knowledge of these variations is necessary to determine the optimal conditions for growing organisms under laboratory conditions. For example, some organisms may not be capable of synthesizing an essential amino acid necessary as a building block for proteins. Without the ability to synthesize the amino acid, the bacterium must obtain the building block from the environment. Thus, if the organism is cultivated in the microbiology laboratory, the amino acid must be provided in the artificial culture medium. 

Among clinically relevant organisms, bacteria are single-cell prokaryotic microorganisms. Fungi and parasites are single-cell or multicellular eukaryotic organisms, as are plants and all higher animals. Viruses are dependent on host cells for survival and therefore are not considered cellular organisms but rather infectious agents. Prions, which are abnormal infectious proteins, are also not considered living cells. A notable characteristic of eukaryotic cells, such as parasites and fungi, is the presence of membrane-enclosed organelles that have specific cellular functions. Examples of these organelles and their respective functions include: • Endoplasmic reticulum—process and transport proteins • Golgi body—modification of substances and transport throughout the cell, including internal delivery of molecules, and exocytosis or secretion of other molecules • Mitochondria—generate energy (ATP) • Lysosomes—provide an environment for controlled enzymatic degradation of intracellular substances • Nucleus—provide a membrane enclosure for chromosomes In addition, eukaryotic cells have an infrastructure, or cytoskeleton, which provides support for cellular structure, organization, and movement. The cytoskeleton in eukaryotic cells also plays an essential role in immunology by mediating phagocytosis for the removal of foreign materials from the host, including bacteria, fungi, and viral agents. Prokaryotic cells, such as bacteria, do not contain organelles. All functions take place in the cytoplasm or cytoplasmic membrane of the cell. Prokaryotic and eukaryotic cell types differ considerably at the macromolecular level, including protein synthesis machinery, chromosomal organization, and gene expression. One notable structure present only in prokaryotic bacterial cells is a cell wall composed of peptidoglycan. This structure has an immeasurable effect on the practice of diagnostic bacteriology and the management of bacterial diseases. 

Polymerization and Assembly

Bacterial Morphology

Various anabolic reactions assemble (polymerize) the building blocks into macromolecules, including lipids, lipopolysaccharides, polysaccharides, proteins, and nucleic acids. This synthesis of macromolecules is driven by energy and enzymatic activity in the cell. Similarly, energy and enzymatic activities also drive the assembly of various macromolecules into the component structures of the bacterial cell. Cellular structures are the product of all the genetic and metabolic processes discussed. 

Most clinically relevant bacterial species range in size from 0.25 to 1 μm in width and 1 to 3 μm in length, thus requiring microscopy for visualization (see Chapter 6 for more information on microscopy). Just as bacterial species and genera vary in their metabolic processes, their cells also vary in size, morphology, and cell-to-cell arrangements and in the chemical composition and structure of the cell wall. The bacterial cell wall differences provide the basis for the Gram stain, a fundamental staining technique used in bacterial identification schemes. This staining procedure separates almost all medically relevant bacteria into two general types: gram-positive bacteria, which stain a deep blue or purple, and gram-negative bacteria, which stain a pink to red (Fig. 6.3). This simple

Biosynthesis

Structure and Function of the Bacterial Cell Based on key characteristics, all cells are classified into two basic types: prokaryotic and eukaryotic. Although these two

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Flagellum

Pilus

Lipopolysaccharide Porin

Capsule (variable) Outer membrane

L-ring

Murein Periplasmic space

Basal body rings

P-ring S-ring M-ring C-ring

Cytoplasmic membrane

Gram-positive

Gram-negative

• Fig. 2.13  General structures of the gram-positive and gram-negative bacterial cell envelopes. The outer

membrane and periplasmic space are present only in the envelope of gram-negative bacteria. In addition to porins, bacterial membranes contain additional proteins involved in stabilizing the layers of the cellular structure, adherence, or sorting and reacting to chemical signals. The murein layer is substantially more prominent in gram-positive envelopes. (Modified from Niedhardt FC, Ingraham JL, Schaechter M, eds. Physiology of the Bacterial Cell: A Molecular Approach. Sunderland, MA: Sinauer Associates; 1990.)

but important color distinction is the result of differences in the constituents of bacterial cell walls that influence the cell’s ability to retain differential dyes after treatment with a decolorizing agent. Common bacterial cellular morphologies include cocci (circular), coccobacilli (ovoid), and bacilli (rod shaped), as well as fusiform (pointed end), curved, or spiral shapes. Cellular arrangements are also noteworthy. Cells may characteristically occur singly, in pairs, or grouped as tetrads, clusters, or in chains (see Fig. 6.4 for examples of bacterial staining and morphologies). The determination of the Gram stain reaction and the cell size, morphology, and arrangement are essential aspects of bacterial identification. 

Bacterial Cell Components Bacterial cell components can be divided into those that make up the outer cell structure and its appendages (cell envelope) and those associated with the cell’s interior. It is important to note that the cellular structures work together to function as a complex and integrated unit.

Cell Envelope As shown in Fig. 2.13, the outermost structure, the cell envelope, comprises: • An outer membrane (in gram-negative bacteria only) • A cell wall composed of the peptidoglycan macromolecule (also known as the murein layer)

• P  eriplasm (in gram-negative bacteria only) • The cytoplasmic or cell membrane, which encloses the cytoplasm Outer Membrane

Outer membranes, which are found only in gram-negative bacteria, function as the cell’s initial barrier to the environment. These membranes serve as the primary permeability barriers to hydrophilic and hydrophobic compounds and contain essential enzymes and other proteins located in the periplasmic space. The membrane is a bilayered structure composed of lipopolysaccharide, which gives the surface of gram-negative bacteria a net negative charge. The outer membrane also plays a significant role in the ability of certain bacteria to cause disease. Scattered throughout the lipopolysaccharide macromolecules are protein structures called porins. These water-filled structures control the passage of nutrients and other solutes, including antibiotics, through the outer membrane. The number and types of porins vary with bacterial species. These differences can substantially influence the extent to which various substances pass through the outer membranes of different bacteria. In addition to porins, other proteins (murein lipoproteins) facilitate the attachment of the outer membrane to the next internal layer in the cell envelope, the cell wall, and may serve as adhesions for attachment to a host cell or as transporters. 

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

NAM NAM NAM NAM NAG NAG NAG NAG Peptide bridge

A

NAM NAM NAM NAM NAG NAG NAG NAG

NAM NAM NAM NAM NAG NAG NAG NAG

CH2 OH O (NAG) OH CH2 OH O NH O (NAM) OH CH2 OH C= O CH2 OH O O O CH3 (NAM) NH O OH (NAG) OH CH2 OH C= O O NH O O NH CH (NAG) 3 O OH C= O C= O HC CH3 NH CH3 CH3 O C= O C= O H C CH3 L-Alanine CH3 Amino D-Glutamate C= O acid L-Alanine Diaminopimelate chain D-Glutamate D-Alanine Peptide Diaminopimelate bridge D-Alanine B

• Fig. 2.14  Peptidoglycan sheet (A) and subunit (B) structure. Multiple

peptidoglycan layers compose the murein structure, and different layers are extensively cross-linked by peptide bridges. Note that amino acid chains are only derived from NAM. NAG, N-acetylglucosamine; NAM, N-acetylmuramic acid. (Modified from Saylers AA, Whitt DD. Bacterial Pathogenesis: A Molecular Approach. Washington, DC: American Society for Microbiology Press; 2010.)

Cell Wall (Murein Layer)

The cell wall, also referred to as the peptidoglycan, or murein layer, is an essential structure found in nearly all clinically relevant bacteria. This structure gives the bacterial cell shape and strength to withstand changes in environmental osmotic pressures that would otherwise result in cell lysis. The murein layer protects against mechanical disruption of the cell and offers some barrier to the passage of larger substances. Because this structure is essential for the survival of bacteria, its synthesis and structure are often the primary target for the development and design of several antimicrobial agents. The structure of the cell wall is unique and is composed of disaccharide-pentapeptide subunits. The disaccharides N-acetylglucosamine and N-acetylmuramic acid are the alternating sugar components (moieties) with the amino acid chain linked to N-acetylmuramic acid molecules (Fig. 2.14). Polymers of these subunits cross-link to one another by means of peptide bridges to form peptidoglycan sheets. In turn, layers of these sheets are cross-linked with one another, forming a multilayered, cross-linked structure of considerable strength. Referred to as the murein sacculus, or sack, this peptidoglycan structure surrounds the entire cell.

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A notable difference between the cell walls of gram-positive and gram-negative bacteria is the substantially thicker peptidoglycan layer in gram-positive bacteria (Fig. 2.13). In addition, the cell wall of gram-positive bacteria contains teichoic acids (i.e., glycerol or ribitol phosphate polymers combined with various sugars, amino acids, and amino sugars). Some teichoic acids are linked to N-acetylmuramic acid, and others (e.g., lipoteichoic acids) are linked to the next underlying layer, the cellular or cytoplasmic membrane. Other bacteria (e.g., Mycobacteria) have waxy substances within the murein layer, such as mycolic acids. Mycolic acids make the cells more refractory to toxic substances, including acids. Bacteria with mycolic acid in the cell wall require unique staining procedures and growth media in the diagnostic laboratory.  Periplasmic Space

The periplasmic space typically is found only in gramnegative bacteria (whether it is present in gram-positive organisms is a subject of debate). The periplasmic space is bounded by the internal surface of the outer membrane and the external surface of the cellular membrane. This area, which contains the murein layer, consists of gel-like substances that assist in the capture of nutrients from the environment. This space also contains several enzymes involved in the degradation of macromolecules and detoxification of environmental solutes, including antibiotics that enter through the outer membrane.  Cytoplasmic (Inner) Membrane

The cytoplasmic (inner) membrane is present in both grampositive and gram-negative bacteria and is the deepest layer of the cell envelope. The cytoplasmic membrane is heavily laced with various proteins, including a number of enzymes vital to cellular metabolism. The cell membrane serves as an additional osmotic barrier and is functionally similar to the membranes of several eukaryotic cellular organelles (e.g., mitochondria, Golgi complexes, lysosomes). The cytoplasmic membrane functions include: • Transport of solutes into and out of the cell • Housing of enzymes involved in outer membrane synthesis, cell wall synthesis, and the assembly and secretion of extracytoplasmic and extracellular substances • Generation of chemical energy (i.e., ATP) • Cell motility • Mediation of chromosomal segregation during replication • Housing of molecular sensors that monitor chemical and physical changes in the environment  Cellular Appendages

In addition to the components of the cell envelope, cellular appendages (i.e., capsules, fimbriae, and flagella) are associated with or proximal to this portion of the cell. The presence of these appendages, which can play a role in the mediation of infection and in laboratory identification, varies among bacterial species and even among strains within the same species.

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The capsule is immediately exterior to the murein layer of gram-positive bacteria and the outer membrane of gramnegative bacteria. The capsule is composed of high-molecular-weight polysaccharides, the production of which may depend on the environment and growth conditions surrounding the bacterial cell. The capsule does not function as an effective permeability barrier or add strength to the cell envelope, but it does protect bacteria from attack by components of the human immune system. The capsule also facilitates and maintains bacterial colonization of biologic (e.g., teeth) and inanimate (e.g., prosthetic heart valves) surfaces through the formation of “slime layers” or biofilms. Both slime layers and biofilms imply the presence of an extracellular polymer matrix that varies in composition and structure in different organisms. A biofilm may consist of a monomicrobic or polymicrobic group of bacteria housed in a complex biochemical matrix. This extracellular matrix stabilizes the cell to protect the organism from hydrodynamic forces in the host and plays a protective role against biocides and agents of the host’s immune system. (See Chapter 3 for further discussion of microbial biofilms.) Fimbriae, or pili, are hairlike, proteinaceous structures that extend from the cell membrane into the external environment; some may be up to 2 μm long. Fimbriae may serve as adhesins that help bacteria attach to animal host cell surfaces, often as the first step in establishing infection. In addition, a pilus may be referred to as a sex pilus; this structure, which is well characterized in the gram-negative bacillus E. coli, serves as the conduit for the passage of DNA from the donor to the recipient during conjugation. The sex pilus is present only in cells that produce a protein referred to as the F factor. F-positive cells initiate mating or conjugation only with F-negative cells, thereby limiting the conjugative process to cells capable of transporting genetic material through the hollow sex pilus. Flagella are complex structures, mostly composed of the protein flagellin, that are intricately embedded in the cell envelope. These structures are responsible for bacterial motility. Although not all bacteria are motile, motility plays an important role in survival and the ability of bacteria to cause disease. Depending on the bacterial species, a single flagellum may be located at one end of the cell (monotrichous flagella), a group of flagella may be located at one end of the cell (lophotrichous flagella), a single flagellum may reside at both ends of the cell (amphitrichous flagella), or the entire cell surface may be covered with flagella (peritrichous flagella). The flagellum acts as a rotary motor containing a complex set of rings that act as bushings to control cellular movement. Gram-negative flagella are equipped with a basal body structure that contains five rings, the L-ring that is embedded in the lipid bilayer, the P-ring in the periplasmic space, a smaller S-ring (stator ring) attached to the M-ring or motor ring, and the C-ring, which anchors the entire complex to the cell. Because grampositive organisms have a much more stable complex cellular structure because of the thick layer of peptidoglycan, the flagella contain only two basal body rings: One is embedded

in the peptidoglycan layer, which is very stable, and the second is embedded in the cell membrane. 

Cell Interior Those structures and substances that are bound internally by the cytoplasmic membrane compose the cell interior and include the cytosol, polysomes, inclusions, nucleoid, plasmids, and endospores. The cytosol, where nearly all other functions not conducted by the cell membrane occur, contains thousands of enzymes and is the site of protein synthesis. The cytosol has a granular appearance caused by the presence of many polysomes (mRNA complexed with several ribosomes during translation and protein synthesis) and inclusions (i.e., storage reserve granules). The number and nature of the inclusions vary depending on the bacterial species and the nutritional state of the organism’s environment. Two common types of granules include glycogen, a storage form of glucose, and polyphosphate granules, a storage form for inorganic phosphates. These granules may be microscopically visible in bacteria stained with specific dyes. Unlike eukaryotic chromosomes, the bacterial chromosome is not enclosed within a membrane-bound nucleus. Instead the bacterial chromosome exists as a nucleoid in which the highly coiled DNA is intermixed with RNA, polyamines, and various proteins that lend structural support. At times, depending on the stage of cell division, more than one chromosome may be present per bacterial cell. Plasmids are the other genetic elements that exist independently in the cytosol, and their numbers may vary from none to several hundred per bacterial cell. The final bacterial structure to be considered is the endospore. Under adverse physical and chemical conditions or when nutrients are scarce, some bacterial genera (Bacillus and Clostridium spp.) are able to form spores (i.e., sporulate). Sporulation involves substantial metabolic and structural changes in the bacterial cell. Essentially, the cell transforms from an actively metabolic and growing state to a dormant state, with a decrease in cytosol and a concomitant increase in the thickness and strength of the cell envelope. The endospore remains in a dormant state until favorable conditions for growth are again encountered. This survival tactic is demonstrated by a number of clinically relevant bacteria and complicates thorough sterilization of materials and food for human use.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

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CHAPTER 2  Bacterial Genetics, Metabolism, and Structure

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Nuss AM, Heroven AK, Waldmann B, et al.: Transcriptomic profiling of Yersinia pseudotuberculosis reveals programming of the Crp regulon by temperature and uncovers Crp as a master regulator of small RNAs, PLoS Genet 11:1–26, 2015. Ryan KJ, editor: Sherris medical microbiology: an introduction to infectious diseases, Norwalk, CT, 2003, McGraw-Hill Medical. Saylers AA, Wilson BA, Whitt DD, Winkler ME: Bacterial pathogenesis: a molecular approach, Washington, DC, 2010, American Society for Microbiology Press. Schomburg D, Gerhard M: Biochemical pathways: an atlas of biochemistry and molecular biology, ed 2, New York, 2012, Wiley. Stortchevoi A, Varshney U, RajBhandary UL: Common location of determinants in initiator transfer RNAs for initiator-elongator discrimination in bacteria and in eukaryotes, J Biol Chem 278(20):17672, 2003. Zhurina MV, Gannesen AV, Zdorovenko EL, Plakunov VK: Composition and functions of the extracellular polymer matrix of bacterial biofilms, Microbiology 83:713–722, 2014.

CHAPTER 2  Bacterial Genetics, Metabolism, and Structure 25.e1

Chapter Review 1. The periplasmic space is required for: a. Nutrient collection in both gram-positive and gramnegative bacteria b. Collection and enzymatic degradation of nutrients in gram-negative bacteria c. Nutrient detoxification and enzymatic degradation in all bacteria d. None of the above 2. Prokaryotic chromosomes: a. Are double-stranded RNA molecules b. Are single-copy, double-stranded DNA molecules c. Are linear double-stranded DNA molecules d. Are unable to replicate independently of plasmids 3. Bacterial cells genetically evolve by: a. Recombination with plasmids, transposons, and other bacterial chromosomes b. Mutation and recombination c. Use of the mechanisms of transduction, transformation, and conjugation d. All of the above 4. Transcription is the: a. Copying of DNA to RNA b. Changing of DNA to RNA c. Production of a complementary DNA d. Completion of a protein sequence 5. A eukaryotic cell: a. Is smaller and less complex than a prokaryotic cell b. Is able to grow only in aerobic conditions c. Contains membrane-bound organelles d. Is unable to grow outside of another cell 6. Matching: Match each term with the correct description. _____ capsule _____ replication _____ repressor _____ tRNA _____ facultative anaerobe _____ gram-negative _____ gram-positive _____ aerobic _____ cell envelope _____ mobilome _____ genome

a. involved in transcriptional regulation b. able to grow in the presence or absence of oxygen c. maintains selective permeability and cell shape d. provides a mechanism to evade the human immune system e. the process of making a new DNA molecule f. involved in protein translation g. mobile genetic elements h. contains a thick layer of peptidoglycan i. final electron acceptor is oxygen j. has an outer and inner membrane k. all genetic elements within a cell

7. Which chemical or physical property is essential for the conservation of genetic information? a. Complementation between base pairs b. Double-stranded c. Antiparallel structure d. All are equally important 8. Expression of a biochemical molecule in an organism requires: a. Replication only b. Transcription only c. Transcription and translation of a protein d. All of the above 9.  True or False _____ All bacteria are considered competent. _____ Conjugation requires cell-to-cell contact. _____ Oxidative phosphorylation occurs across the cell membrane in bacteria. 10.  Short Answer Provide an explanation for why bacteria are capable of rapidly responding to changes in their environment based on molecular and cellular structure. Bacteria are prokaryotes; the genetic material is not contained within a nucleus, allowing replication transcription and translation to occur simultaneously.

3

Host-Microorganism Interactions OBJECTIVES 1. List the various reservoirs (environments) that facilitate host-microorganism interactions. 2. Define direct versus indirect transmission, and provide examples of each. 3. Define and differentiate the interactions between the host and microorganism, including colonization, infection, microbiota, microbiome, pathogens, opportunistic pathogens, and nosocomial (health care–acquired or –associated) and community-acquired infection. 4. List and describe the components involved in specific versus nonspecific immune defenses, including inflammation, phagocytosis, antibody production, and cellular responses. 5. Identify elements involved in the two arms of the immune system: humoral and cell-mediated immunity. 6. Provide specific examples of disease prevention strategies, including preventing transmission, controlling reservoirs, and minimizing risk of exposure. 7. Differentiate between bacterial endotoxins and exotoxins, and provide examples of each. 8. Given a patient history of an infectious process, identify and differentiate a sign versus a symptom. 9. Define and differentiate between an acute infectious process and one that is chronic and/or latent. 10. Describe the three major steps in the formation of a microbial biofilm, and list the advantages of biofilm formation to the microorganism and the disadvantages to the infected host.

I

nteractions between humans and microorganisms are exceedingly complex and far from being completely understood. The interactions between these two living entities plays an important role in the practice of diagnostic microbiology and in the management of infectious disease. Understanding these interactions is necessary for establishing methods to isolate specific microorganisms from patient specimens and for developing effective treatment strategies. This chapter provides the framework for understanding the various aspects of host-microorganism interactions. Box 3.1 lists a variety of terms and definitions associated with hostmicroorganism interactions. Host-microorganism interactions should be viewed as bidirectional in nature. Humans use the abilities and natural 26

products of microorganisms in various settings, including the food and fermentation industry, as biologic insecticides for agriculture; to genetically engineer a multitude of products; and even for biodegrading industrial waste. However, microbial populations share the common goal of survival with humans, using their relationship with humans for food, shelter, and dissemination, and they have been successful at achieving those goals. Which participant in the relationship is the user and which is the used is a fine and intricate balance of nature. This is especially true when considering the microorganisms most closely associated with humans and human disease. In 2008, the National Institutes of Health initiated a project referred to as the Human Microbiome Project (http: //commonfund.nih.gov/hmp/index). The human microbiome consists of microorganisms that are present in and on the human body at any given time without causing harm. Phase I (2008–2012) of the microbiome project focused on four major goals: (1) identify and characterize a core human microbiome in healthy individuals, both male and female; (2) determine whether changes in the human microbiome correlate with health and disease; (3) develop new technology and bioinformatic tools to manage the project data; and (4) address the ethical, legal, and social implications associated with the microbiome project. Interestingly, the study has elucidated that the microbiome complex ecosystem varies significantly across the body and between individuals. Analysis of the human microbiome has demonstrated that it is clearly an emergent property. One hundred and thirteen females were examined for the presence of microorganisms at 18 body sites and 129 males at 15 body sites (excluding vaginal collections). The data demonstrated that dependent on the body site, both low diversity of microorganisms and a high diversity of microorganisms correlates with the development of disease. Phase I also examined the relationships between the microbiome and characteristics of the host, including age, body mass index, and available medical history. The second phase of the project, the Integrative Human Microbiome Project, has begun to analyze data from phase I and apply it to host interactions in healthy and disease states. Established in 2014, Phase II focuses on three major areas: (1) the vaginal

CHAPTER 3  Host-Microorganism Interactions

27

• BOX 3.1 Definitions of Selected Epidemiologic Terms • C  arrier: A person who harbors the etiologic agent but shows no apparent signs or symptoms of infection or disease • C  ommon source: A single source or reservoir from which an etiologic agent responsible for an epidemic or outbreak originates • C  ommunity-associated infection: Infection acquired in an activity or group that is not in a health care setting or environment. • D  isease incidence: The number of new diseases or infected persons in a population • D  isease prevalence: The percentage of diseased persons in a given population at a particular time • E  ndemic: A disease constantly present at some rate of occurrence in a particular location • E  pidemic: A larger-than-normal number of diseased or infected individuals in a particular location • E  tiologic agent: A microorganism responsible for causing infection or infectious disease • H  ealth care–associated infection: Infections acquired as a result of a short- or long-term admission into a health care facility • Iatrogenic: Infection acquired as a result of a medical procedure. • M  icrobiome: An individual’s microbiologic environment, present in or on the human host • M  ode of transmission: The means by which etiologic agents are brought in contact with the human host (e.g., infected blood, contaminated water, insect bite)

• M  orbidity: The state of disease and its associated effects on the host • M  orbidity rate: The incidence of a particular disease state • M  ortality: Death resulting from disease • M  ortality rate: The incidence in which a disease results in death • N  osocomial infection: Infection for which the etiologic agent was acquired in a hospital or long-term health care center or facility • O  utbreak: A larger than normal number of diseased or infected individuals that occurs over a relatively short period • P  andemic: An epidemic that spans the world • R  eservoir: The origin of the etiologic agent or location from which it disseminates (e.g., water, food, insects, animals, other humans) • S  train typing: Laboratory-based characterization of etiologic agents designed to establish their relatedness to one another during a particular outbreak or epidemic • S  urveillance: Any type of epidemiologic investigation that involves data collection for characterizing circumstances surrounding the incidence or prevalence of a particular disease or infection • V  ector: A living entity (animal, insect, or plant) that transmits the etiologic agent • V  ehicle: A nonliving entity that is contaminated with the etiologic agent and as such is the mode of transmission for that agent

microbiome associated with pregnancy and preterm birth; (2) gastrointestinal microbiome and the development of inflammatory bowel disease; and (3) microbiome and the development of type 2 diabetes. Undoubtedly, this research will continue to evolve and potentially provide insight into the characterization, risk, and prevention of disease. The relationship between host and microorganism is ultimately associated with the variation and balance of the normal human microbiome and the appearance of a potentially infectious agent. The complex relationships between the human host and medically relevant microorganisms are demonstrated in the sequential steps associated with microbe-host interactions and the subsequent development of infection and disease. The stages of interaction include (1) the physical encounter between the host and microorganism; (2) colonization or survival of the microorganism on an internal (gastrointestinal, respiratory, or genitourinary tract) or external (skin) surface of the host; (3) microbial entry, invasion, and dissemination to deeper tissues and organs of the human body; and (4) resolution or outcome.

exposure are often direct consequences of a person’s activity or behaviors. Certain activities carry different risks for an encounter. There is a wide spectrum of activities and situations over which a person may or may not have absolute control. For example, acquiring salmonellosis because one fails to cook the holiday turkey thoroughly is avoidable, whereas contracting tuberculosis living in conditions of extreme poverty and overcrowding may be unavoidable. The role that human activities play in the encounter between humans and microorganisms cannot be overstated. Most of the crises associated with infectious disease are preventable or can be greatly reduced with changes in human behavior and living conditions.

The Encounter Between Host and Microorganism The Human Host’s Perspective Because microorganisms are ubiquitous in nature, human encounters are inevitable, but the means of encounter vary widely. Which microbial population and the mechanism of

Microbial Reservoirs and Transmission Humans encounter microorganisms when they enter or are exposed to the same environment in which the microbial agents live or when the infectious agents are brought to the human host by indirect means. The environment or place of origin of the infecting agent is the reservoir. As shown in Fig. 3.1, microbial reservoirs include humans, animals, water, food, air, and soil. The human host may acquire microbial agents by various means or modes of transmission. The mode of transmission is direct when the host directly contacts the microbial reservoir and is indirect when the host encounters the microorganism by an intervening agent of transmission. The agents of transmission that bring the microorganism from the reservoir to the host may be a living entity, such as

28 PA RT I     Basic Medical Microbiology

Microorganism sources (reservoirs) Humans Animals Food (from plant and animal sources) Water Air Soil

Modes of transmission

1. Direct; transmitted by direct contact between reservoir and host 2. Indirect; transmitted to host via intervening agent(s)

Human host

Intervening agents: Vectors — animals, insects, other humans Vehicles — water, food, air, medical devices, various other inanimate objects

• Fig. 3.1  Summary of microbial reservoirs and modes of transmission to humans.

an insect, in which case they are called vectors, or they may be a nonliving entity, referred to as a vehicle or fomite. In addition, some microorganisms may have a single mode of transmission, whereas others may spread by various methods. From a diagnostic microbiology perspective, knowledge about an infectious agent’s mode of transmission is often important for determining optimal specimens for isolation of the organism and for implementing precautions that minimize the risk of laboratory or health care–associated infections (HAIs) (see Chapters 4, 78, and 79 for more information regarding laboratory safety, infection control, and sentinel laboratory responses, respectively). 

Human and Microbe Interactions Humans play a substantial role as microbial reservoirs. In fact, the passage of a neonate from the sterile environment of the mother’s womb through the birth canal, which is heavily colonized with various microbial agents, is a primary example of one human directly (i.e., direct transmission) acquiring a microorganism from another human serving as the reservoir. This is the mechanism that newborns first encounter microbial agents. Other examples in which humans serve as the microbial reservoir include the acquisition of streptococcal pharyngitis through touching; hepatitis through blood transfusions; gonorrhea, syphilis, and acquired immunodeficiency syndrome (AIDS) through sexual contact; and tuberculosis and the common cold through aerosolized droplets associated with coughing or sneezing. Indirect transmission can occur when microorganisms from one individual contaminate a vehicle of transmission, such as water (e.g., cholera), that is then ingested by another person. In the medical setting, indirect transmission of microorganisms from one human host to another by means of a medical procedure (i.e., iatrogenic) and contaminated medical devices helps to disseminate infections in hospitals. Hospital-acquired, health care–associated, or longterm care–associated infections are considered nosocomial

infections. Health care–associated infections (HAIs) include exposure in a variety of settings and not confined to in-patient care in a health care institution. These exposures occur during field containment or transportation of infectious agents as well as in daily contact with infected patients in clinics. In addition, HAIs are not limited to health care professionals and patients, but also include visitors, support staff, and students. In addition, humans are routinely exposed to infectious agents through participation in activities and events throughout their daily lives. These activities include direct and indirect transmission of infectious agents in community settings. These infections are considered communityassociated (CA) infections. 

Animals as Microbial Reservoirs Infectious agents from animal reservoirs are transmissible directly to humans through an animal bite (e.g., rabies) or indirectly through the bite of insect vectors that feed on both animals and humans (e.g., Lyme disease and Rocky Mountain spotted fever). Animals may also transmit infectious agents by acquiring them from or depositing them in water and food supplies. For example, beavers heavily colonized with parasites can cause infection of the human gastrointestinal tract. These parasites may be encountered and subsequently acquired when stream water is contaminated by the beaver and is used by a vacationing camper. Alternatively, animals used for human food carry numerous bacteria (e.g., Salmonella and Campylobacter) that, if not destroyed through appropriate cooking during preparation, can cause severe gastrointestinal illness. Many other infectious diseases can be encountered through direct or indirect animal contact, and information regarding a patient’s exposure to animals is often a key component in the diagnosis of these infections. Some microorganisms primarily infect animal populations and on occasion accidentally encounter and infect humans. When

CHAPTER 3  Host-Microorganism Interactions

a human infection results from such an encounter, it is a zoonotic infection. More specifically, if the human infection is a result of regular interaction with animals for food production, the infection is livestock-associated. 

Insects as Vectors The most common role of insects (arthropods) in the transmission of infectious disease is as vectors rather than as reservoirs. A variety of arthropods can transmit viral, parasitic, and bacterial disease from animals to humans, whereas others transmit microorganisms between human hosts without an intermediate animal reservoir. Malaria, a deadly disease, is a prime example of an infectious disease maintained in the human population by the feeding and survival of an insect vector, the mosquito. Still other arthropods may themselves be agents of disease. These include organisms such as lice and scabies spread directly between humans and cause skin irritations but do not penetrate the body. Because they are able to survive on the skin of the host without gaining access to internal tissues, they are ectoparasites (Chapter 46). In addition, nonfungal infections may result when microbial agents in the environment, such as endospores, are introduced mechanically through the bite of a vector, scratch, or other penetrating wound. 

The Environment as a Microbial Reservoir The soil and natural environmental debris are reservoirs for countless types of microorganisms. It is not surprising that these also serve as reservoirs for microorganisms that can cause infection in humans. Many of the fungal agents (see Part V: Mycology) are acquired by inhalation of soil and dust particles containing microorganisms (e.g., San Joaquin Valley fever). Other, nonfungal infections (e.g., tetanus endospores) may result when microbial agents in the environment are introduced into the human body by a penetrating wound. 

The Microorganism’s Perspective Clearly, numerous activities can result in human encounters with microorganisms. Because humans are engaged in all of life’s complex activities, the tendency is to perceive the microorganism as having a passive role in the encounter process. However, this assumption is a gross oversimplification. Microorganisms are driven by survival, and the environment of the reservoirs they occupy must allow their metabolic and genetic needs to be fulfilled. Reservoirs can be inhabited by hundreds or thousands of different microorganisms. Yet human encounters with the reservoirs, either directly or indirectly, do not result in all microorganisms establishing an association with the human host. Although some microorganisms have evolved strategies that do not require a human host to ensure survival, others have included humans to a lesser or greater extent as part of their survival tactics. These organisms often have mechanisms that enhance their chances for a human encounter.

29

Depending on factors associated with both the human host and the microorganism involved, the encounter may have a beneficial, disastrous, or inconsequential effect on each of the participants. 

Microorganism Colonization of Host Surfaces The Host’s Perspective Once a microbe is in contact with a human host, the outcome of the encounter depends on what happens during each step of the interaction, beginning with colonization. The human host’s role in microbial colonization, defined as the persistent survival of microorganisms on a surface of the human body, is dictated by the defenses that protect vital internal tissues and organs against microbial invasion. The first defenses are the external and internal body surfaces that are in direct contact with the external environment and are the anatomic regions where the microorganisms will initially encounter the human host. These surfaces include: • Skin (including the conjunctival epithelium covering the eye) • Mucous membranes lining the mouth or oral cavity, the respiratory tract, the gastrointestinal tract, and the genitourinary tract Because body surfaces are always present and provide protection against all microorganisms, skin and mucous membranes are constant and nonspecific defense mechanisms. As is discussed later in this text, other protective mechanisms are produced in response to the presence of microbial agents (inducible defenses), and some are directed specifically at particular microorganisms (specific defense mechanisms).

Skin and Skin Structures Skin serves as a physical and chemical barrier to microorganisms; its protective characteristics are summarized in Table 3.1 and Fig. 3.2. The acellular, outermost layer of the skin, along with the tightly packed cellular layers underneath, provides an impenetrable physical barrier to all microorganisms, unless damaged. In addition, these layers continuously shed, thus dislodging bacteria that have attached to the outer layers. The skin is also a dry and cool environment, which is incompatible with the growth requirements of many microorganisms that thrive in a warm, moist environment. The follicles and glands of the skin produce various natural antibacterial substances, including sebum and sweat. However, many microorganisms can survive the conditions of the skin. These bacteria, or the skin microbiome, are skin colonizers, and they often produce substances that may be toxic and inhibit the growth of more harmful microbial agents. The skin human microbiome differs among healthy individuals more than any other body site. Beneath the outer layers of skin are various host cells that protect against organisms that breach the surface barriers. These cells, collectively known as skin-associated lymphoid tissue,

30 PA RT I     Basic Medical Microbiology

TABLE   Protective Characteristics of the Skin and 3.1  Skin Structures

Skin Structure

Protective Activity

Outer (dermal) layers

• A  ct as a physical barrier to microbial penetration • Remove attached bacteria through sloughing of the outer layers • Provide dry, acidic, and cool conditions that limit bacterial growth

Hair follicles, sweat glands, sebaceous glands

• P  roduce acids, alcohols, and toxic lipids that limit bacterial growth

Eyes/conjunctival epithelium

• F  lushing action of tears: removes microorganisms • Lysozyme in tears: destroys the bacterial cell wall • Mechanical blinking of the eyelid: removes microorganisms

Skin-associated lymphoid tissue

• M  ediates specific and nonspecific protection mechanisms against microorganisms that penetrate the outer tissue layers

TABLE   Protective Characteristics of Mucous 3.2  Membranes

Mucous Membrane

Protective Activity

Mucosal cells

• R  apid sloughing for bacterial removal • Tight intercellular junctions: prevent bacterial penetration

Goblet cells

• M  ucus production: protective lubrication of cells; bacterial trapping; contains specific antibodies with specific activity against bacteria • Provision of antibacterial substances to mucosal surface: • Lysozyme (degrades bacterial cell wall) • Lactoferrin (competes for bacterial iron supply) • Lactoperoxidase (production of substances toxic to bacteria)

Mucosaassociated lymphoid tissue

• M  ediates specific responses against bacteria that penetrate the outer layer

Goblet cell Intercellular (mucus production) junctions Sweat pore

Environment Hair

Bacteria

Dead Epidermis layer Cellular layer Duct Dermis

Sebaceous gland

Cell sloughing

Sweat gland Hair follicle

Ciliated cell Subcutaneous tissue (hypodermis)

Deeper tissues and internal organs

External

Bacteria trapped in mucus ball

•

Fig. 3.3 General features of mucous membranes, highlighting the protective features such as ciliated cells, mucus production, tight intercellular junctions, and cell sloughing.

• Fig. 3.2  Skin and skin structures.

mediate specific and nonspecific responses directed at controlling microbial invaders. 

Mucous Membranes Because cells that line the respiratory tract, gastrointestinal tract, and genitourinary tract are involved in numerous functions besides protection, they are not covered with a hardened, acellular layer. However, the cells that compose these membranes still exhibit various protective characteristics (Table 3.2 and Fig. 3.3).

General Protective Characteristics

Mucus is a major protective component of the membranes. This substance serves to trap bacteria before they can reach the outer surface of the cells, lubricates the cells to prevent damage that promotes microbial invasion, and contains specific chemical (i.e., antibodies) and nonspecific antibacterial substances. In addition to the chemical properties and physical movement of the mucus and trapped microorganisms mediated by ciliary action, rapid cellular shedding and tight intercellular connections provide effective barriers to infection. As is the case with the skin, specific cell clusters,

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Mouth Sloughing cells Flow of saliva Lysozyme Resident microflora Lungs Macrophages

31

Nasopharynx Resident microflora Secretions (lysozyme, phagocytes) Ciliated cells

High concentration of resident microflora

Stomach Low pH Proteolytic enzymes Small intestine Fast flow Mucus Sloughing cells Bile salts Peristalsis Colon Slow flow Mucus, sloughing cells Abundant resident microflora Bile salts Peristalsis

Vagina Low pH Resident microflora Bladder Flushing action of urine Low pH Physical barrier of urethra Urethra Urine flow

• Fig. 3.4  Protective characteristics associated with the mucosal linings of different internal body surfaces.

known as mucosa-associated lymphoid tissue, exist below the outer cell layer and mediate specific protective mechanisms against microbial invasion.  Specific Protective Characteristics

Besides the general protective properties of mucosal cells, the mucosal linings throughout the body have characteristics specific to each anatomic site (Fig. 3.4). The mouth, or oral cavity, is protected by the flow of saliva that physically carries microorganisms away from cell surfaces and contains antibacterial substances, such as antibodies (immunoglobulin A [IgA]) and lysozyme that participate in the destruction of bacterial cells. The mouth is heavily colonized with protective microorganisms that produce substances that hinder successful invasion by harmful organisms. In the gastrointestinal tract, the low pH and proteolytic (protein-digesting) enzymes of the stomach prevent the growth of many microorganisms. In the small intestine, bile salts provide protection that disrupts bacterial membranes, and by peristaltic movement and the fast flow of intestinal contents, which hinder microbial attachment to mucosal cells. Although the large intestine also contains bile salts, the movement of bowel contents is slower, permitting a higher concentration of microbial agents the opportunity to attach to the mucosal cells and inhabit the gastrointestinal tract. As in the oral cavity, the high concentration of normal microbial inhabitants in the large bowel also contributes significantly to protection.

In the upper respiratory tract, nasal hairs keep out large airborne particles that may contain microorganisms. The cough-sneeze reflex significantly contributes to the removal of potentially infective agents. The cells lining the trachea contain cilia (hairlike cellular projections) that move microorganisms trapped in mucus upward and away from the delicate cells of the lungs (Fig. 3.3) by the mucociliary escalator. These barriers are so effective that only inhalation of particles smaller than 2 to 3 μm have a chance of reaching the lungs. In the female urogenital tract, the vaginal lining and the cervix are protected by colonization with normal microbial inhabitants and a low pH. A thick mucus plug in the cervical opening is a substantial barrier that keeps microorganisms from ascending and invading the delicate tissues of the uterus, uterine tubes, and ovaries. The anterior urethra of males and females is colonized with microorganisms, and a stricture at the urethral opening provides a physical barrier that, combined with a low urine pH and the flushing action of urination, protects against bacterial invasion of the bladder, ureters, and kidneys. 

The Microorganism’s Perspective As previously discussed, microorganisms that inhabit many surfaces of the human body (Fig. 3.4) are referred to as colonizers, normal flora, normal microbiota, and collectively as the human microbiome. Some are transient colonizers because they are able to survive, but do not multiply, on

32 PA RT I     Basic Medical Microbiology

• BOX 3.2 Microbial Factors That Contribute to

Colonization of Host Surfaces

Survival Against Environmental Conditions • L  ocalization in moist areas • Protection in ingested or inhaled debris • Expression of specific metabolic characteristics (e.g., salt tolerance) 

Achieving Attachment and Adherence to Host Cell Surfaces • • • •

 ili P Adherence proteins Biofilms Various protein adhesins 

Other Factors • M  otility • Production of substances that compete with the host for acquisition of essential nutrients (e.g., siderophores to capture iron) • Ability to coexist with other colonizing microorganisms

the surface and frequently shed with the host cells. Others, called resident microbiota, not only survive but also thrive and multiply; their presence is more persistent. The body’s microbiota varies considerably with anatomic location. For example, environmental conditions, such as temperature and oxygen availability, differ considerably between the nasal cavity and the small bowel. Only microorganisms with the metabolic capability to survive under the physiologic conditions of the anatomic location are inhabitants of those particular body surfaces. Knowledge of the microbiota of the human body is extremely important in diagnostic microbiology, especially for determining the clinical significance of microorganisms isolated from patient specimens. Organisms considered normal microbiota are often in clinical specimens. This may be a result of contamination of normally sterile specimens during the collection process or because the colonizing organism is actually involved in the infection. Microorganisms considered as normal colonizers of the human body and the anatomic locations they colonize are addressed in Part VII.

Microbial Colonization Colonization may be the last step in the establishment of a long lasting, mutually beneficial (i.e., commensal) or harmless relationship between a colonizer and the human host. Alternatively, colonization may be the first step in the process for the development of infection and disease. Whether colonization results in a harmless or damaging infection depends on the characteristics of the host and the microorganism. In either case, successful initial colonization depends on the microorganism’s ability to survive the conditions first encountered on the host surface (Box 3.2). To avoid the dryness of the skin, organisms often seek moist areas of the body, including hair follicles, sebaceous (oil or sebum) and sweat glands, skin folds, underarms, the

genitals or anus, the face, the scalp, and areas around the mouth. Microbial penetration of mucosal surfaces is mediated when an organism embedded in food particles survives oral and gastrointestinal conditions or is contained within airborne particles to aid survival in the respiratory tract. Microorganisms also exhibit metabolic capabilities that assist in their survival. For example, the ability of staphylococci to thrive in relatively high salt concentrations enhances their survival in and among the sweat glands of the skin. Besides surviving the host’s physical and chemical conditions, colonization also requires that microorganisms attach and adhere to host surfaces (Box 3.2). Attachment can be particularly challenging in places such as the mouth and bowel, in which the surfaces are frequently flushed with passing fluids. Pili, the rodlike projections of bacterial envelopes; various molecules (e.g., adherence proteins and adhesins); and biochemical complexes (e.g., biofilm) work together to enhance attachment of microorganisms to the host cell surface. Biofilm is discussed in more detail later in this chapter. (For more information concerning the structure and functions of pili, see Chapter 2.) In addition, microbial motility with flagella allows organisms to move around and actively seek optimum conditions. Finally, because no single microbial species is a lone colonizer, successful colonization also requires that a microorganism be able to coexist with other microorganisms. 

Microorganism Entry, Invasion, and Dissemination The Host’s Perspective In most instances, to establish infection, microorganisms must penetrate or circumvent the host’s physical barriers (i.e., skin or mucosal surfaces); overcoming these defensive barriers depends on both host and microbial factors. When these barriers are broken, numerous other host defensive strategies activate.

Disruption of Surface Barriers Any situation that disrupts the physical barrier of the skin and mucosa, alters the environmental conditions (e.g., loss of stomach acidity or dryness of the skin), changes the functioning of surface cells, or alters the normal microbiota facilitates the penetration of microorganisms past the barriers and into deeper host tissues. Disruptive factors may vary from accidental or intentional (medical) trauma resulting in surface destruction to the use of antibiotics that remove normal, protective, colonizing microorganisms (Box 3.3). A number of these factors are a result of a medical intervention or procedure. 

Responses to Microbial Invasion of Deeper Tissue Once an organism circumvents surface barriers, the host responds to a microbial presence in the underlying tissue in various ways. Some of these responses are nonspecific, because they occur regardless of the type of invading

CHAPTER 3  Host-Microorganism Interactions

organism; other responses are more specific and involve the host’s immune system. Both nonspecific and specific host responses are critical if the host is to survive. Without them, microorganisms would multiply and invade vital tissues and organs, resulting in severe damage to the host.

Nonspecific Responses

Some nonspecific responses are biochemical; others are cellular. Biochemical factors remove essential nutrients, such as iron, from tissues so that it is unavailable for use by invading microorganisms. Cellular responses are central to tissue and organ defenses, and the primary cells responsible are phagocytes.  Phagocytes

• BOX 3.3 Factors That Contribute to the

Disruption of the Skin and Mucosal Surface

Trauma

• • • • •

 enetrating wounds P Abrasions Burns (chemical and fire) Surgical wounds Needle sticks 

Inhalation

• N  oxious or toxic gases • Particulate matter • Smoking 

Implantation of Medical Devices Other Diseases • Malignancies • Diabetes • Previous or simultaneous infections • Alcoholism and other chemical dependencies Childbirth Overuse of Antibiotics

Endocytosis

Bacteria

Nucleus

Phagocytes are cells that ingest and destroy bacteria and other foreign particles. The types of phagocytes are polymorphonuclear leukocytes, also known as neutrophils (PMNs), monocytes (circulating mononuclear white blood cells) or macrophages (mononuclear white blood cells found in tissue), and dendritic cells. Phagocytes ingest bacteria by a process known as phagocytosis and engulf them in a membrane-lined structure called a phagosome (Fig. 3.5). The phagosome fuses with a second structure, the lysosome. When the lysosome, which contains toxic chemicals and destructive enzymes, combines with the phagosome, the bacteria that are trapped within the phagolysosome are neutralized and destroyed. This destructive process occurs inside membrane-lined structures to prevent the noxious substances contained within the phagolysosome from destroying the phagocyte itself. This is evident during the course of rampant infections when thousands of phagocytes exhibit “sloppy” ingestion of the microorganisms and toxic substances spill from the cells, damaging the surrounding host tissue. The two major phagocytes, PMNs and mononuclear cells, differ in viability and anatomic distribution. PMNs develop in the bone marrow and spend their short lives (usually a day

Phagosome–lysosome fusion

Phagosome

Lysosomes Phagocyte

Phagolysosome In phagolysosome, there is release of lysozyme and other toxic substances Outcomes

Bacterial fragments 1 Long-term survival of bacteria in phagocyte

33

2 Bacterial destruction

3 Destruction of phagocyte

• Fig. 3.5  Overview of phagocyte activity and possible outcomes of phagocyte-bacterial interactions.

34 PA RT I     Basic Medical Microbiology

TABLE 3.3    Components of Inflammation

Component

Functions

Phagocytes (polymorphonuclear neutrophils [PMNs], dendritic cells, and monocytes)

• Ingest and destroy microorganisms

Complement system (coordinated group of serum proteins)

• A  ttracts phagocytes to the site of infection (chemotaxis) • Helps phagocytes to recognize and bind to bacteria (opsonization) • Directly kills gram-negative bacteria (membrane attack complex)

Coagulation system (wide variety of proteins and other biologically active compounds)

• A  ttracts phagocytes to the site of infection • Increases blood and fluid flow to the site of infection • Walls off the site of infection, physically inhibiting the spread of microorganisms

Cytokines (proteins secreted by macrophages and other cells)

• M  ultiple effects that enhance the activities of many different cells essential to nonspecific and specific defensive responses

or less) circulating in blood and tissues. Widely dispersed in the body, PMNs usually are the first cells on the scene of bacterial invasion. Mononuclear cells (monocytes) also develop in the bone marrow. When deposited in tissue or at a site of infection, monocytes transform into mature macrophages. In the absence of infection, macrophages usually reside in specific organs, such as the spleen, lymph nodes, liver, or lungs, where they live for days to several weeks, awaiting encounters with invading bacteria. In addition to the ingestion and destruction of bacteria, macrophages play an important role in mediating immune system defenses (see Specific Responses—The Immune System later in this chapter). In addition to the inhibition of microbial proliferation by phagocytes and biochemical substances such as lysozyme, microorganisms are “washed” from tissues during the flow of lymph fluid. The fluid carries infectious agents through the lymphatic system, where they are deposited in tissues and organs (e.g., lymph nodes and the spleen) heavily populated with phagocytes. This process functions as an efficient filtration system.  Inflammation

Because microbes may survive the initial encounters with phagocytes (Fig. 3.5), the inflammatory response plays an extremely important role as a primary mechanism against microbial survival and proliferation in tissues and organs. Inflammation has both cellular and biochemical components that interact in various complex ways (Table 3.3).

The complement system is composed of a coordinated group of proteins activated by the immune system because of the presence of invading microorganisms. On activation of this system, a cascade of biochemical events occurs that attracts (chemotaxis) and enhances the activities of phagocytes. Because PMNs and macrophages are widely dispersed throughout the body, signals attract and concentrate these cells at the point of invasion, and serum complement proteins provide many of these signals. Cytokines are chemical substances, or proteins secreted by a cell, that have effects on the activities of other cells. Cytokines draw more phagocytes toward the infection and activate the maturation of monocytes to macrophages. Additional protective functions of the complement system is enhanced by hemostasis, which works to increase blood flow to the area of infection and can effectively wall off the infection through the production of clots and barriers composed of cellular debris. The manifestations of inflammation are readily evident and are familiar to most adults; they include the following: • Swelling—caused by an increased flow of fluid and cells to the affected body site • Redness—results from vasodilation of blood vessels and increased blood flow at the infection site • Heat—results from increased cellular metabolism and energy production in the affected area • Pain—caused by tissue damage and pressure on nerve endings from an increased flow of fluid and cells On a microscopic level, the presence of phagocytes at the infection site is an important observation in diagnostic microbiology. Microorganisms associated with these host cells are frequently identified as the cause of a particular infection. 

Specific Responses—The Immune System The immune system provides the human host with the ability to mount a specific protective response to the presence of the invading microorganism. In addition to this specificity, the immune system has a “memory.” When a microorganism is encountered a second or third time, an immune-mediated defensive response is immediately available. However, nonspecific (i.e., phagocytes, inflammation) and specific (i.e., the immune system) host defensive systems are interdependent in their efforts to limit the spread of infection. Components of the Immune System

The central molecule of the immune response is the antibody. Antibodies, also referred to as immunoglobulins, are specific glycoproteins produced by plasma cells (activated B cells) in response to the presence of a molecule recognized as foreign to the host (referred to as an antigen). In the case of infectious diseases, antigens are chemicals or toxins secreted by the invading microorganism or components of the organism’s structure and are usually composed of proteins or polysaccharides. Antibodies circulate in the plasma or liquid portion of the host’s blood and are present in secretions such as saliva. These molecules have two active areas: the antigen-binding site (Fab region) and the phagocyte and complement binding sites (Fc region) (Fig. 3.6).

CHAPTER 3  Host-Microorganism Interactions

Variable regions (antigen binding sites)

Heavy chain

V Light chain

Fab fragment Constant regions

FC fragment

Complement binding site Phagocyte binding site

• Fig. 3.6  General structure of the immunoglobulin G (IgG)-class antibody molecule.

Five major classes or isotypes of antibody exist: IgG, IgA, IgM, IgD, and IgE. Each class has distinctive molecular configurations. IgM is the largest and first antibody produced when an invading microorganism is encountered in the host; production of the most abundant antibody, IgG, follows. IgG consists of four subclasses, IgG1 to IgG4, that have variations in their constant regions resulting in different effector functions related to phagocytosis, complement activation, and antibody-dependent cell-mediated cytotoxicity. IgA is secreted in various body fluids (e.g., saliva and tears) and primarily protects body surfaces lined with mucous membranes. IgA also includes subclasses: IgA1 is predominantly located in the blood stream, and IgA2, which is more resistant to proteolytic cleavage, is located predominantly in secretions. Increased IgE is associated with parasitic infections and allergies. IgD is attached to the surface of specific immune system cells and is involved in the regulation of antibody production. As is discussed in Chapter 9, the ability to measure specific antibody production is a valuable tool for the laboratory diagnosis of infectious diseases. Regarding the cellular components of the immune response, there are three major types of cells: B lymphocytes (B cells), T lymphocytes (T cells), and natural killer cells (NK cells) (Box 3.4). B lymphocytes originate from stem cells and develop into B cells in the bone marrow before being widely distributed to lymphoid tissues throughout the body. These cells primarily function as antibody producers (plasma cells). T lymphocytes also originate from bone marrow stem cells, but they mature in the thymus and either directly destroy infected cells (cytotoxic T cells, TC or CTLs) or work with B cells (helper T cells, TH) to regulate antibody production. Regulatory T cells (Tregs) suppress autoimmune responses by other T lymphocytes and mediate immune tolerance. NK cells are a subset of T cells. There are different types of NK cells, with the most prevalent referred to as invariant natural killer T (NKT) cells. NKT cells develop in the thymus from the same precursor cells as

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• BOX 3.4 Cells of the Immune System

B Lymphocytes (B Cells) • L  ocation: Lymphoid tissues (lymph nodes, spleen, gut-associated lymphoid tissue, tonsils) • F  unction: Antibody-producing cells • S  ubtypes: B lymphocytes: Cells waiting to be stimulated by an antigen Plasma cells: Activated B lymphocytes that secrete antibody in response to an antigen B memory cells: Long-living cells preprogramed to an antigen for subsequent exposure 

T Lymphocytes (T Cells) • L  ocation: Circulate and reside in lymphoid tissues (lymph nodes, spleen, gut-associated lymphoid tissue, tonsils) • F  unctions: Multiple (see different subtypes) • S  ubtypes: Helper T cells (TH): Interact with B cells to facilitate antibody production Cytotoxic T cells (TC): Recognize and destroy host cells that have been invaded by microorganisms Suppressor T cells (TS): Mediate regulatory responses within the immune system T memory cells: Long-living cells preprogramed to an antigen for subsequent exposure 

Natural Killer Cells (NK Cells) • F  unction: Similar to that of cytotoxic T cells but do not require the presence of an antigen to stimulate function

other T lymphocytes. NKT cells have a limited repertoire of T-cell receptors that respond to synthetic, bacterial, and fungal glycolipids. NKT cells are activated by the release of cytokines during viral infections. Each of the three cell types is strategically located in lymphoid tissue throughout the body to maximize the chances of encountering invading microorganisms that the lymphatic system drains from the site of infection.  Two Branches of the Immune System

The immune system provides immunity through two main branches: • A  ntibody-mediated immunity, or humoral immunity • C  ell-mediated immunity, or cellular immunity Antibody-mediated immunity involves the activities of B cells and the production of antibodies. When a B cell encounters a microbial antigen, the cell is activated and initiates a series of events. These events are mediated by helper T cells and the release of cytokines. Cytokines mediate clonal expansion and the number of B cells capable of recognizing the antigen increases. Cytokines also activate the maturation of B cells into plasma cells that produce antibodies specific for the antigen. The process results in the production of B memory cells (Fig. 3.7). B memory cells remain quiescent in the body until a second (anamnestic) or subsequent exposure to the original antigen occurs. With secondary exposure, the B memory cells are preprogrammed

36 PA RT I     Basic Medical Microbiology

Antigen receptor + Microbial antigens

B cell

B-cell activation 1. Clonal expansion = multiplication of B cells that specifically recognize antigen that stimulated activation

2. Antigen is taken into B cell, processed, and presented on B-cell surface, which attracts helper T cells

3. Activated helper T cells, in turn, stimulate B cells to undergo maturation to plasma cells for: - Increased production of highly specific antibody - Switching from IgM to IgG antibody production - Production of B-memory cells

•

Fig. 3.7  Overview of B-cell activation, which is central to antibodymediated immunity.

to produce specific antibodies immediately upon encountering the original antigen. Antibodies protect the host in a number of ways: • Helping phagocytes to ingest and kill microorganisms through a coating mechanism, referred to as opsonization • Neutralizing microbial toxins that are detrimental to host cells and tissues • Promoting bacterial clumping (agglutination) that facilitates clearing from the infection site • Inhibiting bacterial motility • V  iral neutralization; blocking the virus from entering the host cell • Combining with microorganisms to activate the complement system and inflammatory response Because a population of activated specific B cells is a developmental process that results from exposure to microbial antigens, antibody production is delayed when the host is first exposed to an infectious agent. This delay in the primary antibody response underscores the importance of nonspecific response defenses, such as inflammation, that work to hold the invading organisms in check while antibody production begins. This also emphasizes the

importance of B memory cell production. By virtue of this memory, any subsequent exposure or secondary antibody response to the same microorganism results in rapid production of protective antibodies avoiding the delays characteristic of the primary exposure. Some antigens, such as bacterial capsules and outer membranes, activate B cells to produce antibodies without the intervention of helper T cells. However, this activation does not result in the production of B memory cells, and subsequent exposure to the same bacterial antigens does not result in a rapid host memory response. The primary cells involved in cell-mediated immunity are T lymphocytes (cytotoxic T cells) that recognize and destroy human host cells infected with microorganisms. This function is extremely important for the destruction and elimination of infecting microorganisms. Cytotoxic T cells activated during the primary immune response also form a subset of memory T cells that are able to respond quickly to a subsequent infection from a previously encountered pathogen. Some pathogens (e.g., viruses, tuberculosis, some parasites, and fungi) are able to survive in host cells, protected from antibody interaction. Antibody-mediated immunity targets microorganisms outside human cells, whereas cell-mediated immunity targets microorganisms inside human cells. However, in many instances, these two branches of the immune system overlap and work together. Like B cells, T cells must become activated in order to be effective. T-cell activation occurs through interactions with other cells that process microbial antigens and present them on their surface (e.g., macrophages, dendritic cells, and B cells). The responses of activated T cells are very different and depend on the subtype of T cell (Fig. 3.8). Activated helper T cells work with B cells for antibody production (Fig. 3.7) and facilitate inflammation by releasing cytokines. Cytotoxic T cells directly interact with and destroy host cells containing microorganisms or other infectious agents, such as viruses. The activated T cell subset, helper or cytotoxic cells, are controlled by an extremely complex series of biochemical pathways and genetic diversity within the major histocompatibility complex (MHC). MHC molecules are present on cells and form a complex with the antigen to present them to the T cells. The two primary classes of major histocompatibility molecules are MHC I and MHC II. MHC I molecules are located on every nucleated cell in the body and are predominantly responsible for the recognition of endogenous proteins expressed from within the cell. MHC II molecules are located on specialized cell types, including macrophages, dendritic cells, and B cells, for the presentation of extracellular molecules or exogenous proteins. In summary, the host presents a spectrum of challenges to invading microorganisms, from physical barriers, including the skin and mucous membranes, to the interactive cellular and biochemical components of inflammation and the immune system. All these systems work together to minimize microbial invasion and prevent damage to vital tissues and organs resulting from the presence of infectious agents. 

CHAPTER 3  Host-Microorganism Interactions

Antigen receptor

T lymphocytes - Helper T cells - Cytotoxic T cells

Antigens

Antigen-presenting cells - Macrophages - B lymphocytes - Dendritic cells

Activation

Activated helper T cells: - Increased in number - Release cytokines that stimulate activities of phagocytes, natural killer cells, and other components of inflammation - Assist B cells in antibody production (Fig. 3.9) or Activated cytotoxic T cells: - Increased in number - Target and destroy host cells that are infected with microorganisms

• Fig. 3.8  Overview of T-cell activation, which is central to cell-­mediated immunity.

The Microorganism’s Perspective Given the complexities of the human host’s defense systems, it is no wonder that microbial strategies designed to survive these systems are equally complex.

Colonization and Infection Many surfaces on the human body are colonized with a wide variety of microorganisms or microbiota without apparent detriment. In contrast, an infection involves the growth and multiplication of microorganisms that result in damage to the host. The extent and severity of the damage depend on many factors, including the microorganism’s ability to cause disease, the site of the infection, and the general health of the individual infected. Disease results when the infection produces notable changes in human physiology associated with damage or loss of function to one or more of the body’s organ systems. 

Pathogens and Virulence Microorganisms that cause infections or disease are considered pathogens, and the characteristics that enable

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them to cause disease are referred to as virulence factors. Most virulence factors protect the organism against host attack or mediate damaging effects on host cells. The terms pathogenicity and virulence reflect the degree to which a microorganism is capable of causing disease. Pathogenicity specifically refers to the organism’s ability to cause disease, whereas virulence refers to the measure or degree of pathogenicity of an organism. An organism of high pathogenicity is very likely to cause disease, whereas an organism of low pathogenicity is much less likely to cause infection. When disease does occur, highly virulent organisms often severely damage the human host. The degree of severity decreases with diminishing virulence of the microorganism. Because host factors play a role in the development of infectious diseases, the distinction between a pathogenic and nonpathogenic organism and colonizer is not always clear. For example, many organisms that colonize the skin usually do not cause disease (i.e., exhibit low pathogenicity) under normal circumstances. However, when damage to the skin occurs (Box 3.3) or when the skin is disrupted in some other way, these organisms can gain access to deeper tissues and establish an infection. Organisms that cause infection when one or more of the host’s defense mechanisms are disrupted or malfunction are known as opportunistic pathogens, and the infections they cause are referred to as opportunistic infections. On the other hand, several pathogens known to cause serious infections can be part of an individual’s microbiome (i.e., carriers) and never cause disease. However, the same organism can cause life-threatening infection when transmitted to other individuals. The reasons for these inconsistencies are not fully understood, but such widely different results undoubtedly involve complex interactions between microorganism and human. Recognizing and separating a pathogenic from a nonpathogenic organism present one of the greatest challenges in interpreting diagnostic microbiology laboratory results. 

Microbial Virulence Factors Virulence factors provide microorganisms with the capacity to avoid host defenses and damage host cells, tissues, and organs in a number of ways. Some virulence factors are specific for certain pathogenic genera, species, or strains of a microorganism, and substantial differences exist in the way bacteria, viruses, parasites, and fungi cause disease. Knowledge of a microorganism’s capacity to cause specific types of infections plays a major role in the development of diagnostic microbiology procedures used for isolating and identifying microorganisms. (See Part VII for more information regarding diagnosis by organ system.) Attachment Whether humans encounter microorganisms in the air, through ingestion, or by direct contact, the first step of infection and disease development, a process referred to as pathogenesis, is microbial attachment to a surface

38 PA RT I     Basic Medical Microbiology

(exceptions being instances in which the organisms are directly introduced by trauma or other means into deeper tissues). Many of the microbial factors that facilitate attachment of pathogens are the same as those used by nonpathogenic colonizers (Box 3.2). Most pathogenic organisms are not part of the normal human microbiota, and attachment to the host requires that they outcompete the microbiota for a place on the body’s surface. Medical interventions, such as the overuse of antimicrobial agents, result in the destruction of the normal microbiota, creating a competitive advantage for the invading pathogenic organism.  Invasion

Once surface attachment has been secured, microbial invasion into subsurface tissues and organs (i.e., infection) is accomplished by disruption of the skin and mucosal surfaces by several mechanisms (Box 3.3) or by the direct action of an organism’s virulence factors. Some microorganisms produce factors that force mucosal surface phagocytes (M cells) to ingest them and then release them unharmed into the tissue below the surface. Other organisms, such as staphylococci and streptococci, are not so subtle. These organisms produce an array of enzymes (e.g., hyaluronidases, nucleases, collagenases) that hydrolyze host proteins and nucleic acids, destroying host cells and tissues. This destruction allows the pathogen to “burrow” through minor openings in the outer surface of the skin and into deeper tissues. Once a pathogen has penetrated the body, it uses a variety of strategies to survive attack by the host’s inflammatory and immune responses. Alternatively, some pathogens cause disease at the site of attachment without further penetration. For example, in diseases such as diphtheria and whooping cough, the bacteria produce toxic substances that destroy surrounding tissues. The organisms generally do not penetrate the mucosal surface they inhabit.  Survival Against Inflammation

If a pathogen is to survive, the action of the phagocytes and the complement components of inflammation must be avoided or controlled (Box 3.5). Some organisms, such as Streptococcus pneumoniae, a common cause of bacterial pneumonia and meningitis, avoid phagocytosis by producing a large capsule that inhibits the phagocytic process. Other pathogens may not be able to avoid phagocytosis but are not effectively destroyed once internalized and are able to survive within phagocytes. This is the case for Mycobacterium tuberculosis, the bacterium that causes tuberculosis. Still other pathogens use toxins and enzymes to attack and destroy phagocytes before the phagocytes attack and destroy them. The defenses offered by the complement system depend on a series of biochemical reactions triggered by specific microorganism molecular structures. Therefore microbial avoidance of complement activation requires that the infecting agent either mask its activating molecules (e.g., via production of a capsule that covers bacterial surface antigens)

• BOX 3.5 Microbial Strategies for Surviving

Inflammation

Avoid Killing by Phagocytes (Polymorphonuclear Leukocytes) • P  roducing a capsule, thereby inhibiting phagocytes’ ability to ingest them • Antigenic variation, changing surface antigens to limit the number of cells recognized by the immune system 

Avoid Phagocyte-Mediated Killing • Inhibiting phagosome-lysosome fusion • Being resistant to destructive agents (e.g., lysozyme) released by lysosomes • Actively and rapidly multiplying within a phagocyte • Releasing toxins and enzymes that damage or kill phagocytes 

Avoid Effects of the Complement System • U  sing a capsule to hide surface molecules that would otherwise activate the complement system, including the formation of a complex protein polysaccharide matrix (biofilm) • Producing substances that inhibit the processes involved in complement activation • Producing substances that destroy specific complement proteins

or produce substances (e.g., enzymes) that disrupt critical biochemical components of the complement pathway. Any single microorganism may possess numerous virulence factors, and several may be expressed simultaneously. For example, while trying to avoid phagocytosis, an organism may also secrete other enzymes and toxins that destroy and penetrate tissue and produce other factors designed to interfere with the immune response. Microorganisms may also use host systems to their own advantage. For example, the lymphatic and circulatory systems used to carry monocytes and lymphocytes to the site of infection may serve to disperse the organism throughout the body.  Survival Against the Immune System

Microbial strategies to avoid the defenses of the immune system are outlined in Box 3.6. Again, a pathogen can use more than one strategy to avoid immune-mediated defenses, and microbial survival does not necessarily require devastation of the immune system. The pathogen may merely need to “buy” time to reach a safe area in the body or to be transferred to the next susceptible host. In addition, microorganisms can avoid much of the immune response if they do not penetrate the surface layers of the body. This strategy is the hallmark of diseases caused by microbial toxins.  Microbial Toxins

Toxins are biochemically active substances released by microorganisms that have a particular effect on host cells. Microorganisms use toxins to establish infections and multiply within the host. Alternatively, a pathogen may be restricted to

CHAPTER 3  Host-Microorganism Interactions

• BOX 3.6 Strategies That Microbial Pathogens

Use to Survive the Immune Response

• R  apid invasion and multiplication resulting in damage to the host before the immune response can be fully activated, or organism’s virulence is so great that the immune response is insufficient • Invasion and destruction of cells involved in the immune response • Survival in host cells and avoiding detection by the immune system • Masking the organism’s antigens with a capsule or biofilm so that an immune response is not activated • Altering the expression and presentation of antigens so that the immune system is constantly fighting a primary encounter (i.e., the memory of the immune system is neutralized) • Production of enzymes (proteases) that directly destroy or inactivate antibodies

• BOX 3.7 Summary of Bacterial Toxins

Endotoxins • G  eneral toxin common to almost all gram-negative bacteria • Composed of the lipopolysaccharide portion of cell envelope • Released when a gram-negative bacterial cell is destroyed • Effects on host include: • Disruption of clotting, causing clots to form throughout the body (i.e., disseminated intravascular coagulation [DIC]) • Fever • Activation of complement and immune systems • Circulatory changes that lead to hypotension, shock, and death 

Exotoxins • M  ost commonly associated with gram-positive bacteria • Produced and released by living bacteria; do not require bacterial death for release • Specific toxins target specific host cells; the type of toxin varies with the bacterial species • Some kill host cells and help spread bacteria in tissues (e.g., enzymes that destroy key biochemical tissue components or specifically destroy host cell membranes) • Some destroy or interfere with specific intracellular activities (e.g., interruption of protein synthesis, interruption of internal cell signals, or interruption of the neuromuscular system)

a particular body site from which toxins are released to cause systemic damage throughout the body. Toxins also can cause human disease in the absence of the pathogens that produced them. This common mechanism of food poisoning involves ingestion of preformed bacterial toxins (present in the food at the time of ingestion) and is referred to as intoxication, a notable example of which is botulism. Endotoxin and exotoxin are the two general types of bacterial toxins (Box 3.7). Endotoxin is a component of the cellular structure of gram-negative bacteria and can have

39

devastating effects on the body’s metabolism, the most serious being endotoxic shock, which often results in death. The effects of exotoxins produced by gram-positive bacteria tend to be more limited and specific than the effects of gram-negative endotoxin. The activities of exotoxins range from enzymes produced by many staphylococci and streptococci that augment bacterial invasion by damaging host tissues and cells to highly specific activities (e.g., diphtheria toxin inhibits protein synthesis, and cholera toxin interferes with host cell signals). Examples of other highly active and specific toxins are those that cause botulism and tetanus by interfering with neuromuscular functions. 

Genetics of Virulence: Pathogenicity Islands Many virulence factors are encoded in genomic regions of pathogens known as pathogenicity islands (PAIs). These mobile genetic elements contribute to the change and spread of virulence factors among bacterial populations of a variety of species. These genetic elements are believed to have evolved from lysogenic bacteriophages and plasmids and are spread by horizontal gene transfer (see Chapter 2 for information about bacterial genetics). PAIs typically comprise one or more virulence-associated genes and “mobility” genes (i.e., integrases and transposases) that mediate movement between various genetic elements (e.g., plasmids and chromosomes) and among different bacterial strains. In essence, PAIs facilitate the dissemination of virulence capabilities among bacteria in a manner similar to the mechanism diagramed in Fig. 2.10; this also facilitates dissemination of antimicrobial resistance genes (Chapter 10). PAIs are widely disseminated among medically important bacteria. For example, PAIs have been identified as playing a role in virulence for each of the following organisms: Helicobacter pylori Pseudomonas aeruginosa Shigella spp. Yersinia spp. Vibrio cholerae Salmonella spp. Escherichia coli (enteropathogenic, enterohemorrhagic or serotoxigenic, verotoxigenic, uropathogenic, enterotoxigenic, enteroinvasive, enteroaggregative, meningitis-sepsis associated; Chapter 19) Neisseria spp. Bacteroides fragilis Listeria monocytogenes Staphylococcus aureus Streptococcus spp. Enterococcus faecalis Clostridioides difficile Biofilm Formation

Microorganisms typically exist as a group or community of organisms capable of adhering to each other or to other surfaces. A variety of bacterial pathogens, along with other microorganisms, are capable of forming biofilms, such as S. aureus, P. aeruginosa, Aggregatibacter spp., Salmonella spp.,

40 PA RT I     Basic Medical Microbiology

A

B

• Fig. 3.9  (A) Biofilm forming isolate of Staphylococcus aureus cultivated on Congo red agar. Biofilm production results in the formation of a black precipitate. (B) Non–biofilm-producing strain of S. aureus cultivated on Congo red agar.

Blood flow

Blood vessel

EPS

EPS

Surface-epithelial cells (1) Attachment

(2) Cell to cell adhesion with yeast pseudohyphae and epithelial cells

Gram-positive cocci

Persister cells

Pseudohyphae

eDNA

(3) Production of extracellular polysaccharide and proliferation

EPS (extracellular polysaccharide)

(4) Some cells near the biofilm surface are released. Some will lyse releasing DNA –– available to recombine with persister cells

(5) Release of bacteria

• Fig. 3.10  Overview of biofilm formation, maturation, and dissemination of infection.

Citrobacter koseri, and Candida albicans. A biofilm is an accumulation of microorganisms embedded in a complex matrix composed of proteins, polysaccharides, extracellular DNA (eDNA), and other molecules. Pathogenic microorganisms use the formation of biofilm to adhere to implants and prosthetic devices. For example, health care–related infections with Staphylococcus spp. (Fig. 3.9) associated with implants have become more prevalent. Biofilm-forming strains have a much more complex antibiotic resistance profile, indicating failure of the antibiotic to penetrate the polysaccharide layer. In addition, some of the cells in a sessile or stationary biofilm may experience nutrient deprivation and therefore exist in a slow-growing or starved state (i.e., persister cells), displaying reduced susceptibility to antimicrobial agents. These organisms also have demonstrated a differential gene expression compared with their planktonic or free-floating counterparts. The biofilm-forming communities are able to adapt and respond to changes in their environment, similar to a multicellular organism. Biofilms may form from the accumulation of a single microorganism (monomicrobic aggregation) or from the accumulation of numerous species (polymicrobic aggregation). The initial stage in biofilm formation begins with the synthesis of

an extracellular polymer matrix accompanied by aggregation and recognition. This process is facilitated by the formation of polysaccharides, proteins, and eDNA. The formation of the biofilm protects the organism from desiccation, forms a barrier against toxic compounds, and prevents the loss of protective organic and inorganic molecules. Once the initial biofilm has developed, a process of maturation of the biofilm occurs, which takes approximately 4 to 6 hours, depending on the growth rate of the microorganism. This includes the complex formation of a three-dimensional architecture, including pores and channels within the polymer matrix. During biofilm accumulation, the cells reach a critical mass that result in the alteration in metabolism and gene expression in the persister cells. This is accomplished through a mechanism of signaling between cells or organisms through chemical signals or inducer molecules, such as acyl homoserine lactone (AHL) in gram-negative bacteria or oligopeptides in gram-positive bacteria. These signals are capable of interspecies and intraspecies communication. In addition, the formation of a complex polymicrobial biofilm provides favorable conditions for the exchange of genetic information and horizontal gene transfer. Fig. 3.10 provides an overview of biofilm formation, maturation, and seeding that results in further dissemination and infection.

CHAPTER 3  Host-Microorganism Interactions

Microbial biofilm formation is important to many disciplines, including environmental science, industry, and public health. Biofilm formation affects the efficient treatment of wastewater; it is essential for the effective production of beer, which requires aggregation of yeast cells; and it affects bioremediation for toxic substances such as oil. It has been reported that approximately 65% of hospital-associated infections are associated with biofilm formation. Box 3.8 provides an overview of pathogenic organisms associated with biofilm formation in human infections. 

Outcome and Prevention of Infectious Diseases Outcome of Infectious Diseases Given the complexities of host defenses and microbial virulence, it is not surprising that the factors determining • BOX 3.8 Biofilms and Human Infections These pathogenic organisms have been associated with biofilm formation in human infections. • Acinetobacter spp. • Aeromonas spp. • C  andida albicans • C  itrobacter spp. • Coagulase-negative staphylococci • C  ronobacter spp. • Enterobacter spp. • E  nterococcus spp. • E  scherichia coli • Klebsiella spp. • Listeria monocytogenes • P  roteus spp. • P  seudomonas aeruginosa • S  erratia spp. • S  taphylococcus aureus • S  treptococcus spp. • L  isteria monocytogenes Not intended to be an all-inclusive list.

Host factors: - General state of health - Integrity of surface defenses - Capacity for inflammatory and immune response - Level of immunity - Impact of medical intervention

Restoration of host to complete health

outcome between these two living entities are also complicated. Outcome depends on the state of the host’s health, the virulence of the pathogen, and whether the host can clear the pathogen before infection and disease cause irreparable harm or death (Fig. 3.11). The time from exposure to an infectious agent and the development of a disease or infection depends on host and microbial factors. Infectious processes that develop quickly are referred to as acute infections, and those that develop and progress slowly, sometimes over a period of years, are known as chronic infections. Some pathogens, particularly certain viruses, can be clinically silent inside the body without any noticeable effect on the host before suddenly causing a severe and acute infection. During the silent phase, the infection is said to be latent. Again, depending on host and microbial factors, acute, chronic, or latent infections can result in any of the outcomes detailed in Fig. 3.11. Medical intervention can help the host to fight the infection but usually is not instituted until after the host is aware that an infectious process is underway. The clues that an infection is occurring are known as the signs and symptoms of disease and result from host responses (e.g., inflammatory and immune responses) to the action of microbial virulence factors (Box 3.9). Signs are measurable indications or physical observations, such as an increase in body temperature (fever) or the development of a rash or swelling. Symptoms are indictors as described by the patient, such as headache, aches, fatigue, and nausea. The signs and symptoms reflect the stages of infection. In turn, the stages of infection generally reflect the stages in host-microorganism interactions (Fig. 3.12). Whether medical procedures contribute to controlling or clearing an infection depends on key factors, including: • The severity of the infection, which is determined by the host and microbial interactions already discussed • Accuracy in diagnosing the pathogen or pathogens causing the infection

Potential outcome

Restoration of host to health with residual effects

41

Microbial factors: - Level of virulence - Number of organisms introduced into host - Body sites pathogen targets for invasion

Survival with host’s health severely compromised

Full spectrum of outcomes

• Fig. 3.11  Possible outcomes of infections and infectious diseases.

Death

42 PA RT I     Basic Medical Microbiology

• W  hether the patient receives appropriate treatment for the infection (which depends on accurate diagnosis) 

by preventing transmission of the infecting agents and by controlling or destroying reservoirs of human pathogens. Interestingly, most of these measures do not really involve medical practices but rather social practices and policies.

Prevention of Infectious Diseases

Immunization

The treatment of an infection is often difficult and not always successful. Because much of the damage may already have been done before appropriate medical intervention is provided, the microorganisms gain too much of a “head start.” Another strategy for combating infectious diseases is to stop infections before they start (i.e., disease prevention). As discussed at the beginning of this chapter, the first step in any host-microorganism relationship is the encounter and exposure to the infectious agent. Therefore, strategies to prevent disease involve interrupting or minimizing the risk of infection when exposures occur. As outlined in Box 3.10, interruption of encounters may be accomplished

Medical strategies exist for minimizing the risk of disease development when exposure to infectious agents occurs. One of the most effective methods is vaccination, also referred to as immunization. This practice • BOX 3.10 Strategies for Preventing Infectious

Diseases

Preventing Transmission • A  void direct contact with infected persons or take protective measures when direct contact will occur (e.g., wear gloves, wear condoms). • Block the spread of airborne microorganisms by wearing masks or isolating persons with infections transmitted by air. • Use sterile medical techniques. 

• BOX 3.9 Signs and Symptoms of Infection and

Infectious Diseases

• • • • • • • • • • • •

 eneral or localized aches and pains G Headache Fever Fatigue Swollen lymph nodes Rashes Redness and swelling Cough and sneezes Congestion of nasal and sinus passages Sore throat Nausea and vomiting Diarrhea

Controlling Microbial Reservoirs • • • • •

 anitation and disinfection S Sewage treatment Food preservation Water treatment Control of pests and insect vector populations 

Minimizing Risk Before or Shortly After Exposure • Immunization or vaccination • Cleansing and use of antiseptics • Prophylactic use of antimicrobial agents

Host-microorganism interactions Encounter and entry

Colonization and entry

Invasion and dissemination

Pathogen encounters and colonizes host surface

Pathogen multiplies and breaches host surface defenses

Pathogen invades deeper tissues and disseminates, encounters inflammatory and immune responses

Outcome Pathogen completes cycle: — Leaves host — Destroys host — Remains in latent state — Is destroyed by host

Corresponding infection-disease stages Incubation stage

Prodromal stage

Clinical stage

Stage of decline

Convalescent stage

No signs or symptoms

First signs and symptoms, pathogen may be highly communicable

Peak of characteristic signs and symptoms of infection or disease

Condition of host deteriorates possibly to death or signs and symptoms begin to subside as host condition improves

Full recovery of surviving host or chronic infection develops, or death

• Fig. 3.12  Host-microorganism interactions and stages of infection or disease.

CHAPTER 3  Host-Microorganism Interactions

takes advantage of the specificity and memory of the immune system. The two basic approaches to immunization are active immunization and passive immunization. With active immunization, modified antigens from pathogenic microorganisms are introduced into the body and cause an immune response. If or when the host encounters the pathogen in nature, the memory of the immune system ensures minimal delay in the immune response, thus affording strong protection. With passive immunization, antibodies against a particular pathogen that have been produced in one host are transferred to a second host, where they provide temporary protection. The passage of maternal antibodies to the newborn is a key example of natural passive immunization. Active immunity is generally longer lasting, because the immunized host’s own immune response has been activated. However, for complex reasons, naturally acquired active immunity has had limited success for relatively few infectious diseases, necessitating the development of vaccines. Successful immunization has proven effective against many infectious diseases, including diphtheria, whooping cough (pertussis), tetanus, influenza, polio, smallpox, measles, hepatitis, and certain S. pneumoniae and Haemophilus influenzae infections. Prophylactic antimicrobial therapy, the administration of antibiotics when the risk of developing an infection is high, is another common medical intervention for preventing infection. 

Epidemiology To prevent infectious diseases, information is required regarding the sources of pathogens, the mode of transmission to and among humans, human risk factors for encountering the pathogen and developing infection, and factors that contribute to good and bad outcomes resulting from the exposure. Epidemiology is the science that characterizes these aspects of infectious diseases and monitors the effect diseases have on public health. Fully characterizing the circumstances associated with the acquisition and dissemination of infectious diseases gives researchers a better chance of preventing and eliminating the diseases. In addition, many epidemiologic strategies developed for use in public health systems also apply in long-term care facilities (e.g., nursing homes, hospitals, assisted-living centers) for the control of HAI infections (i.e., nosocomial infections; for more information on infection control, see Chapter 79). The field of epidemiology is broad and complex. Diagnostic microbiology laboratory personnel and epidemiologists often work closely to investigate problems. Familiarity with certain epidemiologic terms and concepts is important (Box 3.1). Because the central focus of epidemiology is on tracking and characterizing infections and infectious diseases, this field depends heavily on diagnostic microbiology. Epidemiologic investigations cannot proceed unless researchers first know the etiologic or

43

causative agents. Therefore the procedures and protocols used in diagnostic microbiology to detect, isolate, and characterize human pathogens are essential for patient care and also play a central role in epidemiologic studies focused on disease prevention and the general improvement of public health. In fact, microbiologists who work in clinical laboratories are often the first to recognize patterns that suggest potential outbreaks or epidemics.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Akondy RS, Fitch M, Edupuganti S, et  al.: Origin and differentiation of human memory CD8 T cells after vaccination, Nature 552:362–367, 2017. Bennett J, Dolin R, Blaser M: Principles and practice of infectious ­diseases, 9th ed., Philadelphia, PA, 2020, Elsevier Saunders. Brock TD, Madigan M, Martinko J, et al.: Biology of microorganisms, Upper Saddle River, NJ, 2009, Prentice Hall. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, 12th ed., Washington D.C., 2019, ASM. Ding T, Scholoss PD: Dynamics and associations of microbial community types across the human body, Nature 509:357–360, 2014. Dobrindt U: Genomic islands in pathogenic and environmental microorganisms, Nat Rev Microbiol 2:414–424, 2002. Engleberg NC, DiRita V, Dermody TS: schaechter’s mechanisms of microbial disease, Baltimore, MD, 2007, Lippincott Williams & Wilkins. Hu T, Gimferrer I, Alberola-Ila J: Control of early stages in invariant natural killer T-cell development, Immunology 134:1–7, 2011. Huttenhower C, Gevers D, Knight R, et al.: Structure, function and diversity of the Healthy Human Microbiome, Nature 486:207– 214, 2012. Karunakaran E, Mukherjee J, Ramalingam B, Biggs CA: Biofilmology: a multidisciplinary review of the study of microbial biofilms, Appl Microbiol Biotechnol 90:1869–1881, 2011. Lister JL, Horswill AR: Staphylococcus aureus biofilms: recent developments in biofilm dispersal, Front Cell Infect Microbiol 4:178, 2014. Manandhar S, Singh A, Varma A, et  al.: Evaluation of methods to detect in vitro biofilm formation by staphylococcal clinical ­isolates, BMC Res Notes 11:714, 2018. Simões LC, Lemos M, Pereira AM, et al.: Persister cells in a biofilm treated with a biocide, Biofouling 27(4):403–411, 2011. Murray PR, editor: Medical microbiology, 5th ed., St Louis, MO, 2008, Mosby. Schmidt H, Hensel M: Pathogenicity islands in bacterial pathogenesis, Clin Microbiol Rev 17:14–56, 2004. Vaishnavi C, Samanta J, Kochhar R: Characterization of biofilms in biliary stents and potential factors involved in occlusion, World J Gastroenterol 24:112–123, 2018. Vidarsson G, Dekkers G, Rispens T: IgG subclasses and allotypes: from structure to effector functions, Front Immunol 5:520, 2014. Youngblood B, Hale JS, Kissick HT, et  al.: Effector CD8 T cells dedifferentiate into long-lived memory cells, Nature 552:404– 409, 2017. Zhurina MV, Gannesen AV, Zdorovenko EL, Plakunov VK: Composition and functions of the extracellular polymer matrix of bacterial biofilms, Microbiology 83:713–722, 2014.

CHAPTER 3  Host-Microorganism Interactions

43.e1

CASE STUDY 3.1 An 8-year-old boy presents to the emergency department (ED) with right upper abdominal pain associated with vomiting, headache, and fever. The boy had been seen in the ED approximately 1.5 months previously for a sore throat, cough, and headache. After the first visit to the ED, the patient was treated with amoxicillin. The boy was born in northern Africa in a refugee camp. He and his family had emigrated from Africa approximately 8 months ago. Generally, the boy appears to be in good health. His immunizations are current, and he has no allergies. He currently resides with his parents and three siblings, who all appear to be in good health. His mother speaks very little English. The attending physician orders an abdominal computed tomography (CT) scan and identifies a mass in the left hepatic lobe. There appears to be no evidence of gastrointestinal

bleeding. The attending physician orders a complete work-up on the patient, including a complete blood count, microbiology tests, chemistry, coagulation, and a hepatitis panel. The laboratory results indicate some type of infection and inflammatory condition. The patient has an elevated white blood cell (WBC) count that correlates with his erythrocyte sedimentation rate (ESR) and C-reactive protein (CRP) level. The ESR and the CRP level are clear indicators of an inflammatory process.

Questions 1. Identify and differentiate the patient’s signs and symptoms. 2. Explain whether this patient likely has an acute or a chronic infection.   

Chapter Review 1. An infection acquired from working with an animal reservoir is: a. Acquired from a vehicle b. Transmitted by a vector c. A zoonotic infection d. An example of indirect transmission 2. Which of the following is considered an indirect mode of transmission? a. A cut with a dirty knife b. Ingesting contaminated potato salad c. Inhaling a droplet containing a bacterium d. Drinking water from a contaminated source 3. Nonspecific immunity includes all of the following except: a. Inflammation b. Phagocytosis by neutrophils c. B-cell activation to produce antibodies d. Resident normal microbiota 4. Humoral immunity: a. Is activated for all infectious agents b. Is specific for any organism c. Is specifically targeted to an antigen d.  Provides a broad immune response to any microorganism 5. Bacterial endotoxins are: a. All the same b. Part of the gram-negative cell wall c. Capable of causing a systemic shock response d. All of the above 6. A sign is different from a symptom in all of the following ways except: a. It provides measurable data. b. It is believed to be associated with the etiology of the disease. c. It is clearly visible. d. It includes the temperature, respiratory rate, and pulse.

7. A short-lived infection that manifests with a short incubation period and serious illness is considered to be: a. Persistent b. Chronic c. Latent d. Acute 8. A microorganism that colonizes the skin but is capable of causing infection under the appropriate conditions is referred to as: a. A pathogenic organism b. An opportunistic pathogen c. Normal microbiota d. A nosocomial pathogen 9. All of the following are involved in humoral immunity except: a. Cytotoxic T cells b. Complement proteins c. Plasma cells d. Glycoproteins 10. Microorganisms that live in or on the human body without causing damage include: a. Colonizers b. Normal flora c. Microbiota d. Human microbiome e. All of the above 11. Biofilm formation within a host results in: a. Easy clearance of bacterial pathogens b. Availability of organic nutrients to the host c.  Inability of the immune system to remove the pathogen d. Starvation of the microorganisms

43.e2 PA RT I     Basic Medical Microbiology

12.  Matching: Match each term with the correct description. _____ vector _____ nosocomial

____ fomite ____ colonization ____ monocytes ____ complement ____ virulence factor ____ exotoxin ____ immunization

a. injection of antigens or antibodies to provide immunity b. inanimate source of infection c. limited and specific effect d. long-term health care–­ associated infection e. characteristic of a diseasecausing organism f. serum proteins activated in the immune system g. circulate in the blood before activation h. insect that carries an infectious agent i. association between normal microbiota and host

13. Short Answer Compare and contrast the components of the specific and nonspecific immune defenses, including the occurrence and process of inflammation, phagocytic cells, antibody production, cellular response, and natural physical or chemical properties of the human body.

PA RT II   General Principles in Clinical Microbiology S E C T I ON 1    Safety and Specimen Management

4

Laboratory Safety OBJECTIVES 1. Define and differentiate sterilization, disinfection, decontamination, and antiseptic. 2. List the factors that influence the effectiveness of disinfectants in the microbiology laboratory. 3. Describe the methods used for the disposal of hazardous waste, including physical and chemical methods, and the material and/or organisms effectively eliminated by each method. 4. Define a chemical hygiene plan and describe the purpose of the methods and items that are elements of the plan, including proper labeling of hazardous materials, training programs, and safety data sheets (SDS). 5. Name the four types of fire extinguishers and the specific flammables that each is effective in controlling. 6. Describe the process of Standard Precautions in the microbiology laboratory, including handling of infectious materials, personal hygiene, use of personal protective equipment (PPE), handling sharp objects, and handwashing procedures. 7. Define Biosafety Levels 1 through 4, including the precautions required for each and type of facility; identify a representative organism for each. 8. Outline the basic guidelines for packing and shipping infectious substances. 9. Describe the management and response required during a biologic or chemical exposure incident in the laboratory.

M

icrobiology laboratory safety practices were first published in 1913. They included admonitions such as the necessity to (1) wear gloves, (2) wash hands after working with infectious materials, (3) disinfect all instruments immediately after use, (4) use water to moisten specimen labels rather than the tongue, (5) disinfect all contaminated waste before discarding, and (6) report to appropriate personnel all accidents or exposures to infectious agents.

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These guidelines are still incorporated into safety programs in the diagnostic microbiology laboratory. Safety programs have been expanded to include the proper handling of biologic hazards encountered in processing patient specimens and handling infectious microorganisms that include standard precautions and transmission-based precautions, engineering and work place controls and risk assessment; fire and electrical safety; the safe handling, storage, and disposal of chemicals and radioactive substances; and techniques for safely lifting or moving heavy objects. In areas of the country prone to natural disasters (e.g., earthquakes, hurricanes, snowstorms), safety programs include disaster preparedness plans that outline the steps to take in an emergency. Although all microbiologists are responsible for their own health and safety, the institution and supervising personnel are required to provide safety training to familiarize microbiologists with known hazards in the workplace and to prevent exposure. Infection control is also a vital part of laboratory safety and is discussed in detail in Chapter 78. Laboratory safety is considered an integral part of overall laboratory services, and federal law in the United States mandates preemployment safety training, followed by quarterly safety in-services. Safety training regulations are enforced by the United States Department of Labor Occupational Safety and Health Administration (OSHA). Regulations and requirements may vary based on the type of laboratory and updated regulations. It is recommended that the laboratory review these requirements as outlined by OSHA (www.osha.gov). Microbiologists should be knowledgeable, properly trained, and equipped with the proper protective materials, engineering, and working controls while performing duties in the laboratory. Investigation of the causes of accidents indicates that unnecessary exposures to infectious agents occur when individuals become sloppy in performing their duties or when they deviate from standardized safety precautions.

CHAPTER 4  Laboratory Safety

45

TABLE 4.1    Classification Scheme of Items Requiring Sterilization or Disinfection

Classification

Description

Items

Methods

Critical items

Pose a high risk of infection if contaminated with infectious agents.

Surgical instruments Cardiac and urinary catheters Implants Ultrasound probes used in sterile body cavities

Purchased as sterilized Heat-sensitive objects: ethylene oxide, hydrogen peroxide gas plasma Chemical: glutaraldehyde, stabilized hydrogen peroxide with or without peracetic acid in specific concentrations

Semi-critical items

Generally items that are exposed to the mucous membranes or nonintact skin. These items should be free of all infectious agents including vegetative bacteria, fungi, and viruses.

Respiratory therapy and anesthesia equipment, endoscopes, laryngoscope blades, esophageal manometry probes, cystoscopes, anorectal manometry catheters, and diaphragm fitting rings.

Glutaraldehyde, hydrogen peroxide, ortho-phthaladehyde, hydrogen peroxide with peracetic acid

Non-critical items

Items that contact intact skin but not mucous membranes.

Noncritical patient care items such as bedpans, blood pressure cuffs, computers, crutches etc. Noncritical environmental surfaces: bed rails, bedside tables, patient furniture, and floors.

Adapted from Centers for Disease and Control Guideline for Disinfection and Sterilization in Healthcare Facilities; https://www.cdc.gov/infectioncontrol/pdf/guid elines/disinfection-guidelines.pdf

Sterilization, Disinfection, and Decontamination

and presence of a biofilm. These processes may be accomplished by a variety of physical or chemical methods.

The Guideline for Disinfection and Sterilization in Healthcare Facilities provides evidence-based recommendations for all cleaning, disinfection, and sterilization of medical devices and the healthcare environment (Table 4.1). Equipment and services associated with patient care and the healthcare environment are all subject to regulations and recommendations for the level of sterilization and/or disinfection based on the risk of infection to the patient. These items as well as considerations for disinfection in the ambulatory care and home care environment are included in Chapter 78. Sterilization is a process that kills all forms of microbial life, including bacterial endospores. Disinfection is a process that destroys pathogenic organisms, but not necessarily all microorganisms, endospores, or prions. However, some disinfectants will kill endospores with prolonged exposure times (3 to 12 hours). These disinfectants are chemical sterilants. Decontamination is the removal of pathogenic microorganisms so items are safe to handle or dispose of. Many factors limit the success or degree of sterilization, disinfection, or decontamination in a health care setting, such as organic load (organisms and other contaminating materials such as blood or body fluids), the type of organisms present, the concentration and exposure time to the germicide, the physical and chemical nature of the object or surface (hinges, cracks, rough or smooth surfaces), temperature, pH, humidity,

Methods of Sterilization The physical methods of sterilization include: • Incineration • Moist heat • Dry heat • Filtration • Ionizing (gamma) radiation • Chemicals (ethylene oxide [EtO] gas, hydrogen peroxide gas plasma, vaporized hydrogen peroxide, and other liquid chemicals) Incineration is a method of treating infectious waste. Hazardous material is literally burned to ashes at temperatures of 870°C to 980°C. Incineration is the safest method to ensure that no infective materials remain in samples or containers when disposed. Prions (infective proteins) are not eliminated using conventional methods. Therefore incineration is recommended. Toxic air emissions and the presence of heavy metals in ash have limited the use of incineration in the United States. Moist heat (steam under pressure) is used to sterilize biohazardous trash and heat-stable objects; an autoclave is used for this purpose. An autoclave is essentially a large pressure cooker. Moist heat in the form of saturated steam under one atmosphere (15 pounds per square inch [psi]) of pressure causes the irreversible denaturation of enzymes and structural proteins. The most commonly used steam

46 PA RT I I     General Principles in Clinical Microbiology

Steam from jacket to chamber

Steam to jacket Jacket

Jacket

Chamber wall

Outer shell Heat exchanger

Chamber drain screen Water supply Drain

A

B

Water/steam ejector

• Fig. 4.1  Gravity displacement type of autoclave. (A) Typical Eagle Century Series sterilizer for laboratory

applications. (B) Typical Eagle 3000 sterilizer piping diagram. The arrows show the entry of steam into the chamber and the displacement of air. (Courtesy AMSCO International, a subsidiary of STERIS Corp., Mentor, Ohio.)

sterilizer in the microbiology laboratory is the gravity displacement autoclave (Fig. 4.1). Steam enters at the top of the sterilizing chamber; because steam is lighter than air, it displaces the air in the chamber and forces it out the bottom through the drain vent. The two common sterilization temperatures are 121°C and 132°C. Biologic waste that includes broth or solid media is usually autoclaved for 30 minutes at 121°C in a displacement sterilizer or 4 minutes at 132°C in a prevacuum sterilizer. Infectious medical waste containing body fluids or blood, on the other hand, is often sterilized at 132°C for 30 to 60 minutes to allow penetration of the steam throughout the waste and the displacement of air trapped inside the autoclave bag. Prions require a much more extensive sterilization process. Several options are recommended for the removal of prions from surgical instruments or other laboratory materials contaminated with high-risk tissue such as brain, spinal cord, and eye tissue. There are four methods for sterilization: (1) autoclave at 134°C for 18 minutes in a prevacuum sterilizer; (2) autoclave at 132°C for 1 hour in a gravity displacement sterilizer; (3) immerse in 1 N sodium hydroxide for 1 hour, remove and rinse with water, then autoclave at 121°C in a gravity displacement or 134°C in a prevacuum sterilizer for 1 hour; or (4) immerse in 1 N sodium hydroxide for 1 hour and heat in a gravity displacement at 121°C for 30 minutes, then clean and subject to routine equipment sterilization. Moist heat is the fastest and simplest physical method of sterilization. Dry heat requires longer exposure times (1.5 to 3 hours) and higher temperatures than moist heat (160° to

180°C). Dry heat ovens are used to sterilize items such as glassware, oil, petrolatum, or powders. Filtration is the method of choice for antibiotic solutions, toxic chemicals, radioisotopes, vaccines, and carbohydrates, which are all heat sensitive. Filtration of liquids is accomplished by pulling the solution through a cellulose acetate or cellulose nitrate membrane with a vacuum. Filtration of air is accomplished using high-efficiency particulate air (HEPA) filters designed to remove organisms larger than 0.3 μm from isolation rooms, operating rooms, and biologic safety cabinets (BSCs). Although considered a method of sterilization, filtration simply removes microorganisms and particles larger than the pore size; smaller particles will not be removed using this method. The ionizing radiation used in microwaves and radiograph machines is composed of short-wavelength and highenergy gamma rays. Ionizing radiation is used for sterilizing disposables such as plastic syringes, catheters, or gloves before use. The most common chemical sterilant is EtO, which is used in gaseous form for sterilizing heatsensitive objects. The main disadvantages of EtO use are the lengthy cycle times and the potential health hazards it produces. Vapor-phase hydrogen peroxide (an oxidizing agent) has been used to sterilize HEPA filters in BSCs, metals, and nonmetal devices such as medical instruments (e.g., scissors). There are no toxic byproducts produced using vapor-phase hydrogen peroxide. Hydrogen peroxide gas plasma is another method that uses hydrogen peroxide and generates plasma by exciting the gas in an enclosed chamber under deep vacuum with the use of radiofrequency or microwave energy. 

CHAPTER 4  Laboratory Safety

Methods of Disinfection Physical Methods of Disinfection The three physical methods of disinfection are: • Boiling at 100°C for 15 minutes, which kills vegetative bacteria • Pasteurizing at 70°C for 30 minutes, which kills food pathogens without damaging the nutritional value or flavor • Using nonionizing radiation such as ultraviolet (UV) light UV rays are long wavelength and low energy. They do not penetrate well, and organisms must have direct surface exposure, such as the working surface of a BSC, for this form of disinfection to work. 

Chemical Methods of Disinfection Chemical disinfectants comprise many classes, including: • Alcohols • Aldehydes • Halogens (chlorine and chlorine compounds) • Peracetic acid • Hydrogen peroxide • Quaternary ammonium compounds • Phenolics Chemicals used to destroy all life are called chemical sterilants, or biocides; however, these same chemicals, when used for shorter periods, act as disinfectants. Disinfectants used on living tissue (skin) are called antiseptics. Resistance to disinfectants varies with the type of infectious agent. Prions are the most resistant, followed by bacterial endospores (such as Bacillus spp.); mycobacteria (acid-fast bacilli); nonenveloped viruses (e.g., poliovirus); fungi; vegetative (nonsporulating) bacteria (e.g., gramnegative rods); and enveloped viruses (e.g., herpes simplex virus), which are the most susceptible to the action of disinfectants. The Environmental Protection Agency (EPA) registers chemical disinfectants used in the United States and requires manufacturers to specify the activity level of each compound at the working dilution. Therefore, microbiologists who must recommend appropriate disinfectants should check the manufacturer’s cut sheets (product information) for the classes of infectious agents that will be killed. Generally, the time necessary for killing infectious agents increases in direct proportion to the microbial load or bioburden (number of organisms). In the clinical environment, the bioburden generally contains multiple types of infectious agents; it is important to ensure that the exposure time is adequate to kill the most resistant agents within the sample. Organic material such as blood, pus, or mucus also affects killing of the infectious agents by inactivating the chemical disinfectant or preventing contact between the chemical and the infectious agent. The organic material should be mechanically removed before chemical sterilization to decrease the microbial load. This is analogous to removing dried food from utensils before placing them in a dishwasher, and it

47

is important for cold sterilization of instruments such as bronchoscopes. The type of water and its concentration in a solution are also important. Hard water may reduce the rate of killing of microorganisms. In addition, 60% to 90% ethyl or isopropyl alcohol solution (volume/volume) is optimally bactericidal, virucidal, fungicidal, and mycobactericidal, because the increased ability of water (H2O) to hydrolyze bonds in protein molecules makes the killing of microorganisms more effective. Ethyl or isopropyl alcohol is nonsporicidal (does not kill endospores) and evaporates quickly. Therefore, its use is limited to the skin as an antiseptic or on thermometers and injection vial rubber septa as a disinfectant. Stabilized hydrogen peroxide has demonstrated bactericidal, virucidal, sporicidal, and fungicidal activities. Commercially available 3% hydrogen peroxide has been used as a disinfectant on inanimate surfaces. The most common disinfectant in the United States is hypochlorite solutions (NaOCl), 5.25% to 6.15%, referred to as household bleach. The disinfecting capability of bleach is bactericidal, virucidal, fungicidal, mycobactericidal, and sporicidal. It is inexpensive and its effectiveness is not decreased based on the quality of the water used in the solution preparation. One disadvantage is that hypochlorite may cause minor ocular, oropharyngeal, and esophageal irritation if an individual is exposed to high concentrations without proper ventilation. It is also corrosive to metals in high concentrations, discolors fabrics, and can produce a toxic gas if improperly mixed with ammonia or acid in other cleaning agents. The Centers for Disease Control and Prevention (CDC) recommends that tabletops be cleaned after blood spills with a 1:10 dilution of bleach. Because of their irritating fumes, the aldehydes (formaldehyde and glutaraldehyde) are generally not used as surface disinfectants. Glutaraldehyde, which is sporicidal (kills endospores) in 3 to 10 hours, is used for medical equipment such as bronchoscopes because it does not corrode lenses, metal, or rubber. Ortho-phthaladehyde (OPA) has similar effects as glutaraldehyde including the ability to kill endospores. OPA has several advantages over glutaraldehyde. It is considered more stable, requires no activation, does not require exposure monitoring, and is not known to irritate the eyes and nasal passages. Peracetic acid (0.23%) combined with hydrogen peroxide (1.0%) is effective in the presence of organic material and has been used for the surface sterilization of surgical instruments. The use of glutaraldehyde, OPA, or peracetic acid is called cold sterilization. Quaternary ammonium compounds are used to disinfect bench tops or other surfaces in the laboratory. However, high water hardness and gross contamination with organic materials, such as blood, may inactivate heavy metals or quaternary ammonium compounds, thus limiting their utility. They are most often used on noncritical surfaces such as floors, furniture, and walls. Finally, phenolics are derivatives of carbolic acid (phenol). Two phenol derivatives commonly included in hospital

48 PA RT I I     General Principles in Clinical Microbiology

disinfectants are ortho-phenylphenol and ortho-benzyl-parachlorophenol. These products are generally considered bactericidal, fungicidal, virucidal, and tuberculocidal, but not sporicidal. The addition of detergent results in a product that cleans and disinfects at the same time, and at concentrations of 2% to 5%, these products are used for cleaning bench tops. 

Antiseptics In addition to decontamination of inanimate objects or surfaces, personal laboratory safety and preparation of patients for invasive procedures require the use of an antiseptic. A variety of antiseptics are used to prepare a patient’s skin for blood draws or other invasive procedures. Ethyl alcohol solutions, as previously indicated, are considered bactericidal, virucidal, fungicidal, and mycobactericidal. Iodine is prepared either as a tincture with alcohol or as an iodophor coupled to a neutral polymer (e.g., povidone-iodine or poloxamer-iodine). Both iodine compounds are widely used antiseptics. In fact, 70% ethyl alcohol, followed by an iodophor, is the most common combination used for skin disinfection before drawing blood specimens for culture or surgery. Iodophors have also been used as disinfectants for hard surfaces but at higher concentrations. Superoxidized water (SOW), 144 mg/L of hypochlorous acid and chlorine, requires preparation onsite prior to use. The antiseptic is exposed to sodium chloride through a semipermeable membrane and production of oxycholine ions is completed using electrolysis. The solution has proven to be an economical alternative to expensive antiseptics and has been used as a treatment option for the cleaning of chronic wounds prior to administration of antibiotics. The solution may also be used for hand washing as well as a potential disinfectant for equipment and surfaces in the health care setting. Because mercury is toxic to the environment, heavy metals containing mercury are no longer recommended. An eye drop solution containing 1% silver nitrate was placed in the eyes of newborns to prevent infections with Neisseria gonorrhoeae. Silver nitrate, however, is no longer manufactured in the United States. The current chemical treatment is either an ointment containing erythromycin or povidone-iodide. The most important point to remember when working with biocides, antiseptics, or disinfectants is to prepare a working solution of the compound exactly according to the manufacturer’s package insert. Many individuals believe that if the manufacturer says to dilute 1:200, they will get a stronger product if they dilute it 1:10. However, the ratio of water to active ingredient may be critical, and if sufficient water is not added, the chemical for surface disinfection may not be effective. 

Chemical Safety The OSHA Hazard Communication Standard provides for institutional educational practices to ensure that all laboratory personnel have a thorough working knowledge of the

•

Fig. 4.2 National Fire Protection Association diamond indicating a chemical hazard. This information can be customized (as shown here for isopropyl alcohol) by applying the appropriate self-adhesive polyester numbers to the corresponding color-coded hazard area. (Courtesy Lab Safety Supply, Janesville, Wisconsin.)

hazards of the chemicals with which they work. This standard has also been called the “employee right to know.” It mandates that all hazardous chemicals in the workplace be identified and clearly marked with a National Fire Protection Association (NFPA) label stating the health risks, such as carcinogen (cause of cancer), mutagen (cause of mutations in deoxyribonucleic acid [DNA] or ribonucleic acid [RNA]), or teratogen (cause of birth defects), and the hazard class, for example, corrosive (harmful to mucous membranes, skin, eyes, or tissues), poisonous, flammable, or oxidizing (Fig. 4.2). Each laboratory should have a chemical hygiene plan that includes guidelines on proper labeling of chemical containers, manufacturers’ safety data sheets (SDSs, formerly Material Safety Data Sheets [MSDSs]), and the written chemical safety training and retraining programs. Hazardous chemicals must be inventoried annually. In addition, laboratories are required to maintain a file of every chemical they use and a corresponding SDS. The manufacturer provides the SDS for every hazardous chemical; some manufacturers also provide letters for nonhazardous chemicals, such as saline, so that these can be included with the other SDSs. The SDSs are required to present the information in a consistent 16-section format. The sections in the SDS include: • Identification • Chemical name, recommended uses and the name, address, and telephone number of manufacturer or supplier • Hazard(s) identification • Classification of the chemical (e.g., flammable, health hazard, etc.), precautionary statements, hazard symbols or pictures related to the risks, and any other hazards or unknown components included in the chemical.

CHAPTER 4  Laboratory Safety

A

49

B

•

Fig. 4.3 Fume hood schematics. Arrows indicate airflow through the cabinet to the outside vent. (Courtesy the Baker Co., Sanford, Maine.)

• • • •

 omposition/information on ingredients C Specific chemical substances or mixtures First-aid measures Instructions based on exposure, symptoms, or effects of exposure and recommended follow-up treatment or medical care • Firefighting measures • Accidental release measures • Containment, evacuation, and cleanup procedures • Handling and storage • Exposure controls and personal protection • Engineering controls and PPE and procedures • Physical and chemical properties • Stability and reactivity • Toxicological information • Routes of transmission and exposure, effects of that exposure, symptoms and numerical measures related to toxicity such as lethal dose or exposure time • Ecological information (nonmandatory) • Environmental impact statement • Disposal considerations (nonmandatory) • Transport information (nonmandatory) • Shipping and transportation regulations and requirements • Regulatory information (nonmandatory) • Any regional and national regulatory specifications associated with agencies such as OSHA, Department of Transportation, Environmental Protection Agency, or Consumer Product Safety Commission • Other information • Date of preparation or revision of SDS Employees should become familiar with the location and organization of SDS files in the laboratory so that they know where to look in the event of an emergency.

Fume hoods (Fig. 4.3) are provided in the laboratory to prevent inhalation of toxic fumes. Fume hoods protect against chemical odor by exhausting air to the outside, but they are not HEPA-filtered to trap pathogenic microorganisms. It is important to remember that a biosafety cabinet (discussed later in the chapter) is not a fume hood. Work with toxic or noxious chemicals should always be performed while wearing nitrile gloves, in a fume hood or while wearing a fume mask. Spills should be cleaned up using a fume mask, gloves, impervious (impenetrable to moisture) apron, and goggles. Acid and alkaline, flammable, and radioactive spill kits are available to assist in rendering any chemical spills harmless. 

Fire Safety Fire safety is an important component of the laboratory safety program. Each laboratory is required to post fire evacuation plans that are essentially blueprints for finding the nearest exit in case of fire. Fire drills conducted quarterly or annually, depending on local laws, ensure that all personnel know what to do in case of fire. Exit paths should always remain clear of obstructions, and employees should be trained to use fire extinguishers. The local fire department is often an excellent resource for training in the types and use of fire extinguishers. Type A fire extinguishers are used for trash, wood, and paper; type B extinguishers are used for chemical fires; and type C extinguishers are used for electrical fires. Combination type ABC extinguishers are found in most laboratories so that personnel need not worry about which extinguisher to reach for in case of a fire. However, type C extinguishers, which contain carbon dioxide (CO2) or another dry

50 PA RT I I     General Principles in Clinical Microbiology

chemical to smother flames, are also used, because this type of extinguisher does not damage equipment. The important actions in case of fire and the order in which to perform tasks can be remembered with the acronym RACE: 1. Rescue any injured individuals. 2. Activate the fire alarm. 3. Contain (smother) the fire, if feasible (close fire doors). 4. Extinguish the fire, if possible. If you are able to extinguish the fire, it is important to follow four basic steps: PASS. 1. Pull the pin—to release the operating handle. 2. Aim—toward the base of the flames at the ignition and fuel source. 3. Squeeze—the handle to release the contents. 4. Sweep—side to side across the base of the flame to extinguish the fire. 

Electrical cords should be checked regularly for fraying and replaced when necessary. All plugs should be the threeprong, grounded type. All sockets should be checked for electrical grounding and leakage at least annually. No extension cords should be used in the laboratory. 

the microbiology laboratory. For example, Blaser and Feldman noted that 5 of 31 individuals who contracted typhoid fever from proficiency testing specimens did not work in a microbiology laboratory. Two patients were family members of a microbiologist who had worked with Salmonella enteric subsp. enterica Typhi, two were students whose afternoon class was in the laboratory where the organism had been cultured that morning, and one worked in an adjacent chemistry laboratory. In the clinical microbiology laboratory, shigellosis, salmonellosis, tuberculosis, brucellosis, and hepatitis are commonly acquired laboratory infections. Additional infections have been reported from agents such as Coxiella burnetii, Francisella tularensis, Trichophyton mentagrophytes, and Coccidioides immitis. Viral agents transmitted through blood and body fluids cause many health care–associated infections in non–microbiology laboratory workers and in health care workers in general. These include hepatitis B virus (HBV), hepatitis C virus (HCV), hepatitis D virus (HDV), and human immunodeficiency virus (HIV). Laboratory-associated infection is not a new phenomenon, but data are based primarily on voluntary reporting. Therefore, such incidents are widely underreported because of fears of repercussions associated with such events. 

Handling of Compressed Gases

Exposure Control Plan

Compressed gas cylinders (CO2, anaerobic gas mixture) contain pressurized gases and must be properly handled and secured. When leaking cylinders have fallen, tanks have become missiles, resulting in loss of life and destruction of property. Therefore, gas tanks should be properly chained and stored in well-ventilated areas. The metal cap, which is removed when the regulator is installed, should always be in place when a gas cylinder is not in use. Cylinders should be transported chained to special dollies. 

The laboratory director and supervisor are legally responsible for ensuring that an Exposure Control Plan has been implemented and that the mandated safety guidelines are followed. The plan identifies tasks that are hazardous to employees and promotes employee safety through use of the following: • Employee education and orientation • Appropriate disposal of hazardous waste • Standard (formerly Universal) Precautions • Engineering controls and safe work practices, as well as appropriate waste disposal and use of BSCs • PPE, such as laboratory coats, shoe covers, gowns, gloves, and eye protection (goggles, face shields) • A post-exposure plan for investigating all accidents and a plan to prevent recurrences 

Electrical Safety

Biosafety Individuals are exposed in various ways to health care–associated infections, transporting specimens and in public areas such as elevators or cafeterias, by: • Rubbing the eyes or nose with contaminated hands • Inhaling aerosols produced during centrifugation, mixing with a vortex, or spills of liquid cultures • Accidentally ingesting microorganisms by putting pens or fingers in the mouth • Receiving percutaneous inoculation (i.e., through puncture from an accidental needle stick) • Manipulating or opening bacterial cultures in liquid media or on plates, creating potentially hazardous aerosols, outside of a biosafety hood • Failure to wash hands upon leaving the restroom or other public areas before entering the laboratory Risks from a microbiology laboratory may extend to adjacent laboratories and to the families of those who work in

Employee Education and Orientation Each institution should have a safety manual that is reviewed by all employees and a safety officer who is knowledgeable about the risks associated with health care–associated infections. The safety officer should provide orientation for new employees and quarterly continuing education updates for all personnel. Initial training and all retraining should be documented in writing. This training should include all items in the laboratory exposure control plan as well as fire, chemical, hazardous materials management (use, storage, and disposal), and blood borne pathogens. 

CHAPTER 4  Laboratory Safety

•

Fig. 4.4 Autoclave bags. (Courtesy Allegiance Healthcare, McGaw Park, Illinois.)

Disposal of Hazardous Waste All materials contaminated with potentially infectious agents must be decontaminated before disposal. These include unused portions of patient specimens, patient cultures, stock cultures of microorganisms, and disposable sharp instruments, such as glass microscope slides, glass or plastic tubes, scalpels, syringes, and needles. It is recommended that syringes with needles not be accepted in the laboratory; staff members should be required to submit capped syringes to the laboratory. Infectious waste may be decontaminated by use of an autoclave, incinerator, or any one of several alternative waste-treatment methods. Some state or local municipalities permit urine and feces to be carefully poured into a sanitary sewer. Infectious waste from microbiology laboratories is usually autoclaved on site or sent for incineration. Infectious waste (agar plates, plastic tubes, and reagent bottles) should be placed into two leak-proof, plastic bags for sturdiness (Fig. 4.4); this is known as double bagging. Sharp objects, including pipettes, microscope slides, broken glass, glass tubes or bottles, scalpels, and needles, are placed in sharps containers (Fig. 4.5), then autoclaved or incinerated. 

Standard Precautions The CDC guidelines known as Standard Precautions (previously Universal Precautions) require that blood and body fluids from every patient be treated as potentially infectious. The essentials of Standard Precautions and safe laboratory work practices are as follows: • Do not eat, drink, smoke, or apply cosmetics (including lip balm). • Do not insert or remove contact lenses. • Do not bite nails or chew on pens. • Do not mouth-pipette. • Limit access to the laboratory to trained personnel only.

51

• A  ssume all patients are infectious for all blood-borne pathogens. • Use appropriate barrier precautions to prevent skin and mucous membrane exposure, including wearing gloves at all times and masks, goggles, gowns, or aprons if splash or droplet formation is a risk. • Thoroughly wash hands and other skin surfaces after removing gloves and immediately after any contamination. • Take special care to prevent injuries with sharp objects, such as needles and scalpels. Standard Precautions should be followed for handling blood and body fluids, including all secretions and excretions submitted to the microbiology laboratory (e.g., serum, semen, all sterile body fluids, saliva from dental procedures, and vaginal secretions). Standard Precautions applies to blood and all body fluids, except sweat. Practice of Standard Precautions by health care workers handling all patient material lessens the risks associated with such specimens. Among the Standard Precautions, hand washing is one of the single most useful techniques to prevent the transmission and acquisition of infection in a health care setting. Hand washing using running water with plain or antimicrobial soaps does not disrupt the normal microbiota but has demonstrated a reduction in transient microorganisms and viral agents. Studies have indicated that effective hand washing with plain soap versus antimicrobial products are both equally efficient and directly correlates with the duration of the hand washing. All personnel should wash their hands with soap and water after removing gloves, after handling infectious material, and before leaving the laboratory area. When hand washing is not available, waterless alcoholbased (60% to 62%) products provide a rapid and convenient means of controlling transmission of many organisms. These alcohol-based products have been used worldwide to control infections and restrict the transmission of pathogens. However, some pathogens such as Enterococcus faecium have now become tolerant or resistant to these products. Alcohol-based products are also not useful when hands are soiled or contaminated with other organic material such as blood and body fluids. Mouth-pipetting is strictly prohibited. Mechanical devices must be used for drawing all liquids into pipettes. Eating, drinking, and applying cosmetics are strictly forbidden in work areas. Food and drink must be stored in refrigerators in areas separate from the work area. All health care workers should follow Standard Precautions whether working inside or outside the laboratory. When collecting specimens outside the laboratory, individuals should follow these guidelines: • Wear gloves and a laboratory coat. • Deal carefully with needles and lancets. • Discard sharps in an appropriate, puncture-resistant container. • Never recap needles by hand, if necessary, special safety devices are available. (Needles are available with built-in safety devices to prevent accidental needle sticks). 

52 PA RT I I     General Principles in Clinical Microbiology

• Fig. 4.5  Sharps containers. (Courtesy Lab Safety Supply, Janesville, Wisconsin.)

Laboratory Design and Engineering Controls Laboratory Environment Microbiology laboratory environments are classified according to the Biosafety Level (BSL) of the classification of microorganisms or risk groups that are approved for testing in the laboratory. The BSL determines the design of the laboratory environment; procedures, access, and laboratory safety controls are based on the potential for transmission of the infectious agents to prevent laboratory-acquired infections and exposure to others. The laboratory should be designed to facilitate a one-way workflow from initial specimen handling of biologically contaminated specimens to areas where pure cultures and additional testing is performed to prevent cross-contamination of specimens. The BSL should be prominently displayed on laboratory doors to restrict access, and the biohazard symbol should be displayed on any equipment (refrigerators, incubators, centrifuges) that contains infectious material The air-handling system of a microbiology laboratory should move air from lower- to higher-risk areas, never the reverse. Ideally, the microbiology laboratory should be under negative pressure, and air should not be recirculated. The selected use of the appropriate BSC for procedures that generate infectious aerosols is critical to laboratory safety. The microbiology laboratory poses many hazards to unsuspecting and untrained people; therefore, access should be limited to employees and other necessary personnel (biomedical engineers, housekeepers). Visitors, especially young children, should be discouraged. Certain areas of high risk, such as the mycobacteriology and virology laboratories, should be closed to visitors. Custodial personnel should be trained to discriminate among the waste containers, dealing only with those that contain noninfectious material. Care

should be taken to prevent insects from infesting any laboratory area. Mites, for example, can crawl over the surface of media, carrying microorganisms from colonies on a plate to other areas. Houseplants can also serve as a source of insects and should be excluded from the laboratory environment. A pest control program should be in place to control rodents and insects. 

Biological Safety Levels The Biosafety in Microbiological and Biomedical Laboratories Manual published by the CDC serves as a reference for laboratory design, work-practice and environmental controls, personnel requirements, disposal, and personnel according to the relative risks of working with various biologic agents. The risk assessment used to determine the recommended Biological Safety Level is based on the characterization of the agent in the following categories: 1.  Route of transmission—direct exposure, inoculation, ingestion, or inhalation 2. Infective dose—amount of organism required to cause an infection. 3. Stability in the environment—temperature, desiccation, decontamination, or sterilization of surfaces. 4. Host range—human, insects (vectors), animals 5. Endemic nature—indigenous versus exotic non-indigenous; wild type versus genetically modified agents. The manual is available on the CDC website (www.cdc.g ov/biosafety/publications/bmbl5/BMBL.pdf ). Biosafety Level 1 (BSL-1) agents include those that have no known potential for infecting healthy adult individuals and are well defined and characterized. These agents are used in laboratory teaching exercises for undergraduate students, secondary educational training, and teaching laboratories for students in microbiology. Precautions for

CHAPTER 4  Laboratory Safety

working with BSL-1 agents include standard good laboratory technique such as hand washing, but do not require the use of engineering or work practice controls that are considered primary or secondary barriers. BSL levels and practices are summarized in Table 4.2. BSL-2 agents are those most commonly being sought in clinical specimens and used in diagnostic, teaching, and other laboratories. BSL-2 precautions are sufficient for the handling of clinical specimens suspected of harboring any indigenous pathogens that pose moderate risk of infection. Specimens expected to contain prions (PrPSc), abnormal proteins associated with neurodegenerative diseases, including spongiform encephalitis, should be handled using BSL-2 procedures. This level of safety includes the principles outlined previously, provided the potential for splash or aerosol is low. If splash or aerosol is probable, the use of primary containment equipment is recommended, as are limiting access to the laboratory during working procedures, training laboratory personnel in handling pathogenic agents, direction by competent supervisors, and performing aerosol-generating procedures in a BSC. BSL-3 procedures have been recommended for the handling of material suspected of harboring organisms unlikely to be encountered in a routine clinical laboratory. However, although rarely encountered in a routine clinical laboratory, agents included in this group may be identified or utilized in a clinical, diagnostic, teaching, research, or production facility. These precautions, in addition to those undertaken for BSL-2 agents, consist of laboratory design and engineering controls that contain potentially dangerous material by careful control of air movement and the requirement that personnel wear protective clothing and gloves. Those working with BSL-3 agents should have baseline sera specimens stored for comparison with acute sera that can be drawn in the event of unexplained illness. BSL-3 organisms are primarily transmitted by infectious aerosol. BSL-4 agents are exotic agents that are considered high risk and cause life-threatening disease. Personnel and all materials must be decontaminated before leaving the facility, and all procedures are performed under maximum containment (special protective clothing, class III BSC). There are two general types of BSL-4 facilities, a cabinet laboratory that utilizes a BSC-3 and a suit laboratory where employees wear a positive-pressure-supplied air-protective suit. Most of the facilities that deal with BSL-4 agents are public health or research laboratories. As mentioned, BSL-4 agents pose life-threatening risks and are transmitted via aerosols; in addition, no vaccine or therapy is available for these organisms.

Biologic Safety Cabinets A BSC is a device that encloses a workspace in such a way as to protect workers from aerosol exposure to infectious disease agents. Either air that contains the infectious material is sterilized, by heat, ultraviolet light, or, most commonly, by passage through a HEPA filter that removes most particles

53

larger than 0.3 μm in diameter. These cabinets are designated as class I through III, according to the effective level of biologic containment. Class I cabinets allow room (unsterilized) air to pass into the cabinet and around the area and material within, sterilizing only the air to be exhausted (Fig. 4.6). They have negative pressure, may be ventilated to the outside or exhausted to the work area, and are usually operated with an open front. Class II cabinets sterilize air that flows over the infectious material, as well as air to be exhausted. The air flows in “sheets,” which serve as barriers to particles from outside the cabinet and direct the flow of contaminated air into the filters (Fig. 4.7). Such cabinets are called vertical laminar flow BSCs. Class II cabinets have a variable sash opening through which the operator gains access to the work surface. Depending on their inlet flow velocity and the percent of air that is HEPA filtered and recirculated, class II cabinets are further differentiated into type A or B. A class IIA cabinet is self-contained, and 70% of the air is recirculated into the work area. The exhaust air in class IIB cabinets is discharged outside the building. A class IIB cabinet is selected if radioisotopes, toxic chemicals, or carcinogens will be used. Because they are completely enclosed and have negative pressure, Class III cabinets afford the most protection to the worker. Air coming into and going out of the cabinet is filter sterilized, and the infectious material within is handled with rubber gloves that are attached and sealed to the cabinet (Fig. 4.8). Most hospital clinical microbiology laboratory scientists use class IIA or IIB cabinets. Routine inspection and documentation of adequate function of these cabinets are critical factors in an ongoing quality assurance program. It is important to the proper operation of laminar flow cabinets that an open area of 3 feet around the cabinet be maintained during operation of the air-circulating system; this ensures that infectious material is directed through the HEPA filter. BSCs must be certified initially, whenever moved more than 18 inches, and annually thereafter. 

Personal Protective Equipment OSHA regulations require that health care facilities provide employees with all personal protective equipment (PPE) necessary to protect them from hazards encountered during the course of work (Fig. 4.9). PPE usually includes plastic shields or goggles to protect workers from droplets, disposal containers for sharp objects, trays in which to carry smaller hazardous items (e.g., culture tubes), handheld pipetting devices, impervious gowns, laboratory coats, disposable gloves, masks, safety carriers for centrifuges (especially those used in the acid fast bacteriology [AFB] laboratory), and HEPA respirators. HEPA respirators are required for all health care workers, including phlebotomists, who enter the rooms of patients with tuberculosis, as well as workers who clean up spills of pathogenic microorganisms (Chapter 78). All respirators should be fit-tested for each individual so that each person

Biosafety Level

Risk Group

Representative Organisms

BSL-1

Agents not associated with disease in healthy individuals. Low individual and community risk.

BSL-2

BSC

Primary Barriers

Secondary Barriers

Additional Practices

Bacillus subtilis Naegleria gruberi Exempt organisms according to NIH guidelines.

None

None

General good laboratory practice, handwashing.

Limit laboratory access. Decontamination procedures in addition, surfaces that are easily cleaned, non-porous.

Agents associated with disease that is preventable and treatable and disease is not considered serious. Moderate individual and low community risk.

Broad spectrum, indigenous moderate risk agents. Representative agents include: HIV Hepatitis B virus Salmonella spp. Toxoplasmosis sp.

BSC-2 or other primary barrier when procedures may produce aerosols or splash.

Primary BSC-2 as indicated or other equipment that prevents aerosolization during manipulation such as enclosed centrifuge safety cups. Requires the use of PPE as appropriate.

Same as above In addition, waste decontamination and disposal of contaminated sharps. Laboratory decontamination procedures and practices. Eye wash station. Pest management program.

Same as above. Biohazard symbol posted on entrance. Immunization requirements and medical surveillance for exposure provided for personnel. Specific personnel training requirements. Biosafety manual including incident and exposure procedures.

BSL-3

Agents associated with serious or lethal disease and where it may be preventable and treatable. High individual risk and low community risk.

Indigenous or exotic agents with potential respiratory transmission. Representative agents include: Mycobacterium tuberculosis, St. Louis encephalitis virus, Coxiella burnetii Mold stages of systemic fungi and organisms grown in quantities greater than found in clinical specimens.

BSC-2 or 3

Same as above and all equipment should be enclosed for laboratory manipulations such as the use of a gas-tight aerosol generation chamber. PPE is solid-front or wrap around, scrub suits or coveralls that are not removed from the laboratory.

Same as above and the addition of ventilation requirements that minimize release of infectious aerosols from the laboratory. Separated from the main laboratory with self-closing doors; all windows must be sealed. Access limited using an anteroom to prevent exchange with outside areas of the laboratory

Same as above. Serum samples of personnel routinely checked for seroconversion comparisons to monitor exposure.

BSL-4

Agents highly likely to cause serious or lethal disease and is not generally known to be preventable or treatable. High individual and community risk.

Dangerous and exotic agents. Representative agents include Marburg virus or Congo-Crimean hemorrhagic fever.

BSC-3 or in a full-body, air-supplied positive pressure personnel suite

Same as above.

Same as above, with specialized zone ventilation systems. Laboratory is generally a separate building. Items are processed through a fumigation chamber or airlock system.

Same as above. Personnel decontamination procedures included such as showering or chemical treatment. Log of all personnel or items entering and leaving laboratory. Generally includes complex waste management procedures.

BSC, Biologic safety cabinet; HIV, human immunodeficiency virus; NIH, National Institutes of Health; PPE, personal protective equipment. Adapted from information included in the United States Department of Health and Human Services, Centers for Disease and Control. Biosafety in Microbiological and Biomedical Laboratories. 5th ed. Washington, DC: US Government Printing Office; 2009. Accessed January 13, 2019.

54 PA RT I I     General Principles in Clinical Microbiology

TABLE 4.2    Laboratory Biosafety Levels

CHAPTER 4  Laboratory Safety

Exhaust

Total exhaust

55

Exhaust collar with air-tight damper

Exhaust filter

HEPA filter

Air intake Supply filter Airflow plenum Work area access

View screen

Fixed view screen Negative pressure system Glove ports

Work area

Access

• Fig. 4.8  Schematic of a Class III biologic safety cabinet with arrows

Air flow

showing airflow through cabinet. The cabinet is self-contained providing the maximum amount of protection to the laboratory from any aerosolized particles. (Courtesy the Baker Co., Sanford, Maine.)

• Fig. 4.6  Schematic of Class I biologic safety cabinet. Room air flows into the cabinet and is circulated out through a high-efficiency particulate air (HEPA) filter and the exhaust portals.

Typical class II biological safety cabinet Exhaust HEPA filter Supply HEPA filter View screen Access opening, typically 8 inches

Airflow plenum Work area

• Fig. 4.7  Schematic of a Class II biologic safety cabinet indicating the

airflow. Air is pulled into the cabinet and circulated through an airflow plenum through two levels of HEPA filter prior to exiting through the exhaust portals. (Courtesy the Baker Co., Sanford, Maine.)

is assured that his or hers is working properly. Men must shave their facial hair to achieve a tight fit. Respirators are evaluated according to guidelines of the National Institute for Occupational Safety and Health (NIOSH), a branch of the CDC. N95 or P100 disposable masks are commonly used in the clinical laboratory and are available from a variety of manufacturers.

• Fig. 4.9  Personal protective equipment. A microbiologist wearing a laboratory coat, gloves, and hood with a shield attached to a HEPA filter pack with a high efficiency particulate air system.

Microbiologists should wear laboratory coats over their uniform scrubs or street clothes, and these coats should be removed before leaving the laboratory. Most exposures to blood-containing fluids occur on the hands or forearms, so gowns with closed wrists or forearm covers and gloves that cover all potentially exposed skin on the arms are most beneficial. Most laboratories utilize disposable gowns and lab coats. If the laboratory protective clothing becomes contaminated with body fluids or potential pathogens, it should be removed and disposed of in the biohazard waste. If washable laboratory attire is provided, the institution or a uniform agency should clean laboratory coats; it is no longer permissible for microbiologists to launder their own coats. Obviously, laboratory workers who plan to enter an area of the hospital where patients at special risk of acquiring

56 PA RT I I     General Principles in Clinical Microbiology

A

B • Fig. 4.10  (A) The Bio-Pouch (lower right) is made of laminated, low-density polyethylene, which is virtually unbreakable. (B) The Bio-Bottle is made of high-density polyethylene and is used as the secondary container. This packaging is used for both types of infectious substances. (Courtesy Air Sea Containers, Miami, Florida.)

infection are present (e.g., intensive care units, the nursery, operating rooms, or areas in which immunosuppressive therapy is being administered) should take every precaution to cover their uniform scrubs or street clothes with clean or sterile protective clothing appropriate to the area. Special impervious protective clothing is advisable for certain activities, such as working with radioactive substances or caustic chemicals. Solid-front gowns are indicated for those working with specimens being cultured for mycobacteria. Unless large-volume spills of potentially infectious material are anticipated, impervious laboratory gowns are not necessary in most microbiology laboratories. 

Postexposure Control All laboratory accidents and potential exposures must be reported to the supervisor and safety officer, who will immediately arrange to send the individual to employee health or an outside occupational health physician. Immediate medical care is of foremost importance; investigation of the accident should take place after the employee has received appropriate care. If the accident is a needle stick injury, for example, the patient should be identified and the risk of the laboratorian acquiring a blood-borne infection should be assessed. The investigation helps the physician determine the need for prophylaxis, such as hepatitis B virus immunoglobulin (HBIG) or an HBV booster immunization in the event of exposure to hepatitis B. The physician also is able to discuss the potential for disease transmission to family members, such as after exposure to a patient with Neisseria meningitidis. Postexposure prophylaxis should be administered, and additional sera should be collected at intervals of 6 weeks, 3 months, and 6 months for HIV testing. Finally, the safety committee, or at least the laboratory director and safety officer, should review the events of the accident to determine whether it could have been prevented and to delineate measures to prevent future accidents. The

investigation of the accident and corrective action should be documented in an incident report. 

Mailing Biohazardous Materials The requirements for packaging and shipping biologic materials, dangerous goods, or infectious substances are highly regulated by the Department of Transportation in the United States along with the International Air Transport Association (IATA) and the International Civil Aviation Organization (IACO). Infectious substances now are classified as category A, B, or C organisms. A category A specimen is an infectious substance capable of causing disease in healthy humans and animals. Category B includes infectious substances that are not included in category A. Only the category A organisms or specimens listed in Table 4.1 must be shipped as dangerous goods. If the laboratory director is unsure whether a patient has symptoms of a category A agent, it is prudent to ship the specimen as an infectious substance rather than a biologic substance. Fig. 4.10A shows triple packaging for diagnostic, clinical, or infectious substances in a pouch; Fig. 4.10B shows triple packaging for diagnostic, clinical, or infectious substances in a rigid bottle. Packaging instructions are available in the annual IATA regulations under section 620 (dangerous goods). All air and ground shippers, such as the US Postal Service (USPS), the US Department of Transportation (DOT), and Federal Express (FedEx), have adopted IATA standards. Training in the proper packing and shipping of infectious material is a key feature of the regulations. Every institution that ships infectious materials, whether a hospital or physician office laboratory (POL), is required to have appropriately trained individuals; training may be obtained through carriers, package manufacturers, and special safety training organizations. The shipper is the individual (institution) ultimately responsible for safe and appropriate packaging. Any fines or penalties are the shipper’s responsibility.

CHAPTER 4  Laboratory Safety

Infectious specimens or isolates should be wrapped with absorbent material and placed inside a plastic biohazard bag, called a primary receptacle. The primary receptacle is then inserted into a secondary container, most often a watertight, hard plastic mailer. The secondary container is capped and placed inside an outer, tertiary container that protects it from physical and water damage (Fig. 4.10B). A label on the outer box confirms that the packaging meets all the required standards. The package must be labeled with a specific hazard label as an infectious substance. A packing list and a Shippers Declaration for Dangerous Goods Form must accompany the air bill or ground form. Diagnostic or clinical specimens are packaged similarly. The shipper should note that some carriers have additional requirements for coolant materials, such as ice, dry ice, or liquid nitrogen. Because the shipper is liable for appropriate packaging, it is best to check with individual carriers in special circumstances and update the instructions yearly when the new IATA Dangerous Goods Regulations are published. IATA regulations can be found at the website www.iata.org. International importation or exportation of biologic agents requires a permit from the CDC. Information on importing and exporting a variety of materials may be found at http://www.cdc.gov/laboratory/speci men-submission/shipping-packing.html. Visit the Evolve site for a complete list of procedures, review questions, and case studies.

57

Bibliography Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, PA, 2020, Elsevier Saunders. Blaser MJ, Feldman RA: Acquisition of typhoid fever from proficiency testing specimens, N Engl J Med 303:1481, 1980. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASCM Press. Centers for Disease Control and Prevention: Guideline for Disinfection and Sterilization in Healthcare Facilities. 2008, revised edition February 2017. Available at: https://www.cdc.gov/infectioncontr ol/pdf/guidelines/disinfection-guidelines.pdf. Centers for Disease Control and Prevention: Guidelines for safe work practices in human and animal medical diagnostic laboratories, Morb Mortal Wkly Rep 61:1–101, 2012. Accessed 13 January 2019. Eftekharizadeh F, Dehnavieh R, Hekmat SN, et al.: Health technology assessment on superoxidized water for treatment of chronic wounds, Med J Islam Repub Iran 30:384, 2016. Fleming DO, Hunt DL: Biological safety: principles and practices, ed 3, Washington, DC, 2000, ASM Press. Occupational Safety and Health Administration: Hazard Communication Standard: Safety Data Sheets: 29 CFR Part 1910, 1200(g), Revised, 2012. Pidot SJ, Gao W, Buultjens AH, et al.: Increasing tolerance of hospital Enterococcus faecium to handwash alcohols, Sci Transl Med 10(452), 2018. https://doi.org/10.1126/scitranslmed.aar6115. United States Department of Health and Human Services, Centers for Disease and Control: Biosafety in microbiological and biomedical laboratories, ed 5, Washington, DC, 2009, US Government Printing Office. Accessed 13 January 2019.

CHAPTER 4  Laboratory Safety

57.e1

Chapter Review 1. Disinfection may be defined as a process that: a. Removes all forms of microbial life b. Is accomplished by physical means c. Removes pathogenic organisms but not spores d. Is used on living tissue 2. All of the following factors affect the outcome when using a disinfectant except: a. Using water from a soft water system b. Disinfecting a surface that contains large pores c. The type of cloth used to wipe the surface d. Temperature and pH 3. In a chemical hygiene plan, the SDS must include: a. The facility’s name b. The laboratory director’s credentials c. The substance’s stability and reactivity d. The purchase date 4. Most laboratories use which type of fire extinguisher? a. Type A b. Type B c. Type C d. Combination ABC 5. When putting out a fire in the laboratory, personnel should be sure to: a. Grab anything that is not replaceable b. Aim directly at the core of the fire c. Turn off the computer system d. Activate the fire alarm and contain and extinguish the fire, if possible 6. Means of exposure to agents in the laboratory include all of the following except: a. Percutaneous inoculation b. Inhalation c. Drinking water d. Placing objects in the mouth 7. Removal of resistant bacterial spores can be accomplished using all of the following methods except: a.  Moist heat at 121°C at 15 psi b. Incineration c.  Moist heat at 132°C for 1 hour d. Dry heat at 180°C for 3 hours 8. Pasteurization is used to disinfect food products for all of the following reasons except: a.  To prevent the killing of flavor-adding microorganisms b. To prevent the destruction of essential vitamins and minerals in the food

c. To completely sterilize food from contaminating organisms d. To remove food pathogens 9.  When disinfecting a contaminated surface, which of the following would provide the most efficient decontamination? a. 100% bleach for 10 minutes b. 80% bleach for 10 minutes c. 70% ethyl alcohol for 8 minutes d. 10% bleach for 5 minutes 10. An engineering control includes: a. Positive air pressure capable of removing all hazardous contaminants b. Gloves, laboratory coats, and face shields c. Safety caps provided on tubes d.  The use of biologic safety cabinets for sample processing 11. The laboratory received a stool culture from a patient experiencing severe abdominal cramps and bloody diarrhea. The sample should be minimally processed using: a. BSL-1 safety practices b. BSL-2 safety practices c. BSL-3 safety practices d. Insufficient information is available to determine the correct answer. 12.  Matching: Match each term with the correct description. _____ standard

precautions _____ sterilization _____ antiseptics _____ fume hood _____ incineration _____ biologic safety cabinet _____ 10% bleach

a. effective on laboratory benches b. uses a filtering system c. hazardous chemicals d. kills all life forms e. CDC guidelines f. living tissue g. temperatures exceed 870°C

13.  Short Answer A fire has started inside a piece of equipment in the laboratory. Describe the steps the laboratory personnel should follow to minimize the damage and cost from the fire.

5

Specimen Management OBJECTIVES 1. State four critical parameters that should be monitored in the laboratory from specimen collection to set up and describe the effects each may have on the quality of the laboratory results (e.g., false negatives or positives, inadequate specimen type, incorrect sample). 2. Identify the proper or improper labeling of a specimen, and determine the adequacy of a specimen given a patient scenario. 3. Define and differentiate backup broth, nutritive media, and differential and selective media. 4. Describe the oxygenation states (atmospheric conditions) associated with anaerobic and aerotolerant, facultative anaerobic, aerobic, and microaerobic (microaerophilic and capnophilic) organisms. Provide an example for each. 5. Determine specimen acceptability and the proper procedure for rejection or recollection. 6. List the critical parameters associated with the reporting of direct and indirect organism detection.

M

icrobiologists work in public health laboratories, hospital laboratories, reference or independent laboratories, and physician office laboratories (POLs). The current trend in the diagnostic setting is changing the landscape of laboratory services. Many health care systems are consolidating microbiology to a single laboratory. This creates a potential for an increase in the time between specimen collection and processing. The result may be a delay in reporting critical results and compromised integrity of the specimen. Depending on the level of service and type of testing at each facility, in general, a microbiologist will perform one or more of the following functions: • Cultivation (growth), identification, and antimicrobial susceptibility testing of microorganisms • Direct detection of infecting organisms by microscopy • Direct detection of specific products of infecting organisms using chemical, immunologic, or molecular techniques • Detection of antibodies produced by the patient in response to an infecting organism (serology) This chapter presents an overview of the issues involved with infectious disease diagnostic testing. Many of these issues are covered in detail in separate chapters. 58

General Concepts for Specimen Collection and Handling Specimen collection and transportation are critical considerations because the results generated by the laboratory are limited by the quality and condition of the specimen upon arrival in the laboratory. Specimens should be obtained to preclude or minimize the possibility of introducing contaminating microorganisms that are not involved in the infectious process and can either interfere with the growth of or outgrow the pathogen. This is a particular problem, for example, in specimens collected from mucous membranes that are already colonized with an individual’s endogenous or “normal” microbiota; these organisms are usually contaminants but may also be opportunistic pathogens. For example, the throats of hospitalized patients on ventilators may be colonized with Klebsiella pneumoniae; although K. pneumoniae is not usually involved in cases of community-acquired pneumonia, it can cause a hospital-acquired respiratory infection in this subset of patients. Using special techniques that bypass areas containing normal microbiota when feasible (e.g., covered-brush bronchoscopy in critically ill patients with pneumonia) prevents many problems associated with false-positive results. Likewise, careful skin preparation before procedures, such as blood cultures and spinal taps, decreases the chance that organisms normally present on the skin will contaminate the specimen.

Appropriate Collection Techniques Specimens should be collected during the acute (early) phase of an illness (or within 2 to 3 days for viral infections) and before antimicrobials, antifungals, or antiviral medications are administered, if possible. Swabs generally are poor specimens if tissue or needle aspirates can be obtained. It is the microbiologist’s responsibility to provide clinicians with a collection manual or instruction cards listing optimal specimen collection techniques and transport information. Information for the nursing staff and clinicians should include the following: • Safety considerations • Selection of the appropriate anatomic site and specimen • Collection instructions, including the type of swab or transport medium

CHAPTER 5  Specimen Management

• T  ransportation instructions, including time and temperature constraints • Labeling instructions, including patient demographic information (minimum of two patient identifiers) • Special instructions, such as patient preparation • Sterile versus nonsterile collection devices • Minimal acceptable quality and recommended quantity Instructions should be written so that specimens collected by the patient (e.g., urine, sputum, or stool) are handled properly. Most urine or stool collection kits contain instructions in several languages, but nothing substitutes for a concise set of verbal instructions. Similarly, when distributing kits for sputum collection, the microbiologist should be able to explain to the patient the difference between spitting in a cup (saliva) and producing good lower respiratory secretions from a deep cough (sputum). General collection information is shown in Table 5.1. An in-depth discussion of each type of specimen is found in Part VII. 

Specimen Transport Ideally, most specimens should be transported to the laboratory within 2 hours of collection. There are instances where the time from collection to laboratory processing should not exceed 15 minutes if not refrigerated or placed in specific transport media (Table 5.2). All specimen containers should be leak-proof, and the specimens should be transported within sealable, leak-proof plastic bags with a separate section for paperwork; resealable bags or bags with a permanent seal are common for this purpose. Bags should be marked with a biohazard label (Fig. 5.1). Many microorganisms are susceptible to environmental conditions such as the presence of oxygen (anaerobic bacteria), changes in temperature (Neisseria meningitidis), or changes in pH (Shigella). Thus the use of special preservatives or temperature-controlled or holding media for the transportation of specimens is important to ensure organism viability (survival). 

Specimen Preservation Preservatives, such as boric acid for urine or polyvinyl alcohol (PVA) and buffered formalin for stool for ova and parasite (O&P) examination, are designed to maintain the appropriate colony counts (urines) or the integrity of trophozoites and cysts (O&P), respectively. Other transport or holding media maintain the viability of microorganisms present in a specimen without supporting the growth of the organisms. This maintains the organisms in a state of suspended animation so that no organism overgrows another or dies out. Stuart’s medium and Amie’s medium are two common holding media. Sometimes charcoal is added to these media to absorb fatty acids present in the specimen that could result in pH changes in the media and the killing of fastidious (fragile) organisms such as Neisseria gonorrhoeae or Bordetella pertussis.

59

Anticoagulants are used to prevent clotting of specimens such as blood, bone marrow, and synovial fluid because microorganisms will otherwise be bound up in the clot. The type and concentration of anticoagulant are very important because many organisms are inhibited by some of these chemicals. Sodium polyanethol sulfonate (SPS) at a concentration of 0.025% (w/v) is usually used because Neisseria spp. and some anaerobic bacteria are particularly sensitive to higher concentrations. Because the ratio of specimen to SPS is so important, it is necessary to have both large (adultsize) and small (pediatric-size) tubes available, so organisms in small amounts of bone marrow or synovial fluid are not overwhelmed by the concentration of SPS. SPS is also included in blood culture collection systems. Heparin is also a commonly used anticoagulant, especially for viral cultures, although it may inhibit the growth of gram-positive bacteria and yeast. Citrate, ethylenediaminetetraacetic acid (EDTA), or other anticoagulants should not be used for microbiology because their efficacy has not been demonstrated for most organisms. It is the microbiologist’s job to make sure media containing the appropriate anticoagulant is used for each procedure. The laboratory generally should not specify a color (“yellow-top”) tube for collection without specifying the anticoagulant (SPS) because at least one popular brand of collection tube (Vacutainer, Becton, Dickinson and Company) has a yellow-top tube with either SPS or trisodium citrate/citric acid/dextrose (ACD); ACD is not appropriate for use in microbiology.

Specimen Storage If specimens cannot be processed as soon as they are received, they must be stored (Table 5.1). Several storage methods are used (refrigerator temperature [4°C], ambient [room] temperature [22–25°C], body temperature [35–37°C], and freezer temperature [either −20°C or −70°C]), depending on the type of transport media (if applicable) and the etiologic (infectious) agents suspected. Urine, stool, viral specimens, sputa, swabs, and foreign devices such as catheters should be stored at 4°C. Serum for serologic studies may be frozen for up to 1 week at −20°C, and tissues or specimens for long-term storage should be frozen at −70°C. 

Specimen Labeling Specimens should be labeled with the patient’s name, identifying number (hospital or sample number), birth date, date and time of collection, source, and the initials of the individual that collected the sample. Enough information must be provided on the specimen label so that the specimen can be matched with the test requisition when it is received in the laboratory. 

Specimen Requisition The specimen (or test) requisition is an order form that is sent to the laboratory along with a specimen. Often the

Specimen

Container

Patient Preparation

Special Instructions

Transportation to Laboratory

Storage Before Processing

Primary Plating Media

Anaerobic Media

Direct Examination

Comments

Abscess (Also Lesion, Wound, Pustule, Ulcer) Superficial

Recommend E-swab transport system or aerobic swab moistened with Stuart’s or Amie’s medium

Wipe area with sterile saline or 70% alcohol.

Aspirate or tissue are preferred if possible, pass swab deeply into the lesion along leading edge of wound.

≤2 h

24 h/RT

BA, CA, Mac, CNA optional

BBA, LKV, BBE

Gram

Contamination of surface material may introduce normal microbiota. Add CNA if smear suggests mixed gram-positive and gram-negative flora.

Deep

Anaerobic transporter; ≥1 mL if sample

Wipe area with sterile saline or 70% alcohol.

Aspirate material from wall or excise tissue.

≤2 h

24 h/RT

BA, CA, Mac, CNA

BBA, LKV, BBE

Gram

Wash any granules and “emulsify” in saline.

Blood culture media set (aerobic and anaerobic bottle) Disinfect the container with 70% isopropyl alcohol or chlorhexidine, wait 30 s

Disinfect venipuncture site with chlorhexidinealcohol.

Draw blood at time of febrile episode; draw two sets from right and left arms; do not draw more than four sets in a 24-h period; draw ≥20 mL/set (adults) or 1–20 mL/set (pediatric) depending on patient’s weight; or per manufacturer’s instructions.

Within 2 h/RT

≤2 h/RT Must be incubated at 37°C on receipt in laboratory.

Blood culture bottles, aerobic; consider isolator tubes fungi and other intracellular agents.

Blood culture bottles, anaerobic.

Direct Gram stain from positive blood culture bottles.

C or BCYE if F. tularensis is suspected. Other considerations: brucellosis, tularemia, cell wall–deficient bacteria, leptospirosis, or AFB; blood cultures should be collected before administration of antibiotics when possible.

Blood

60 PA RT I I     General Principles in Clinical Microbiology

TABLE a 5.1    Collection, Transport, Storage, and Processing of Specimens Commonly Submitted to a Microbiology Laboratory

Specimen

Container

Patient Preparation

Special Instructions

Storage Before Processing

Primary Plating Media

≤24 h, RT if in culture bottle or tube

24 h, RT

BA, CA May use blood culture bottles if volume is sufficient.

BBA

May use an aerobic and anaerobic blood culture bottle set for body fluids. BA, CA, thio, CNA, Mac

BBA, BBE, LKV

Gram

May need to concentrate by centrifugation or filtration—stain and culture sediment.

BBA, BBE, LKV

Gram

May need to homogenize.

Gram—best sensitivity by cytocentrifugation (may also want to do AO if cytocentrifuge not available).

If only 1 tube, submit to microbiology first to avoid contamination; otherwise tube 2. Add thio for CSF collected from shunt. Recommended to also collect blood culture.

Transportation to Laboratory

Anaerobic Media

Direct Examination

Comments

Bone Marrow Aspirate Bone marrow

Blood culture bottles or 1.5 mL lysiscentrifugation tube

Puncture site prepared by primary care provide for surgical incision.

Amniotic abdominal, ascites (peritoneal), bile, joint (synovial), pericardial, pleural

Sterile, screwcap tube or anaerobic transporter or direct inoculation into blood culture bottles, or capped syringe

Disinfect skin with iodine preparation before aspirating specimen.

Needle aspiration

≤15 min

2

−

−

V

+

Comamonas spp.

>2

−

−

V

+

Roseomonas spp.a

1–2b

V

Pink

V

v

Organism

+, >90% of strains are positive; −, >90% of strains are negative; V, variable. aRepresents composite of several species and genomospecies. bGenomospecies 5 is nonmotile. Data compiled from Holt JG, Krieg NR, Sneath PH, et al, eds. Bergey’s Manual of Determinative Bacteriology. 9th ed. Baltimore: Williams & Wilkins: 1994; and Carroll KC, Pfaller MA, Landry ML, et al. Manual of Clinical Microbiology. 12th ed. Washington, DC: ASM; 2019.

TABLE   Key Biochemical and Physiologic Characteristics for Rod-Shaped Motile Species With Peritrichous 24.5  Flagella

Organism

Urea Hydrolysis

Nitrate Reduction

Gas from Nitrate

Growth on Cetrimide

Jordan’s Tartratea

Alcaligenes faecalisb

−

−

−

V

−

Alcaligenes piechaudii

−

+

−

+

+

+

+

+

+

+

−

Alcaligenes xylosoxidans Advenella

V

−

+, >90% of strains are positive; −, >90% of strains are negative; V, variable. aJordan’s tartrate agar deep is a medium used to differentiate gram-negative enteric microorganisms based on the utilization of tartrate. bReduces nitrite. Data compiled from Holt JG, Krieg NR, Sneath PH, et al, eds. Bergey’s Manual of Determinative Bacteriology. 9th ed. Baltimore: Williams & Wilkins; 1994; and Carroll KC, Pfaller MA, Landry ML, et al. Manual of Clinical Microbiology. 12th ed. Washington, DC: ASM; 2019.

Approach to Identification The ability of most commercial identification systems to accurately identify the organisms discussed in this chapter is limited or uncertain. Strategies for identification of these genera are based on the use of conventional biochemical tests and special staining for flagella. Although most clinical microbiology laboratories do not routinely perform flagella stains, motility and flagella placement are the easiest ways to differentiate among these organisms. Many microbiologists groan at the mere mention of having to perform a flagella stain, but the method described in Procedure 12.17 is a wet mount that is easy to perform. At the very least, a simple wet mount to observe cells for motility helps distinguish between the motile and nonmotile genera. The motile organisms described in this chapter have peritrichous flagella (e.g., Alcaligenes spp.) or polar flagella (e.g., Delftia, Comamonas spp.). Organisms are first categorized on the basis of Gram stain morphology and rod shape (Tables 24.4 and 24.5).

They are then further characterized based on whether the organisms have peritrichous flagella (Table 24.4), or flagellated by polar tufts (Table 24.5).

Comments Regarding Specific Organisms In order to distinguish the genera included in this chapter, 16S rRNA sequence analysis is recommended. Alcaligenes piechaudii reduces nitrate to nitrite. A. faecalis has a fruity odor and reduces nitrite to gas. Urea hydrolysis is a key test for Myroides spp., which is also distinguished by production of a characteristic fruity odor. D. acidovorans is unique in producing an orange color when Kovac’s reagent is added to tryptone broth (indole test). Advenella can be separated from related species by the ability to assimilate phenyl acetate. K. gyiorum is positive for the oxidation of D-galacturonic acid and negative for D-serine, whereas K. similis is negative for the oxidation of D-galacturonic acid and positive for D-serine.

402 PA RT I I I    Bacteriology

Roseomonas spp. must be separated from other pinkpigmented, gram-negative (e.g., Methylobacterium spp.) and gram-positive (e.g., certain Rhodococcus spp. or Bacillus spp.) organisms. Roseomonas spp. differ from Rhodococcus and Bacillus spp. by being resistant to vancomycin, as determined by using a 30-μg vancomycin disk on an inoculated 5% blood agar plate. Unlike Methylobacterium spp., Roseomonas spp. grow on MacConkey agar and at 42°C. All Roseomonas species strongly hydrolyze urea but not esculin and are beta-galactosidase negative. 

Serodiagnosis Serodiagnostic techniques are not generally used for the laboratory diagnosis of infections caused by the organisms discussed in this chapter. 

Antimicrobial Susceptibility Testing and Therapy Validated susceptibility testing methods do not exist for these organisms. Although they will grow on the media and under the conditions recommended for testing the more commonly encountered bacteria, this does not necessarily mean that interpretable and reliable results will be produced. Chapter 11 should be reviewed for preferable strategies that can be used to provide susceptibility information when validated testing methods do not exist for a clinically important bacterial isolate. The lack of validated in vitro susceptibility testing methods does not allow definitive treatment and testing guidelines to be given for most organisms listed in Table 24.6. If antimicrobial sensitivity testing is required for Alcaligenes

TABLE 24.6    Antimicrobial Therapy and Susceptibility Testing

Validated Testing Methodsa

Species

Therapeutic Options

Potential Resistance to Therapeutic Options

Advenella

No definitive guidelines

Unknown

Alcaligenes faecalis

No definitive guidelines Potentially active agents include combinations of amoxicillin or ticarcillin with clavulanic acid, various cephalosporins, and ciprofloxacin

Capable of beta-lactamase production Commonly resistant to ampicillin, amoxicillin, ticarcillin, aztreonam, kanamycin, gentamicin, and nalidixic acid

Not available

Alcaligenes piechaudii

No definitive guidelines

Resistant to ampicillin, cefpodoxime, and gentamicin

Not available

Alcaligenes xylosoxidans

No definitive guidelines Potentially active agents include imipenem, piperacillin, ticarcillin/clavulanic acid, ceftazidime, and trimethoprim-sulfamethoxazole

Aminoglycosides, expanded spectrum cephalosporins other than ceftazidime, and quinolones demonstrated no activity Resistant to tobramycin, azithromycin, and clarithromycin

Not available

Comamonas acidovorans, Comamonas testosteroni, Comamonas spp.

No definitive guidelines Potentially active agents include extended- to broad-spectrum cephalosporins, carbapenems, quinolones, and trimethoprimsulfamethoxazole

Unknown

Not available

Delftia acidovorans

No definitive guidelines

Frequently resistant to aminoglycosides

Not available

Kerstersia spp.

No definitive guidelines

Unknown

Roseomonas spp.

No definitive guidelines Potentially active agents include aminoglycosides, imipenem, and quinolones

Generally resistant to cephalosporins and penicillins

aValidated

Comments

Susceptible to combinations of amoxicillin or ticarcillin with clavulanic acid; cephalosporins and ciprofloxacin

C. acidovorans tends to be more resistant than the other two species, especially to aminoglycosides

Susceptible to ciprofloxacin and cefotaxime Not available

testing methods include standard methods recommended by the Clinical and Laboratory Standards Institute (CLSI) and commercial methods approved by the US Food and Drug Administration (FDA).

CHAPTER 24  Alcaligenes, Comamonas, and Similar Organisms

spp., methods include broth macrodilution and microdilution, agar dilution, breakpoint methods, and Etest. Although standardized methods have not been established for the other species discussed in this chapter, in vitro susceptibility studies have been published, and antimicrobial agents that have potential activity are noted, where appropriate, in Table 24.6. 

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there are no recommended vaccination or prophylaxis protocols. For those organisms occasionally identified in health care–associated infections, prevention of infection is best accomplished by following appropriate sterile techniques and infection control guidelines.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Prevention Because the organisms may be encountered throughout nature and do not generally pose a threat to human health,

Bibliography Balows A, Truper HG, Dworkin M, et al.: The prokaryotes: a handbook on the biology of bacteria—ecophysiology, isolation, identification, applications, ed 2, New York, NY, 1981, Springer-Verlag. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Castagnola E, Tasso L, Conte M, Nantron M, Barretta A, Giacchino R: Central venous catheter–related infection due to Comamonas acidovorans in a child with non-Hodgkin’s lymphoma, Clin Infect Dis 19:559–560, 1994. Deutscher M, Severing J, Balada-Llasat JM: Kerstersia gyiorum isolated from a bronchoalveolar lavage in a patient with a chronic tracheostomy, Case Rep Infect Dis 2014;2014:479581. Doi Y, Poirel L, Paterson DL, Nordmann P: Characterization of a naturally occurring class D beta-lactamase from Achromobacter xylosoxidans, Antimicrob Agents Chemother 52:1952–1956, 2008. Dunne Jr WM, Maisch S: Epidemiological investigation of infections due to Alcaligenes species in children and patients with cystic fibrosis: use of repetitive-element–sequence polymerase chain reaction, Clin Infect Dis 20:836–841, 1995. Holt JG, Krieg NR, Sneath PH, et al.: Bergey’s manual of determinative bacteriology, ed 9, Baltimore, MD, 1994, Williams & Wilkins. Lan Y, Yan Q, Yan Y, Liu W: First case of Kerstersia gyiorum isolated from a patient with chronic osteomyelitis in China, Front Lab Med 1:141–143, 2017.

Rihs JD, Brenner DJ, Weaver RE, Steigerwalt AG, Hollis DG, Yu VL: Roseomonas: a new genus associated with bacteremia and other human infections, J Clin Microbiol 31:3275–3283, 1993. Saiman L, Chen Y, Tabibi S, et al.: Identification and antimicrobial susceptibility of Alcaligenes xylosoxidans isolated from patients with cystic fibrosis, J Clin Microbiol 39:3942–3945, 2001. Vancanneyt M, Segers P, Torck U, et  al.: Reclassification of Flavobacterium odoratum (Stutzer 1929) strains to a new genus, Myroides, as Myroidesodoratus comb nov and Myroidesodora timimus spnov, Int J Syst Bacteriol 46:926, 1996. Vandamme P, Heyndrickx M, Vancanneyt M, et al.: Bordetella trematum spnov, isolated from wounds and ear infections in humans, and reassessment of Alcaligenes denitrificans (Rüger and Tan, 1983), Int J Syst Bacteriol 46:849–858, 1996. Wen A, Fegan M, Hayward C, Chakraborty S, Sly LI: Phylogenetic relationships among members of the Comamonadaceae, and description of Delftia acidovorans (den Dooren de Jong, 1926 and Tamaoka et al, 1987) gen nov, comb nov, Int J Syst Bacteriol 49:567–576, 1999. Yabuuchi E, Kawamura Y, Kosako Y, Ezaki T: Emendation of the genus Achromobacter and Achromobacter xylosoxidans (Yabuuchi and Yano) and proposal of Achromobacter ruhlandii (Packer and Vishniac) comb nov, Achromobacter piechaudii (Kiredjian et  al) comb nov, and Achromobacter xylosoxidans subsp denitrificans (Rüger and Tan) comb nov, Microbiol Immunol 42:429–438, 1998.

Chapter Review 1. Which organism has been isolated from CF patients and is Alcaligenes-like? a. D. acidovorans b. Advenella spp. c. Comamonas spp. d. C. pauculus 2. No mode of transmission has been identified for which organism? a. A. denitrificans b. A. faecalis c. Advenella spp. d. Comamonas spp. 3. The organisms in this chapter are difficult to identify. Strategies for identification should include: a. Biochemical tests b. Growth on MacConkey agar c. Flagella staining d. a and c e. All of the above 4. These organisms’ characteristic Gram stain morphology appear as medium to long straight rods. a. Alcaligenes, Myroides, and Roseomonas b. Alcaligenes, Roseomonas, and Kerstersia c. Alcaligenes, Advenella, and Kerstersia d. Myroides, Advenella, and Roseomonas 5. Indole production is typically characterized by the formation of a pink color on addition of Kovac’s reagent; which of the following organisms is unique in its response to Kovac’s reagent?

a. Alcaligenes xylosoxidans produces no color. b. Rhodococcus spp. produce a red color. c. Roseomonas spp. produce a bright magenta color. d. D. acidovorans produces an orange color. 6. The organisms presented in this chapter are grouped together because they are all predominantly: a. Capable of growth on MacConkey agar, oxidase positive, non–glucose utilizers b. Capable of growth on MacConkey agar, oxidase negative, non–glucose utilizers c. Capable of growth on MacConkey agar, oxidase positive, non–lactose fermenters d. Capable of growth on MacConkey agar, oxidase negative, non–lactose fermenters 7. Matching: Match each term with the corresponding term or phrase. _____ Alcaligenes spp. _____ Kerstersia spp. _____Alcaligenes faecalis _____ Myroides spp. _____ Roseomonas spp. _____ K. similis _____ Comamonas spp. _____ K. gyiorum _____ Rhodococcus spp.

a. mucoid b. polar flagella and pleomorphic c. limited to a single species d. wound and abscesses e. sensitive to vancomycin f. feather-edged colonies g. spreading colonies h. positive for D-serine i. positive for D-galacturonic acid

403.e1

25

Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum OBJECTIVES 1. Describe the general characteristics of the organisms discussed in this chapter, including natural habitat, route of transmission, Gram stain reactions, and cellular morphology. 2. Describe the media used to isolate Vibrio spp. and the organisms’ colonial appearance. 3. Explain the physiologic activity of the cholera toxin and its relationship to the pathogenesis of the organism.

4. Describe the clinical significance of Aeromonas spp., Chromobacterium spp., and Vibrio spp. other than Vibrio cholerae. 5. Correlate the patient’s signs and symptoms and laboratory data to identify an infectious agent.

GENERA AND SPECIES TO BE CONSIDERED Current Name

Previous Name

Aeromonas caviae complex Aeromonas caviae Aeromonas eucrenophila Aeromonas media Aeromonas rivipollensis Aeromonas hydrophila complex Aeromonas hydrophila subsp. hydrophila Aeromonas hydrophilia subsp. ranae Aeromonas bestiarum Aeromonas dhakensis Aeromonas salmonicida subsp. salmonicida Aeromonas salmonicida subsp. achromogenes Aeromonas salmonicida subsp. masoucida Aeromonas salmonicida subsp. smithia Aeromonas salmonicida subsp. pectinolytica Aeromonas veronii complex Aeromonas veronii biovar sobria Aeromonas veronii biovar veronii Aeromonas diversa Aeromonas encheleia Aeromonas jandaei Aeromonas schubertii Chromobacterium haemolyticum Chromobacterium violaceum Photobacterium damselae

Vibrio damsela

404

Chapter 25  Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum

Current Name

Previous Name

Grimontia hollisae Vibrio alginolyticus Vibrio cholerae Vibrio cincinnatiensis Vibrio fluvialis Vibrio furnissii Vibrio harveyi Vibrio metschnikovii Vibrio mimicus Vibrio parahaemolyticus Vibrio vulnificus

CDC group EF-13, Vibrio hollisae Vibrio parahaemolyticus biotype 2

405

CDC group EF-6 Vibrio carchariae CDC enteric group 16 Vibrio cholerae (sucrose negative) Pasteurella parahaemolyticus CDC group EF-3

General Characteristics The organisms discussed in this chapter are considered together because they are all oxidase-positive, glucose-fermenting, gram-negative bacilli capable of growth on MacConkey agar. Their individual morphologic and physiologic features are presented later in this chapter in the discussion of laboratory diagnosis. Other halophilic (salt-loving) organisms, such as Shewanella algae, require salt but do not ferment glucose, as do the halophilic Vibrio spp. Aeromonas spp. are gram-negative straight rods with rounded ends or coccobacillary facultative anaerobes that

occur singly, in pairs, or in short chains. They are typically oxidase- and catalase-positive and produce acid from oxidative and fermentative metabolism. Chromobacterium violaceum is a facultative anaerobic, motile, gram-negative rod or cocci. The family Vibrionaceae includes six genera, three of which are discussed in this chapter. The Photobacterium and Grimontia genera each include a single species. The genus Vibrio consists of 10 species of gram-negative, facultative anaerobic, curved or comma-shaped rods. Most Vibrio spp. require sodium for growth and glucose fermentation; with the exception of Vibrio metschnikovii, all species are motile and catalase- and oxidase-positive. 

TABLE 25.1    Epidemiology

Species

Habitat (Reservoir)

Mode of Transmission

Vibrio cholerae

Niche outside of human gastrointestinal tract between occurrence of epidemics and pandemics is uncertain; may survive in a dormant state in brackish or saltwater; human carriers also are known, particularly in endemic regions

Fecal-oral route, by ingestion of contaminated washing, swimming, cooking, or drinking water; also by ingestion of contaminated shellfish or other seafood

Vibrio alginolyticus

Brackish or saltwater

Exposure to contaminated water

Vibrio cincinnatiensis

Unknown

Unknown

Photobacterium damsela

Brackish or saltwater

Exposure of wound to contaminated water

Vibrio fluvialis

Brackish or saltwater

Ingestion of contaminated water or seafood

Vibrio furnissii

Brackish or saltwater

Ingestion of contaminated water or seafood

Grimontia hollisae

Brackish or saltwater

Ingestion of contaminated water or seafood

Vibrio metschnikovii

Brackish, salt, and freshwater

Ingestion of contaminated water or seafood

Vibrio mimicus

Brackish or saltwater

Ingestion of contaminated water or seafood

Vibrio parahaemolyticus

Brackish or saltwater

Ingestion of contaminated water or seafood

Vibrio vulnificus

Brackish or saltwater

Ingestion of contaminated water or seafood

Aeromonas spp.

Aquatic environments around the world, including freshwater, polluted or chlorinated water, brackish water, and, occasionally, marine water; may transiently colonize gastrointestinal tract; often infect various warm- and cold-blooded animal species

Ingestion of contaminated food (e.g., dairy, meat, produce) or water; exposure of disrupted skin or mucosal surfaces to contaminated water or soil; traumatic inoculation of fish fins or fishing hooks

Chromobacterium violaceum

Environmental, soil and water of tropical and subtropical regions Not part of human microbiota

Exposure of disrupted skin to contaminated soil or water

406 PA RT I I I    Bacteriology

Epidemiology Many aspects of the epidemiology of Vibrio spp., Aeromonas spp., and C. violaceum are similar (Table 25.1). The primary habitat for most of these organisms is water; generally, brackish or marine water for Vibrio spp., freshwater for Aeromonas spp., and soil or water for C. violaceum. Aeromonas spp. may also be found in brackish water or marine water with a low salt content. None of these organisms are considered part of the normal human microbiota. Transmission to humans is by ingestion of contaminated water, fresh produce, meat, dairy products, or seafood or by exposure of disrupted skin and mucosal surfaces to contaminated water. The epidemiology of the most notable human pathogen in this chapter, Vibrio cholerae, is far from being fully understood. This organism causes epidemics and pandemics (i.e., worldwide epidemics) of the diarrheal disease cholera. Since 1817, the world has witnessed seven cholera pandemics. In 2010, after a devastating earthquake in Haiti, more than 604,634 infections and 7436 deaths occurred as a result of V. cholerae O1 infections within a 24-month period. During these outbreaks the organism was spread among people by the fecal-oral route due to poor sanitation.

Most V. cholerae infections are asymptomatic, and therefore humans are likely the reservoir for infection in endemic regions. The form of the organism shed from infected humans is somewhat fragile and cannot survive long in the environment. However, evidence suggests that the bacillus has dormant stages that allow its long-term survival in brackish water or saltwater environments during interepidemic periods. These dormant stages are considered viable but nonculturable. Asymptomatic carriers of V. cholerae have been documented, but they are not thought to be a significant reservoir for maintaining the organism between outbreaks. 

Pathogenesis and Spectrum of Disease As a notorious pathogen, V. cholerae elaborates several toxins and factors that play important roles in the organism’s virulence. Cholera toxin (CT) is primarily responsible for the key features of cholera (Table 25.2). Release of this toxin causes mucosal cells to hypersecrete water and electrolytes into the lumen of the gastrointestinal tract. The result is profuse, watery diarrhea, leading to dramatic fluid loss. The fluid loss results in severe dehydration and hypotension that, without medical intervention, frequently leads to death.

TABLE 25.2    Pathogenesis and Spectrum of Diseases

Species

Virulence Factors

Spectrum of Disease and Infections

Vibrio cholerae

Cholera toxin (CT); zonula occludens (Zot) toxin (enterotoxin); accessory cholera enterotoxin (Ace) toxin; O1 and O139 somatic antigens, hemolysin/cytotoxins, motility, chemotaxis, mucinase, and toxin coregulated (TCP) pili.

Cholera: profuse, watery diarrhea leading to dehydration, hypotension, and often death; occurs in epidemics and pandemics that span the globe May also cause nonepidemic diarrhea and, occasionally, extra intestinal infections of wounds, respiratory tract, urinary tract, and central nervous system.

Vibrio alginolyticus

Specific virulence factors for the non–V. cholerae species uncertain.

Ear infections, wound infections; rare cause of septicemia; involvement in gastroenteritis is uncertain.

Vibrio cincinnatiensis

Rare cause of septicemia.

Photobacterium damsela

Wound infections and rare cause of septicemia.

Vibrio fluvialis

Gastroenteritis.

Vibrio furnissii

Rarely associated with human infections.

Grimontia hollisae

Gastroenteritis; rare cause of septicemia.

Vibrio metschnikovii

Rare cause of septicemia; involvement in gastroenteritis is uncertain.

Vibrio mimicus

Gastroenteritis; rare cause of ear infection.

Vibrio vulnificus

Wound infections and septicemia; involvement in gastroenteritis is uncertain.

Aeromonas spp.

Produces various toxins and factors, but their specific role in virulence is uncertain.

Gastroenteritis, wound infections, bacteremia, and miscellaneous other infections, including endocarditis, meningitis, pneumonia, conjunctivitis, and osteomyelitis.

Chromobacterium violaceum

Endotoxin, adhesins, invasins, and cytolytic proteins have been described.

Rare but dangerous infection. Begins with cellulitis or lymphadenitis and can rapidly progress to systemic infection with abscess formation in various organs and septic shock.

Chapter 25  Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum

This toxin-mediated disease does not require the organism to penetrate the mucosal barrier. Therefore blood and the inflammatory cells typical of dysenteric stools are notably absent in cholera. Instead, “rice water stools,” composed of fluids and mucous flecks, are the hallmark of CT activity. V. cholerae is divided into three major subgroups: V. cholerae O1, V. cholerae O139, and V. cholerae non-O1/non-O139. The somatic antigens O1 and O139 associated with the V. cholerae cell envelope are positive markers for strains capable of epidemic and pandemic spread of the disease. Strains carrying these markers almost always produce CT, whereas non-O1/non-O139 strains do not produce the toxin and hence do not produce cholera. Therefore, although these somatic antigens are not virulence factors per se, they are important virulence and epidemiologic markers that provide important information about V. cholerae isolates. The non-O1/non-O139 strains are associated with nonepidemic diarrhea and extraintestinal infections. V. cholerae produces several other toxins and factors that aid in the pathogen’s ability to colonize, but the exact role of each in disease is still uncertain (Table 25.2). To effectively release toxin, the organism first must infiltrate and distribute itself along the cells lining the mucosal surface of the gastrointestinal tract. Motility and chemotaxis mediate the distribution of organisms, and mucinase production allows penetration of the mucous layer. Toxin coregulated pili (TCP) provide the means by which bacilli attach to mucosal cells for release of CT. The enterotoxin zona occludens toxin (Zot) has been shown to disrupt the tight junctions of the intestinal cells, effectively decreasing tissue resistance. Depending on the species, other vibrios are variably involved in three types of infection: gastroenteritis, wound infections, and bacteremia. Although some of these organisms have not been definitively associated with human infections, others, such as Vibrio vulnificus, are known to cause fatal septicemia, especially in patients suffering from an underlying liver disease. Aeromonas spp. are similar to Vibrio spp. in terms of the types of infections they cause. Although these organisms can cause gastroenteritis, especially in children, their role in intestinal infections is not always clear. Therefore, the significance of their isolation in stool specimens should be interpreted with caution. Severe watery diarrhea has been associated with Aeromonas strains that produce a heat-labile enterotoxin and a heat-stable enterotoxin. In addition to diarrhea, complications of infection with Aeromonas spp. include hemolytic-uremic syndrome (HUS) and kidney disease. C. violaceum is not associated with gastrointestinal infections, but acquisition of this organism by contamination of wounds can lead to fulminant, life-threatening systemic infections. 

Laboratory Diagnosis Specimen Collection and Transport Because no special considerations are required for isolation of these genera from extra intestinal sources, the general specimen collection and transport information provided in Table 5.1 is

407

• Fig. 25.1  Gram stain of Vibrio parahaemolyticus.

applicable. However, stool specimens suspected of containing Vibrio spp. should be collected and transported in Cary-Blair medium. Buffered glycerol saline is not acceptable, because glycerol is toxic to vibrios. Feces is preferable, but rectal swabs are acceptable during the acute phase of diarrheal illness. 

Specimen Processing No special considerations are required for processing of the organisms discussed in this chapter. Refer to Table 5.1 for general information on specimen processing. 

Direct Detection Methods V. cholerae toxin can be detected in stool using an enzymelinked immunosorbent assay (ELISA) or a commercially available latex agglutination test (Oxoid, Inc., Odgensburg, NY), but these tests are not widely used in the United States. A variety of rapid antigen tests are available worldwide, including the SMART II Cholera O1 and Bengal SMART O139 (investigational use only) (New Horizon Diagnostics Corporation, Columbia, MD) and immunochromatographic assays such as SD Bioline Cholera Ag O1/O139 (Standard Diagnostics, Gyeonggi-do, Republic of Korea) for the detection of V. cholerae in stool. The sensitivities and specificities vary for each assay. Microscopically, vibrios are gram-negative, straight or slightly curved rods (Fig. 25.1). When stool specimens from patients with cholera are examined using dark-field microscopy, the bacilli exhibit characteristic rapid darting or shooting-star motility. However, direct microscopic examination of stools by any method is not commonly used for laboratory diagnosis of enteric bacterial infections. A variety of multiplex nucleic acid–based testing methods have been developed for the detection of V. cholerae. The xTAG gastrointestinal panel (Luminex Corporation, Austin, TX) has been evaluated for the diagnosis of infectious disease for the isolation and differentiation of a variety of enteric pathogens including Escherichia coli, Campylobacter, Yersinia, and V. cholerae. In addition, the FilmArray (BioFire Diagnostics, Salt Lake City, UT) offers a multiplex polymerase chain reaction (PCR) system with an FDA-cleared gastrointestinal panel for V.

408 PA RT I I I    Bacteriology

cholerae and Vibrio (parahaemolyticus, vulnificus, and cholerae). These assays are able to provide a quick diagnosis that appears sensitive and specific to pathogens capable of causing diarrhea. Aeromonas spp. are gram-negative, straight rods with rounded ends or coccobacilli. Direct specimens may contain few or no white blood cells even though an infection is present. No molecular or serologic methods are available for direct detection of Aeromonas spp. Cells of C. violaceum are slightly curved, medium to long, gram-negative rods with rounded ends. A PCR amplification assay has been developed for the identification of C. violaceum. 

Cultivation Media of Choice Stool cultures for Vibrio spp. are plated on the selective medium thiosulfate citrate bile salts sucrose (TCBS) agar. TCBS contains 1% sodium chloride, bile salts that inhibit the growth of Gram-positive organisms, and sucrose for the differentiation of the various Vibrio spp. Bromothymol blue and thymol blue pH indicators are added to the medium. The high pH of the medium (8.6) inhibits the growth of other intestinal microbiota. Although some Vibrio spp. grow very poorly on this medium, those that grow well produce either yellow or green colonies, depending on whether they are able to ferment sucrose (which produces yellow colonies). Alkaline peptone water (pH 8.4) may be used as an enrichment broth for obtaining growth of vibrios from stool. After inoculation, the broth is incubated for 5 to 8 hours at 35°C and then subcultured to TCBS. Oxidase testing is unreliable when performed on colonies grown on TCBS media. The organisms will also grow on MacConkey or salmonella shigella (SS) agar. All species will appear as nonfermenters with the exception of V. vulnificus. Other species may be indistinguishable from other rapid sucrose-fermenting enterics when grown on Hektoen enteric (HE) or xylose-lysine-deoxycholate (XLD) media. Refer to Chapter 19 for a description of the enteric media included here. Chromogenic Vibrio agar (CHROMagar Microbiology, Paris, France), which was developed for the recovery of Vibrio parahaemolyticus from seafood, supports the growth of other Vibrio spp. Colonies on this agar range from white to pale blue and violet. Aeromonas spp. are indistinguishable from Yersinia enterocolitica on modified cefsulodin-irgasan-novobiocin (CIN) agar (4 μg/mL of cefsulodin); therefore, it is important to perform an oxidase test to differentiate the two genera; Aeromonas spp. are oxidase positive. Aeromonas agar is a relatively new alternative medium that uses D-xylose as a differential characteristic. These organisms typically grow on a variety of differential and selective agars used for the identification of enteric pathogens. They are also beta-hemolytic on blood agar. C. violaceum grows on most routine laboratory media. The colonies may be beta-hemolytic and have an almondlike odor. Most strains produce violacein, an ethanolsoluble violet pigment. All of the genera considered in this chapter grow well on 5% sheep blood, chocolate, and MacConkey agars. They

•

Fig. 25.2  Colonies of Chromobacterium violaceum on DNase agar. Note violet pigment.

also grow well in the broth of blood culture systems and in thioglycollate or brain-heart infusion broths. 

Incubation Conditions and Duration These organisms produce detectable growth on 5% sheep blood and chocolate agars when incubated at 35°C in carbon dioxide or ambient air for a minimum of 24 hours. MacConkey and TCBS agars should be incubated at 35°C in ambient air. The typical violet pigment of C. violaceum colonies (Fig. 25.2) is optimally produced when cultures are incubated at room temperature (22°C). 

Colonial Appearance Table 25.3 describes the colonial appearance and other distinguishing characteristics (e.g., hemolysis and odor) of each genus on 5% sheep blood and MacConkey agars. The appearance of Vibrio spp. on TCBS is described in Table 25.4 and shown in Fig. 25.3. 

Approach to Identification The colonies of these genera resemble those of the Enterobacterales but can be distinguished notably by their positive oxidase test result (except for V. metschnikovii, which is oxidase-negative). The oxidase test must be performed from 5% sheep blood or another medium without a fermentable sugar (e.g., lactose in MacConkey agar or sucrose in TCBS), because fermentation of a carbohydrate results in acidification of the medium, and a false-negative result may occur if the surrounding pH is below 5.1. Likewise, if the violet pigment of a suspected C. violaceum isolate interferes with performance of the oxidase test, the organism should be grown under anaerobic conditions (where it cannot produce pigment) and retested. The reliability of commercial identification systems has not been widely validated for identification of these organisms, although most are listed in the databases of several systems. The API 20E system (bioMérieux, St. Louis, MO) is one of the best systems available for the identification of

Chapter 25  Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum

409

TABLE 25.3    Colonial Appearance and Characteristics

Organism

Medium

Appearance

Aeromonas spp.

BA Mac

Large, round, raised, opaque; most pathogenic strains are beta-hemolytic except Aeromonas caviae, which is usually nonhemolytic Both NLF and LF

Chromobacterium violaceum

BA Mac

Round, smooth, convex; some strains are beta-hemolytic; most colonies appear black or very dark purple; cultures smell of ammonium cyanide (almondlike) NLF

Vibrio spp. and Grimontia hollisae

BA Mac

Medium to large, smooth, opaque, iridescent with a greenish hue; V. cholerae, V. fluvialis, and V. mimicus can be beta-hemolytic NLF except V. vulnificus, which may be LF

Photobacterium damsela

BA Mac

Medium to large, smooth, opaque, iridescent with a greenish hue; may be beta-hemolytic NLF

BA, 5% sheep blood agar; LF, lactose fermenter; Mac, MacConkey agar; NLF, non–lactose fermenter.

vibrios because the inoculum is prepared in 0.85% saline; the amount of salt is sufficient to allow growth of the halophilic organism. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) could provide a rapid identification method for the identification of vibrios isolated from clinical specimens. The VITEK MS (bioMérieux, St. Louis, MO) database includes V. cholerae, V. parahaemolyticus, V. vulnificus, as well as V. alginolyticus, fluvialis, metschnikovii, and mimicus. In a recent study, Cheng et al. (2015) evaluated the identification of V. vulnificus, V. parahaemolyticus, V. fluvialis, and 33 species of O1 and nonO139 V. cholerae. The system was not able to identify the serogroup types O1 and non-O139 to the species level. Although MALDI-TOF MS does allow for the accurate identification of some species of Aeromonas hydrophila/ punctate, sobria, and jandaei, it should be noted that the ability of most commercial identification systems to accurately identify Aeromonas organisms to the species level is limited and uncertain, and with some kits, difficulty arises in separating Aeromonas spp. from Vibrio spp. Identification of potential pathogens should be confirmed using conventional biochemical tests or serotyping. Tables 25.4 and 25.5 show several characteristics that can be used to presumptively group Vibrio spp., Aeromonas spp., and C. violaceum. New technologies such as nucleic acid–based testing and MALDI-TOF MS for additional isolates discussed in this chapter are typically not available because of clinical efficacy and database limitations.

Comments Regarding Specific Organisms V. cholerae and V. mimicus are the only Vibrio spp. that do not require salt for growth. Therefore, a key test for distinguishing the halophilic species from V. cholerae, V. mimicus, and Aeromonas spp. is growth in nutrient broth with 6% salt. Furthermore, the addition of 1% NaCl to conventional biochemical tests is recommended to allow growth of halophilic species.

The string test can be used to differentiate Vibrio spp. from Aeromonas spp. In this test, organisms are emulsified in 0.5% sodium deoxycholate, which lyses Vibrio cells but not those of Aeromonas spp. Cell lysis releases deoxyribonucleic acid (DNA), which can be pulled up into a string with an inoculating loop (Fig. 25.4). A vibrostatic test using 0129 (2,4-diamino-6,7-diisopropylpteridine)–impregnated disks also has been used to separate vibrios (susceptible) from other oxidase-positive glucose fermenters (resistant) and to differentiate V. cholerae O1 and non-O1 (susceptible) from other Vibrio spp. (resistant). However, recent strains of V. cholerae O139 and V. cholera O1 have demonstrated resistance; therefore, the dependability of this test is questionable. Serotyping should be performed immediately to further characterize V. cholerae isolates. Toxigenic strains of serogroup O1 and O139 can be involved in cholera epidemics. Strains that do not type in either antiserum are identified as non-O1. Although typing sera are commercially available, isolates of V. cholerae are usually sent to a reference laboratory for serotyping. Identification of V. cholerae or V. vulnificus should be reported immediately because of the life-threatening nature of these organisms. Aeromonas spp. and C. violaceum can be identified using the characteristics shown in Table 25.5. Aeromonas spp. identified in clinical specimens should be identified as A. hydrophilia, A. caviae complex, or A. veronii complex. Pigmented strains of C. violaceum are so distinctive that a presumptive identification can be made based on colonial appearance, oxidase, and Gram staining. Nonpigmented strains (approximately 9% of isolates) may be differentiated from Pseudomonas, Burkholderia, Brevundimonas, and Ralstonia organisms based on glucose fermentation and a positive test result for indole. Negative lysine and ornithine reactions are useful criteria for distinguishing C. violaceum from Plesiomonas shigelloides. In addition to the characteristics listed in

Species

Oxidase

Indole

Gas From Glucose

Grimontii hollisae

+

+

−

−

−

−

−

−

−

+

Very poor

Green

Vibrio alginolyticus

+

V

−

−

+

+

−

V

−

+

Good

Yellow

Vibrio cholerae

+

+

−

V

+

+

−

+

+

V

Good

Yellow

Vibrio cincinnatiensisd

+

V

−

−

+

V

−

−

−

+

Very poor

Yellow

Photobacterium damsela

+

−

−

−

−

V

+

−

−

+

Reduced at 36°C

Greene

Vibrio fluvialis

+

V

−

−

+

−

+

−

−

+

Good

Yellow

Vibrio furnissii

+

V

+

−

+

−

+

−

−

+

Good

Yellow

Vibrio harveyi

+

+

−

−

V

+

−

−

−

+

Good

Yellow

Vibrio metschnikovii

−

V

−

V

+

V

V

−

−

V

May be reduced

Yellow

Vibrio mimicus

+

+

−

V

−

+

−

+

+

V

Good

Green

Vibrio parahaemolyticus

+

+

−

−

−

+

−

+

−

+

Good

Greenf

Vibrio vulnificus

+

+

−

(+)

−

+

−

+

−

+

Good

Greeng

a1%

Lactose

Sucrose

Lysine Decarboxylasea

Arginine Dihydrolasea

Ornithine Decar‑ boxylasea

Growth in 0% NaClb

Growth in 6% NaClb

TCBSc Growth

Colony on TCBSc

NaCl added to enhance growth. broth with 0% or 6% NaCl added. cThiosulfate citrate bile salts sucrose agar. dFerments myoinositol. e5% yellow. f1% yellow. g0% yellow. +, >90% of strains are positive; −, >90% of strains are negative; (+), delayed; V, variable. bNutrient

410 PA RT I I I    Bacteriology

TABLE 25.4    Key Biochemical and Physiologic Characteristics of Vibrio spp. and Grimontii hollisae

Chapter 25  Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum

A

B

411

Antimicrobials reduce the severity of the illness and shorten the duration of organism shedding. The Clinical and Laboratory Standards Institute (CLSI) has established methods for testing for V. cholerae, and the CLSI document should be consulted for this purpose. The need for antimicrobial intervention for gastrointestinal infections caused by other Vibrio spp. and Aeromonas spp. is less clear. However, extraintestinal infections with these organisms and with C. violaceum can be life threatening, and directed therapy is required. C. violaceum is often resistant to beta-lactams and colistin. Antimicrobial agents with potential activity are listed, where appropriate, in Table 25.6. It is important to note these organisms’ ability to show resistance to therapeutic agents; especially noteworthy is the ability of Aeromonas spp. to produce various beta-lactamases. 

Prevention • Fig. 25.3  Colonies of Vibrio cholerae (A) and Vibrio parahaemolyticus (B) on thiosulfate citrate bile salts sucrose agar.

No cholera vaccine is available in the United States. However, the World Health Organization (WHO) now maintains a global stockpile of cholera vaccine to use in endemic hot spots and in emergency situations. Because of recent outbreaks, WHO developed a Global Task Force on Cholera Control. Information is available at http://www.who.int/cholera/task_ force/en/. No approved vaccines or chemoprophylaxis exists for the other organisms discussed in this chapter.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography

• Fig. 25.4  String test used to differentiate Vibrio spp. (positive) from Aeromonas spp. and Plesiomonas shigelloides (negative).

Table 25.5, failure to ferment either maltose or mannitol also differentiates C. violaceum from Aeromonas spp. 

Serodiagnosis Agglutination, vibriocidal, or antitoxin tests are available for diagnosing cholera using acute and convalescent sera. However, these methods are most commonly used for epidemiologic purposes. Serodiagnostic techniques are not generally used for laboratory diagnosis of infections caused by the other organisms discussed in this chapter. 

Antimicrobial Susceptibility Testing and Therapy Two components of the management of patients with cholera are rehydration and antimicrobial therapy (Table 25.6).

Barzilay EJ, Schaad N, Magloire R, et al.: Cholera surveillance during the Haiti epidemic—the first 2 years, N Engl J Med 368:599–609, 2013. Caroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Cheng WC, Jan IS, Chen JM, et  al.: Evaluation of the Bruker Biotyper matrix-assisted laser desorption ionization-time of flight mass spectrometry system for the identification of blood isolates of Vibrio species, J Clin Microbiol 53:1741–1744, 2015. Clark RB, Lister PD, Arneson-Rotert L, Janda JM: In vitro susceptibilities of Plesiomonas shigelloides to 24 antibiotics and antibiotic-beta-lactamase-inhibitor combinations, Antimicrob Agents Chemother 34:159–160, 1990. Colwell RR: Global climate and infectious disease: the cholera paradigm, Science 274:2025–2031, 1996. Committee on Infectious Diseases: 2006 red book: report of the Committee on Infectious Diseases, ed 27, Elk Grove Village, IL, 2006, American Academy of Pediatrics. Erler R, Wichels A, Heinemeyer EA, et al.: VibrioBase: a MALDI-TOF MS database for fast identification of Vibrio spp. that are potentially pathogenic in humans, Syst Appl Microbiol 38:16–25, 2015. Feng W, Gu X, Sui W, et  al.: The application and epidemiological research of xTAG GPP multiplex PCR in the diagnosis of infectious diarrhea, Zhonghua Yi Xue Za Zhi 95:435–439, 2015. Jensen J, Jellinge ME: Severe septic shock and cardiac arrest in a patient with Vibrio metschnikovii: a case report, J Med Case Rep 8:348, 2014.

412 PA RT I I I    Bacteriology

TABLE 25.5    Key Biochemical and Physiologic Characteristics of Aeromonas spp. and Chromobacterium violaceum

Species

Oxidase

Indole

Gas From Glucose

Esculin Hydrolysis

Fermentation of Sucrose

Lysine Decarboxylase

Arginine Dihydrolase

Ornithine Decarbo‑ xylase

Growth in 0% NaCla

Growth in 6% NaCla

TCBSb Growth

Aeromonas caviae complex

+

V

−

+

+

−

+

−

+

−

−

Aeromonas hydrophila complex

+

+

V

V

V

V

+

−

+

V

−

Aeromonas jandaei (A. veronii complex)

+

+

+

−

−

+

+

−

+

−

−

Aeromonas schubertii (A. veronii complex)

+

V

−

−

−

+

+

−

+

−

−

Aeromonas veronii biovar sobria

+

+

+

−

+

+

+

−

+

−

−

Aeromonas veronii biovar veronii

+

+

+

+

+

+

−

+

+

−

−

Chromobacterium violaceumc

V

V

−d

−

V

−

+

−

+

−

ND

+, >90% of strains are positive; –, >90% of strains are negative; ND, no data; V, variable. aNutrient agar with 0% or 6% NaCl added. bThiosulfate citrate bile salts sucrose agar. c91% produce an insoluble violet pigment; often, nonpigmented strains are indole-positive. dGas-producing strains have been described.

Chapter 25  Vibrio, Aeromonas, Plesiomonas shigelloides, and Chromobacterium violaceum

413

TABLE 25.6    Antimicrobial Therapy and Susceptibility Testing

Potential Resistance to Therapeutic Options

Validated Testing Methodsa

Species

Therapeutic Options

Vibrio cholerae

Adequate rehydration plus antibiotics. In vitro susceptibility guidelines indicate susceptibilities to aminoglycosides, azithromycin, fluoroquinolones, extended-spectrum cephalosporins, carbapenems, and monobactams.

Known resistance to tetracycline, chloramphenicol, and trimethoprimsulfamethoxazole

Refer to CLSI guidelines.

Other Vibrio spp.

No definitive guidelines. For gastroenteritis, therapy may not be needed; for wound infections and septicemia; same as for V. cholerae.

Similar to resistance reported for V. cholerae.

Refer to CLSI guidelines.

Aeromonas spp.

No definitive guidelines. For gastroenteritis, therapy may not be needed; for soft tissue infections and septicemia, potentially active agents include ceftriaxone, cefotaxime, ceftazidime, imipenem, aztreonam, amoxicillin-clavulanate, quinolones, and trimethoprim-sulfamethoxazole.

Capable of producing various beta-lactamases that mediate resistance to penicillins and certain cephalosporins.

Refer to CLSI guidelines.

Chromobacterium violaceum

No definitive guidelines. Potentially active agents include cefotaxime, ceftazidime, imipenem, and aminoglycosides.

Variable activity of penicillins; poor activity of first- and second-generation cephalosporins.

Not available.

Comments

Grows on MuellerHinton agar, but interpretive standards do not exist.

aValidated

testing methods include standard methods recommended by the Clinical and Laboratory Standards Institute (CLSI) and commercial methods approved by the US Food and Drug Administration (FDA).

Kimura B, Hokimoto S, Takahasi H: Photobacterium histaminum (Okuzumi et  al, 1994) is a later subjective synonym for Photobacterium damselae subsp damselae (Love et al, 1981; Smith et al, 1991), Int J Syst Evol Microbiol 50:1339–1342, 2000. Spina A, Kerr KG, Cormican M, et  al.: Spectrum of enteropathogens detected by FilmArray GI Panel in a multicentre study of communityacquired gastroenteritis, Clin Microbiol Infect 21:719–728, 2015. Thompson FL, Hoste B, Vandemeulebroecke K, Swings J: Reclassification of Vibrio hollisae as Grimontia hollisae gen nov, comb nov, Int J Syst Evol Microbiol 53:1615–1617, 2003.

Ti TY, Tan CW, Chong AP, Lee EH: Nonfatal and fatal infections caused by Chromobacterium violaceum, Clin Infect Dis 17:505– 507, 1993. Wei S, Zhao H, Xian Y, Hussain MA, Wu X: Multiplex PCR assays for the detection of Vibrio alginolyticus, Vibrio parahaemolyticus, Vibrio vulnificus and Vibrio cholerae with an internal amplification control, Diagn Microbiol Infect Dis 79:115–118, 2014.

CASE STUDY 25.1 After vacationing in San Diego, a 21-year-old male surfer sees his physician complaining of severe left ear pain. He is afebrile, but the auditory canal and tympanic membrane are erythematous. Amoxicillin is prescribed for presumed otitis media. Over the next 4 days, the symptoms persist and a bloody discharge develops. The patient returns to his physician, who cultures the drainage and prescribes gentamicin eardrops. The patient’s symptoms improve over the next 7 days. On culture, a non–lactose fermenter was isolated from MacConkey agar.

Questions 1. The isolate is indole- and oxidase-positive. A biochemical identification system had positive reactions for lysine and ornithine but not arginine. What genus and species of bacteria are in the differential and how would you identify this microorganism? 2. How do you think the patient acquired this infection? 3. Commercial systems are known to misidentify the Vibrio spp. as Aeromonas spp. and vice versa. What is the reason for such a critical error? 4. Susceptibility testing using the disk method is not problematic for Vibrio spp. as long as which extra step is taken with testing?   

Chapter Review 1. A patient presents with diarrhea after spending 2 weeks in Haiti after the country’s devastation by an earthquake. A stool specimen is collected and inoculated to enrichment broth before subculturing to TCBS. After 48 hours of incubation on TCBS, no growth is identified on the media. What should the laboratory scientist do next? a. Request a new specimen. b. Run quality control organisms to check the integrity of the TCBS media. c. Report the culture as no growth with a comment that indicates the organism may be viable but nonculturable, and the result does not rule out the presence of an infection. d. Report all cultures as no growth. 2. A stool specimen is submitted for culture. The results are beta-hemolytic on blood agar, NLF on MacConkey, oxidase positive, bull’s-eye appearance on CIN agar. This organism is most likely: a.  A. hydrophilia b. Y. enterocolitica c.  C. violaceum d. G. hollisae 3. A suspected isolate of Vibrio spp. is isolated from a young child with diarrhea. The organism is identified as a curved gram-negative rod, oxidase- and lactosepositive, sucrose-negative, that produces yellow colonies on TCBS and is NaCl tolerant. This organism is most likely: a.  V. mimicus b. V. furnissii c.  V. cholerae d. V. fluvialis

4.  Matching: Match each term with the corresponding term or description. _____ halophilic _____ Zot _____ C. violaceum _____ Aeromonas spp. _____ V. alginolyticus _____ V. cholerae

a. ear infections b. cellulitis and abscess formation c. salt-loving d. enterotoxin e. gastroenteritis and endocarditis f. profuse watery diarrhea

5.  Short Answer (1) Describe the string test and how it is used to differentiate Vibrio spp. from Aeromonas spp. (2) What simple biochemical test can be used to differentiate V. cholerae, V. mimicus, and Aeromonas spp. from the other organisms discussed in this chapter? (3) Explain the chemical principle for the selective and differential properties of TCBS.

413.e1

S E C T I ON 9    Gram-Negative Bacilli and Coccobacilli (MacConkey-Negative, Oxidase-Positive)

26

Sphingomonas and Similar Organisms OBJECTIVES 1. Identify cultivation methods and colonial characteristics for Sphingomonas paucimobilis and similar organisms. 2. State the initial clues that alert clinical laboratorians to the presence of this group of organisms in a patient specimen. 3. Explain the classification and identification algorithm for this group of organisms. 4. Describe the susceptibility testing methods appropriate for this group of organisms. 5. Explain the pathogenicity of organisms in this group.

ORGANISMS TO BE CONSIDERED Current Name Sphingobacterium mizutaii Sphingomonas parapaucimobilis Sphingomonas paucimobilis

Previous Name

Pseudomonas paucimobilis, CDC IIk-1

genus Sphingobacterium is ubiquitous in nature in soil and aquatic environments. The only species discussed here, Sphingobacterium mizutaii, is the only species that is indole-positive and fails to grow on MacConkey agar. The remaining species are indole-negative and generally grow on MacConkey agar (Chapter 23). Sphingomonas spp. are associated with natural aquatic sources. Sphingomonas spp. have been isolated in cases of keratitis associated with contact lenses, bacteremia, and artificial medical device implants. These organisms are opportunistic pathogens, and when encountered in clinical specimens, their clinical significance and potential as contaminants should be carefully considered. 

Laboratory Diagnosis Specimen Collection and Transport No special considerations are required for specimen collection and transport of the organisms discussed in this chapter. Refer to Table 5.1 for general information on specimen collection and transport. 

General Considerations

Specimen Processing

The organisms discussed in this chapter are considered together because they usually fail to grow on MacConkey agar, are oxidase-positive, and oxidatively utilize glucose. 

No special considerations are required for processing of the organisms discussed in this chapter. Refer to Table 5.1 for general information on specimen processing. 

Epidemiology, Spectrum of Disease, and Antimicrobial Therapy

Direct Detection Methods

As demonstrated in Table 26.1, these organisms are rarely or only occasionally isolated from human specimens and have limited roles as agents of infection. Because they are infrequently encountered in the clinical setting, little information is available on their epidemiology, ability to cause human infections, and potential for antimicrobial resistance. The 414

No specific procedures other than microscopy are required for direct detection of these organisms in clinical material. 

Serodiagnosis Serodiagnostic techniques are not generally used for the laboratory diagnosis of infections caused by the organisms discussed in this chapter. 

CHAPTER 26  Sphingomonas and Similar Organisms

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TABLE 26.1    Epidemiology, Spectrum of Disease, and Antimicrobial Therapy

Organism

Epidemiology

Disease Spectrum

Antimicrobial Therapy

Sphingobacterium mizutaii

Sphingobacteria are ubiquitous in nature.

Rarely involved in human infections; has been associated with blood, cerebrospinal fluid, and wound infections.

Erythromycin, trimethoprimsulfamethoxazole, and pefloxacin.

Sphingomonas paucimobilis Sphingomonas parapaucimobilis

S. paucimobilis inhabits environmental niches and can exist in hospital water systems. Not part of human microbiota. Mode of transmission is uncertain but probably involves patient exposure to contaminated medical devices or solutions.

S. paucimobilis virulence factors are unknown. It has been implicated in community- and health care–associated infections, specifically in blood and urine infections.

No definitive guidelines; potentially active agents include trimethoprimsulfamethoxazole, chloramphenicol, ciprofloxacin, and aminoglycosides; resistance to beta-lactams is known, but validated susceptibility testing methods do not exist.

TABLE 26.2    Colonial Appearance and Characteristics

Organism

Medium

Appearance

Sphingobacterium mizutaii

BA

Yellow pigmented colonies

Sphingomonas paucimobilis Sphingomonas parapaucimobilis

BA

Small, circular, smooth, convex; bright yellow growth pigment

BA, 5% sheep blood agar.

Cultivation Media of Choice Sphingomonas spp. and S. mizutaii grow well on routine laboratory media, such as 5% sheep blood and chocolate agars; however, they fail to grow on MacConkey agar. They usually grow well in thioglycollate and brain-heart infusion broths and in broths used in blood culture systems. 

Incubation Conditions and Duration Within 24 to 48 hours of inoculation and incubation, most of these organisms produce detectable growth on media incubated at 35°C to 37°C in 5% carbon dioxide (CO2) or ambient air. 

Colonial Appearance Table 26.2 describes the colonial appearance and distinguishing characteristics (e.g., pigment) of each organism on 5% sheep blood agar. 

Approach to Identification The ability of many commercial identification systems, including matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS), to

accurately identify the organisms discussed in this chapter may be limited or uncertain. Tables 26.3 and 26.4 show some biochemical tests that are helpful for presumptive differentiation among the various organisms in this group.

Comments Regarding Specific Organisms Sphingobacterium mizutaii

S. mizutaii exhibits II-forms. It can produce a yellow pigment, and it does not grow on MacConkey agar. Frequently classified as nonmotile, the organism can be motile by gliding movement. It is able to grow in the presence of 40% bile, and it is oxidase-positive, catalase-positive, esculinpositive, indole-negative, and urease-negative (although a report exists that 20% are positive for Christensen urease). Key characteristics are shown in Table 26.3. Reported infections in humans have included septicemia (blood culture), meningitis (CSF specimen), and cellulitis (wound source). This bacterium has been reported to be susceptible to erythromycin, trimethoprim-sulfamethoxazole, and pefloxacin.  Sphingomonas paucimobilis

S. paucimobilis is a medium-size, straight, gram-negative rod with a single polar flagellum; growth requires at least 48 hours’ incubation on sheep blood agar (Fig. 26.1). Optimal growth occurs at 30°C in 5% CO2 or ambient air; it does grow at

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TABLE 26.3    Key Biochemical and Physiologic Characteristics

Insoluble Pigment

Glucose Oxidized

Xylose Oxidized

Sucrose Oxidized

Esculin Hydrolysis

Motility

Sphingobacterium mizutaii

Va

＋

(＋)

＋

＋

nm

Sphingomonas spp.b

Yellow

＋

＋

＋

＋

＋c

Organism

aYellow

pigment production may be enhanced by incubation at room temperature. S. paucimobilis and S. parapaucimobilis. cUsually nonmotile in motility medium, but motility is present on wet mount. ＋, ＞90% strains positive; (＋), delayed; nm, nonmotile; V, variable. From Weyant RS, Moss CW, Weaver RE, et al, eds. Identification of Unusual Pathogenic Gram-Negative Aerobic and Facultatively Anaerobic Bacteria. 2nd ed. Baltimore: Williams & Wilkins; 1996. bIncludes

TABLE 26.4    Specific Biochemical Characteristics for Differentiation of the Sphingomonas spp.

Biochemical Test

S. paucimobilis

S. parapaucimobilis

Oxidation of glucose

Positive

Positive

Oxidation of xylose

Positive

Positive

Oxidation of maltose

Positive

Positive

Esculin hydrolysis

Positive

Positive

Motility

Positivea

Positive

Indole

Negative

Negative

Susceptibility to polymyxin B

Susceptible

Variable

H2S (lead acetate paper suspended over KIA)

Negative

Positive

Citrate

Negative

Positive

DNase

Positive

Negative

aMotility

positive by wet mount or in motility medium incubated at 18°C to 22°C, but organism is nonmotile when incubated at 37°C. H2S, Hydrogen sulfide; KIA, Kligler iron agar. Data compiled from Winn WC, Allen SD, Janda WM, et al. Koneman’s Color Atlas and Textbook of Diagnostic Microbiology. 6th ed. Philadelphia: Lippincott Williams & Wilkins; 2006.

37°C but not at 42°C. It grows as a deep yellow colony on tryptic soy and blood agars. It is obligately aerobic, it grows in broth (e.g., brain-heart infusion, thioglycollate, blood culture media), and 90% of isolates do not grow on MacConkey agar (10% grow on MacConkey agar and appear as non– lactose fermenters). S. paucimobilis oxidatively utilizes glucose, xylose, and sucrose. Biochemical test results of interest include the following: esculin hydrolysis–positive, motile by wet mount or in motility medium when incubated at 18°C to 22°C (nonmotile when incubated at 37°C), oxidase-positive (90% to 94% positive), catalase-positive, urease-negative, and indole-negative. S. paucimobilis is susceptible to polymyxin B, a trait that distinguishes it from Sphingobacterium spp. Key characteristics are included in Table 26.4. Antimicrobial susceptibility testing indicates that S. paucimobilis is susceptible to tetracycline, chloramphenicol, trimethoprim-sulfamethoxazole, and aminoglycosides.

Susceptibility to vancomycin has been noted when the organism is grown on sheep blood agar with a vancomycin disk (30 μg). S. paucimobilis is ubiquitous in soil and water and has been isolated environmentally from swimming pools, hospital equipment, and water and laboratory supplies. It has been associated with human infections and found in a variety of clinical specimens, specifically, peritonitis associated with wound infections (chronic ambulatory peritoneal dialysis, leg ulcer, empyema, splenic abscess, brain abscess); blood cultures; and CSF, urine, vaginal, and cervical samples. Recent literature indicates that S. paucimobilis is regarded as having minor clinical significance; however, community-acquired infection, diabetes mellitus, and alcoholism are significant risk factors for primary bacteremia. A retrospective study suggests that the prevalence of S. paucimobilis infection in humans seems to have increased in recent times, and although it

CHAPTER 26  Sphingomonas and Similar Organisms

•

Fig. 26.1  Sphingomonas paucimobilis growth on blood agar. (From Seo SW, Chung IY, Kim E, et  al. A case of postoperative Sphingomonas paucimobilis endophthalmitis after cataract extraction. Kor J Ophthalmol. 2008;22:63.)

has low virulence, infection can lead to septic shock, particularly in immunocompromised patients. Another report indicates that although this bacterium has low mortality associated with infection, it commonly causes complications in hospitalized patients.  Sphingomonas parapaucimobilis

417

as indicated by blackening of lead acetate paper suspended over Kligler iron agar (KIA), it is Simmons citrate positive (S. paucimobilis is negative), and it is negative for extracellular DNAse (S. paucimobilis is positive). Like S. paucimobilis, S. parapaucimobilis is positive for OF glucose, OF xylose, and OF maltose but negative for OF mannitol. It has been distinguished from Sphingobacterium spp. by its susceptibility to polymyxin B; however, S. parapaucimobilis may demonstrate variable susceptibility to polymyxin B. Key characteristics are included in Table 26.4. Antimicrobial susceptibility testing indicates that S. parapaucimobilis displays variable resistance but is usually susceptible to tetracycline, chloramphenicol, sulfamethoxazole, aminoglycosides, third-generation cephalosporins, and fluoroquinolone. S. parapaucimobilis has been associated with human infections; specifically, it has been isolated from sputum, urine, and the vagina. 

Antimicrobial Susceptibility Antimicrobial susceptibility for this group of bacteria ranges from variable resistance to identifiable patterns of susceptibility. Standardized guidelines are not available. However, when clinically necessary, susceptibility testing should be completed using an overnight MIC or E-test method. 

Prevention

S. parapaucimobilis is similar to S. paucimobilis in many ways. It is a medium-sized, straight, gram-negative rod that produces a deep yellow pigment. It is obligately aerobic, motile, and does not grow on MacConkey agar. S. parapaucimobilis can be distinguished from S. paucimobilis by several characteristics. S. parapaucimobilis is H2S positive,

Because these organisms are rarely implicated or identified in human infections, no vaccines or prophylactic measures are available.

Bibliography

Hollis DG, Moss CW, Daneshvar MI, Wallace-Shewmaker PL: CDC group IIc: phenotypic characteristics, fatty acid composition, and isoprenoid quinone content, J Clin Microbiol 34:2322–2324, 1996. Lambiase A, Rossano F, Del Pezzo M, et al.: Sphingobacterium respiratory tract infection in patients with cystic fibrosis, BMC Res Notes 2:262, 2009. Lemaitre D, Elaichouni A, Hundhausen M, et  al.: Tracheal colonization with Sphingomonas paucimobilis in mechanically ventilated neonates due to contaminated ventilator temperature probes, J Hosp Infect 32:199–206, 1996. Lin JN, Lai CH, Chen YH, et al.: Sphingomonas paucimobilis bacteremia in humans: 16 case reports and a literature review, J Microbiol Immunol Infect 43:35–42, 2010. Refaat M, Zakka P, Khoury M, et al.: Cardiac implantable electronic device infections: observational data from a tertiary care center in Lebanon, Medicine (Baltimore) 98(16).e14906, 2019.

Boken DJ, Romero JR, Cavalieri SJ: Sphingomonas paucimobilis bacteremia: four cases and review of the literature, Infect Dis Clin Pract 7:286, 1998. Caroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Charity RM, Foukas AF: Osteomyelitis and secondary septic arthritis caused by Sphingomonas paucimobilis infection, 33. 2005, pp 93–95. Daneshvar MI, Hill B, Hollis DG, et al.: CDC group O-3: phenotypic characteristics, fatty acid composition, isoprenoid quinone content, and in  vitro antimicrobic susceptibilities of an unusual gram-negative bacterium isolated from clinical specimens, J Clin Microbiol 36:1674–1678, 1998. Freney J, Hansen W, Ploton C, et  al.: Septicemia caused by Sphingobacterium multivorum, J Clin Microbiol 25:1126–1128, 1987.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

418 PA RT I I I    Bacteriology

Reina J, Bassa A, Llompart I, Portela D, Borrell N: Infections with Pseudomonas paucimobilis: report of four cases and review, Rev Infect Dis 13:1072–1076, 1991. Roca M, Garcia A, Peñas-Pardo L, Bosch-Aparicio N, Agustí J: Sphingomonas paucimobilis keratitis in a patient with neurotrophic keratopathy and severe neurosensory hypoacusis: treatment with penetrating keratoplasty and amniotic membrane grafting, Oman J Ophthalmol 11(3):291–293, 2018. Saboor F, Amin F, Nadeem S: Community acquired Sphingomonas paucimobilis in a child - A rare case, J Pak Med Assoc 68(11):1714– 1716, 2018. Salazar R, Martino R, Sureda A, Brunet S, Subirá M, Domingo-Albós A: Catheter-related bacteremia due to Pseudomonas paucimobilis in neutropenic cancer patients: report of two cases, Clin Infect Dis 20:1573–1574, 1995.

Toh HS, Tay HT, Kuar WK, Weng TC, Tang HJ, Tan CK: Risk factors associated with Sphingomonas paucimobilis infection, J Microbiol Immunol Infect 44:289–295, 2011. Weyant RS, Moss CW, Weaver RE, et  al.: Identification of unusual pathogenic gram-negative aerobic and facultatively anaerobic bacteria, ed 2, Baltimore, 1996, Williams & Wilkins. Willems A, Falsen E, Pot B, et  al.: Acidovorax, a new genus for Pseudomonas facilis, Pseudomonas delafieldii, E. Falsen (EF) group 13, EF group 16, and several clinical isolates, with the species Acidovorax facilis comb.nov., Acidovorax delafieldii comb.nov., and Acidovorax temperans sp.nov, Int J Syst Bacteriol 40:384–398, 1990. Winn WC, Allen SD, Janda WM, et  al.: Koneman’s color atlas and textbook of diagnostic microbiology, ed 6, Philadelphia, 2006, Lippincott Williams & Wilkins.

CASE STUDY 26.1 A 16-year-old male patient with acute lymphoblastic leukemia presents to his oncologist with pain and swelling of the left knee. He recently received a course of chemotherapy and radiotherapy, and he is taking oral steroids. Straw-colored fluid with 2＋ white blood cells is aspirated from his knee. No microorganisms are seen on the smear, and none grow in culture. Unfortunately, only a few drops of the fluid are cultured on plate media. Over the next 6 months, the patient is in and out of the hospital, receiving antibiotics and having more cultures done, with no positive findings to explain his pain and swelling. He is admitted to the hospital, where an arthroscopic procedure is performed to evaluate the problem. Widespread synovitis is seen. Culture samples obtained from the surgery grow a yellow-pigmented, gram-negative rod on blood agar, but no growth is observed on MacConkey agar. Indole and urease testing are negative, but the oxidase test and wet mount motility

are positive. The bacterium is identified as Sphingomonas paucimobilis. The patient is treated with a 6-week course of intravenous amikacin and ceftazidime. Despite the effectiveness of treatment, the patient is left with residual knee pain and stiffness because of articular cartilage destruction.

Questions 1. Which microorganisms are in the differential diagnosis for the patient? 2. What tests can be done to provide differential evidence for bacterial identification? 3. What method or methods should be used to test for susceptibility of the pathogens identified in this case?   

From Charity R, Foukas A. Osteomyelitis and secondary septic arthritis caused by Sphingomonas paucimobilis. Infection. 2005;33:93.

  

CASE STUDY 26.2 A 20-month-old female was diagnosed with cystic fibrosis at the age of 6 months. She is taken to the hospital on her second day of respiratory difficulty and presents with cough, abundant mucus expectoration, and a temperature of 37.9°C. Because she has a history of Pseudomonas aeruginosa infections, treatment is started with ceftazidime and amikacin. Bronchial aspirates are obtained for culture; plated on blood, chocolate, and MacConkey agars; and incubated (37°C, 48 hours). A medium specific for isolation of slow-growing Burkholderia organisms also is inoculated and incubated appropriately. Abundant growth of oxidase-positive colonies that are nonmotile, catalase-positive, gram-negative rods is identified as Sphingobacterium multivorum by means of a Vitek GNI card and API 20NE. Definitive identification is provided by biochemical tests that show the following positive results: growth on MacConkey agar; urease; esculin hydrolysis; beta-D-galactosidase production; assimilation of glucose, arabinose, mannose, N-acetylglucosamine, and maltose; and acidification of glucose, lactose, maltose, sucrose, and xylose. Negative results are identified for the following: motility at 23°C (room temperature), 37°C, and 42°C; nitrate and nitrite reduction; indole production;

arginine dihydrolase; lysine and ornithine decarboxylase; gelatin hydrolysis; hydrogen sulfide production; and assimilation of mannitol, gluconate, malate, and citrate. Antimicrobial susceptibility testing identifies susceptibility to carbenicillin, ceftazidime, ceftriaxone, cefuroxime, chloramphenicol, azlocillin, cefotaxime, ticarcillin, ciprofloxacin, imipenem, piperacillin, and amikacin. Resistance to aztreonam, mezlocillin, gentamicin, tobramycin, and cotrimoxazole also is identified. A Burkholderia-specific medium shows no growth. The patient responds well to fluid and antimicrobial therapy and is discharged from the hospital.

Questions 1. Which microorganisms are in the differential diagnosis for this patient? 2. What tests can be done to provide differential evidence for bacterial identification? 3. What method or methods should be used to test for susceptibility of the pathogens identified in this case?   

From Reina J, Borrell N, Figuerola J. Sphingobacterium multivorum isolated from a patient with cystic fibrosis. Eur J Clin Microbiol Infect Dis. 1992;11:81.

  

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Chapter Review 1. Of all the bacteria discussed in this chapter, which one has been most highly implicated in disease? a. S. parapaucimobilis b. S. mizutaii c. S. paucimobilis d. None of the organisms listed 2. Initial clues of the presence of this group of bacteria for clinical laboratorians are: a.  They produce yellow pigment, do not grow on MacConkey agar, and oxidize glucose. b. They produce no pigment, do not grow in thioglycollate broth, and oxidize glucose. c. They produce tan/buff pigment, do not grow on blood agar, and oxidize xylose. d. They produce yellow pigment, do not grow on blood agar, and ferment glucose. 3.  Identification approaches useful for speciation of Sphingomonas are: a. H2S production b. Citrate utilization c. DNase production d. a and c e. a, b, and c

4. True or False _____ Growth on 5% sheep blood, chocolate, and MacConkey agar plates is a common trait of Sphingomonas spp. _____ Most bacteria discussed in this chapter are common causes of pathogenicity. _____ Although antimicrobial susceptibility results of bacteria discussed in this chapter have been reported in the literature, there is currently no validated method available. 5. Matching: Match the bacterium with the appropriate test result. (Results can be used more than once.) _____S. paucimobilis _____S. multivorum _____S. parapaucimobilis

a. yellow pigment b. oxidize glucose c. hydrolyze esculin d. produce urease e. produce DNase

27

Moraxella and Neisseria spp. OBJECTIVES

General Characteristics

1. Identify the distinguishing characteristics of the species within the genera Moraxella and Neisseria. 2. Identify what species within this group of bacteria are typically isolated as opportunistic pathogens. 3. Explain the procedure the microbiologist can use to determine whether the bacteria in this grouping exist as true cocci, and name these organisms. 4. Identify the species of Moraxella and Neisseria that may be isolated from human wounds resulting from a dog or cat bite. 5. Identify the species of Moraxella commonly isolated from cases of human conjunctivitis. 6. Explain the media used for culture for this group of organisms, including the chemical principle and composition. 7. List some of the conventional biochemical tests that can be used to distinguish these organisms from other bacteria, and explain the principle for each. 8. Correlate patient signs and symptoms with laboratory data, and identify the most likely etiologic agent.

The organisms discussed in this chapter are either cocci, coccobacilli, or short- to medium-sized, gram-negative rods. This group of bacteria consists of several species within the genera Moraxella and Neisseria, other than the three frequently isolated pathogens, Moraxella catarrhalis, Neisseria gonorrhoeae, and Neisseria meningitidis (Chapter 39). Most of these organisms rarely cause opportunistic infection and should be considered as potential contaminants. The genus Moraxella includes approximately 20 species. Moraxella osloensis, M. nonliquefaciens, and M. lincolnii are considered normal human respiratory microbiota with low virulence. Many other Moraxella spp. are normal mucosal microbiota of a variety of animals including cattle, horses, goats, dogs, cats, camels, guinea pigs, rabbits, pigs, and sheep. Neisseria weaveri, Neisseria ­animalis, N. bacilliformis, N. oralis, Neisseria animaloris, Neisseria ­zoodegmatis, and Moraxella canis, are oropharyngeal flora in dogs and cats and are sometimes seen in humans following a bite wound. Subinhibitory concentrations of penicillin, such as occurs in the presence of a 10-unit penicillin disk, cause the coccoid forms of these bacteria to elongate to bacilli morphology. In contrast, true cocci, such as most Neisseria spp. and M. catarrhalis, with which these organisms may be confused, maintain their original cocci shape in the presence of penicillin. In addition, the organisms discussed in this chapter do not use glucose (except N. animaloris, N. oralis, N. elongata subsp. glycolytica, and some strains of N. zoodegmatis), do not ferment carbohydrates (i.e., are oxidizers or asaccharolytic), and most do not grow on MacConkey agar but will grow well on blood and chocolate agar, as well as in commercial blood culture systems. Specific morphologic and physiologic features are presented later in this chapter in the discussion of laboratory diagnosis. 

GENERA AND SPECIES TO BE CONSIDERED Current Name Moraxella atlantae Moraxella canis Moraxella lacunata Moraxella lincolnii Moraxella nonliquefaciens Moraxella osloensis Neisseria animalis Neisseria animaloris Neisseria bacilliformis Neisseria elongata subspecies elongata Neisseria elongata subspecies glycolytica Neisseria elongata subspecies nitroreducens Neisseria oralis Neisseria weaverii Neisseria zoodegmatis

Previous Name

CDC EF-4a CDC group M6

Epidemiology, Spectrum of Disease, and Antimicrobial Therapy CDC group M5 CDC EF-4b

Infections caused by Moraxella spp. and Neisseria spp. most likely result when a breakdown of the patient’s mucosal or epidermal defensive barriers allows subsequent invasion of sterile sites by an organism that is part of the patient’s normal microbiota (i.e., an endogenous 419

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TABLE 27.1    Epidemiology, Pathogenesis, and Spectrum of Disease

Organism

Habitat (Reservoir)

Moraxella Normal human micrononliquefaciens, biota that inhabit Moraxella lacunata, mucous membranes Moraxella osloensis, covering the nose, Moraxella lincolnii, throat, other parts of Moraxella canis, the upper respiratory and tract, conjunctiva, and, Moraxella atlantae for some species (i.e., M. osloensis), the urogenital tract; may also colonize the skin

Spectrum of Disease and Infections

Mode of Transmission

Virulence Factors

Infections are rare; when they occur, they are probably caused by the patient’s endogenous strains; person-to-person transmission may be possible, but this has not been documented

Unknown; because they are rarely associated with infections, they are considered opportunistic organisms of low virulence

M. lacunata has historically been associated with eye infections, but these infections also may be caused by other Moraxella spp.; other infections include bacteremia, ­endocarditis and meningitis, ­septic arthritis, sinusitis, and ­respiratory infections

Neisseria elongata

Normal microbiota of upper respiratory tract

When infections occur, they are probably caused by the patient’s endogenous strains

Unknown; this is considered an opportunistic organism of low virulence

Rarely implicated in infections; has been documented as a cause of bacteremia, endocarditis, and osteomyelitis

Neisseria animaloris, Neisseria weaveri, and Neisseria zoodegmatis

Oral microbiota of dogs

Dog bite

Unknown

Commonly found in infections of dog bite wounds

strain; Table 27.1). The fact that these organisms are rarely the cause of infection indicates that they have low virulence. No person-to-person transmission has been noted with the organisms included in this chapter. Whenever these organisms are encountered in clinical specimens, the possibility that they are contaminants should be seriously considered. This is especially the case when the specimen source may have been exposed to a mucosal surface. Moraxella spp. have been isolated from cases of endocarditis, bacteremia, septic arthritis, and endophthalmitis. M. catarrhalis is the species most commonly associated with human infections, primarily of the respiratory tract. However, because the cellular morphology of this species is more similar to that of Neisseria spp. than that of the other Moraxella spp., details of this organism’s characteristics are discussed in Chapter 39. Data collected from the Centers for Disease Control and Prevention (CDC) show that these rare isolates may also be a cause of infection. In a study of the bacterium, N. elongata subsp. nitroreducens, one fourth of the isolates received at the CDC for analysis were from cases of bacterial endocarditis. Data collected during a 16-year period found that most of these isolates were from blood, but they were also recovered from wounds, respiratory secretions, and peritoneal fluid. Individuals at risk had preexisting heart damage or had undergone dental manipulations. 

Laboratory Diagnosis Specimen Collection and Transport No special considerations are required for specimen collection and transport of the organisms discussed in this chapter. Refer to Table 5.1 for general information on specimen collection and transport. 

Specimen Processing No special considerations are required for processing of the organisms discussed in this chapter. Refer to Table 5.1 for general information on specimen processing. 

Direct Detection Methods Other than a Gram stain of patient specimens, there are no specific procedures for the direct detection of these organisms in clinical material. Moraxella atlantae, Moraxella nonliquefaciens, and M. osloensis may appear either as coccobacilli or as short, broad rods that tend to resist decolorization and may appear gram-variable. This is also true for M. canis, which appears as cocci in pairs or short chains. Moraxella lacunata is a coccobacilli or medium-sized rod, and Moraxella lincolnii is a coccobacilli that may appear in chains. N. animaloris and N. zoodegmatis are either coccobacilli or short, straight rods. N. weaveri and N. bacilliforms are medium-length, straight bacillus. All other species are cocci that appear in either singles or pairs. 

CHAPTER 27  Moraxella and Neisseria spp.

Cultivation Media of Choice

hours. For those species that may grow on MacConkey agar, the medium should be incubated at 35°C in ambient air. 

Moraxella spp. and the elongated Neisseria spp. grow well on 5% sheep blood and chocolate agars. Most strains grow slowly on MacConkey agar and resemble the non–lactose-fermenting Enterobacterales. Both genera also grow well in the broth of commercial blood culture systems and in common nutrient broths, such as thioglycollate and brain-heart infusion. 

Colonial Appearance Table 27.2 describes the colonial appearance and other distinguishing characteristics (e.g., pitting) of each species on 5% sheep blood and MacConkey agars. The ability of most commercial identification systems to accurately identify the organisms discussed in this chapter is limited or uncertain. Table 27.3 lists some conventional biochemical tests that can be used to presumptively differentiate the species in this chapter. This is a simplified scheme; clinically important

Incubation Conditions and Duration Chocolate and 5% sheep blood agars should be incubated at 35°C in carbon dioxide or ambient air for a minimum of 48

TABLE 27.2    Colonial Appearance and Characteristics

Organism

Medium

Appearance

Moraxella atlantae

BA

Small (90% of strains positive; −, >90% of strains negative; V, indicates a variable reaction; W, indicates a weak reaction; B, reactions based on biotypes; see Table 31.2. ND, Not determined; ODC, ornithine decarboxylase. bProduces

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Serodiagnosis An enzyme-linked immunosorbent assay (ELISA) has been developed to detect antibodies to H. ducreyi. ELISA has been used to show seroconversion after Hib vaccination. None of the assays are commonly used for diagnostic purposes because of individual patient variability in the production of antibodies, antibody avidity, and persistence. 

Antimicrobial Susceptibility Testing and Therapy

• Fig. 31.3  Haemophilus Quad plate.

expected to become the future standard of care in the diagnostic microbiology laboratory. Turnaround time and annotation of the sequences can be completed in ≤24 hours, making outbreak investigations more effective for treatment and prevention of additional infections. 

Serotyping Serologic typing of H. influenzae may be used to establish an isolate as being any one of the six serotypes (i.e., a, b, c, d, e, and f ) and should be completed as soon as possible after isolation and identification. The amount of capsular antigen produced by the organisms decreases over time, in particular on repeated subculturing in the laboratory. All H. influenzae from cases of invasive infections should be serotyped to determine what H. influenzae type is the cause of the infection. Testing can be performed using a slide agglutination test (Chapter 9); a saline control without the reagent antibodies should always be tested simultaneously alongside the patient’s specimen to detect autoagglutination (i.e., the nonspecific agglutination of the test organism without homologous antiserum). Molecular methods have also been used to type H. influenzae. The assays are based on the amplification of the outer membrane protein D gene (glpQ) from the capsule (cap) locus, the capsule producing gene (bexA), the 16S rRNA, and the insertion-like sequence. PCR amplification has demonstrated an increased sensitivity compared with traditional serotyping. 

Standard methods have been established for performing in vitro susceptibility testing with clinically relevant isolates of Haemophilus spp. (see Chapter 11 for details on these methods). Although widespread H. influenzae is capable of producing beta-lactamase (penicillin resistance), cephalosporins and carbapenems as well as combination agents that contain a betalactamase inhibitor, such as clavulanate, sulbactam, or tazobactam combined with a beta-lactam, may be effective therapeutic agents. Other antimicrobials that remain useful for the treatment of Haemophilus infections include cephalosporins, macrolides, fluoroquinolones, and tetracyclines. Therefore, routine susceptibility testing of clinical isolates as a guide to therapy may not be necessary. Care should be taken when preparing inoculum concentrations (0.5 McFarland) for Haemophilus spp., in particular, beta-lactamase–producing strains of H. influenzae, because higher suspensions may lead to false-resistant results. In addition, beta-lactamase strains of H. influenzae that demonstrate elevated minimal inhibitory concentrations to ampicillin and amoxicillin have been identified. Cephalosporin activity is also decreased in these isolates. Rapid beta-lactamase testing using a chromogenic cephalosporin disk or spot test is sufficient for the majority of clinically significant isolates of Haemophilus spp. without further susceptibility testing. Standardized and reliable susceptibility testing for H. ducreyi has not been developed and therefore should not be offered in a routine clinical microbiology laboratory. In addition, automated susceptibility testing for H. influenzae is unreliable and is not recommended. 

Prevention Several multiple-dose, protein-polysaccharide conjugate vaccines are licensed in the United States for H. influenzae type b. These vaccines have substantially reduced the incidence of severe invasive infections caused by type b organisms, and vaccination of children starting at 2 months of age is strongly recommended. The antibody to the Hib capsule and activation of the complement pathway within the host play a primary role in clearance and protection from infection. Newborns are protected for a short period after birth because of the presence of maternal antibodies.

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CHAPTER 31  Haemophilus

451

Bibliography Alfa M: The laboratory diagnosis of H. ducreyi, Can J Infect Dis Med Microbiol 16:31–34, 2005. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Committee on Infectious Diseases: 2006 Red book: report of the committee on infectious diseases, ed 27, Elk Grove Village, IL, 2006, American Academy of Pediatrics. Darville T, Jacobs RF, Lucas RA, Caldwell B: Detection of Haemophilus influenzae type b antigen in cerebrospinal fluid after immunization, Pediatr Infect Dis J 11:243–244, 1992. Deshmukh D, Joseph J, Chakrabarti M, et  al.: New insights into culture negative endophthalmitis by unbiased next generation sequencing, Sci Rep 9:844, 2019. Falla TJ, Crook DW, Brophy LN, Maskell D, Kroll JS, Moxon ER: PCR for capsular typing of Haemophilus influenzae, J Clin Microbiol 32:2382–2386, 1994. Lagergård T: Haemophilus ducreyi: pathogenesis and protective immunity, Trends Microbiol 3:87–92, 1995.

Leber AL: Aerobic cultures, clinical microbiology procedures handbook, ed 4, Washington, DC, 2016, ASM Press. Leber AL, Everhart K, Balada-Llasat JM, et al.: Multicenter evaluation of BioFire filmarray meningitis/encephalitis panel for detection of bacteria, viruses, and yeast in cerebral spinal fluid specimens, J Clin Microbiol 54:2251–2261, 2016. Nørskov-Lauritsen N: Classification, identification, and clinical significance of Haemophilus and Aggregatibacter species with host specificity for humans, Clin Microbiol Rev 29:214–240, 2014. St Geme III JW: Nontypeable Haemophilus influenzae disease: epidemiology, pathogenesis, and prospects for prevention, Infect Agents Dis 2:1–16, 1993. Van Dyck E, Bogaerts J, Smet H, Tello WM, Mukantabana V, Piot P: Emergence of Haemophilus ducreyi resistance to trimethoprim-sulfamethoxazole in rwanda, Antimicrob Agents Chemother 38:1647– 1648, 1994.

CASE STUDY 31.1 A 20-year-old male presented to the emergency department with history of a temperature up to 103°F and mild respiratory distress. He reported that he had the worst sore throat of his life and was having difficulty swallowing. On physical examination, the patient was found to have a “cherry-red” epiglottis. Blood and throat cultures were obtained, and the patient was treated with cefotaxime. An endotracheal tube was placed for 48 hours until the inflammation of the epiglottis subsided. The throat culture grew normal respiratory microbiota, but a gramnegative rod was isolated from the blood culture in 24 h only on chocolate agar.

Questions 1. What is the genus of the organism that was isolated from this patient’s blood? 2. The organism grew on blood agar only around a colony of Staphylococcus (Fig. 31.2) but produced porphyrins from delta-aminolevulinic acid and fermented lactose. What is the species of this organism? 3. What is the importance of identification of Haemophilus to the species level from specimens isolated from sterile sites?

Chapter Review 1. All species of the genus Haemophilus require which of the following for in vitro growth? a. Nicotine adenine dinucleotide (NAD) b. Cystine c. Hemin d. a and c 2. Which of the following Haemophilus spp. is an agent of a sexually transmitted disease? a. H. parainfluenzae b. H. influenzae c. H. ducreyi d. H. hemolyticus 3. Which capsule type of H. influenzae is most common? a. Type a b. Type b c. Type c d. NTHi 4. All of the following organisms require X and V factors, except: a. H. influenzae b. H. haemolyticus c. H. influenzae biotype aegyptius d. A. aphrophilus 5. What routine antimicrobial susceptibility testing should be completed on Haemophilus spp.? a. Automated microdilution b. Kirby Bauer Disk Diffusion on all clinical isolates c. Beta-lactamase screening d. Susceptibility testing is not required on any isolate

6. True or False _____ H. influenzae can be found as normal microbiota of the upper respiratory tract of humans. _____ H. influenzae type a vaccine has been developed to decrease infection in children. _____ Five percent sheep blood agar provides the factors necessary for the growth of all Haemophilus spp. _____ Haemophilus spp. are able to grow on MacConkey agar. _____ Most strains of Haemophilus can grow anaerobically and aerobically. 7. Matching: Match the correct term with the appropriate description. ____ hemin ____ NAD ____ H. influenza ____ H. aegyptius ____ H. ducreyi ____ ALA-porphyrin test ____ satellite phenomenon ____ acridine orange

a. conjunctivitis b. V factor c. chancroid d. determines X factor requirement e. Staphylococcus streak technique f. X factor g. type b h. detects smaller numbers of organisms

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Bartonella OBJECTIVES 1. Explain the routes of transmission for Bartonella infections, and describe the organism’s interaction with the host. 2. Discuss the clinical manifestations of trench fever, including signs, symptoms, and individuals at risk of acquiring the disease. 3. Explain the criteria used to diagnose Bartonella henselae. 4. Describe the methods for culturing Bartonella, including growth rates, media, incubation temperature, and other relevant conditions. 5. Describe the strategies to prevent exposure and infection by these organisms in immunocompromised individuals.

GENERA AND SPECIES TO BE CONSIDERED Bartonella alsatica Bartonella bacilliformis (type species) Bartonella clarridgeiae Bartonella elizabethae Bartonella grahamii Bartonella henselae Bartonella koehlerae Bartonella quintana Bartonella rochalimae Bartonella schoenbuchensis Bartonella tribocorum Bartonella vinsonii subsp. arupensis Bartonella vinsonii subsp. berkhoffii

The Bartonella spp. are able to grow on chocolate agar and, albeit very slowly, on routine blood (trypticase soy agar with 5% sheep blood agar), typically appearing after 12 to 14 days and sometimes requiring as long as 45 days; neither organism grows on MacConkey agar. Presently, there is no optimal procedure for the isolation of these organisms from clinical specimens. 452

Bartonella General Characteristics The genus Bartonella spp. currently includes 35 named species. Fourteen species have been associated with human disease (Table 32.1) and four have been identified as presumptive human pathogens. Other members of the genus have been found in animal reservoirs such as rodents, ruminants, and moles. Bartonella spp. are most closely related to Brucella abortus and are short, gram-negative, pleomorphic coccobacillary or bacillary, facultative, intracellular bacteria that parasitize mammalian cells. They are fastidious organisms that are oxidase and catalase negative and grow best on blood-enriched media. 

Epidemiology and Pathogenesis Organisms belonging to the genus Bartonella cause numerous infections in humans; most of these infections are thought to be zoonoses. Interest in these organisms has increased because of their recognition as causes of an expanding array of clinical syndromes in immunocompromised and immunocompetent patients. For example, Bartonella species have been recognized with increasing frequency as a cause of culture-negative endocarditis. Humans acquire infection either naturally (infections caused by Bartonella quintana or Bartonella bacilliformis) or accidentally (other Bartonella species) via insect vectors or potentially by animal scratches or bites. Human infections can generally be divided into anthroponotic bartonellosis (humans as reservoirs) or zoonotic (animals as reservoirs) based on the normal host for the insect vector. Nevertheless, questions remain regarding the epidemiology of these infections; some epidemiologic information is summarized in Table 32.1. Bartonella is a facultative intracellular bacterium that closely interacts with the host cells and has unique abilities to cause either acute or chronic infection as well as the proliferation of microvascular endothelial cells and angiogenesis (forming new capillaries from preexisting ones) or suppurative manifestations. Three Bartonella species (Bartonella quintana, Bartonella

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TABLE 32.1    Bartonella spp. and Clinical Relevance in Human Accidental Hosts

Organism

Habitat (Main Reservoir)

Mode of Transmission

Clinical Manifestation(s)

Bartonella alsatica

Rabbits

Fleas and ticks

Bacteremia and endocarditis

Bartonella bacilliformis

Humans

Sandflies

Carrión’s diseasea Chronic disease: Verruga peruana, bacteremia

Bartonella clarridgeiae

Cats

Fleas and ticks

Bacteremia Cat-scratch disease

B. elizabethae

Rats and gerbils

Fleas

Endocarditis

B. grahamii

Voles, mice, rats, and deer

Fleas, ticks, and leeches

Cat-scratch disease and neuroretinitis

B. henselae

Cats

Cats and dogs; bites or scratches, fleas, and ticks

Bacteremia Endocarditis Cat-scratch disease Bacillary angiomatosis Peliosis hepatitis or splenic peliosis Neuroretinitis

B. koehlerae

Cats

Fleas and ticks

Chronic bacteremia and endocarditis; implicated in arthritis peripheral neuropathies or tachyarrhythmias

B. quintana

Humans

Human body louse

Trench fever Chronic bacteremia Endocarditis Bacillary angiomatosis Chronic lymphadenopathy Pericarditis

B. rochalimae

Foxes, coyotes, dogs, rats, and skunks

Fleas and ticks

Fever and bacteremia

B. schoenbuchensis

Cattle, roe deer, and moose

Deer keds, biting flies, and ticks

Dermatitis

B. tribocorum

Rats and mice

Fleas and mites

Bacteremia

B. vinsonii subsp. arupensis

White-footed mice

Fleas and ticks

Bacteremia and endocarditis

B. vinsonii subsp. berkhoffii

Coyotes, dogs, and foxes

Ticks

Bacteremia, endocarditis, neurologic disorders, and rheumatic symptoms

aDisease

confined to a small endemic area in South America; characterized by a septicemic phase with anemia, malaise, fever, and enlarged lymph nodes in the liver and spleen, followed by a cutaneous phase with bright red cutaneous nodules; usually self-limited.

bacilliformis, and B. henselae) are capable of causing angiogenic lesions. Research has demonstrated that some species are capable of interacting with host red blood cells, endothelial cells, and possibly bone marrow progenitor cells. Colonization of vascular endothelium is considered a crucial step in the establishment and maintenance of Bartonella-triggered angioproliferative lesions. Within several hours after infection of cultured human umbilical vein endothelial cells, Bartonella species adhere to and enter these cells by an actin-dependent process resembling other bacterial-directed phagocytosis or uptake into host cells. B. henselae possess nine outer membrane proteins (OMP), one of which is able to bind to endothelial cells. Typically, Bartonella species multiply and persist in the red blood cells in the reservoir host and share common

persistence and dissemination strategies. In addition to angioproliferation, bartonellae can inhibit endothelial cell apoptosis (programmed cell death); these organisms also activate monocyte and macrophage cells capable of producing potent angiogenic factors. Although more research is needed regarding the pathogenesis of infections caused by Bartonella, it is evident these organisms possess unique pathogenic strategies to expand their bacterial niche to sustain survival within the human host. It is evident that the pathologic response to these infections varies substantially with the status of the host immune system. For example, infection with the same Bartonella species, such as B. henselae, can cause a focal suppurative reaction (e.g., catscratch disease [CSD]) in immunocompetent patients or

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a multifocal angioproliferative lesion (e.g., bacillary angiomatosis) in immunocompromised patients. B. quintana, the causative agent for trench fever, also causes bacillary angiomatosis in immunocompromised patients. 

Spectrum of Disease The diseases caused by Bartonella species are listed in Table 32.1. Infection with B. bacilliformis presents as either an acute hemolytic bacteremia (Oroya fever) or a chronic vasoproliferative disease. The chronic form of infection is often referred to as Carrion disease, named after a medical student, Daniel Carrion, who died after self-inoculation with material from a wartlike lesion (verruga). The acute form of disease is a progressive, severe, febrile anemia. This is a result of intravascular hemolysis of erythrocytes infected with B. bacilliformis. Chronic bacteremia occurs in individuals who live in endemic areas. The chronic infection appears as cutaneous nodular angioproliferative lesions previously described as verruga. Depending on the status of the host and the disease presentation, mortality rates associated with B. bacilliformis range from 40% to 90% prior to the use of antibiotics. Trench fever, caused by B. quintana, was largely considered a disease of the past. Clinical manifestations of trench fever range from a mild influenza-like headache and bone pain to splenomegaly (enlarged spleen) and a short-lived maculopapular rash. During the febrile stages of trench fever, infection may persist long after the disappearance of all clinical signs; some patients may have six or more recurrences. B. quintana has been reported in cases of bacteremia, blood-culture negative endocarditis, chronic lymphadenopathy, and bacillary angiomatosis. Bacillary angiomatosis is a vascular proliferative disease involving the skin (other organs such as the liver, spleen, and lymph nodes may also be involved). Prolonged bacteremia with B. quintana and B. henselae can cause epithelioid angiomatosis, a vasculoproliferative disease of the skin. B. henselae is associated with bacteremia, endocarditis, and bacillary angiomatosis. Of note, observations indicate that B. henselae infections appear to be subclinical and are markedly underreported, because problems with current diagnostic approaches are recognized (see Laboratory Diagnosis). In addition, B. henselae is the primary cause of CSD with B. grahamii implicated in a few human cases. B. henselae also causes rheumatic manifestations and bacillary peliosis hepatitis or splenic peliosis. About 12,500 cases of CSD occur annually in the United States; about 33% of these occur in children ≤14 years of age. The infection begins as a papule or pustule at the primary inoculation site; regional tender lymphadenopathy develops in approximately 3 weeks. The spectrum of disease ranges from chronic, self-limited adenopathy to a severe systemic illness affecting multiple body organs in about 25% of CSD cases. Localized infections are common in immunocompetent patients, including ocular bartonellosis, but are rare in transplant patients. In addition, rare cases of optical neuropathy and unilateral loss of vision have been characterized in acute cases of CSD. Rheumatic manifestations may include myositis, arthritis with skin nodules in children, leukocytoclastic vasculitis,

erythema nodosum, or fever of unknown origin. Although complications such as a suppurative (draining) lymph nodes or encephalitis are reported, fatalities are rare. Zoonotic disease is caused by a variety of species and may be associated with blood culture–negative endocarditis or myocarditis. Diagnosis of CSD requires three of the following four criteria: • History of animal contact plus site of primary inoculation (e.g., a scratch) • Negative laboratory studies for other causes of lymphadenopathy • Characteristic histopathology of the lesion • A positive skin test using antigen prepared from heattreated pus collected from another patient’s lesion Peliosis hepatitis caused by B. henselae may occur independently or in conjunction with cutaneous bacillary angiomatosis or bacteremia. Patients with bacillary peliosis hepatitis or splenic peliosis demonstrate gastrointestinal symptoms. Symptoms include fever, chills, and an enlarged liver and spleen that contain blood-filled cavities. This systemic disease primarily develops in patients infected with HIV and other immunocompromised individuals. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing Clinical specimens submitted to the laboratory for direct examination and culture include blood collected in a lysis-centrifugation blood culture tube (Isolator; Alere, Inc., Waltham, MA), sodium citrate or plastic EDTA tubes, aspirates, and tissue specimens (e.g., lymph node, spleen, or cutaneous biopsies). If a delay occurs when processing tissue specimens for culture of Bartonella spp., samples should be frozen at −20°C. Notably, attempts to culture Bartonella spp. in routine microbiology laboratories is not recommended due to low recovery rates. Fresh tissue samples and formalin-fixed, paraffin-embedded tissues from lymph nodes, lesions, infected organs and aspirated vitreous fluid, and cerebrospinal fluid can be processed for direct detection by nucleic acid testing. There are no special requirements for specimen collection, transport, or processing that enhances organism recovery. Refer to Table 5.1 for general information on specimen collection, transport, and processing. 

Direct Detection Methods Microscopy Detection of Bartonella spp. during the histopathologic examination of tissue biopsies is enhanced using the Warthin-Starry silver stain or immunohistochemical techniques. Because of the fastidious nature of the organisms and slow growth associated with low levels of bacteremia in patients, these techniques lack sensitivity and specificity. 

Nucleic Acid Detection Tissue, body fluids, and blood can be successfully used for identification of Bartonella species from clinical samples by nucleic acid amplification detection. The 16S rRNA gene is extremely

CHAPTER 32  Bartonella

homologous within the genus and does not provide effective discrimination of Bartonella spp. Several other gene targets have been successfully used for identification, including citrate synthase (gltA), heat shock protein (groEL), riboflavin synthase (ribC), a cell division protein (ftsZ), and the 17-kDa antigen. In addition, the 16S–23S ribosomal ribonucleic acid (rRNA) gene intergenic transcribed spacer region has been used as a reliable method for the detection and the classification of Bartonella deoxyribonucleic acid (DNA) in clinical samples.

Cultivation Bartonella spp. are slow-growing fastidious organisms that require heme for growth. Various agar bases such as Columbia agar, trypticase soy, brucella, or heart infusion agar supplemented with hemoglobin, 5% rabbit blood or hemin have been successfully used for the cultivation of the organisms. Lysed, centrifuged sediment of blood collected in an isolator tube or minced tissue is directly inoculated onto fresh chocolate and heart infusion agar plates containing 5% rabbit blood. Cultures should be incubated at 35°C to 37°C in 5% CO2, except B. bacilliformis and B. clarridgeiae, which grow optimally at 25°C to 30°C in ambient air. Some species demonstrate improved growth on specific media: B. henselae, heart infusion agar; B. quintana, chocolate agar; B. koehlerae requires fresh chocolate agar. Due to the slow growth of these organisms, they should be incubated for at least 4 weeks in high humidity. Biphasic or broth culture systems may be used for the isolation of Bartonella spp.; however, the organisms rarely reach a high level of turbidity or activate the CO2 detector in automated systems. Subculturing and staining with acridine orange on negative cultures before discarding improves identification. Biopsy material is cultivated with an endothelial cell culture system; cultures are incubated at 35°C in 5% to 10% CO2 for 15 to 20 days. Lymph node tissue, aspirates, or swabs can be inoculated onto laked horse blood agar slants supplemented with hemin; plates are sealed and incubated in 5% CO2 up to 6 weeks at 37°C with 85% humidity. Heparinized plasma may also be cultured on T84 bladder carcinoma cells in a centrifugation shell-vial. 

A

455

Approach to Identification Bartonella spp. should be suspected when colonies of small, gram-negative bacilli are recovered after prolonged incubation (Fig. 32.1). Macroscopic colonial morphology is often variable. B. henselae may appear as irregular, dry, white “cauliflower-like” colonies that pit the agar or small, circular, tan, moist colonies. The majority of Bartonella spp. will appear tan and smooth on repeated subculture. The Gram-stain appearance of Bartonella as small, slightly curved gram-negative rods is similar to Campylobacter, Helicobacter, or Haemophilus spp. These organisms are all oxidase, urease, nitrate reductase, and catalase negative. Most of the Bartonella spp. are biochemically inert, making phenotypic methods for identification unreliable for identification from clinical specimens. Prolonged incubation requiring more than 7 days, microscopic and macroscopic morphologies in conjunction with negative oxidase, and catalase testing may be sufficient for preliminary identification. Bartonella spp. are not included in MALDI-TOF Reference databases. Nucleic acid amplification and sequencing is recommended for confirmation and identification of Bartonella species and strains. 

Serodiagnosis There are no US Food and Drug Administration-cleared serologic tests for the diagnosis of Bartonella infections. 

Antimicrobial Susceptibility Testing and Therapy Although there are no current standards or guidelines for antimicrobial susceptibility testing by the Clinical and Laboratory Standards Institute (CLSI) or the European Committee on Antimicrobial Susceptibility Testing (EUCAST), testing can be performed by agar dilution using blood or chocolate agar and microdilution methods with broth supplemented with blood. Treatment recommendations for Bartonella diseases, including CSD, depend on the specific disease presentation and have demonstrated sensitivity to β-lactams, aminoglycosides, chloramphenicol, tetracyclines, macrolides,

B • Fig. 32.1  (A) Colonies of Bartonella henselae on blood agar. (B) Gram stain of a colony of B. henselae from blood agar.

456 PA RT I I I    Bacteriology

rifamycins, fluoroquinolones, and trimethoprim-sulfamethoxazole. The efficacy of various antibiotics for CSD is difficult to assess because of the self-limiting nature of the disease and the decrease in symptoms in the absence of therapy. In addition to the clinical presentation, the treatment must be specifically adapted to the correct Bartonella sp. Moreover, results of in vitro testing may not correlate with clinical efficacy; for example, the administration of penicillin is not effective therapy despite susceptibility in  vitro. Suggested therapy for endocarditis, suspected or documented, is gentamicin with or without doxycycline, respectively. However, this treatment has been associated with kidney failure. Rifampin in combination with doxycycline

is the current recommended treatment due to the intracellular penetration and bactericidal activity of rifampin. 

Prevention There are no vaccines available to prevent infections caused by Bartonella spp. Exposure to cats or cat fleas has been implicated in the transmission of B. henselae to humans. It is recommended that immunocompromised individuals avoid contact with cats, especially kittens, and control flea infestation.

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Bibliography Ak R, Doganay F, Akoglu EU, Ozturk TC: A challenging differential diagnosis of optic neuropathy in ED: CSD, BMJ Case Rep 2015; 2015:bcr2015210252. Avidor B, Graidy M, Efrat G, et al.: Bartonella koehlerae, a new catassociated agent of culture-negative human endocarditis, J Clin Microbiol 42:3462–3468, 2004. Berger P, Papazian L, Drancourt M, La Scola B, Auffray JP, Raoult D: Ameba-associated microorganisms and diagnosis of nosocomial pneumonia, Emerg Infect Dis 12:248–255, 2006. Breitschwerdt EB, Kordick DL: Bartonella infection in animals: carriership, reservoir potential, pathogenicity, and zoonotic potential for human infection, Clin Microbiol Rev 13:428–438, 2000. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Chan D, Geiger JA, Vasconcelos EJR, Oakley B, Diniz PPVP: Bartonella rochalimae detection by a sensitive and specific PCR platform, Am J Trop Med Hyg 99:840–843, 2018. De Bruin A, Van Leeuwen AD, Jahfari S, et  al.: Vertical transmission of Bartonella schoenbuchensis in Lipoptena cervi, Parasit Vectors 8:176, 2015. Dehio C: Recent progress in understanding Bartonella-induced vascular proliferation, Curr Opin Microbiol 6:61–65, 2003. Fournier PE, Robson J, Zeaiter Z, McDougall R, Byrne S, Raoult D: Improved culture from lymph nodes of patients with cat scratch disease and genotypic characterization of Bartonella henselae isolates in Australia, J Clin Microbiol 40:3620–3624, 2002. Garcia-Caceres U, Garcia FU: Bartonellosis: an immunosuppressive disease and the life of Daniel Alcides Carrión, Am J Clin Pathol 95(Suppl 1):S58–S66, 1991. Giladi M, Avidor B, Kletter Y, et al.: Cat scratch disease: the rare role of Afipia felis, J Clin Microbiol 36:2499–2502, 1998. Greub G, Raoult D: Bartonella: new explanations for old diseases, J Med Microbiol 51:915–923, 2002. Gutierrez R, Vayssier-Taussat M, Buffet JP, Harrus S: Guidelines for the isolation, molecular detection, and characterization of Bartonella species, Vector Borne Zoonotic Dis 17:42–50, 2017. Jacomo V, Raoult D: Human infections caused by Bartonella spp. Parts 1 and 2, Clin Microbiol Newsl 22(1–5):9–13, 2000. Jacomo V, Kelly PJ, Raoult D: Natural history of Bartonella infections (an exception to Koch’s postulate), Clin Diagn Lab Immunol 9:8–18, 2002. Kordick DL, Hilyard EJ, Hadfield TL, et al.: Bartonella clarridgeiae: a newly recognized zoonotic pathogen causing inoculation papules,

fever, and lymphadenopathy (cat scratch disease), J Clin Microbiol 35:1813–1818, 1997. Kosoy M, McKee C, Albayrak L, Fofanov Y: Genotyping of Bartonella bacteria and their animal hosts: current status and perspectives, Parasitology 145:543–562, 2018. LaScola B, Raoult D: Culture of Bartonella quintana and Bartonella henselae from human samples: a 5-year experience (1993-1998), J Clin Microbiol 37:1899, 1999. Lawson PA, Collins MD: Description of Bartonella clarridgeiae sp nov isolated from the cat of a patient with Bartonella henselae septicemia, Med Microbiol Lett 5:640, 1996. Lee RA, Ray M, Kasuga DT, et al.: Ocular bartonellosis in transplant recipients: two case reports and review of the literature, Transpl Infect Dis 17:723–727, 2015. Maggi RG, Breitschwerdt EB: Potential limitations of the 16S-23S rRNA intergenic region for molecular detection of Bartonella species, J Clin Microbiol 43:1171–1176, 2005. Manfredi R, Sabbatani S, Chiodo F: Bartonellosis: light and shadows in diagnostic and therapeutic issues, Clin Microbiol Infect 11:167– 169, 2004. Mazur-Melewska K, Mania A, Kemnitz P, Figlerowicz M, Służewski W: Cat-scratch disease: a wide spectrum of clinical pictures, Postepy Dermatol Alergol 32:216–220, 2015. Mozayeni BR, Maggi RG, Bradley JM, Breitschwerdt EB: Rheumatological presentation of Bartonella koehlerae and Bartonella henselae bacteremias: a case report, Medicine (Baltim) 97(17).e0465, 2018, https://doi.org/10.1097/MD.0000000000010465. Okaro U, Addisu A, Casanas B, Anderson B: Bartonella species, an emerging cause of blood-culture-negative endocarditis, Clin Microbiol Rev 30:709–746, 2017. Oksi J, Rantala S, Kilpinen S, et  al.: Cat scratch disease caused by Bartonella grahamii in an immunocompromised patient, J Clin Microbiol 51:2781–2784, 2013. Oteo JA, Maggi R, Portillo A, et al.: Prevalence of Bartonella spp. by culture, PCR and serology, in veterinary personnel from Spain, Parasit Vectors 10:553–561, 2017. Rolain JM, Brouqui P, Koehler JE, Maguina C, Dolan MJ, Raoult D: Recommendations for treatment of human infections caused by Bartonella species, Antimicrob Agents Chemother 48:1921–1933, 2004. Silva BTGD, Souza AM, Campos SDE, et al.: Bartonella henselae and Bartonella clarridgeiae infection, hematological changes and associated factors in domestic cats and dogs from an Atlantic rain forest area, Brazil, Acta Trop 193:163–168, 2019.

Chapter Review 1. Humans acquire Bartonella infection in the following way: a. Arthropod-borne transmission b. Rodents c. Naturally d. a and c 2. Most Bartonella infections are thought to be: a. Nosocomial infections b. Zoonotic infections c. Normal flora d. All of the above 3. Bartonella is characterized by all of the following, except: a. Gram negative b. Oxidase negative c. MacConkey negative d. Chocolate positive 4. B. quintana has been known to cause: a. Carrion disease b. Trench fever c. CSD d. Lyme disease

5. Bartonella species can be detected by which of the following? a. Warthin-Starry silver stain b. Polymerase chain reaction c. Immunofluorescence d. All of the above 6. Bartonella spp. can be cultivated on artificial media that is enriched with: a. Iron b. Dextrose c. Erythrocytes d. a and b 7. Which of the following aid in Bartonella prevention for the immunocompromised patient? a. Vaccination b. Avoiding contact with cats c. Controlling flea infestation d. b and c

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33

Campylobacter, Arcobacter, and Helicobacter OBJECTIVES 1. List the Campylobacter species most often associated with infections in humans, and explain how they are transmitted. 2. Identify the culture methods for optimum recovery of Campylobacter spp., including agar, temperatures, oxygenation, and length of incubation. 3. Describe how to isolate Campylobacter from blood, including special stains, atmospheric conditions, and length of incubation. 4. List the colonial morphology, microscopic characteristics, and biochemical reactions of Campylobacter and Helicobacter. 5. List the key biochemical test to identify Helicobacter pylori in specimens. 6. Describe how H. pylori colonize in the stomach and how motility plays an important role in the pathogenesis of the organism. 7. Differentiate the isolation and identification of Campylobacter, Arcobacter, and Helicobacter species, including H. pylori and enterohepatic helicobacters. 8. Describe why therapy is often problematic for H. pylori.

GENERA AND SPECIES TO BE CONSIDERED Campylobacter coli Campylobacter concisus Campylobacter curvus Campylobacter fetus subsp. fetus Campylobacter fetus subsp. venerealis Campylobacter gracilis Campylobacter hominis Campylobacter hyointestinalis subsp. hyointestinalis Campylobacter jejuni subsp. doylei Campylobacter jejuni subsp. jejuni Campylobacter lari subsp. lari Campylobacter pyloridis Campylobacter rectus Campylobacter showae Campylobacter sputorum subsp. sputorum Campylobacter upsaliensis Campylobacter ureolyticus Arcobacter cryaerophilus Arcobacter butzleri Arcobacter skirrowii Helicobacter pylori

Helicobacter bizzozeronii Helicobacter cinaedi Helicobacter felis Helicobacter fennelliae Helicobacter heilmannii Helicobacter suis Helicobacter cinaedi Helicobacter fennelliae Helicobacter heilmannii Helicobacter salomonis Helicobacter suis Helicobacter bilis Helicobacter felis Helicobacter canis Helicobacter canadensis Helicobacter pullorum

Because of their morphologic similarities and an inability to recover these organisms using routine laboratory media for primary isolation, the genera Campylobacter, Arcobacter, and Helicobacter are considered together in this chapter (Fig. 33.1). All organisms belonging to these genera are small, curved or straight, motile, gram-negative bacilli. When cultures are old or the organisms are exposed to air for long periods, they may form spherical or coccoid bodies. With few exceptions, most of these bacteria also have a requirement for a microaerobic (5% to 10% O2) atmosphere with some species requiring the addition of 5% to 7% H2.

Campylobacter and Arcobacter General Characteristics Campylobacter and Arcobacter spp. are relatively slow growing, fastidious, and, in general, asaccharolytic; organisms known to cause disease in humans are listed in Table 33.1. 

Epidemiology and Pathogenesis Most Campylobacter species are pathogenic and associated with a wide variety of diseases in humans and other animals. These organisms demonstrate considerable ecologic diversity. Campylobacter spp. are microaerobic (5% to 10% O2) inhabitants of the gastrointestinal tracts of various animals, 457

458 PA RT I I I    Bacteriology

Clinical specimen plate to the following media:

Blood agar

Anaerobic blood agar

Translucent, moist colonies (Helicobacter spp. may appear hemolytic on blood agar)

Campy-BA or CCDA (Campylobacter agar base, blood free)

Gray to white, moist colonies

Gram stain Curved gram-negative rods Oxidase Positive

Positive Campylobacter spp. Helicobacter spp. Arcobacter spp. (weak)

Positive Helicobacter spp. (Table 33.4)

Catalase

Urease

Negative

Negative Campylobacter spp. (Table 33.2)

Restreak to BA, recheck at 24 h. Discard if negative.

Negative Helicobacter spp. (Table 33.4) Arcobacter spp. (Table 33.2)

•

Fig. 33.1 Identification scheme for the differentiation of the genera Helicobacter, Campylobacter, and Arcobacter. BAP, Blood agar plate.

including poultry, dogs, cats, sheep, and cattle, as well as the reproductive organs of several species. In general, Campylobacter spp. produce three syndromes in humans: febrile systemic disease, periodontal disease, and, most commonly, gastroenteritis. Humans are the only recognized reservoir for C. concisus, C. rectus, C. curvus, and C. showae and implicated in periodontal disease. Arcobacter spp. are aerotolerant and are also inhabitants of the gastrointestinal tracts of various animal species. Three of the eighteen species of Arcobacter have been identified in human infections. Arcobacter species appear to be associated with gastroenteritis. Studies have indicated that Arcobacter butzleri, one of the most common Campylobacter-like organisms isolated from stool, is often associated with persistent, watery diarrhea. The organism has also been isolated from patients with bacteremia, endocarditis, and peritonitis. The organism is found in the environment in untreated water. It is also prevalent in commercially prepared meats including chicken, beef, pork, lamb, and poultry. Arcobacter cryaerophilus has been isolated from patients with bacteremia

and diarrhea. A. skirrowii has been isolated from a patient with chronic diarrhea; however, the clinical significance of this isolate is unknown. Within the genus Campylobacter, Campylobacter jejuni subsp. jejuni and Campylobacter coli are associated with clinically indistinguishable infections in humans and are transmitted via contaminated food, milk, or water. Outbreaks have been associated with contaminated drinking water and improperly pasteurized milk, among other sources. In contrast to other agents of foodborne gastroenteritis, including Salmonella and staphylococci, Campylobacter spp. does not multiply in food. Other Campylobacter spp. have been isolated from patients following consumption of untreated water as well as from immunocompromised patients or patients recently returned from international travel. In addition to food outbreaks, Campylobacter spp. may be present in poultry, cattle, sheep, pigs, and domestic pets. C. lari and C. upsaliensis have been identified in cases of gastrointestinal and urinary tract infections and bacteremia. C. jejuni subsp. doylei has been isolated from children with diarrhea and

CHAPTER 33  Campylobacter, Arcobacter, and Helicobacter

459

TABLE   Source and Spectrum of Disease in Humans of Representative Species of Campylobacter and Arcobacter 33.1 

Organism

Source

Spectrum of Disease in Humans

Campylobacter concisus, Campy­ lobacter curvus, Campylobacter rectus, Campylobacter showae

Humans

Periodontal disease; gastroenteritis

Campylobacter gracilis

Humans

Deep-tissue infections: head, neck, and viscera; gingival crevices Peritonitis, bacteremia, and pulmonary infections

Campylobacter coli

Pigs, poultry, sheep, bulls, birds

Gastroenteritisa Septicemia

Campylobacter jejuni subsp. jejuni

Poultry, pigs, bulls, dogs, cats, birds, and other animals

Gastroenteritisa Septicemia Meningitis Proctitis

Campylobacter jejuni subsp. doylei

Humans

Gastroenteritisa Gastritis Septicemia

Campylobacter lari subsp. concheus, Campylobacter lari subsp. lari

Birds, poultry, other animals; river and seawater

Gastroenteritisa Septicemia Prosthetic joint infection Urinary tract infections

Campylobacter hyointestinalis subsp. hyointestinalis

Pigs, cattle, hamsters, deer

Gastroenteritis Proctitis

Campylobacter upsaliensis

Dogs, cats

Gastroenteritis Septicemia abscesses

Campylobacter fetus subsp. fetus

Cattle, sheep

Septicemia Gastroenteritis Abortion Meningitis

Campylobacter fetus subsp. venerealis

Cattle

Septicemia

Campylobacter pyloridis

Humans, shell fish

None reported

Campylobacter sputorum biovar sputorum

Humans, cattle, pigs

Abscesses Gastroenteritis

Campylobacter ureolyticus

Humans

Gastroenteritis

Arcobacter cryaerophilus

Pigs, bulls, and other animals

Gastroenteritisa Septicemia

Arcobacter butzleri

Pigs, bulls, humans, other animals; water

Gastroenteritisa Septicemia

Arcobacter skirrowii

Pigs, sheep, cattle, poultry, and humans

Gastroenteritis

aMost

common clinical presentation.

from gastric biopsies in adults. In developed countries, most C. jejuni infections are transmitted by direct contact during the preparation and eating of chicken. Person-to-person transmission of Campylobacter infections plays only a minor role in the transmission of disease. There is a marked seasonality with the rates of C. jejuni infection in the United States; the highest rates of infection occur in late summer and early fall. Campylobacter spp. has been recognized as

the most common causative agent of gastroenteritis in the United States. Although infections with C. jejuni are evident in acute inflammatory enteritis of the small intestine and colon, the pathogenesis remains unclear. However, multiplication of organisms in the intestine leads to cell damage and an inflammatory response. Blood and polymorphonuclear neutrophils are often observed in stool specimens. Most

460 PA RT I I I    Bacteriology

strains of C. jejuni are susceptible to the nonspecific bactericidal activity of normal human serum; this susceptibility probably explains why C. jejuni bacteremia is uncommon. 

Spectrum of Disease As previously mentioned, Campylobacter spp. are the causative agent of gastrointestinal or extraintestinal infections. Extraintestinal infections including abscesses, bacteremia, cholecystitis, myocarditis, meningitis, abortion and neonatal sepsis, nephritis, pancreatitis, peritonitis, prostatitis, and septic arthritis have been reported following Campylobacter gastroenteritis. The different campylobacters and their associated diseases are summarized in Table 33.1. Gastroenteritis associated with Campylobacter spp. is usually a self-limiting illness and does not require antibiotic therapy. Recently, postinfection complications with C. jejuni have been recognized and include reactive arthritis and Guillain-Barré syndrome, an acute demyelination (removal of the myelin sheath from a nerve) of the peripheral nerves. Studies indicate that 20% to 40% of patients with this syndrome were infected with C. jejuni 1 to 3 weeks before the onset of neurologic symptoms. Reactive arthritis, Reiter syndrome, and irritable bowel syndrome are also known to follow campylobacter infections. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing There are no special requirements for the collection, transport, and processing of clinical specimens for the detection of campylobacters; the two most common clinical specimens submitted to the laboratory are feces (rectal swabs are also acceptable for culture for infants and children) and blood. Specimens should be processed as soon as possible. Delays of more than 2 hours require the stool specimen to be placed either in Cary-Blair transport medium, modified Cary-Blair, Fecal Enteric Plus, or other equivalent transport medium. Cary-Blair or Fecal Enteric Plus transport medium is suitable for other enteric pathogens; specimens received in transport medium should be processed immediately or stored at 4°C until processed. C. fetus, C. jejuni, and C. upsaliensis have been successfully recovered from patient samples using automated blood culture systems. 

Direct Detection Upon Gram staining, Campylobacter spp. display a characteristic microscopic morphology as small, curved or seagullwinged, faintly staining, gram-negative rods (Fig. 33.2). Campylobacter spp. are difficult to visualize using the standard secondary safranin counterstain in the Gram stain procedure. Carbol-fuchsin or 0.1% aqueous basic fuchsin may be used to improve visualization from smears of stools or pure cultures. The presence of fecal white cells is not a recommended test for predicting bacterial infection with Campylobacter; the absence of fecal leukocytes does not rule out

infection. Other screening tests for inflammation, including fecal lactoferrin and calprotectin, lack sensitivity and specificity for the diagnosis of gastroenteritis. 

Antigen Detection The use of culture independent tests (CIDT) for the diagnosis of Campylobacter infections has increased. Several commercial antigen detection systems are available for the direct detection of Campylobacter in stool specimens. These enzyme immunoassays (EIAs) can be used to detect antigens in stool samples for several days if stored at 4°C. These include the Premiere Campylobacter assay and ImmunoCard Stat! (Meridian Bioscience, Cincinnati, OH) and the ProSpecT microplate assay (Thermo Fisher Scientific, Waltham, MA), which detect both C. jejuni and C. coli but are not able to differentiate them. In addition, some EIAs have been reported to cross-react with Campylobacter upsaliensis. Because of the poor specificity and positive predictive value of the antigen detection tests, it is recommended that these tests not be used independently to diagnose Campylobacter infections.

Nucleic Acid Detection Nucleic acid amplification provides an alternative to culture methods for the detection of Campylobacter spp. from clinical specimens. The detection of Campylobacter deoxyribonucleic acid (DNA) in stools from a large number of patients with diarrhea suggests that Campylobacter spp. other than C. jejuni and C. coli may account for a proportion of cases of acute gastroenteritis in which no causative agent is identified. Five nucleic acid–based test methods are US Food and Drug Administration (FDA)–approved for use in the United States. These include the xTAG Gastrointestinal Pathogen Panel and the Verigene enteric pathogens test (Luminex Corporation, Toronto, CA), BioFire Film Array Gastrointestinal Panel (bioMérieux Co, Durham, NC), Verigene (Nanosphere Inc., Northbrook, IL), and Prodesse ProGastro SSCS (Hologic, Marlborough, MA). These assays are highly sensitive and often used as the primary diagnostic tool for the identification of Campylobacter infections. Although CIDT are rapid and provide same-day diagnosis in patients with Campylobacter gastroenteritis, it is recommended that the laboratory continue to cultivate these organisms. Culturing is necessary for antimicrobial susceptibility and monitoring outbreaks and may be required by the manufacturer of the nucleic acid or other methods.

Media Campy-BA is an enriched selective blood agar plate used to isolate C. jejuni. The medium is composed of a Brucella agar base; sheep red blood cells; and vancomycin, trimethoprim, polymyxin B, amphotericin B, and cephalothin. Campy medium (CVA) contains cefoperazone, vancomycin, and amphotericin B. The antibiotics in both media suppress the growth of normal fecal flora. Campylobacter agar base

CHAPTER 33  Campylobacter, Arcobacter, and Helicobacter

A

461

B • Fig. 33.2  (A) Gram stain appearance of Campylobacter jejuni subsp. jejuni from a colony on a primary isolation plate. Note seagull and curved forms (arrows). (B) Appearance of C. jejuni subsp. jejuni in a direct Gram stain of stool obtained from a patient with campylobacteriosis. Arrows point to the seagull form.

blood free (CCDA) is a modified agar that does not include blood. The blood is replaced with charcoal, sodium pyruvate, and ferrous sulfate. The medium supports growth of most Campylobacter spp. An additional blood-free media is charcoal-based selective medium (CSM). 

Cultivation Stool

Successful isolation of Campylobacter spp. from stool requires selective media and optimal incubation conditions. Cultures should not be performed from formed stool for diagnostic purposes. Recommended inoculation of two selective agars is associated with increased recovery of the organisms. Reports indicate that CVA and CCDA demonstrate a higher recovery rate than blood-based media. Because Campylobacter and Arcobacter spp. have different optimum temperatures and atmospheric requirements, two sets of selective plates should be incubated, one at 42°C and one at 37°C with increased hydrogen. Extended incubation may be required (48 to 72 hours) before there is evidence of visible growth. In addition, Campylobacter spp. such as C. concisus, C. upsaliensis, C. ureolyticus, and C. sputorum, are not routinely isolated due to the nonthermophilic nature and/or inhibition by antibiotics included in the commonly used selective media. A filtration method can also be used in conjunction with a nonselective medium to enhance recovery of Campylobacter and Arcobacter spp. A filter (0.45 to 0.65-μm pore-size polycarbonate or cellulose acetate) is placed on the agar surface, and a drop of stool is placed on the filter. The plate is incubated upright. After 60 minutes at 37°C, the filter is removed and the plates are reincubated in a microaerobic atmosphere. The organisms are motile and capable of migrating through the filter, producing isolated colonies on the agar surface and effectively removing contaminating stool microbiota. C. concisus, A. butzleri, A.

cryaerophilus, and Helicobacter cinaedi have been isolated after 5 to 6 days of incubation using the filter technique. An enrichment broth may be used for the recovery of Arcobacter or Campylobacter species from stool such as Preston enrichment, Campy-thio, and Campylobacter enrichment broth. The clinical advantage and cost-effectiveness of the use of enrichment broth has not been evaluated.  Blood

Campylobacter spp. are capable of growth in less than 5 days in most blood culture media, although they may require extended incubation periods of up to 2 weeks for detection. Subcultures should be incubated in 5% O2, 10% CO2, and 80% N2 (microaerobic) environment. Turbidity may not be visible in blood culture media; therefore subcultures at 24 to 48 hours to a nonselective blood agar medium or microscopic examination using acridine orange stain may be necessary. The presence of Campylobacter spp. in blood cultures is effectively detected through carbon dioxide monitoring. Isolation from sources other than blood or feces is extremely rare. Recovery of the organisms is enhanced by inoculation (minced tissue, wound exudate) to a nonselective blood or chocolate agar plate and incubation at 37°C in a carbon dioxide–enriched, microaerobic atmosphere. Selective agars containing cephalosporin, rifampin, and polymyxin B may inhibit growth of some strains and should not be used for isolation from sterile sites.  Atmosphere

Campylobacter spp. require a microaerobic environment as previously indicated; however, not all species will grow in this environment. Some species, including Campylobacter sputorum, C. concisus, C. mucosalis, C. curvus, C. rectus, and C. hyointestinalis require increased hydrogen concentration for optimal growth. A gas mixture of 10% CO2, 6% H2, and 84% N2 using an evacuation-replacement system such

462 PA RT I I I    Bacteriology

• Fig. 33.3  Colonies of Campylobacter jejuni after 48 hours of incubation on a selective medium in a microaerobic atmosphere.

as the Anoxomat, Advanced Instruments, Norwood, MA (Chapter 40) is sufficient for growth of these organisms. 

Approach to Identification Plates should be examined for characteristic colonies, which are gray to pink or yellow gray and slightly mucoid; some colonies may exhibit a tailing effect along the streak line (Fig. 33.3). Colony morphology varies with the type of medium used for isolation. Suspicious-looking oxidase-positive colonies observed on selective media incubated at 42°C may be presumptively identified as Campylobacter spp., usually C. jejuni or C. coli. A wet preparation of the organism in broth can be examined for characteristic darting motility and curved morphology on Gram stain. Most isolates of C. jejuni subsp. jejuni can be distinguished from other Campylobacter species by the presence of sodium hippurate hydrolysis. Campylobacter fetus is incapable of growth at 42°C, and optimal growth is 37°C. Susceptibility or resistance to cephalothin and nalidixic-acid are no longer recommended for species identification due to the increasing fluoroquinolone resistance present in clinical isolates of Campylobacter spp. Table 33.2 provides characteristics for biochemical differentiation of the common clinically relevant organisms included in this chapter. Almost all the pathogenic Campylobacter spp. are oxidase positive and catalase positive. Laboratories will commonly report stool isolates as “Campylobacter spp.” Because C. coli and C. jejuni are similar, biochemically, molecular methods or MALDI-TOF is required to differentiate hippurate negative C. jejuni. Most Campylobacter spp. are asaccharolytic, unable to grow in 3.5% NaCl, although strains of Arcobacter appear more resistant to salt and, except for A. cryaerophilus, unable to grow in ambient air requiring incubation in 10% O2, 10% CO2, and 80% N2 at 37°C. Growth in 1% glycine is variable. Other tests useful for identifying these species are the rapid hippurate hydrolysis test, production of hydrogen sulfide (H2S) in triple sugar iron agar slants, nitrate reduction, and hydrolysis of indoxyl acetate. Indoxyl acetate disks are available commercially. Cellular fatty acid analysis is useful for species identification. This method is not available in routine clinical microbiology laboratories.

Because Campylobacter and Arcobacter are difficult to identify using biochemical and phenotypic tests, nucleic acid (NAATS)–based amplification of the 16S and 23S ribosomal ribonucleic acid (rRNA) gene along with a variety of other targets and direct sequencing of the PCR products have successfully been used to identify most Campylobacter species. The assays accurately discriminate related taxa, including Campylobacter, Arcobacter, or Helicobacter species. In addition to molecular methods, matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) is increasingly being used to identify Campylobacter spp. Identification has been reported with 99% to 100% accuracy for C. jejuni, C. coli, C. fetus subsp. fetus, Campylobacter lari, and A. butzleri. 

Serodiagnosis Serodiagnosis is not widely applicable for the diagnosis of infections caused by these organisms.  

Antimicrobial Susceptibility Testing and Therapy C. jejuni and C. coli demonstrate variable susceptibility to many antimicrobial agents, including macrolides, tetracyclines, chloramphenicol, aminoglycosides, and quinolones. Erythromycin has been the drug of choice for patients with severe gastroenteritis (severe dehydration, bacteremia), with ciprofloxacin as an alternative therapeutic option. Ciprofloxacin resistance has been reported as high as 26.7% to 35.6% in C. jejuni and C. coli, with 25% to 50% resistance to erythromycin. Previously, fluoroquinolones were the antibiotic therapy most commonly prescribed for Campylobacter infection; however, a rapidly increasing proportion of Campylobacter strains worldwide have been identified as fluoroquinolone resistant. Parenteral therapy (not taken through the alimentary canal but by an alternate route, such as intravenously) is used to treat systemic C. fetus infections. Arcobacter spp. demonstrate variable resistance patterns to macrolides and fluoroquinolones. The Clinical Laboratory Standards Institute (CLSI) recognizes broth microdilution and disk diffusion and ETEST (bioMérieux, Inc.) screening methods for susceptibility testing. Methodologies and breakpoints may vary; it is important to review the criteria used in the laboratory’s region. Because of emerging resistance to fluoroquinolones, susceptibility testing is recommended for patients with Campylobacter spp. gastroenteritis. Susceptibility testing should be performed on all isolates from sterile fluids and body sites. 

Prevention No vaccines are available for Campylobacter spp. Infections caused by Campylobacter spp. are acquired by ingesting contaminated foodstuffs or water. Proper preparation and cooking of all foods derived from animal sources, particularly poultry, will decrease the risk of transmission. All milk

TABLE 33.2    Differential Characteristics of Clinically Relevant Campylobacter and Arcobacter spp.

Growth at 25°C

H2 Required

Hippurate Hydrolysis

Catalase

H2S in Triple Sugar Iron Agar

Indoxyl Acetate Hydrolysis

Selenite Reduction

Urease

Campylobacter coli

−

−

−

+

−

+

+

−

Campylobacter concisus

−

+

−

−

+

−

V

−

Campylobacter curvusa

−

+

−

−

+

V

−

−

Campylobacter fetus subsp. fetus

+

−

−

+

−

−

V

−

Campylobacter fetus subsp. venerealis

+

−

−

V

−

−

V

−

Campylobacter gracilis

ND

ND

−

V

−

V

−

−

−

+

−

−

−

−

−

ND

Campylobacter hyointestinalis subsp. hyointestinalis

V

V

−

+

+

−

+

−

Campylobacter jejuni subsp. jejuni

−

−

+

+

−

+

V

−

Campylobacter jejuni subsp. doylei

−

−

V

V

−

+

−

−

Campylobacter lari

−

−

−

+

−

−

V

V

Campylobacter pyloridis

ND

ND

−

+

ND

ND

ND

ND

−

+

−

V

−

+

−

−

Campylobacter showae

−

+

−

+

−

V

−

−

Campylobacter sputorum bv. sputorum

−

+

−

−

+

−

V

−

Campylobacter upsaliensis

−

−

−

−

−

+

+

−

ureolyticusa

−

+

−

V

−

−

−

+

Arcobacter butzlerib

V

−

−

V

−

+

−

−

Arcobacter cryaerophilusc

+

−

−

V

−

+

−

−

Arcobacter skirrowii

+

−

−

+

−

+

ND

−

Campylobacter

Campylobacter

Campylobacter

hominisa

rectusa

+, positive; −, negative; ND, test not determined; V, variable. aAnaerobic, not microaerobic. bGrows at 40°C. cAerotolerant, not microaerobic; except for a few strains, A. cryaerophilus cannot grow on MacConkey agar, but A. butzleri does.

CHAPTER 33  Campylobacter, Arcobacter, and Helicobacter

Genus and Species

463

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should be pasteurized and drinking water chlorinated. Care must be taken during food preparation to prevent crosscontamination from raw poultry to other food items. 

Helicobacter spp. General Characteristics Approximately 39 species are included in the genus Helicobacter, the majority of which colonize mammalian stomachs or enterohepatic regions (intestine, liver, and biliary tract). The genus Helicobacter consists of curved, helical or spiral, or fusiform microaerophilic, gram-negative rods with or without periplasmic fibers, with the majority of species exhibiting urease activity. The organisms may appear coccoid or spheroidal if cultivated for long periods or in suboptimal growth conditions. 

Epidemiology and Pathogenesis Helicobacter pylori’s primary habitat is the human gastric mucosa. The organism is distributed worldwide and although acquired early in life in underdeveloped countries, the exact mode of transmission is unknown. An oral-oral, fecal-oral, and a common environmental source have been proposed as possible routes of transmission, with familial transmission associated with H. pylori infections. Research studies suggest mother-to-child transmission as well as among siblings and other household contacts as the most probable cause of intrafamilial spread. In industrialized nations, antibody surveys indicate that approximately 50% of adults older than 60 years are infected by H. pylori. Gastritis incidence increases with age. H. pylori has occasionally been cultured from feces and dental plaque, thereby suggesting a fecal-oral or oral-oral transmission. The habitat for H. heilmannii and closely related species (H. heilmannii-like or HHLO) appears to be the human gastrointestinal tract with one or more species (H salomonis, H. felis, H. suis) of zoonotic origin. H. heilmannii, HHLO, and H. pylori may be normal microbiota of the human host. Additional enterohepatic species inhabit the intestinal and hepatobiliary tract of birds and mammals, such as H. bilis, H. canadensis, H. canis, H. cinaedi, H. fennelliae, and H. pullorum. H. pylori is capable of colonizing the mucous layer of the antrum, cardia, and corpus of the stomach and areas of the gastric metaplasia of the proximal duodenum but fails to invade the epithelium. Motility allows H. pylori to escape the acidity of the stomach and burrow through and colonize the gastric mucosa in close association with the epithelium. In addition, the organism produces urease that hydrolyzes ureaforming ammonia (NH3), significantly increasing the pH around the site of infection. The change in pH protects the organism from the acidic environment produced by gastric secretions. H. pylori also produces a protein called CagA and injects the protein into the gastric epithelial cells. The protein subsequently affects host cell gene expression, inducing cytokine release and altering cell structure, and interactions with

TABLE   Genes and Their Possible Role in Enhancing 33.3  the Virulence of Helicobacter pylori

Gene

Possible Role

VacA

Exotoxin (VacA) Creates vacuoles in epithelial cells, decreases apoptosis, and loosens cell junctions

CagA

Pathogenicity island Encodes a type IV secretion system for transferring CagA proteins into host cells

BabA

Encodes outer membrane protein: mediates adherence to blood group antigens on the surface of gastric epithelial cells

IceA

Presence associated with peptic ulcer disease in some populations

neighboring cells, enabling H. pylori to successfully invade the gastric epithelium. Individuals who demonstrate positive antibody response to the CagA protein are at increased risk of developing both peptic ulcer disease and gastric carcinoma. Other possible virulence factors include adhesins for colonization of mucosal surfaces, mediators of inflammation, and a cytotoxin capable of causing damage to host cells (Table 33.3). Although H. pylori is noninvasive, untreated colonization persists despite the host’s immune response. 

Spectrum of Disease H. pylori causes gastritis, peptic ulcer disease, and gastric cancer. However, most individuals tolerate the presence of H. pylori for decades with few, if any, symptoms. Infection with this organism is also a risk factor for the development of atrophic gastritis, gastric ulcer disease, gastric adenocarcinomas, and gastric mucosa–associated lymphoid tissue (MALT) lymphomas. Enterohepatic helicobacters have been identified in association with human disease including H. fennelliae, H. canis, H. cinaedi, and H. pullorum. These isolates are transmitted from animals to humans and may be isolated from human blood or fecal samples. H. cinaedi and H. fennelliae have been isolated in cases of proctocolitis, gastroenteritis, neonatal meningitis, skin rashes, and bacteremia in immunocompromised patients. HHLO have been isolated in association with human cases of mild to moderate gastritis, peptic ulcer, and gastric MALT. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing Tissue biopsy material of the stomach for detection of H. pylori should be placed directly into transport media such as Stuart’s transport medium, Brucella broth with 20% glycerol, or Portagerm pylori media (bioMérieux, Inc., Durham, NC) to prevent drying. Specimens for biopsy may be refrigerated up to 24 hours before processing; tissues should be minced and gently homogenized. If longer storage is required, samples should be frozen at −70°C in a 10% glycerol–containing medium.

CHAPTER 33  Campylobacter, Arcobacter, and Helicobacter

465

Fecal specimens may be used for stool antigen tests but cannot be used for routine culture of gastric helicobacters. These samples should be tested immediately and stored at −20°C or according to the manufacturer’s recommendations. Repeated freezing and thawing of specimens should be avoided. Enterohepatic helicobacters may be processed for culture using campylobacter protocols as previously described in this chapter. Blood samples for serologic diagnosis of Helicobacter spp. infection may be collected, transported, and processed by standard methods. Gastric juice has been used for nucleic acid detection and culture of H. pylori. 

Direct Detection Gastric biopsy specimens that are preserved in 10% f­or­ maldehyde or paraffin-imbedded tissue are often used for the histopathologic diagnosis of H. pylori infection. Pathologists use the Warthin-Starry or other silver stains to examine biopsy specimens. Squash preparations of biopsy material can be performed and stained with rapid Giemsa stain, fluorescent acridine orange stain, or Gram stain. Gram stain using an alternate secondary stain, 0.5% carbolfuchsin or 0.1% basic fuchsin, enhances recognition of the bacteria’s typical morphology. However, because of the presence of bacterial atypical morphologies, the results may not be interpreted correctly. Sampling error may occur during processing, resulting in no identification of the organisms. Presumptive evidence of the presence of H. pylori in biopsy material may be obtained by placing a portion of crushed tissue biopsy material directly into urease broth, onto commercially available urease agar kits, or on a paper strip containing a pH indicator. These tests are collectively referred to as rapid urease tests (RUTs). A positive test is considered indicative of the organism’s presence. The CLOtest (Kimberly-Clark, Neenah, Murray Hill, NG) is an example of an agar-gel based RUT (Fig. 33.4). Another noninvasive indirect test to detect H. pylori is the urea breath test (UBT). This test relies on the presence of H. pylori urease. The patient ingests nonradioactively labeled natural isotope (13C) urea, and if the organism is present, the urease produced by H. pylori hydrolyzes the urea to form ammonia and labeled bicarbonate that is exhaled as 13CO2; the 13CO2 is detected by a special spectrometer. This test has excellent sensitivity and specificity. EIA H. pylori stool antigen tests (Premier Platinum HpSA, Meridian Diagnostics, Inc., Cincinnati, OH, or IDEIA Hp StAR, Oxoid Ltd., Basingstoke, United Kingdom) and rapid immunochromatographic point of care assays using monoclonal antibodies (ImmunoCard STAT! HpSA, Meridian Bioscience, Cincinnati, OH, or RAPID Hp StAR, Oxoid Ltd., Basingstoke, United Kingdom) have been introduced to directly detect H. pylori. 

Nucleic Acid Detection A variety of nucleic acid–based methods have been developed to directly detect H. pylori and HHLO in clinical specimens and to identify bacterial strains and host genotype characteristics, bacterial density in the stomach, and antimicrobial

•

Fig. 33.4 CLOtest Rapid Urease Test indicating a positive reaction (top test) and a negative reaction (bottom test).

resistance patterns. Polymerase chain amplification and fluorescence in situ hybridization using species-specific probes has enhanced the diagnosis and detection in gastric biopsy specimens. Amplification targets include the 16S RNA, 23S rRNA ureA, glmM, vacA, and cagA genes. Interpretation of nucleic acid–based methods should be used in conjunction with other diagnostics methods. There are no methods available for the detection of enterohepatic Helicobacter species of clinical relevance at the time of this writing. Varieties of methods are under development for the direct detection of H. pylori nucleic acids in feces. These techniques are problematic in that DNA is often degraded when passing through the gastrointestinal tract as well as the presence of a variety of amplification enzyme inhibitors in feces. Potential new methods are expected to be placed in clinical trials and submitted for FDA approval that would not only detect the organism but also the presence of mutations that confer macrolide resistance. 

Cultivation Stool specimens submitted for culture of enterohepatic helicobacters such as H. bilis, H. canadensis, H. canis, H. cinaedi, H. fennelliae, and H. pullorum are inoculated onto selective media used for Campylobacter isolation but without cephalothin, such as Campy-CVA. H. cinaedi, H. canis, and H. fennelliae have been occasionally isolated from commercial blood culture systems in patients suspected of bacteremia. The recovery of H. pylori from tissue biopsy specimens, including gastric antral biopsies; nonselective agar media, including brain heart infusion agar, and Brucella agar with 5% sheep blood; Wilkins Chalgren agar; and trypticase soy agar have resulted in successful recovery of the organisms. The combination of a selective agar (Columbia agar with an egg yolk emulsion, supplements, and antibiotics) and a nonselective

466 PA RT I I I    Bacteriology

TABLE 33.4    Differential Characteristics of Helicobacter Species

Species

Catalase

Urease

Indoxyl Acetate Hydrolysis

Nitrate

Alkaline Phosphatase

Gamma-glutamyl Transpeptidase

H. bilis

+

+

−

+

−

+

H. canis

−

−

+

−

+

+

H. canade­ nsis

+

−

+

+

−

−

H. cinaedi

+

−

−

+

−

−

H. felis

+

+

−

+

+

+

H. fennelliae

+

−

+

−

+

−

H. heilmanni

ND

+

ND

ND

ND

ND

H. salomonis

+

+

+

+

+

+

H. suis

+

+

−

−

+

+

H. pullorum

+

−

−

+

−

ND

+, Positive; −, negative; ND, test not determined.

agar (modified chocolate agar with Columbia agar, 1% Vitox, and 5% sheep blood) was reported as the optimal combination for recovering H. pylori from antral biopsies. Incubation up to 1 week in a humidified, microaerobic environment (4% O2, 5% CO2, 5% H2, and 86% N2 atmosphere) at 35°C to 37°C may be required before growth is visible. 

Approach to Identification Colonies of Helicobacter spp. may require 4 to 7 days of incubation before growth is observed. Colonies may appear as small, translucent, circular colonies or swarming phenotypes from some gastric isolates. Culture plates should be reviewed daily for a minimum of 10 days before a negative culture is reported. Organisms are identified presumptively as H. pylori by the typical cellular morphology and positive results for oxidase, catalase, and RUT. H. pylori and some enterohepatic species may be definitively identified by using a similar approach to Campylobacter spp. However, on subculture these organisms may lose their classic morphology, making identification difficult. MALDI-TOF has been successfully used to identify H. pylori from cultures of gastric biopsy specimens (Table 33.4). 

Serodiagnosis Serologic diagnosis is also available for H. pylori. The immune response typically presents with a rise in immunoglobulin M (IgM), followed by immunoglobulin G (IgG) and immunoglobulin A (IgA), although 2% of patients fail to seroconvert. Numerous serologic EIAs designed to detect IgG and IgA antibodies to H. pylori are commercially available. Reported performance of these assays varies depending on the reference method used to confirm H. pylori infection, the antigen source for the assay, and the population studied. In addition to variability in assay performance, the clinical

utility of IgA testing in these assays is controversial. In some cases, IgA has demonstrated a much lower sensitivity and specificity than IgG for the detection of H. pylori infections. These assays are incapable of differentiation of active versus past H. pylori infections. As of this writing, there are no serologic assays for the routine diagnosis of HHLO or enterohepatic Helicobacter spp. 

Antimicrobial Susceptibility Testing and Therapy Except for metronidazole and clarithromycin, most laboratory susceptibility assays are unsuccessful in predicting clinical outcome. Routine testing of H. pylori isolates’ susceptibility to metronidazole is recommended using the ETEST and agar or broth dilution methods. Therapy for H. pylori infection is problematic. H. pylori readily becomes resistant when metronidazole, clarithromycin, azithromycin, rifampin, or ciprofloxacin is prescribed as a single agent. Current regimens recommend triple-drug therapy including a proton pump inhibitor, clarithromycin, and either amoxicillin or metronidazole. An alternative treatment for H. pylori infection includes fluoroquinolone, levofloxacin, and a rifamycin (rifabutin). Similar treatments have demonstrated successful therapy in patients with gastritis and peptic ulcer disease associated with HHLO. No guidelines exist for the treatment of enterohepatic helicobacters. 

Prevention No vaccines are available for H. pylori.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

CHAPTER 33  Campylobacter, Arcobacter, and Helicobacter

Bibliography Allos BM: Campylobacter jejuni infections: update on emerging issues and trends, Clin Infect Dis 32:1201–1206, 2001. Blaser MJ: The biology of cag in the Helicobacter pylori-human interaction, Gastroenterology 128:1512–1515, 2005. Bullman S, Lucid A, Corcoran D, Sleator RD, Lucey B: Genomic investigation into strain heterogeneity and pathogenic potential of the emerging gastrointestinal pathogen Campylobacter ureolyticus, PloS One 8:e71515, 2013. Butzler JP: Campylobacter, from obscurity to celebrity, Clin Microbiol Infect 10:868–876, 2004. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Crowe SE: Helicobacter infection, chronic inflammation, and the development of malignancy, Curr Opin Gastroenterol 21:32–38, 2005. Day AS, Jones NL, Lynetl JT, et al.: cagE is a virulence factor associated with Helicobacter pylori-induced duodenal ulceration in children, J Infect Dis 181:1370–1375, 2000. Engberg J, On SL, Harrington CS, Gerner-Smidt P: Prevalence of Campylobacter, Arcobacter, Helicobacter, and Sutterella spp. in human fecal samples as estimated by a reevaluation of isolation methods for campylobacters, J Clin Microbiol 38:286–291, 2000. Fitzgerald C, Patrick M, Gonzalez A, et al.: Multicenter evaluation of clinical diagnostic methods for detection and isolation of Campylobacter spp. from stool, J Clin Microbiol 54:1209–1215, 2016. Gemmell MR, Berry S, Mukhopadhya I, et al.: Comparative genomics of Campylobacter concisus: analysis of clinical strains reveals genome diversity and pathogenic potential, Emerg Microbes Infect 7:116, 2018. Gonzalez MD, Wilen CB, Burnham CA: Markers of intestinal inflammation for the diagnosis of infectious gastroenteritis, Clin Lab Med 35:333–344, 2015. Gorkiewicz G, Feierl G, Schober C, et al.: Species-specific identification of campylobacters by partial 16S rRNA gene sequencing, J Clin Microbiol 41:2537–2546, 2003. Hsieh YH, Wang YF, Moura H, et al.: Application of MALDI-TOF MS systems in the rapid identification of Campylobacter spp. of public health importance, J AOAC Int 101:761–768, 2018. Han S, Zschausch H, Meyer HW, et al.: Helicobacter pylori: clonal population structure and restricted transmission within families revealed by molecular typing, J Clin Microbiol 38:3646–3651, 2000.

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Konno M, Fujii N, Yakota S, et  al.: Five-year follow-up study of mother-to-child transmission of Helicobacter pylori infection detected by a random amplified polymorphic DNA fingerprinting method, J Clin Microbiol 43:2246–2250, 2005. Leber AL: Fecal and other gastrointestinal cultures and toxin assays; fecal cultures for campylobacter and related organisms and gastroenteritis panels. In Clinical microbiology procedures handbook, ed 4, Washington, DC, 2016, ASM Press. Maher M, Finnegan C, Collins E, et al.: Evaluation of culture methods and a DNA probe-based PCR assay for detection of Campylobacter species in clinical specimens of feces, J Clin Microbiol 41:2980– 2986, 2003. Matsui H, Takahashi T, Murayama SY, et al.: Development of new PCR primers by comparative genomics for the detection of Helicobacter suis in gastric biopsy specimens, Helicobacter 19:260–271, 2014. Nachamkin I, Nguyen P: Isolation of Campylobacter species from stool samples by use of a filtration method: assessment from a United States-based population, J Clin Microbiol 7:2204–2207, 2017. O’Donovan D, Corcoran GD, Lucey B, Sleator RD: Campylobacter ureolyticus: a portrait of the pathogen, Virulence 5:498–506, 2014. Pereira V, Abraham P, Nallapeta S, Shetty A: Gastric bacterial flora in patients harbouring Helicobacter pylori with or without chronic dyspepsia: analysis with matrix-assisted laser desorption ionization time-of-flight mass spectroscopy, BMC Gastroenterol 18:20, 2018. Rieder G, Fischer W, Haas R: Interaction of Helicobacter pylori with host cells: function of secreted and translocated molecules, Curr Opin Microbiol 8:67–73, 2005. Sakar SR, Hossain MA, Pual SK, Mahmud MC, Ray NC, Haque N: Use of modified blood agar plate for identification of pathogenic campylobacter species at Mymensingh Medical College, Mymensingh Med J 23:667–671, 2014. Simala-Grant J, Taylor DE: Molecular biology methods for the characterization of Helicobacter pylori infections and their diagnosis, APMIS 112:886–897, 2004. Vandenberg O, Dediste A, Houf K, et  al.: Arcobacter species in humans, Emerg Infect Dis 10:1863–1867, 2004.

CASE STUDY 33.1 A 10-year-old male became ill a few days after a Fourth of July picnic where fried chicken was served. He complained of diarrhea, abdominal pain, and fever. Symptoms continued over the next week, and he was seen at the local clinic. Blood was found in his stool, and cultures were ordered. He was treated with ampicillin but switched to azithromycin (a macrolide similar to erythromycin) for 5 days when the culture results were reported.

Questions 1. At 42°C in a microaerobic environment, water droplet– type oxidase- and catalase-positive colonies were isolated. A Gram stain showed gram-negative rods with seagull-shaped morphology. What rapid test is used to confirm the identity of this bacterium? 2. What follow-up testing would be required if the hippurate hydrolysis is negative? 3. What is the most likely route of transmission to the patient in this incident?

Chapter Review 1.  Campylobacter are: a. Small, curved, motile, gram-positive bacilli b. Small, curved, motile, gram-negative bacilli c. Small, curved, nonmotile, gram-negative bacilli d. Small, curved, nonmotile, gram-positive bacilli 2.  Campylobacter species produce the following syndromes in immunocompetent patients except: a. Endocarditis b. Febrile systemic disease c. Periodontal disease d. Gastroenteritis 3.  C. jejuni and C. coli are usually transmitted by: a. Food b. Milk c. Water d. All of the above 4. Which Campylobacter species has been recognized as the most common causative agent of gastroenteritis in the United States? a.  C. lari b. C. fetus c.  C. coli d. C. jejuni 5. Which of the following has been recognized in postinfection complications of a C. jejuni infection? a. Guillain-Barré syndrome b. Chronic pulmonary disease c. Encephalitis d. Endocarditis 6.  Campylobacter species should be grown at what temperature? a. 25°C b. 37°C c. 42°C d. 37°C and 42°C 7. A positive hippurate hydrolysis is a characteristic of: a.  C. coli b. C. jejuni c.  C. lari d. C. fetus 8.  Campylobacter infection may be prevented by which of the following? a. Thoroughly cooking all foods b. Pasteurized milk c. Chlorinated water d. All of the above

9.  All of the following agars may be used to isolate Helicobacter except: a. Campy-CVA b. CCDA c. Thiosulfate-citrate-bile salts-sucrose (TCBS) d. CSM 10.  H. pylori may be identified presumptively by which positive tests? a. NO3, hippurate, catalase b. Oxidase, indoxyl acetate, NO3 c. Oxidase, catalase, rapid urea d. NO3, catalase, growth at 25°C 11.  True or False _____ Hippurate hydrolysis can be used to differentiate C. jejuni from all other Campylobacter species. _____ Campylobacter causes febrile systemic disease, periodontal disease, and gastroenteritis in humans. _____ Campylobacter is most commonly transmitted via respiratory droplets. _____ Stool specimens should be plated on MacConkey, Hektoen enteric (HE), and sorbitol-MacConkey (SMAC) agar for optimal recovery of Campylobacter species. _____ Curved, microaerophilic, gram-negative rods showing strong urease activity are suggestive of Campylobacter species. _____ Enterohepatic helicobacters may be isolated using the standard laboratory procedures for the isolation of Campylobacter spp. _____ H. pylori causes gastritis, peptic ulcer disease, and gastric cancer. 12.  Matching: Match the correct term with the appropriate description. _____ microaerobic _____ Guillain-Barré _____ Cary-Blair _____ seagull-winged _____ darting motility _____ H. pylori _____ CCDA _____ demyelination

a. removal of the myelin sheath from a nerve b. long, polar flagella c. requires less oxygen d. neurologic syndrome e. a selective agar f. urea diagnostic test in broth g. helix-shaped morphology h. transport medium 467.e1

34

Legionella OBJECTIVES 1. Define the general characteristics of Legionella spp. 2. List sources for Legionella in the environment, including those that are naturally occurring and those that are manmade. 3. State the means of transmission for Legionella spp. 4. Describe the life cycle of Legionella, and explain how it avoids destruction by the host. 5. Compare and contrast the three primary clinical manifestations of Legionella, including signs and symptoms. 6. List the appropriate specimen collection, transport, and processing techniques for Legionella spp. 7. Outline the different types of testing for Legionella, including sensitivity and specificity. 8. Explain the chemical principle for buffered charcoal–yeast extract (BCYE) with and without inhibitory agents and the proper use for each. 9. Describe the morphology of the Legionella spp. when grown under optimal growth conditions, including oxygenation, temperature, and length of incubation. 10. State the drugs of choice for effective therapy.

GENUS AND SPECIES TO BE CONSIDERED Legionella dumoffii Legionella micdadei Legionella longbeachae Legionella pneumophila L. pneumophila subsp. pneumophila L. pneumophila subsp. fraseri L. pneumophila subsp. pascullei Legionella spp.

Legionella belongs to the family Legionellaceae, which includes a single genus, Legionella, comprising approximately 59 species and 3 subspecies. Legionella pneumophila is the causative agent of Legionnaires’ disease, a febrile and pneumonic illness with numerous clinical presentations. Legionella was discovered in 1976 by scientists at the Centers for Disease Control and Prevention (CDC) who were investigating an epidemic of pneumonia among Pennsylvania State American Legion members attending a convention in Philadelphia. There is retrospective serologic evidence of Legionella infection as far back as 1947. Bacteria resembling Legionella that are capable of living in amoebae 468

have been designated as Legionella-like amoebal pathogens (LLAPs). Legionella lytica, one of the LLAPs, has been shown to cause human disease.

General Characteristics All Legionella spp. are mesophilic (20°C to 42°C), obligately aerobic, faintly staining, thin, gram-negative, fastidious bacilli. Legionella do not grow on routine media and require a medium supplemented with iron, L-cysteine, branchedchain fatty acids, and ubiquinones and buffered to pH 6.9 for optimal growth. The organisms are asaccharolytic and utilize protein for energy generation. The overwhelming majority of Legionella spp. are motile. There are approximately 26 species of Legionella documented as human pathogens in addition to L. pneumophila. There have been more than 500 strains of L. pneumophila that have been completely sequenced, which indicates that the species is genetically diverse. The gene sequence for the lipopolysaccharide core and the O side chain for serogroup 1 demonstrates a predominance in clinical isolates of L. pneumophila. Box 34.1 is an abbreviated list of some of the species of Legionella.

Epidemiology Legionellae are ubiquitous and widely distributed in the environment. As a result, most individuals are exposed to Legionella spp.; however, few develop symptoms. In nature, legionellae are found primarily in aquatic habitats and thrive at warmer temperatures; these bacteria can survive extreme ranges of environmental conditions for long periods; studies have shown that Legionella spp. are capable of survival for several months in free-flowing water. The organisms exist in microbial biofilms as intracellular parasites of freeliving amoebae including Acanthamoeba and Naegleria spp. Legionella spp. have been isolated from most natural water sources investigated, including lakes, rivers, and marine waters, as well as moist soil. Organisms are also widely distributed in man-made facilities, including air-conditioning ducts and cooling towers; potable water; large, warm-water plumbing systems; humidifiers; whirlpools; and technicalmedical equipment in hospitals. L. longbeachae, first isolated in Long Beach, California, is found predominantly in potting soil and compost.

CHAPTER 34  Legionella

Legionella infections are acquired exclusively from environmental sources; no person-to-person spread has been documented. Inhalation and aspiration of infectious aerosols (1–5 μm in diameter) are considered the primary means of transmission. Exposure to these aerosols can occur in the workplace or in industrial or health care settings; for example, nebulizers filled with tap water and showers have been implicated. 

• BOX 34.1 Legionella spp. Isolated From Human

Sources

Legionella pneumophila Legionella anisa Legionella birminghamensis Legionella cardiaca Legionella cincinnatiensis Legionella micdadei Legionella bozemanae Legionella dumoffii Legionella feeleii Legionella gormanii Legionella hackeliae Legionella jordanis Legionella lansingensis Legionella londiniensis Legionella lytica Legionella maceachernii Legionella nagasakiensis Legionella norrlandica Legionella longbeachae Legionella oakridgensis Legionella parisiensis Legionella rubrilucens Legionella sainthelensi Legionella steelei Legionella tucsonensis Legionella wadsworthii

Pathogenesis and Spectrum of Disease The virulence mechanisms of Legionella spp. are an important factor in the ability to infect and subsequently multiply within amoebae (Acanthamoeba and Naegleria spp.); Tetrahymena spp., a ciliated protozoan; and certain host cells. The organism can also multiply within biofilms, well-organized microcolonies of bacteria usually enclosed in polymer matrices that are separated by water channels that remove wastes and deliver nutrients. This contributes to the organism’s survival in the environment. In addition, L. pneumophila exists in two well-defined, morphologically distinct forms in Hela cells: (1) a highly differentiated, cystlike form that is highly infectious, metabolically dormant, and resistant to antibiotics and detergent-mediated lysis and (2) a replicative intracellular form that is structurally similar to agar-grown bacteria. The existence of the cystlike form may account for the ability of L. pneumophila to survive for long periods between hosts (amoebae or humans). L. pneumophila is considered a facultative intracellular pathogen. The organism causes disease by infecting human monocytes, predominantly alveolar macrophages. Once inside the macrophage, the organism is able to avoid destruction by the host’s phagocytic cells and multiply. Molecular analysis of L. pneumophila has demonstrated that the organism’s genome contains eukaryotic-like gene sequences that in effect subvert the normal eukaryotic cellular functions for intracellular survival. The pathogenesis of L. pneumophila has been extensively studied; however, little is known about the pathogenic mechanisms associated with other Legionella spp. Legionella have a similar life cycle in both protozoa and human macrophages, as illustrated in Fig. 34.1: • Binding of microorganisms to receptors on the surface of eukaryotic cells • Penetration of microorganisms into phagocytes • Escape from bactericidal attack • Formation of a replicative vacuole (a compartment within the cell where bacterial replication occurs)

5 min

L. pneumophila

30 min 4-6 h

Dot/Icm T4SS Effector proteins 14 h Nucleus Golgi complex

469

Endoplasmic reticulum Macrophage

Infection of neighboring cells

• Fig. 34.1  Legionella pneumophila cellular trafficking and growth mechanism inside a human macrophage. (Modified from 2009 annualreport.nichd.nih.gov/ump.html.)

470 PA RT I I I    Bacteriology

• I ntracellular multiplication and killing of the host cell There are differences in the mechanisms used to enter and exit from the respective type of host cell. After infection, organisms are taken up by phagocytosis primarily in alveolar macrophages. Almost immediately following phagocytosis, L. pneumophila injects hundreds of pathogenicity effector molecules into the host macrophage using a type IVB secretion system (Dot or Icm). Strains that do not have the type IVB secretion system are avirulent. A type IVB secretion system has been identified in Legionella dumoffii. Once inside the host macrophage, the microorganism survives and replicates within a specialized, membrane-bound vacuole by resisting acidification and evading fusion with lysosomes. After replication, the organisms will kill the phagocytes, releasing them into the lungs. The organisms will again be phagocytized by a mononuclear cell, and multiplication of the organism will increase. Many bacterial pathogens use secretion systems as a part of how they cause disease. L. pneumophila utilizes the type IVB secretion system, referred to as defective organelle trafficking (Dot) or intracellular multiplication (Icm), to “trick” the eukaryotic macrophage into transporting the organism to the endoplasmic reticulum. In eukaryotic cells, most proteins secreted or transported inside vesicles to other cellular compartments are synthesized at the endoplasmic reticulum (ER) (Fig. 34.1). This Dot/Icm secretion system in L. pneumophila consists of 27 genes. Bacterial type IV secretion systems are bacterial devices that deliver macromolecules such as proteins across and into cells. After entry but before bacterial replication, L. pneumophila, residing in a membrane-bound vacuole, is surrounded by a ribosome-studded membrane derived from the host cell’s ER and mitochondria. Thus, by exploiting host cell functions, L. pneumophila is able to gain access to the lumen of the ER, which supports its survival and replication where the environment is rich in peptides. A second type II secretion system has also been implicated in the virulence of some strains of Legionella. The type II secretion system carries numerous genes for enzymatic degradation, including lipases, proteinases, and a number of proinflammatory or

tissue destructive proteins. The type II secretion system produces a number of novel proteins that permit growth of the organism in low temperatures, is required for biofilm establishment, sliding motility, and intracellular infection. Mutations within the type II secretion system result in decreased infectivity of the organism. Legionella is the only intracellular pathogen known to possess a functional type II secretion system. Several additional bacterial factors have also been identified as crucial for intracellular infection; some of these are listed in Box 34.2. Finally, several cellular components and extracellular products of L. pneumophila, such as an extracellular cytotoxin that impairs the ability of phagocytic cells to use oxygen and various enzymes (e.g., phospholipase C), have been purified and proposed as virulence factors. The sequestering of Legionella within macrophages also makes it difficult to treat the infection effectively with antimicrobial agents. Therefore, a competent cell-mediated immune response is also important for recovery from Legionella infections. Humoral immunity appears to play an insignificant role in the defense against this organism. Legionellosis is a spectrum of clinical presentations, ranging from asymptomatic infection to severe, life-threatening diseases. Serologic evidence exists for the presence of asymptomatic disease, because many healthy people tested possess antibodies to Legionella spp. Table 34.1 provides a • BOX 34.2 Examples of Legionella pneumophila

Factors Crucial for Intracellular Infection

• • • •

 eat shock protein 60 H Outer membrane protein Macrophage infectivity potentiator Genes encoding for the type II secretion systems required for intracellular growth • Type IV pili • Flagella • Dot/Icm type IVB secretion system

TABLE 34.1    Disease Spectrum Associated With Legionella spp.

Epidemiology

Disease

Pneumonia (Legionnaires’ disease)

Community and nosocomial, health care–associated transmission (inhalation of aerosolized particles); immunocompromised patients, particularly in cellmediated immunity; rarely occurs in children

Acute pneumonia indistinguishable from other bacterial pneumonias; clinical syndrome may include nonproductive cough, myalgia, diarrhea, hyponatremia, hypophosphatemia, and elevated liver enzymes.

Pontiac fever

Community setting associated with employment (industrial or recreational) or other group

Self-limiting, febrile illness; symptoms may include cough, dyspnea, abdominal pain, fever, and myalgia; pneumonia does not occur.

Extrapulmonary

Rare, metastatic complications from underlying pneumonia; incidents of inoculation into sites via punctures have been identified; therapeutic bathing; highly associated with immunocompromised patients

Abscesses have been identified in the brain, spleen, lymph nodes, muscles, surgical wounds, and a variety of tissues and organs.

From Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders.

CHAPTER 34  Legionella

more detailed description of the following three primary clinical manifestations: • Legionnaires’ disease is a severe pneumonia with a case fatality rate of 10% to 20%. It is a global public health issue. Legionnaires’ disease occurs in sporadic, endemic, and epidemic forms. The incidence of the disease varies greatly and appears to depend on the geographic area, but it is estimated that Legionella spp. cause less than 1% to 5% of cases of pneumonia. • A mild, self-limited, nonfatal, influenza-like (e.g., fever, headache, malaise) respiratory infection known as Pontiac fever. • Other rare extrapulmonary sites, such as wound abscesses, encephalitis, or endocarditis. The disease can affect anyone but principally affects those who are susceptible because of age, illness, immunosuppression, or other risk factors, such as heavy smoking. The clinical manifestations after infection with a particular species are primarily caused by differences in the host’s immune response and perhaps by inoculum size; the same Legionella spp. gives rise to different expressions of disease in different individuals. L. longbeachae is the second most common cause of legionellosis. Infections associated with L. longbeachae are particularly common in Australia, but cases have been documented in other countries including the United States. The infection is most likely due to inhalation or aspiration of contaminated dust compost or soil and can be very serious, often leading to hospitalization and sometimes death. Legionella micdadei is believed to be associated with approximately 60% of Legionnaire’s disease not caused by L. pneumophila or L. longbeachae. This organism has been predominantly isolated from immunocompromised patients. There are a few bacteria that grow within amoebae and are closely related phylogenetically based on 16S ribosomal ribonucleic acid (rRNA) gene sequencing to Legionella species: LLAPs. Several LLAPs have been assigned to the Legionella genus. One LLAP has been isolated from the sputum of a patient with pneumonia after the specimen was incubated with the amoeba Acanthamoeba polyphaga. Serologic surveys of patients with community-acquired pneumonia suggest LLAPs may be occasional human pathogens. 

Laboratory Diagnosis Although culture remains the gold standard for the diagnosis of Legionnaires’ disease, it is currently diagnosed with a urinary antigen test that is highly accurate for L. pneumophila serogroup 1. The use of selective culture media yields a high sensitivity and specificity. Other diagnostic tests, including serotyping and nucleic acid sequencing are used to fully characterize clinical isolates of Legionella spp.

Specimen Collection and Transport Specimens from which Legionella can be isolated include respiratory tract secretions of all types, including expectorated

471

sputum, additional lower respiratory specimens including pleural fluid, bronchoscopy specimens, and lung biopsy specimens. Rare sources associated with infection include other sterile body fluids, such as pericardial fluid and specimens from kidney, liver, spleen, myocardium, respiratory sinuses, skin, soft tissues, wounds, peritoneal fluid, joint fluids, bone marrow, and intestine. Because sputum from patients with Legionnaires’ disease is usually nonpurulent and may appear bloody or watery, the grading system used for screening sputum for routine cultures may not be applicable. Patients with Legionnaires’ disease usually have detectable numbers of organisms in their respiratory secretions, even for some time after antibiotic therapy has been initiated. If the disease is present, the initial specimen is often likely to be positive. However, additional specimens should be processed if the first specimen is negative, and suspicion of the disease persists. Pleural fluid has not yielded many positive cultures in studies performed in several laboratories, but it may contain organisms and should be processed if available. Urine for antigen collection should be collected in a sterile container. The sample should be transported to the laboratory and refrigerated if a delay in processing occurs. Specimens should be transported without holding media, buffers, or saline, which may inhibit the growth of Legionella. The organisms are hardy and are best preserved by maintaining specimens in a small, tightly closed container to prevent desiccation and transporting them to the laboratory within 30 minutes of collection. If a longer delay is anticipated, specimens should be refrigerated. If moisture of the specimens cannot be ensured, 1 mL of sterile broth may be added. Samples collected for Legionella spp. nucleic acid testing do not require special collection or processing. 

Specimen Processing All specimens for Legionella culture should be handled and processed in a class II biologic safety cabinet (BSC). When specimens from nonsterile body sites are submitted for culture, selective media or treatment of the specimen to reduce the numbers of contaminating organisms is proposed. Brief treatment of sputum specimens with hydrochloric acid before culture has been shown to enhance the recovery of legionellae. However, this technique is time consuming and is recommended for specimens from patients with cystic fibrosis. Respiratory secretions may be held for up to 48 hours at 5°C before culture; if culturing is delayed longer, then the specimen may be frozen. Tissues are homogenized before smears and cultures are performed, and clear, sterile body fluids are centrifuged for 30 minutes at 4000 × g. The sediment is then vortexed and used for culture and smear preparation. Blood for culture of Legionella may be processed with the lysis-centrifugation tube system (Isolator, Alere, Inc., Waltham, MA) and plated directly to buffered charcoal–yeast extract (BCYE) agar. Specimens collected by bronchoalveolar lavage are quite dilute and therefore should be concentrated at least tenfold by centrifugation before culturing. 

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• Fig. 34.2  Fluorescent antibody–stained Legionella pneumophila.

Direct Detection Methods Several laboratory methods are used to detect Legionella spp. directly in clinical specimens.

Microscopy Legionella spp. are small, gram-negative coccobacilli or short rods when directly observed in a clinical specimen. However, in indirect microscopy when examining an isolate from artificial culture media, the organism appears as long, filamentous bacilli. Legionella spp. have a thin cell wall and stain poorly in the Gram procedure if neutral red or safranin is used as the counterstain. This characteristic is related to the composition of the cell walls, which have large amounts of branched-chain cellular fatty acids. Because of their faint staining, intracellular form in human tissues and large amounts of proteinaceous material in clinical specimens, Legionella spp. are not usually detectable directly in clinical material by Gram stain. The use of 0.1% fuchsin substituted for safranin in the Gramstain procedure may enhance the visibility of the organisms. Organisms can be observed on histologic examination of tissue sections using silver stains. However, these stains generally lack sensitivity and produce significant artifacts making interpretation highly complex. L. micdadei stains with acid-fast stain in fresh and formalin fixed tissue. Direct immunofluorescent microscopy is the most sensitive and specific method for the detection of L. pneumophila in respiratory samples and tissue. The sensitivity of the DFA test ranges from 25% to 75%, and its specificity is greater than 95%. If positive, organisms appear as brightly fluorescent rods (Fig. 34.2). The high complexity of the test and lack of highly reproducible sensitivity discourage many laboratories from offering DFA testing, and it is rarely used except during autopsies. 

Antigens Microtube-based enzyme immunoassay (EIA) methods can be used to serologically diagnose L. pneumophila serogroup 1 in tissues. Several EIA serologic diagnostic kits are commercially available, with sensitivity ranging from 80% to 90% and a specificity of about 98%. The sensitivity of kits for testing antibody from other serotypes is still unknown.

Rapid detection of Legionella antigen in urine and other body fluids has been accomplished by commercially available EIA kits and by immunochromatography (ICT); however, the ICT have demonstrated up to a 40% decrease in sensitivity compared to the microtube-based immunoassays. Detecting the antigen in urine allows for early diagnosis of the infection. The antigen is detectable in most patients between 1 and 3 days after the onset of symptoms and may persist for some weeks or months. The urine antigen assays are 90% to 99.9% specific, which is similar to the traditional culture method, but may have greater sensitivity than culture. Compared with other diagnostic methods, the advantages of urinary antigen detection are striking. Specimens are easily obtained, the antigen is detectable very early in the course of disease, and the test is rapid and specific. A rapid immunochromatographic assay for detecting L. pneumophila serogroup 1 antigen in urine is also available. This assay detects urinary antigen within a very short time and does not require laboratory equipment. Concentration of urine improves the sensitivity of both the EIA and immunochromatographic assays without decreasing their specificity. A drawback of the immunochromatographic urine antigen assay is that it only detects the presence of antigen of L. pneumophila serogroup 1, which constitutes 80% to 90% of all Legionella infections. In addition, false positives may occur in urine in the presence of rheumatoid-like factors, urinary sediment, and freeze thawing of urine. All positive urine antigen tests should be confirmed. The urine sample should be clarified by brief centrifugation and boiled for 5 to 15 minutes (dependent on protocol) to remove rheumatoid-like factors. 

Nucleic Acid Detection Polymerase chain reaction (PCR) assays have been extensively used to detect deoxyribonucleic acid (DNA) from environmental and clinical samples, particularly those from the respiratory tract. Traditionally, the rRNA (16S and 23S) genes and the macrophage infectivity protein (mip) gene have been used for assays targeting the Legionella genus and the serogroup 1 target (wzm) gene for L. pneumophila–specific assays. Assays for Legionella and L. pneumophila using real-time PCR platforms have also been described. With respiratory samples, Legionella PCR has a reported specificity of more than 99% and sensitivity of 80% to 100%. An important feature of Legionella PCR is that the method can potentially detect all serogroups of L. pneumophila and other species and is therefore useful in the early diagnosis of infections, particularly in health care–associated cases and outbreak investigations. Although no FDA-cleared nucleic acid–based tests are available in the United States, they are available in Europe. Nucleic acid amplification tests have demonstrated an 11% increase in positive yields over urine antigen testing. This may be a result of the predominance of serogroup I and the Pontiac subgroup in infections. NAAT may also demonstrate improved sensitivity in immunocompromised patients because of the low sensitivity of urine testing in these patients. 

CHAPTER 34  Legionella

A

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B • Fig. 34.3  (A) Colonies of Legionella pneumophila demonstrating the cut-glass internal granular speckling.

(B) Colonies of Legionella pneumophila on buffered charcoal–yeast extract agar demonstrating blue-green glistening. (Photo courtesy Brooks Murillo-Kennedy, Houston, TX.)

Cultivation The specimen should be processed for cultivation by diluting the specimen 1:10 in tryptic soy broth or distilled water to reduce inhibition by tissue and serum factors or decontamination for sputum and respiratory specimens to remove contaminating and inhibitory normal microbiota. Decontamination to reduce contaminating microbiota in sputum and respiratory samples should be completed using a 1:10 dilution of a low pH KCl-HCl buffer (pH 2.2) and incubating at room temperature for 4 minutes prior to plating. An alternative method for decontamination is heading at 50°C for 30 minutes. Primary isolation of Legionella spp. should be carried out by inoculating a nonselective media and two different selective media. Specimens for recovery of Legionella should be inoculated to at least one BCYEα agar plate containing L-cysteine and α-ketoglutarate without inhibitory agents. This medium contains charcoal to detoxify the medium, remove carbon dioxide (CO2), and modify the surface tension to allow the organisms to proliferate more easily. BCYEα is also prepared with ACES buffer (N-[2-Acetoamido]-2-aminoethanesulfonic acid) and the growth supplements cysteine (required by Legionella), yeast extract, alpha-ketoglutarate, and iron. Additional selective media, such as the BCYEα with polymyxin B, anisomycin (to inhibit fungi), and cefamandole, is recommended for specimens, such as sputum, that are likely to be contaminated with other human microbiota. Less selective media containing polymyxin B, anisomycin, and vancomycin should be used to isolate Legionella spp. other than L. pneumophila. L. micdadei has been recovered from BCYEα supplemented with bovine serum albumin from a guinea pig spleen, but no data is available for direct culture from human specimens. BCYEα with natamycin, aztreonam, and vancomycin has been used to isolate L. longbeachae from environmental samples. Culture plates are incubated at 35°C to 37°C in a humid atmosphere for up to 14 days and are examined every 3 or 4 days. Even the detection of one or a few colonies is sufficient to confirm the diagnosis. Some Legionella spp. may be stimulated by increased 2% to 5% concentration of CO2,

• Fig. 34.4  Gram stain of a colony of Legionella pneumophila showing thin, gram-negative bacilli (arrows).

including Legionella sainthelensi and Legionella oakridgensis. The low level of CO2 will not prevent the growth of L. pneumophila. If this concentration is not possible, incubation in air is preferable to 5% to 10% CO2, which may inhibit some legionellae, specifically L. pneumophila. At 5 days, colonies are 3 to 4 mm in diameter, gray-white to blue-green, glistening, convex, and circular and may exhibit a cut-glass type of internal granular speckling (Fig. 34.3). A Gram stain yields thin, gram-negative bacilli (Fig. 34.4). 

Approach to Identification Because Legionella spp. are biochemically inert, and many tests produce equivocal results, extensive biochemical testing is of little use. Definitive identification generally requires the facilities of a specialized reference laboratory. Using a long-wave UV light in a dark room, suspect colonies of L. pneumophila will appear pale yellow-green with diffusion of the pigment into the medium. It is important to note that some young colonies will not fluoresce. Other species of Legionella will fluoresce a brilliant bluish white or a brilliant red. L. pneumophila colonies should be Gram stained, using 0.1% fuchsin as the counterstain, to determine whether bacteria are small to filamentous, gram-negative rods. Colonies should be plated to two media, including a BYCEα plate

474 PA RT I I I    Bacteriology

Serodiagnosis

Suspicious colonies on BCYEα Gram stained Small to filamentous, gram-negative rods

Growth

BYCEα containing L-cysteine

BYCEα deficient L-cysteine

Yes

No

L. pneumophila Further serotyping identification

•

Fig. 34.5 Presumptive identification of Legionella pneumophila in culture.

containing L-cysteine and one made without. L. pneumophila will only grow on the BYCEα L-cysteine media, providing a more definitive identification. In addition, if only a small amount of growth is present on the primary medium, the growth may be emulsified in sterile water and used for subculturing, staining, and serologic identification. F. tularensis is the only other gram-negative bacterium that demonstrates L-cysteine growth dependence; however, the colonial morphology is opaque and homogenous. In addition, some serotyping reagents for Legionella spp. will cross-react with F. tularensis. Once the isolate has been determined to be L-cysteine dependent, further identification is completed using serotyping. Many different bacteria, including Bordetella pertussis have been reported to cross-react with serologic reagents used for the characterization and identification of Legionella spp. It is important to correlate all diagnostic testing with cellular and colonial morphology to avoid misidentification of other bacteria as Legionella spp. Gram-negative rods that demonstrate the colonial morphology and L-cysteine growth dependency may be presumptively reported as Legionella spp. If the organism is reactive in the serogroup 1 typing assay, the organism may be presumptively identified as L. pneumophila (Fig. 34.5). Further identification of isolates can be completed using matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) or DNA sequencing. MALDI-TOF MS databases include either no Legionella spp. or only L. pneumophila. This method is unable to distinguish serogroup designations. The research use only database in MALDI-TOF MS systems has been successfully used to identify 80% to 90% of isolates being tested in reference laboratories. Upon completion of sufficient data collection, the MALDI-TOF MS databases may be used to identify the organism upon FDA clearance. 

Most patients with legionellosis have been diagnosed retrospectively by detection of a fourfold rise in anti-Legionella antibody with an indirect fluorescent antibody (IFA) test. Serum specimens should be tested no closer than 2 weeks apart. Diagnostic efficacy associated with serologic testing increases with the collection and testing of acute and convalescent paired sera. Convalescent sera should be collected at 4, 6, and 12 weeks after the appearance of the disease. Disease is confirmed by a fourfold rise in titer to more than 128. A single serum with a titer of more than 256 and a characteristic clinical picture may be presumptive for legionellosis; however, because as many as 12% of healthy persons yield titers as high as 1:256, this practice is strongly discouraged. Unfortunately, individuals with Legionnaires’ disease may not exhibit an increase in serologic titers until as long as 10 weeks after the primary illness, or they may never display significant antibody titer increases. It is essential to correlate serologic findings with the patient’s clinical presentation because of the variation in antibody response associated with legionellosis. Most patients will develop a classic IgM, IgG, and IgA response. However, some patients may develop antibodies for a single class (in other words, IgG, IgM, or IgA only). Commercially prepared antigen-impregnated slides for IFA testing are available from numerous suppliers. 

Antimicrobial Susceptibility Testing and Therapy In  vitro susceptibility studies are not predictive of clinical response and should not be performed for individual isolates of legionellae. Antimicrobial agents such as quinolones, tetracycline, and the macrolides (e.g., clarithromycin and azithromycin) are active against L. pneumophila. Penicillins, cephalosporins of all generations, and aminoglycosides are not effective and should not be used. Low-level resistance to ciprofloxacin was recently identified; however, the clinical significance is unclear. Macrolides and quinolones have also demonstrated effective treatment for infections with L. micdadei, L. dumoffii, and L. longbeachae. 

Prevention Although under development, a vaccine against Legionella infections is not currently available. The effectiveness of other approaches to the prevention of Legionella infections, such as the elimination of its presence from cooling towers and potable water, is uncertain. Legionellosis is a notifiable disease in most industrialized countries. An important piece of information for surveillance of the disease is the history of exposure. The incubation period for legionellosis is normally between 2 and 10 days. Thus, during an outbreak, an exposure history for 2 weeks before the onset of illness should be obtained from the patient. In the presence of a pneumonic illness, a laboratory diagnosis will support clinical suspicion of

CHAPTER 34  Legionella

the infection and help classify the case. Many countries have developed guidelines or regulations for the control of Legionella in water systems and for the prevention of legionellosis.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Amemura-Maekawa J, Kura F, Chida K, et al.: Legionella pneumophila and other Legionella species isolated from Legionellosis patients in Japan between 2008 and 2016, Appl Environ Microbiol 84(18) :e00721-18, 2018. Becton, Dickinson & Company: BD ProbeTec ET legionella pneumophila (LP) amplified DNA assay, package insert, Sparks, MD, 2006, Becton, Dickinson & Company. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Centers for Disease Control and Prevention. Legionnaires’ disease surveillance summary report, 2014–2015. Published October 2018. Garduño RA, Garduño E, Hiltz M, Hoffman PS: Intracellular growth of Legionella pneumophila gives rise to a differentiated form dissimilar to stationary-phase forms, Infect Immun 70:6273–6283, 2002. Greenberg D, Chiou CC, Famigilleti R, Lee TC, Yu VL: Pediatric legionellosis: implications for underdiagnoses, Lancet Infect Dis 6:529–535, 2006. Hlavsa MC, Cikesh BL, Roberts VA, et al.: Outbreaks associated with treated recreational water – United States, 2000–2014, MMWR Morb Mortal Wkly Rep 67(19):547–551, 2018. Harb OS, Kwaik YA: Interaction of Legionella pneumophila with protozoa provides lessons, ASM News 66:609, 2000. Helbig JH, Uldum SA, Bernander S, et al.: Clinical utility of urinary antigen detection for diagnosis of community-acquired, travelassociated, and nosocomial Legionnaires’ disease, J Clin Microbiol 41:838–840, 2003. Herpers BL, de Jongh BM, van der Zwaluw K, van Hannen EJ: Realtime PCR assay targets the 23S-5S spacer for direct detection and differentiation of Legionella spp. and Legionella pneumophila, J Clin Microbiol 41:4815–4816, 2003. Jarraud S, Descours G, Ginevra C, et al.: Identification of Legionella in clinical samples, Methods Mol Biol 954:27–56, 2013.

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Korevaar E, Khoo CA, Newton HJ: Genetic manipulation of nonpneumophila Legionella: protocols developed for Legionella longbeachae, Methods Mol Biol 1921:145–157, 2019. Marrie TJ, Raoult D, LaScola B, et  al.: Legionella-like and other amoebal pathogens as agents of community-acquired pneumonia, Emerg Infect Dis 7:1026, 2001. McDade JE, Shepard CC, Fraser DW, Tsai TR, Redus MA, Dowdle WR: Legionnaires’ disease: isolation of a bacterium and demonstration of its role in other respiratory disease, N Engl J Med 297(22):1197–1203, 1977. Mercante JW, Winchell JM: Current and emerging Legionella diagnostics for laboratory and outbreak investigations, Clin Microbiol Rev 28:95–133, 2015. Muder RR, Yu VL: Infection due to Legionella species other than Legionella pneumophila, Clin Infect Dis 35:990, 2002. Murdock DR: Diagnosis of Legionella infection, Clin Infect Dis 36:64–69, 2003. Osborne AJ, Jose BR, Perry J, et  al.: Complete genome sequences of two geographically distinct Legionella micdadei clinical isolates, Genome Announc 5(22):1–2, 2017. Pasculle W: Update on Legionella, Clin Microbiol Newsl 22:97, 2000. Pedro-Botet L, Yu VL: Legionella: macrolides or quinolones, Clin Microbiol Infect 12(Suppl 3):25–30, 2006. Qin T, Zhou H, Ren H, et al.: Distribution of secretion systems in the genus Legionella and its correlation with pathogenicity, Front Microbiol 8:388, 2017. Qui J: Effector translocation by the Legionella Dot/Icm type IV secretion system, Curr Top Microbiol Immunol 376:103, 2013. Roy CR, Tilney LG: The road less traveled: transport of Legionella to the endoplasmic reticulum, J Cell Biol 158:415, 2002. Salcedo SP, Holden DW: Bacterial interactions with the eukaryotic secretory pathway, Curr Opin Microbiol 8:92, 2005. Stout JE, Sens K, Mietzner S, et al.: Comparative activity of quinolones, macrolides and ketolides against Legionella species using in vitro broth dilution and intracellular susceptibility testing, Int J Antimicrob Agents 25:302–307, 2005. Yu VL, Stout JE: Rapid diagnostic testing for community-acquired pneumonia: can innovative technology for clinical microbiology be exploited? Chest 136:1618–1621, 2009.

CASE STUDY 34.1 A 6-month-old female was diagnosed clinically with pneumonia. She was treated with intramuscular ceftriaxone followed by an oral cephalosporin for 3 days. The next day she was found to be unresponsive and rushed to the hospital. She was afebrile but tachypneic (increased breathing) and tachycardic (increased heart rate); she had an elevated white blood cell (WBC) count predominated with lymphocytes. A bronchoalveolar lavage was collected and was positive for Legionella by direct fluorescent antibody. A culture grew Legionella pneumophila serogroup 6 after 8 days. Despite appropriate therapy with erythromycin and rifampin, the infant’s pulmonary disease was fatal. No underlying disease was found in the baby.

Questions 1. The Legionella urine antigen test was negative in this baby. What is the explanation for this finding? 2. The baby appeared to be a normal healthy infant. List as many risk factors as possible for acquiring Legionella pneumonia. 3. List the factors that hamper the laboratory diagnosis of Legionella. 4. In general, sputum sent to the laboratory to diagnose pneumonia will be purulent with mucus and increased polymorphonuclear WBCs. Describe the type of sputum observed with Legionella infection.  

ADVANCED CASE STUDY 34.2 A 78-year-old male retired executive presented to the clinic with a 3- to 5-day illness that consisted of a headache and diarrhea. Over-the-counter remedies and hydration were recommended as empiric therapy for viral gastroenteritis. Two days later his wife drove him to the clinic, because the man was too weak to drive. She reported that her husband had become confused and had a high fever for the past 24 hours. In addition, he had developed a dry cough and was complaining of feeling short of breath. The patient has a previous medical history of hypertension and dyslipidemia. He is a former smoker but has no structural lung disease and no history of heart failure. Physical examination revealed a temperature of 102.5°F, blood pressure of 110/65 mm Hg, a pulse of 110 bpm, a respiratory rate of 26 breaths per minute, and an oxygen saturation of 86%. A lung examination revealed crackles bilaterally in both left and right lungs. There were no signs of heart murmur or cyanosis. The patient had no other significant physical findings. Chest x-ray revealed diffuse pulmonary infiltrates bilaterally. The patient was admitted to the hospital with a provisional diagnosis of pneumonia and potentially H1N1 influenza. The following laboratory results were obtained:

Laboratory Results Chemistry Arterial pH Pco2 (carbon dioxide partial pressure) Po2 (oxygen partial pressure) HCO3− (bicarbonate) CRP (C-reactive protein) BNP (B-type natriuretic peptide) Hematology WBC (white blood cells) RBC (red blood cells) Hgb (hemoglobin) Hct (hematocrit)

Patient 7.35 40

Reference Range 7.35–7.45 35–45 mm Hg

60

75–85 mm Hg

24 12 100

20–25 mmol/L ＜1 mg/dL 9–86 pg/mL (male 75–83 years)

14 5.11 15.5 0.46

5–10 × 109/L 5–6 × 1012/L 13.5–17.5 0.41–0.53 L/L

Chemistry

Patient

Reference Range

Platelets Segmented neutrophils Lymphocytes Monocytes Eosinophils Basophils

120 85% 13% 2% 0% 0%

150–400 × 109/L 25%–60% 20%–50% 2%–11% 0%–8% 0%–2%

 Questions

1. Evaluate the laboratory results as presented. Are there any unusual indicators here or in the patient’s history that would signify a predisposition for unusual respiratory infections? 2. What additional laboratory tests would be indicated at this time? After hospitalization, the patient continued to demonstrate a low sodium level of 120 to 123 mEq/L (reference range 135–145 mEq/L) despite rehydration efforts. His diarrhea and headache resolved by day 6 of his hospitalization, but his hypoxia increased. He was subsequently intubated and placed on mechanical ventilation for respiratory failure and continued to have a markedly elevated temperature of up to 104.3°F despite antibiotic treatment with cefotaxime and azithromycin. Repeat chest x-ray indicated an increase in infiltrates. His antibiotic treatment was broadened to include fungal and anaerobic antimicrobial agents, and the patient was placed in respiratory isolation. The following serologic test results were obtained: Influenza Rapid Antigen Test—negative; TB (tuberculin) skin test—negative; sputum by endotracheal suction for acid-fast bacteria—negative; mycoplasma IgM—＜1:16 (negative); Legionella Urine Antigen Test—positive. A diagnosis of Legionella pneumonia was made. The azithromycin was maximized to optimal dosing of 500 mg/day, and the additional antibiotic treatment was discontinued. Eventually the patient was extubated and made a full recovery from the pneumonia. 3. What are the major factors in the patient’s clinical history that would point to an atypical pneumonia related to Legionella pneumophila infection?

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Chapter Review 1.  Legionella can be spread by all of the following except: a. Cooling towers b. Person-to-person contact c. Lakes d. Humidifiers 2.  All of the following are primary manifestations of Legionnaires’ disease except: a. Trench fever b. Pontiac fever c. Endocarditis d. Pneumonia 3. Which of the following agars should be used for culturing Legionella? a. CIN b. BCYE c. SMAC d. XLD 4. What is the specimen of choice for isolating Legionella? a. Stool b. Urine c. Cerebrospinal fluid (CSF) d. Respiratory secretions 5. Which of the following is acceptable for therapy? a. Fluoroquinolones b. Penicillin c. Cephalosporin d. Aminoglycosides 6.  Legionella injects proteins into the host cell by: a. Dot/Icm b. Viral multiplication c. Damaging the epithelial cell lining the blood vessels d. Inhibiting the host defense 7.  Legionella can be definitively diagnosed by a: a. Twofold rise in anti-Legionella antibody with an IFA b. Single serum with a titer of 128 c. Monoclonal immunofluorescent stain d. Gram stain

8.  Which of the following is not a characteristic of Legionella? a. Faintly staining, thin, gram-negative bacilli b. Requires iron and L-cysteine supplements c. Growth enhanced by microaerophilic conditions d. Medium buffered to pH 6.9 for optimal growth 9. All of the following are true of Legionella except: a. Thrives at warmer temperatures b. Can survive up to 5 years in water c. Has been isolated in rivers and lakes d. Is acquired from environmental sources 10.  True or False _____ Humoral immunity plays an important role in the defense against Legionella. _____ Sputum specimens should be treated with sulfuric acid before culturing. _____ Respiratory secretions may be held for 24 hours at room temperature before culturing. _____ Specimens collected by bronchial alveolar lavage should be concentrated tenfold before culturing. 11.  Matching: Match the correct term with the appropriate description. _____ Legionnaires’ _____ Pontiac fever _____ BCYEα _____ urine Ag test _____ Dot _____ Icm

a. self-limiting, nonfatal respiratory infection b. defective organelle trafficking c. intracellular multiplication d. pneumonic illness e. agar recommended for Legionella isolation f. rapid detection by EIA

35

Brucella OBJECTIVES 1. Identify the primary routes of transmission for Brucella spp. 2. List the occupations at risk for developing brucellosis. 3. Identify the signs and symptoms associated with brucellosis. 4. State two reasons why the microbiology laboratory should be notified when Brucella spp. infection is suspected. 5. Describe the media and incubation requirements to isolate Brucella spp. 6. Describe the differential characteristics of Brucella abortus, Brucella melitensis, Brucella suis, and Brucella canis. 7. Identify the species classified as select agents and discuss the reporting requirements as nationally notifiable conditions with respect to the Laboratory Response Network (LRN).

GENERA AND SPECIES TO BE CONSIDERED Classified as Select Agents Brucella abortus Brucella melitensis Brucella suis

Other Species Brucella canis Brucella ceti Brucella microti Brucella neotomae Brucella ovis Brucella papionis Brucella pinnipedialis

The family Brucellaceae comprises three genera: Ochrobactrum (Chapter 22), Mycoplana, and Brucella. Brucella spp. are discussed in this chapter. Brucellae are free-living organisms that are subcategorized into 10 recognized species. Eight of the species are terrestrial, and four of those have been associated with human disease: Brucella abortus (seven biovars), Brucella melitensis (three biovars), Brucella suis (five biovars), and Brucella canis. Two marine species, B. ceti and B. pinnipedialis, are also known to cause human disease. Of the four terrestrial species known to cause human infection, all but B. canis are considered Category B select agents. Brucellosis is a reportable disease in all 57 states and territories; it is mandatory that disease cases be reported to state and territorial 476

jurisdictions when identified by a health provider, hospital, or laboratory. Reporting requirements vary by jurisdiction. Brucellosis is also a nationally notifiable condition. Notification of brucellosis cases (without direct personal identifiers) to the Centers for Disease Control and Prevention (CDC) by state and territorial jurisdictions is voluntary for nationwide aggregation and monitoring of disease data. Clinical or diagnostic laboratories and other entities that have identified B. suis, B. melitensis, or B. abortus are required to immediately (within 24 hours) notify the CDC Division of Select Agents and Toxins (DSAT) (http://selectagents.gov).

General Characteristics Brucellae are small, facultative, intracellular, nonmotile, aerobic, gram-negative coccobacilli or short bacilli that stain poorly by conventional Gram stain. Many isolates require supplementary carbon dioxide (CO2) for growth, especially on primary isolation. Brucella spp. are closely related to Bartonella, Rhizobium, and Agrobacterium spp. 

Epidemiology and Pathogenesis The disease brucellosis occurs worldwide, especially in Mediterranean and Persian Gulf countries, India, and parts of Mexico and Central and South America. The organisms are capable of survival for extended periods (e.g., soil, 10 weeks; aborted fetuses, 11 weeks; bovine stool, 17 weeks; milk and ice cream, 3 weeks); they can survive in fresh cheese for several months. Animals are carriers and do not typically demonstrate symptoms of disease. The most prominent symptom in animals is the infectious abortion of the fetus. Brucellosis is a zoonosis and is recognized as a cause of devastating economic loss among domestic livestock. Each of the Brucella spp. that are pathogenic for humans has a limited number of preferred animal hosts (Table 35.1). In the host, Brucella spp. tends to localize in tissues rich in erythritol (e.g., placental tissue), a four-carbon alcohol that enhances their growth. Humans become infected by four primary routes: • Ingestion of infected unpasteurized animal milk products (most common means of transmission) • Inhalation of infected aerosolized particles (laboratoryacquired infection is the most important source of transmission)

CHAPTER 35  Brucella

TABLE   Brucella spp. and Their Respective Natural 35.1  Animal Hosts

Organism

Preferred Animal Host

Brucella abortus

Cattle and buffalos

Brucella melitensis

Sheep, goats, or camels

Brucella suis

Swine and a variety of wild animals

Brucella canis

Dogs

Brucella ovis

Rams (not associated with human infection)

Brucella microti

Red foxes (not associated with human infection)

Brucella neotomae

Desert and wood rats (not associated with human infection)

Brucella papionis

Baboons and bull frogs (not associated with human infection)

Brucella pinnipedialis

Marine mammals, seals (rare human infection)

Brucella ceti

Dolphins, cetaceans (rare human infection)

• D  irect contact with infected animal parts through ruptures of skin and mucous membranes • Accidental inoculation of mucous membranes by aerosolization Rare cases of transmission by blood and bone marrow transplantation and by sexual intercourse, in addition to neonatal brucellosis, have been reported. Individuals considered at risk for contracting brucellosis include dairy farmers, livestock handlers, slaughterhouse employees, veterinarians, and laboratory personnel (public health, medical, and research). The organism has a very low infectious dose (100 organisms or fewer). Mishandling and misidentification of the organism is often associated with laboratory transmission of the organism. Brucella spp. are facultative, intracellular parasites that are able to exist in both intracellular and extracellular environments. After infecting a host, brucellae are ingested by neutrophils, within which they replicate, causing cell lysis. Neutrophils containing viable organisms circulate in the bloodstream and are subsequently phagocytized by mononuclear phagocytic cells in the spleen, liver, and bone marrow. If the infection goes untreated, granulomas develop in these organs, and the brucellae survive in monocytes and macrophages. Brucellae show a tendency to invade and persist in the human host by inhibiting apoptosis (programmed cell death). Resolution of the infection depends on the host’s nutritional and immune status, the size of the inoculum and route of infection, and the Brucella species causing the infection; in general, B. melitensis and B. abortus are more virulent for humans.

477

Survival and multiplication of Brucella in phagocytic cells are features essential to the establishment, development, and chronicity of the disease. Brucella spp. can change from a smooth to a rough colonial morphology based on the composition of the cell wall lipopolysaccharide O-side chain (LPS); those with a smooth LPS are more resistant to intracellular killing by neutrophils than those with a rough LPS. The smooth phenotype has been identified in B. abortus and B. melitensis. Brucellae ensure intracellular survival by interfering with the phagosome-lysosome fusion in macrophages and epithelial cells. In addition, like Legionella spp. (Chapter 34), brucellae use a type IV secretion system, VirB, for intracellular survival and replication. Unlike Legionella spp., however, brucellae modulate phagosome transport to avoid being delivered to lysosomes. Essentially, VirB is involved in controlling the maturation of the Brucella vacuole into an organelle that allows replication. In the mouse model, if nucleic acid mutations occur in this region, B. abortus is unable to establish chronic infections. In addition, Brucella spp. produce urease, which provides protection during passage through the digestive system when the organism is ingested in food products. Urease breaks down urea, producing ammonia, and neutralizes the gastric pH. Despite our current knowledge, many questions remain about the pathogenesis of disease caused by Brucella spp. 

Spectrum of Disease The clinical manifestations of brucellosis vary greatly, ranging from asymptomatic infection to serious, debilitating disease. For the most part, brucellosis is a systemic infection that can involve any organ of the body. Symptoms, which are nonspecific, include fever, chills, weight loss, night sweats, headache, muscle aches, fatigue, and depression. Lymphadenopathy and splenomegaly are common physical findings. After an incubation period of about 2 to 4 weeks, the onset of disease is commonly insidious. Complications can occur, such as arthritis, spondylitis (inflammation of the vertebrae), genital, pulmonary, and renal complications, and endocarditis. Neurobrucellosis occurs in approximately 3% to 5% of infections and can result in mild symptoms such as fever and headache to meningitis, coma, and paralysis. Relapse is considered an important feature of brucellosis; it is associated with delayed initiation of treatment, ineffective antibiotic therapy, and positive blood culture findings during the initial presentation. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing Samples for the diagnosis and identification of infections associated with Brucella spp. may be used for culture, serology, or nucleic acid–based testing. The gold standard for the definitive diagnosis of brucellosis is isolation of the organisms in cultures of blood; bone marrow; cerebrospinal fluid

478 PA RT I I I    Bacteriology

(CSF); pleural, abdominal, and synovial fluids; urine; spleen or liver abscesses; or other tissues. Bone marrow or blood are considered the best specimens for culture. If processing is delayed, the specimen may be held in the refrigerator. Some research laboratories offer nucleic acid–based testing on blood (serum or whole blood), CSF, and bone marrow specimens for the identification of the organism. It is essential that the clinical microbiology laboratory be notified whenever brucellosis is suspected: • To ensure that specimens are cultivated in an appropriate manner for optimal recovery from clinical specimens • To prevent accidental exposure of laboratory personnel handling the specimens, because Brucella spp. are considered category B select agents (specimen labels should indicate that Brucella spp. are a potential pathogen). All specimens should be handled using a BS-3 or BSL-2 with BSL-3 precautions. Blood for culture can be collected routinely (Chapter 67) into most commercially available blood culture bottles and the lysis-centrifugation system (Isolator, Alere, Waltham, MA). For other clinical specimens, no special requirements must be met for collection, transport, or processing. Within 2 hours of obtaining a high-confidence presumptive or confirmatory result, a Laboratory Response Network (LRN) Laboratory Director or a designee must notify: • the State Public Health Laboratory Director, • the State Epidemiologist, • the Health Officer for the State Public Health Department, • the CDC Emergency Operations Center (EOC), and • the FBI Weapons of Mass Destruction (WMD) POC. For emergency and non-emergency situations, LRN laboratories will submit data for all samples, including positive and negative results related to the event, within 12 hours of obtaining each result. The LRN is a national network of local, state, federal, military, and international public health, food testing, veterinary diagnostic, and environmental testing laboratories that provides laboratory infrastructure and capacity to respond to biological and chemical public health emergencies. More information on the LRN is included in Chapter 79. 

ribosomal ribonucleic acid (rRNA), and transposon insertion sequence 711(IS711). Although nucleic acid–based testing can be used for rapid identification, standardization is needed to improve the use in routine laboratories. 

Direct Detection Methods

Cultivation

Direct stains of clinical specimens are not particularly useful for the diagnosis of brucellosis. 

Cultivation and identification of Brucella spp. remain the primary method used for laboratory diagnosis of brucellosis. Although most isolates of Brucella spp. grow on blood and chocolate agars (some isolates are also able to grow on MacConkey agar), more enriched agars and special incubation conditions generally are needed to achieve optimal recovery of these fastidious organisms from clinical specimens. Brucella agar or infusion base is recommended for specimen types other than blood. The addition of 5% heated horse or rabbit serum enhances growth on all media. ThayerMartin or Martin-Lewis medium may be used to isolate the organisms from mixed cultures or contaminated specimens. Cultures should be incubated in 5% to 10% CO2 in

Nucleic Acid Detection Conventional and real-time polymerase chain reaction (PCR) assays are reliable and specific means of directly detecting Brucella organisms in clinical specimens. Sensitivity varies among assays, ranging from 50% to 100%. This variation is related to the differences in nucleic acid extraction procedures, specimen type, and detection formats. Several gene targets have been used, including a cell surface protein (BCS P31), a periplasmic protein (BP26), 16S

Serodiagnosis Because isolating brucellae is difficult, a serologic test is widely used (e.g., serum agglutination test [SAT] or microplate agglutination [MAT]). This technique detects antibodies to B. abortus, B. melitensis, and B. suis; however, the SAT does not detect B. canis antibodies. An indirect Coombs test is performed after the SAT. This test detects nonagglutinating or incomplete antibodies in complicated and chronic cases of brucellosis. The serology associated with Brucella infection follows the classic antibody response: IgM appears initially, followed by IgG. A titer of 1:160 or greater in the SAT is considered diagnostic if this result fits the clinical and epidemiologic findings. The SAT can cross-react with class M immunoglobulins with a variety of bacteria, such as Francisella tularensis and Vibrio cholerae. Enzyme-linked immunosorbent assays (ELISAs) also have been developed. Purified LPS or protein extracts are primarily used in ELISAs. However, currently no reference antigen exists; therefore, it is important to identify the antigen used in the commercial system when evaluating test results. In patients with neurobrucellosis, ELISA offers significant diagnostic advantages over conventional agglutination methods. Additional serologic assays are commercially available, including a lateral flow dipstick for screening outbreaks and an immunocapture agglutination method. The immunocapture assay demonstrates sensitivity and specificity similar to a Coombs test and is less cumbersome to perform. The dipstick test has a high degree of sensitivity (>90%). Microarrays that contain numerous antigens are being evaluated for their efficacy in elucidating differential antibody responses from patients during different stages of disease. Serologic testing should be interpreted with care, as methods lack standardization of antigen preparation, methodologies, and detection. 

CHAPTER 35  Brucella

A

479

B • Fig. 35.1.  Growth of Brucella spp. on chocolate agar after incubation for 2 days (A) and 4 days (B).

A

B • Fig. 35.2.  Brucella melitensis with traditional Gram stain (A) and Gram stain with 2-minute safranin counterstain (B) to allow easier visualization of the organism.

a humidified atmosphere; inoculated plates are incubated for up to 3 weeks before they are considered negative and discarded. Commercial blood culture systems (e.g., BacT/Alert [bioMérieux, Durham NC], BACTEC [Becton Dickinson, Franklin Lakes, NJ], and lysis-centrifugation systems) all have successfully detected brucellae in blood. Other blood culture bottles, such as those with brain-heart infusion and trypticase soy broth support the growth of brucellae if the bottles are continuously vented and placed in a CO2 incubator. Most isolates can be detected within 5 to 7 days using commercial systems. Bottles need not be incubated longer than 10 to 14 days. Culture bottles may not become turbid. All subculture plates should be held for a minimum of 10 days. On culture, colonies appear small, convex, smooth, translucent, gamma-hemolytic, and slightly yellow and opalescent after at least 48 hours of incubation (Fig. 35.1). Rough variants may be seen with B. canis. The colonies may become brownish with age. 

Approach to Identification Brucellosis is the most commonly reported laboratoryacquired bacterial infection; therefore, all handling and manipulations of suspected Brucella spp. should be performed using BSL-3 precautions in a BSL-2 or higher biologic safety cabinet. Gram stain of the organisms reveals small coccobacilli that resemble fine grains of sand (Fig. 35.2). Brucella spp. are catalase and urease positive, and most strains are oxidase positive. B. canis may be oxidase variable. Other nonfermentative, urea-positive, oxidase- and catalase-positive, gramnegative coccobacilli that may be confused with brucellae are Bordetella, Haemophilus, Psychrobacter, Paracoccus, Methylobacterium, Cupriavidus, and Oligella spp. (Table 35.2). Brucella spp. are differentiated by the rapidity with which the organism hydrolyzes urea, its relative ability to produce hydrogen sulfide (H2S), its requirements for CO2, and its susceptibility to the aniline dyes thionine and basic fuchsin (Table 35.3). For determination of the CO2 requirement, identical plates of Brucella agar or brain-heart infusion agar

Brucella spp.

Bordetella spp.

Cupriavidus pauculus

Haemophilus spp.a

Methylobacterium spp.b

Gram stain

Faintly staining, tiny coccobacilli

Bacilli

Bacilli

Tiny coccobacilli Vacuolated bacilli

Motility

−

+

+

−

Urea

+

+

+

V

Oligella ureolyticusc

Paracoccus yeeid

Psychrobacter immobilis

Psychrobacter phenylpyruvicus

Tiny coccobacilli cocci

Coccobacilli to short rods

Coccobacilli and bacilli; decolorize poorly

−

+

−

−

−

V

+

+

V

+

aDemonstrates

no growth on blood agar; will satellite Staphylococcus aureus. bProduces a pink pigment and appears mucoid on blood agar. cPrimarily a urinary tract pathogen. dMucoid on blood agar. +, positive; −, negative; V, variable. Modified from Leber, AL. Aerobic Bacteriology, Clinical Microbiology Procedures Handbook. 4th ed. Washington, DC: ASM Press; 2016.

TABLE 35.3    Characteristics of Human Pathogenic Brucella spp. Inhibition by Dye

Species

CO2 Required for Growth

Time to Positive Urease

H2S Produced

Thioninea

Fuchsina

Brucella abortus

±

2 h (rare 24 h)

+ (most strains)

+

−

Brucella melitensis

−

2 h (rare 24 h)

−

−

−

Brucella suis

−

15 min

±

−

+ (most)

Brucella canis

−

15 min

−

−

+

aDye

tablets (Key Scientific Products, Round Rock, TX). ＋, ＞90% of strains positive; −, ＞90% of strains negative; ±, variable results.

480 PA RT I I I    Bacteriology

TABLE 35.2    Characteristics of Organisms That Resemble Brucella spp.

CHAPTER 35  Brucella

should be given equal inocula (e.g., with a calibrated loop) of a broth suspension of the organism to be tested. One plate should be incubated in CO2 and the other plate in ambient air. Most strains of B. abortus do not grow in ambient air but show growth in CO2. Presumptive identification can be reported based on the colonial morphology and a positive catalase, oxidase, urease, and slide agglutination reaction with specific B. abortus or B. melitensis antisera. Brucella spp. isolates should be sent to state or other reference laboratories for confirmation or definitive identification because most clinical laboratories lack the necessary media and containment facilities required for further analysis. Subtyping of biovars may be performed using a variety of molecular techniques, including pulsed-field electrophoresis, random amplification of polymorphic DNA, amplified fragment length polymorphism, various PCR techniques, and multilocus DNA target sequence typing. MALDI-TOF MS is an important and increasingly available tool in clinical microbiology laboratories because it allows a rapid and accurate identification of bacteria. MALDI Biotyper software is a convenient molecular method that can be used for diagnosing brucellosis. This method requires strict culture conditions and sample preparation to ensure correct identification. Accurate identification of Brucella species using MALDITOF MS has been achieved by constructing a Brucella reference library based on multilocus variable-number tandem repeat analysis (MLVA) data. MLVA is a technique that utilizes multiple repeat nucleic acid sequences that are distributed in different variations across an organism’s genome. Comparing MS-spectra from Brucella species against a custom-made MALDI-TOF MS reference library can be used as a rapid identification method for Brucella species. This method was able to identify 99.3% of 152 isolates tested to the species level, and B. suis biovar 1 and 2 were identified as the correct biovar. This demonstrates that for Brucella, even minimal genomic differences between the biovars translate to specific proteomic differences and clearly identifiable spectra using MALDI-TOF MS. 

Antimicrobial Susceptibility Testing and Therapy Because of the fastidious nature of the brucellae and their intracellular localization, in vitro susceptibility testing is not reliable. In addition, the organisms rarely develop antibiotic resistance, and laboratory safety is a consideration. To prevent relapse of infection, patients with brucellosis undergo prolonged treatment (6 weeks) with antimicrobials that can penetrate macrophages and act in the acidic intracellular environment. Clear guidelines for treatment and duration of therapy remain unclear. Combination therapy with two or more antimicrobials results in fewer relapses than using a single antimicrobial. Doxycycline, rifampin, streptomycin, gentamicin,

481

aminoglycosides, or ceftriaxone are some of the antimicrobials that are used to treat brucellosis. In some instances, surgical drainage may also be required to treat the localized foci of infection and prevent the development of disseminated infection. 

Prevention Successful vaccines against Brucella infection have been developed for livestock. However, the development of human vaccines has met with serious medical contraindications and low efficacy. The prevention of brucellosis in humans depends on elimination of the disease in domestic livestock as well as heating dairy products and related food to reduce disease transmission.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Boschiroli ML, Ouahrani-Betlache S, Foulongne V, et  al.: Type IV secretion and Brucella virulence, Vet Microbiol 90:341–348, 2002. Caroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Centers for Disease Control and Prevention: Brucellosis reference guide: exposures, testing and prevention, 2017. Available at: https://www.cdc.gov/brucellosis/pdf/brucellosi-reference-guide.pdf. Ferreira L, Vega Castaño S, Sánchez-Juanes F, et  al.: Identification of Brucella by MALDI-TOF mass spectrometry. Fast and reliable identification from agar plates and blood cultures, PloS One 5(12):e14235, 2010, https://doi.org/10.1371/journal. pone.0014235. Guzmán-Verri C, González-Barrientos R, Hernández-Mora G, et al.: Brucella ceti and brucellosis in cetaceans, Front Cell Infect Microbiol 2:3, 2012. Jimenez de Bagues MP, Dudal S, Dornand J, Gross A: Cellular bioterrorism: how Brucella corrupts macrophage physiology to promote invasion and proliferation, Clin Immunol 114:227–238, 2004. Karger A, Melzer F, Timke M, et al.: Interlaboratory comparison of intact-cell matrix-assisted laser desorption ionization-time of flight mass spectrometry results for identification and differentiation of Brucella spp., J Clin Microbiol 51:3123–3126, 2013. Leber AL: Aerobic bacteriology. In Clinical microbiology procedures Handbook, ed 4, Washington, DC, 2016, ASM Press. Lista F, Reubsaet FA, De Santis R, et  al.: Reliable identification at the species level of Brucella isolates with MALDI-TOF-MS, BMC Microbiol 11:267, 2011, https://doi.org/10.1186/1471-2180-11267. Pappas G, Akritidis N, Bosilkovski M, Tsianos E: Brucellosis, N Engl J Med 352:2325–2336, 2005. Roy CR: Exploitation of the endoplasmic reticulum by bacterial pathogens, Trends Microbiol 10:418, 2002. Scholz HC, Nockler K, Gollner C, et al.: Brucella inopinata sp. nov., isolated from a breast implant infection, Int J Syst Evol Microbiol 60:801–808, 2010. Smith LD, Ficht TA: Pathogenesis of Brucella, Crit Rev Microbiol 17:209–230, 1990. Yagupsky P: Detection of Brucellae in blood cultures, J Clin Microbiol 37:3437–3742, 1999.

CASE STUDY 35.1 A 67-year-old female from the Middle East has total arthroplasty of the right knee, and 3 years later the same procedure is performed in the left knee. She seeks medical attention because of pain in her left knee. Her knee is aspirated, and a WBC count of 3.6 × 109/L is reported, but no organisms are seen on Gram stain. Coagulase-negative staphylococci are grown from joint fluid cultured in blood culture bottles after 3 days of incubation. A few tiny, poorly staining gram-negative bacilli are present on the direct blood and chocolate agar plates after 5 days of incubation, but not in the blood culture. The bacilli are oxidase and catalase positive. A repeat culture 2 weeks later grows only the

gram-negative bacilli. Surgical debridement with appropriate antimicrobial therapy results in control of the infection.

Questions 1. When fastidious, gram-negative coccobacilli are isolated from a normally sterile site, what is the first step that should be taken in the laboratory? 2. What rapid test can expedite the identification of this fastidious coccobacillus? Describe the limitations associated with this method. 3. How did this patient acquire the infection with this organism? 4. How is the diagnosis confirmed?   

Chapter Review 1. Organisms belonging to the genus Brucella are: a.  Motile b.  Gram-positive bacilli c.  Anaerobic d.  Facultative intracellular 2. Humans become infected with Brucella spp. by all of the following means except: a.  Direct contact with infected animal parts b.  Ingestion of unpasteurized milk c.  Person-to-person contact d.  Inhalation of infected aerosolized particles. 3. Which Brucella species may require CO2 for growth, is urease positive in 2 hours, and is inhibited by thiamine dye? a.  B. abortus b.  B. melitensis c.  B. suis d.  B. canis 4. What titer in the SAT is considered diagnostic and correlates to the clinical findings? a.  1:1 b.  1:4 c.  1:80 d.  1:160 5.  Why should the microbiology laboratory be notified when brucellosis is suspected? a.  It is pathogenic for humans. b.  It is a vaccine-preventable disease. c.  Brucella organisms are category B select agents. d.  It is a zoonotic disease. 6. Which of the following Brucella spp. is not considered a bioterrorism agent? a.  B. abortus b.  B. melitensis c.  B. suis d.  B. canis 7. Resolution of a Brucella infection depends on what? a.  Host’s nutritional and immune status b. Size of inoculum c.  Route of infection d.  All of the above

8. When a specimen other than blood is tested, which agar is recommended? a.  Blood agar b.  Chocolate agar c.  Brucella agar d. MacConkey agar 9. Which test should be used for screening patients in an outbreak of possible brucellosis? a. ELISA b. Laminate flow dipstick c. Immunocapture agglutination d. Coombs test 10.  True or False _____ Brucella spp. are capable of survival in soil for longer than 2 months. _____ Brucella organisms tend to localize in tissue such as placental tissue. _____ B. canis and B. suis are the most virulent species for humans. _____ Isolation of Brucella organisms is a definitive diagnosis of brucellosis. _____ Most strains of B. abortus show equal growth in air and in a candle jar. 11.  Matching: Match each term with the appropriate description. _____ facultative a. infection with intracellular Brucella spp. _____ category B select b. contains additive of agent horse/rabbit serum _____ brucellosis c. rapid test for _____ Brucella agar presumptive Brucella _____ particle infection agglutination d. transmitted primarily _____ unpasteurized by aerosols animal milk e. most common _____ apoptosis means of Brucella transmission f. can exist in extracellular and intracellular environments g. programmed cell death 481.e1

36

Bordetella pertussis, Bordetella parapertussis, and Related Species OBJECTIVES 1. Describe the general characteristics of the Bordetella spp. 2. State the normal habitat and routes of transmission for Bordetella pertussis and Bordetella parapertussis. 3. Describe the three stages of pertussis, including the duration and symptoms. 4. Describe the proper collection and transport of specimens for the detection of bordetellae. 5. Explain the limitations of nucleic acid-based methods for detecting B. pertussis, including assay specificity and sensitivity. 6. Describe the optimal conditions for culturing B. pertussis, including specimens of choice for optimal recovery. 7. Outline the major tests used to identify and differentiate B. pertussis and B. parapertussis. 8. Correlate the patient’s signs, symptoms, and laboratory results to identify the etiologic agent associated with infection.

GENERA AND SPECIES TO BE CONSIDERED Bordetella ansorpii (putative species) Bordetella avium Bordetella bronchialis Bordetella bronchiseptica (Chapter 24) Bordetella flabilis Bordetella hinzii Bordetella holmesii Bordetella muralis Bordetella pertussis Bordetella parapertussis (Chapter 20) Bordetella petrii Bordetella pseudohinzii (considered a proposed species) Bordetella sputigena Bordetella trematum (Chapter 20)

The genus Bordetella includes four primary human pathogens: B. bronchiseptica, B. holmesii, B. pertussis, and B. parapertussis. Three new species have been identified from human respiratory specimens: B. bronchialis, 482

B. flabilis, and B. sputigena. These organisms have been identified in patients with cystic fibrosis (CF). B. bronchiseptica is reviewed in Chapter 24 because it grows on MacConkey agar. Although B. parapertussis and B. holmesii can also grow on MacConkey agar, they are discussed here with B. pertussis because they are associated with human upper respiratory tract infections, with almost identical symptoms, epidemiology, and therapeutic management. Additional Bordetella species may cause rare asymptomatic infections in immunocompromised patients; these include B. hinzii, B. holmesii, B. petrii, B. trematum, and B. ansorpii (described species that remains putative, not validly named). (See the chapter cross-references in the preceding table for information on organisms not discussed in this chapter.)

General Characteristics General features of Bordetella spp. other than B. pertussis and B. parapertussis are summarized in Chapter 24. In contrast to B. bronchiseptica, B. pertussis and B. parapertussis are nonmotile and infect only humans. In the evolutionary process, these exclusive human pathogens have a close genetic relationship. They remain separate species based on their chemotaxonomic differences, pathogenesis, and host range.

Epidemiology and Pathogenesis Before the introduction of the vaccine (and currently in nonimmunized populations), pertussis (whooping cough) periodically appears as an epidemic disease that cycles approximately every 2 to 5 years. Transmission occurs person-to-person through inhalation of respiratory droplets. Humans are the only known reservoir. Pertussis is a highly contagious, acute infection of the upper respiratory tract caused primarily by B. pertussis and less commonly by B. parapertussis. The latter agent generally has a less severe clinical presentation both in duration of symptoms and in the percentage of identified cases. B. holmesii causes a pertussis-like illness, but little is known

CHAPTER 36  Bordetella pertussis, Bordetella parapertussis, and Related Species

about the biology, virulence mechanisms, and pathogenic significance. Additional species of bordetellae, B. bronchiseptica, B. petrii, B. avium, B ansorpii, and B. hinzii are rarely isolated from the respiratory secretions of patients with respiratory illness. Pertussis occurs worldwide, with millions of cases reported annually. Although the incidence has decreased significantly since vaccination became widespread, outbreaks of pertussis occur periodically. B. pertussis infections appear to be endemic in adults and adolescents because of a natural decrease in immune protection from natural infections and vaccine-induced immunity; these infections may serve as the source of the epidemic cycles involving unvaccinated or partially immunized infants and children. CF patients often present with polymicrobial respiratory infections and may serve as a source for periodic outbreaks. The microbiome of CF patients demonstrates a high degree of variability; however, only six genera, including Bordetella spp., are routinely identified from CF specimens.

TABLE   Major Virulence Determinants of Bordetella 36.1  pertussis

Function

Factor/Structure

Adhesion (auto transporters)

Fimbriae (FIM), types 2 and 3: Serotypespecific agglutinins for colonization of respiratory mucosa Filamentous hemagglutinin (FHA): Mediates adhesion to the ciliated upper respiratory tract Pertactin (PRN): Mediates eukaryotic cell binding and is highly immunogenic Tracheal colonization factor Brk Aa

Toxicity

Pertussis toxin (encoded by the ptx gene, an A/B toxin related to cholera toxin): Induces lymphocytosis and suppresses chemotaxis and oxidative responses in neutrophils and macrophages Adenylate cyclase hemolysin: Hemolyzes red cells and activates cyclic adenosine monophosphate, thereby inactivating several types of host immune cells Dermonecrotic toxin (exact role unknown) Tracheal cytotoxin (ciliary dysfunction and damage) Endotoxin (lipopolysaccharide) Type III secretionb Bordetella-secreted protein regulator (BSPR)c

Overcome host defenses

Outer membrane: Inhibits host lysozyme Siderophore production: Prevents host lactoferrin and transferrin from limiting iron

Pathogenesis B. pertussis, the primary pathogen of whooping cough, uses several mechanisms to overcome the immune defenses of healthy individuals. The mechanisms are complex and involve the interplay of several virulence factors (Table 36.1). Some factors help establish infection, others are toxigenic to the host, and still others override specific components of the host’s mucosal defense system. For example, when B. pertussis reaches the host’s respiratory tract, the surface adhesins attach to respiratory ciliated epithelial cells and paralyze the beating cilia by producing a tracheal cytotoxin. The organism produces a major virulence factor, pertussis toxin (PT). PT enters the bloodstream, subsequently binding to specific receptors on host cells. After binding, PT disrupts several host cell functions, such as initiation of host cell translation; the inability of host cells to receive signals from the environment causes a generalized toxicity. The center membrane of B. pertussis blocks access of the host’s lysozyme to the bacterial cell wall via its outer membrane. B. pertussis and B. parapertussis share two nearly identical virulence control systems encoded by the bags (Bordetella virulence gene) and plrSR (persistence in the lower respiratory tract) loci that are responsive to variation in environmental conditions. Because of this very complex system, Bordetella organisms appear to be able to alter phenotypic expression, enhancing transmission, colonization, and survival. 

Spectrum of Disease Several factors influence the clinical manifestations of B. pertussis (Box 36.1). Classic pertussis is usually a disease of children and can be divided into three symptomatic stages: catarrhal, paroxysmal, and convalescent. During the catarrhal stage, symptoms are the same as for a mild cold with a runny nose and mild cough; this stage may last several weeks. Episodes of severe and violent coughing increase in number, marking the beginning of the paroxysmal

483

aPlays

a role in pathogenesis by conferring serum resistance. type of secretion allows Bordetella organisms to transport proteins directly into host cells; it is required for persistent tracheal colonization. cThis transcriptional regulator is involved in mediation of the type III secretion system during iron-starved conditions. bThis

• BOX 36.1 Factors Known to Affect the Clinical

Manifestation of Bordetella pertussis Infection

• • • •

 atient’s age P Previous immunization or infection Presence of passively acquired antibody Antibiotic treatment

stage. As many as 15 to 25 paroxysmal coughing episodes can occur in 24 hours; these are associated with vomiting and with “whooping,” which is the result of air rapidly inspired into the lungs past the swollen glottis. Lymphocytosis occurs, although, typically, the patient has no fever and no signs and symptoms of systemic illness. This stage may last 1 to 6 weeks.

484 PA RT I I I    Bacteriology

In addition to classic pertussis, B. pertussis can cause mild illness and asymptomatic infection, primarily in household contacts and in a number of unvaccinated and previously vaccinated children. Since the 1990s, a shift in the age distribution of pertussis cases to adolescence and adults has been observed in highly vaccinated populations. Adults and adolescents are now recognized as a reservoir for transmitting infection to vulnerable infants. Among these immunized individuals, a prolonged cough may be the only manifestation of pertussis; a scratchy throat, other pharyngeal symptoms, and episodes of sweating commonly occur in adults with pertussis. A number of studies have documented that 13% to 32% of adolescents and adults with an illness involving a cough of 6 days’ duration or longer have serologic or culture evidence of B. pertussis infection. Organisms that may produce pertussis-like symptoms include adenoviruses, respiratory syncytial virus, human parainfluenza viruses, influenza viruses, and Mycoplasma pneumoniae. Other Bordetella species have been associated with infection in immunocompromised patients. B. bronchiseptica, B. holmesii, and B. hinzii produce a pertussis-like respiratory illness. B. trematum has been isolated from individuals working with poultry, and Bordetella ansorpii has been associated with septicemia. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing Confirming the diagnosis of pertussis is challenging. Culture, which is most sensitive early in the illness, has been the traditional diagnostic standard for pertussis and shows nearly 100% specificity but varied sensitivity. Organisms may become undetectable by culture 2 weeks after the start of paroxysms. Nasopharyngeal aspirates (vacuum assisted), nasopharyngeal wash (syringe method), or a nasopharyngeal swab (calcium-alginate or Dacron on a wire handle) are acceptable specimens, because B. pertussis colonizes the ciliated epithelial cells of upper respiratory tract. Calcium-alginate swabs with aluminum shafts are not recommended for polymerase chain reaction (PCR) because they may inhibit the polymerase enzyme in PCR detection. Flocked swabs may be used but have not been validated for PCR or culture of B. pertussis. In addition, cotton swabs may be inhibitory to specimen growth and are not recommended. If possible, two nasopharyngeal swabs should be collected, one from each nostril. Specimens obtained from the throat, sputum, or anterior nose are unacceptable because these sites are not lined with ciliated epithelium. For collection, the swab is bent to conform to the nasal passage and held against the posterior aspect of the nasopharynx. If coughing does not occur, another swab is inserted into the other nostril to initiate the cough. The swab is left in place during the entire cough, removed, and immediately inoculated onto a selective medium at the bedside (Table 36.2) or placed in appropriate transport media. Samples should be collected before the administration of antibiotics. Transport time is critical and should not exceed 48 hours. B. pertussis and B. parapertussis are highly sensitive to metabolites and other toxic substances present in microbiological

TABLE   Examples of Selective Media for Primary 36.2  Isolation of Bordetella pertussis and

Bordetella parapertussis

Agar Media

Description

Bordet-Gengou

Potato infusion agar with glycerol and sheep blood with methicillin or cephalexina (short shelf-life)

Regan-Loweb

Charcoal agar with 10% horse blood and cephalexin (4- to 8-week shelf-life)

Stainer-Scholte

Synthetic agar lacking blood products

aCephalexin

is superior to methicillin and penicillin for inhibiting normal respiratory flora. bRegan-Lowe agar has been found to work best for recovery of B. pertussis from nasopharyngeal swabs.

media, requiring special fluid transport media. Half-strength Regan-Lowe agar enhances recovery when used as a transport and enrichment medium. Cold casein hydrolysate medium and casamino acid broth (available commercially) have proved to be effective transport media, particularly for the preparation of slides for direct fluorescent antibody staining. Dry swabs may be transported in ambient air for PCR testing. 

Direct Detection Methods Because of the limitations associated with culture and serologic diagnostic methods, significant effort has been put into developing nucleic acid amplification methods. Nucleic acidbased diagnostic tests for the direct detection of B. pertussis and B. parapertussis genes by various PCR procedures, including real-time PCR, have replaced DFA in the clinical laboratory. These assays have a diagnostic sensitivity at least comparable (and in most cases superior) to that of culture. Sensitivity of the assays appears to decrease with the duration of the cough, but they may be useful in diagnosis for up to 4 to 6 weeks. A word of caution: Positive results have been obtained with samples containing B. holmesii and B. bronchiseptica (Chapter 24) depending on the sequence targeted in conventional and real-time PCR assays. Most laboratories use transposon insertion sequence IS481 for B. pertussis and IS1001 for B. parapertussis; however, strains of B. holmesii, B. parapertussis, and B. bronchiseptica that carry IS481 have been identified, and thus, careful interpretation of results and correlation with the clinical presentation are required. The addition of transposon insertion sequence IS1002, which is closely related to IS481, and recA significantly improves the specificity of the nucleic acid amplification and species identification. (Table 36.3). Additional PCR assays are available for the detection of the PT promoter, PT, recA, filamentous hemagglutinin and the porin genes. However, because these are single-copy genes and not multicopy insertion sequences, assay sensitivity is reduced. Nasopharyngeal swabs (rayon or Dacron swabs on plastic shafts) and aspirates are the two types of samples primarily used for pertussis PCR; calcium-alginate swabs are unacceptable, as previously mentioned, because they inhibit PCR-based detection. 

CHAPTER 36  Bordetella pertussis, Bordetella parapertussis, and Related Species

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TABLE   Sequences Used to Differentiate Bordetella 36.3  spp.

Organism

IS481

B. pertussis

+

B. parapertussis B. bronchiseptica

+

B. holmesii

+

IS1001 IS1002

+

+

+

+

RecA

+

From Martini H, Detemmerman, Soetens O, et al. Improving specificity of Bordetella pertussis detection using a four target real-time PCR. PLoS One. 2017;12:e0175587.

• Fig. 36.2  Typical Gram stain appearance of Bordetella pertussis.

spp. characteristics are presented in Table 36.4. Whole-cell agglutination reactions in specific antiserum can be used for species identification. 16S rRNA gene sequencing or ribotyping and matrix-assisted laser desorption ionization timeof-flight mass spectrometry (MALDI-TOF MS) are often used to identify these organisms. 

Serodiagnosis

• Fig. 36.1  Growth of Bordetella pertussis on Regan-Lowe agar.

Cultivation Cultivation is generally 100% specific; however, it has been replaced with PCR in most laboratories for routine diagnosis. Plates are incubated at 35°C in a humidified atmosphere without elevated carbon dioxide for up to 12 days. Most isolates are detected in 3 to 7 days; B. parapertussis appears in 2 to 3 days. Colony morphology is not distinct for the identification of other Bordetella spp. Regan-Lowe agar, Bordet-Gengou agar, and Stainer-Scholte synthetic medium are suitable culture media. Regan-Lowe agar contains beef extract, starch, casein digest, and charcoal supplemented with horse blood. Bordet-Gengou agar is a potato fusion base containing glycerol and either sheep or horse blood. Most media contain cephalexin as an additive for suppression of contaminating organisms. Young colonies of B. pertussis and B. parapertussis are small and shiny, resembling mercury drops; colonies become whitish gray with age (Fig. 36.1). Sensitivity of culture approaches 100% in the best of hands and depends on the stage of illness at the time of specimen collection, the technique used for specimen collection, specimen adequacy and transport, and culture conditions. 

Approach to Identification A Gram stain of the organism reveals minute, faintly staining coccobacilli singly or in pairs (Fig. 36.2). Use of a 2-minute safranin “O” counterstain or a 0.2% aqueous basic fuchsin counterstain enhances their visibility. Bordetella

Although several serologic tests are available for the diagnosis of pertussis, including agglutination and enzyme immunoassay, the enzyme-linked immunosorbent assay (ELISA) or bead-based assays are recommended for serologic diagnosis at this time. The current most reliable serologic test available for diagnosis is an anti-PT (antibody to pertussis toxin) ELISA that has been used with acute and paired convalescent sera successfully in older children, adolescents, and adults. A titer greater than 100 to 125 IU/mL has been reported as a reliable indicator of exposure of patients to PT-producing bacteria. Serological assays cannot distinguish infection versus vaccine-induced immunity for B. pertussis and is unable to detect infection with B. parapertussis. 

Antimicrobial Susceptibility Testing and Therapy Laboratories currently do not perform routine susceptibility testing of B. pertussis and B. parapertussis because the organisms remain susceptible to some penicillins or the macrolides (clarithromycin, azithromycin), ketolides, quinolones, and other antibiotics, such as tetracyclines, chloramphenicol, and trimethoprim-sulfamethoxazole. Three erythromycin-resistant isolates of B. pertussis have been discovered; therefore, continued surveillance of B. pertussis is advised. Both B. pertussis and B. parapertussis are resistant to most oral cephalosporins and B. bronchiseptica is resistant to many penicillins, cephalosporins, and trimethoprimsulfamethoxazole. Routine antimicrobial susceptibility testing for Bordetella pertussis and B. parapertussis is not recommended due to the lack of standardization; however, testing for other Bordetella isolates should be reported based on the laboratories’ procedures for other fastidious gram-negative bacilli. 

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TABLE 36.4    Characteristics That Differentiate the Major Isolates of Bordetella spp.

Characteristic

B. hinzii

B. holmesii

B. pertussis

B. parapertussis

B. bronchiseptica

Oxidase

+

(+)

＋

−

＋

Motility

+

−

−

−

＋

Nitrate

−

−

−

−

＋

Urease

−

−

−

＋ (24 h)

＋ (4 h)

Regan-Lowe agar

ND

2–3 days

3–4 days

2–3 days

1–2 days

Blood agar

−

+

−

＋

＋

MacConkey agar

+

(+)

−

−

＋

Growth

ND, not determined; (+), 10 colonies. and DNase positive. cN. cinerea may be differentiated from N. flavescens by a positive reaction with the amylosucrase test. dSome strains of N. cinerea may appear glucose-positive in some rapid systems and be mistaken for N. gonorrhoeae. However, N. cinerea grows on nutrient agar at 35°C and reduces nitrite, unlike N. gonorrhoeae. eN. elongata subsp. glycolytica is positive for acid production from glucose. fN. subflava biovar flava is positive for acid production from fructose and N. subflava biovar perflava is positive for acid production from sucrose and fructose. gOnly 2 of 10 strains were tested. hKingella denitrificans may grow on modified Thayer-Martin agar and be mistaken for N. gonorrhoeae on microscopic examination. However, K. denitrificans can reduce nitrate and is catalase negative, unlike N. gonorrhoeae. iNeisseria subflava produces a yellow pigment on Loeffler agar; N. sicca does not. +, >90% of strains positive; (+), >90% of strains positive but reaction may be delayed (i.e., 2–7 days); −, >90% of strains negative; V, variable. Data compiled from Centers for Disease and Control; Carroll KC, Pfaller MA, Landry ML, et al. Manual of Clinical Microbiology. 12th ed. Washington, DC: ASM; 2019; and Weyant RS, Moss CW, Weaver RE, et al, editors. Identification of Unusual Pathogenic Gram-negative Aerobic and Facultatively Anaerobic Bacteria. 2nd ed. Baltimore: Williams & Wilkins; 1996. bButyrate

CHAPTER 39  Neisseria and Moraxella catarrhalis

Fructose

Nitrate Reduction

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amplified nucleic acid–based testing or by matrix-assisted laser desorption ionization time-of-flight mass spectroscopy (MALDI-TOF MS). Isolates from normally sterile body fluids should also be completely identified. However, isolates from genital sites in adults at risk of STI can be identified presumptively (i.e., oxidase-positive, gram-negative diplococci that grow on gonococcal selective agar). Likewise, an oxidase-positive, gram-negative diplococcus that hydrolyzes tributyrin using the enzyme butyrate esterase from an eye or ear culture can be identified as M. catarrhalis (Fig. 12.7). 

Matrix-Assisted Laser Desorption Ionization Timeof-Flight Mass Spectrometry MALDI-TOF MS is an automated mass spectrometry system that can identify organisms based on their protein structure (Chapter 7). Two major mass spectrometry systems are available for and have demonstrated the ability to identify Neisseria and Moraxella species. There have been reports of misidentification of commensal Neisseria species as N. meningitidis. Only organisms that are FDA-cleared for the specific MALDI-TOF MS system may be reported; however, other species may be validated by the individual laboratory. 

Comments About Specific Organisms Determination of carbohydrate utilization patterns historically has been performed in cysteine trypticase soy agar (CTA) with 1% glucose (dextrose), maltose, lactose, and sucrose (Evolve Procedure 39.1). This medium is no longer widely used because it does not work well for oxidative Neisseria spp., specifically N. gonorrhoeae and N. meningitidis. Carbohydrate utilization patterns are currently determined by inoculating an extremely heavy suspension of the organism to be tested in a small volume of buffered, lowpeptone substrate with the appropriate carbohydrate. These methods do not require subculture or growth, and results are available in approximately 4 hours. Other rapid identification kits that are commercially available include the Neisseria Preformed Enzyme Test (PET) (BioConnections, Knypersley, United Kingdom), the RapID NH (ThermoFisher Scientific, Waltham, MA), or the API NH (bioMérieux, Marcy-l’Étoile, France). Each of these different kits has varying levels of sensitivity for correctly identifying N. gonorrhoeae, especially if the strain is proline iminopeptidase (Pip) negative. Saprophytic Neisseria spp. are not routinely identified in the clinical laboratory. Neisseria cinerea may be misidentified as N. gonorrhoeae if the isolate produces a weak positive glucose (dextrose) reaction. However, it grows on nutrient agar at 35°C, whereas the gonococcus does not. Moreover, N. cinerea is inhibited by colistin, whereas N. gonorrhoeae is not. M. catarrhalis can be differentiated from the gonococci and meningococci based on its growth on blood agar at 22°C and on nutrient agar at 35°C, the reduction of nitrate to nitrite, its inability to use carbohydrates, and its production of DNase. Severe M. catarrhalis is the only member of this group of organisms that hydrolyzes DNA.

Chromogenic substrate enzyme tests for beta-galactosidase, gamma-glutamyl aminopeptidase, and prolyl-hydroxyl prolyl aminopeptidase are available for the differentiation of N. gonorrhoeae, N. meningitidis, N. lactamica, and M. catarrhalis. M. catarrhalis lacks all three of these enzymes. The presence of prolyl-hydroxyl prolyl aminopeptidase alone identifies an organism as N. gonorrhoeae. The presence of beta-galactosidase and gamma-glutamyl aminopeptidase indicates N. meningitidis. Two commercial chromogenic substrate kits are the Gonocheck II (EY Laboratories, San Mateo, CA) and BactiCard Neisseria (Remel Laboratories, Lenexa, KS). A limitation of these methods is misidentification of various nonpathogenic strains of Neisseria spp. In addition, isolate colonies on selective media should be used to prevent misidentification of contaminants as a Neisseria spp. Modified chromogenic substrate kits, such as the BactiCard Neisseria, can be used to identify and speciate Neisseria and Haemophilus organisms from selective and nonselective media. These modified tests use a combination of enzyme substrate tests and additional biochemical tests. N. lactamica may grow on selective media and may be confused with N. meningitidis. The ONPG test (Procedure 12.32) is used to determine an organism’s ability to produce beta-galactosidase, which is an indicator of lactose utilization. N. lactamica is ONPG positive, and N. meningitidis is ONPG negative. The eugonic fermenter N. animaloris propagates well on routine laboratory media and ferments glucose (dextrose); this distinguishes it from dysgonic fermenters that grow poorly on blood and chocolate agars (Chapter 27). N. animaloris ferments no carbohydrates other than glucose and is indole negative and arginine dihydrolase positive. 

Immunoserologic Identification A single particle agglutination method is available for immunoserologic identification of N. gonorrhoeae in the United States. The GonoGen II test (Becton Dickinson, Sparks, MD) can be performed from colonies growing on primary plates in which isolates are typed with specific monoclonal antibodies. The GonoGen II is a colorimetric test that uses antibodies adsorbed to metal sol particles. False-positives have occurred with N. lactamica and N. meningitidis, as well as false-negatives with other isolates. Extended protein extraction methods can be used to improve the sensitivity and specificity of the test. 

Serotyping Twelve different serogroups are distinguishable for N. meningitidis. Antisera are commercially available for identifying N. meningitidis serogroups A, B, C, H, I, K, L, W135, X, Y, and Z. Some manufacturers produce polyvalent antisera containing combinations of serogroups for identification. Serologic identification is usually performed by slide agglutination. A, B, C, W135, and Y are the serotypes that most commonly cause systemic disease in the United States. Serotyping is an important tool that reference or public health laboratories should possess to help aid in the identification

CHAPTER 39  Neisseria and Moraxella catarrhalis

of clusters of cases that may be related during outbreak investigations. Nucleic acid amplification methods referred to as genogrouping are more reliable for the identification of commensal strains of Neisseria species. Whole genome sequencing is expected to replace serotyping. 

Antimicrobial Susceptibility Testing and Therapy Although beta-lactamase production is common among M. catarrhalis isolates, many beta-lactam antimicrobials maintain activity against these isolates. Because several other agents are also effective, susceptibility testing to guide therapy is not routinely required. Standard methods have been established for performing in  vitro susceptibility testing with N. gonorrhoeae and N. meningitidis (Chapter 11). The Clinical and Laboratory Standards Institute (CLSI) recommends the use of agar dilution for minimum inhibitory concentration (MIC) measurements and GC agar containing 1% growth supplement for N. gonorrhoeae disk diffusion methods. In addition, various agents can be considered for testing and therapeutic use. Historically, fluoroquinolones were widely used to treat gonorrhea; however, resistance to these agents has emerged (i.e., fluoroquinolone-resistant N. gonorrhoeae [FRNG]), and they are no longer recommended for treatment of gonococcal infections. The Gonococcal Isolate Surveillance Project (GISP) is a national collaborative surveillance project in the United States. The program was established in 1986 to analyze and monitor antimicrobial resistance trends among N. gonorrhoeae. Participating regional labs test isolates for resistance to antimicrobials that were commonly prescribed to treat gonococcal urethritis, including azithromycin, cefixime, ceftriaxone, ciprofloxacin, gentamicin, penicillin, and tetracycline. Because of increasing resistance of fluoroquinolones reported in 2007, the CDC stopped recommending these agents (e.g., ciprofloxacin, levofloxacin) for treatment of gonococcal urethritis. This left cephalosporins as the only treatment option. In 2010, the CDC recommended dual therapy of a cephalosporin with azithromycin or tetracycline. Between 2006 and 2011, MICs to the oral cephalosporin cefixime have increased, and more treatment failures were reported throughout the world. As a result, in 2015 the CDC issued an update to their treatment guidelines that suggests a single-dose intramuscular injection of 250 mg of ceftriaxone taken along with either a 1-g oral dose of azithromycin or 100 mg oral doxycycline twice daily for 7 days. However, because many patients may be co-infected with C. trachomatis, dual therapy with azithromycin is recommended for eliminating both infections, as C. trachomatis is susceptible to a 1-g oral dose of azithromycin. Patients with severe penicillin

503

immunoglobulin E (IgE)-mediated allergies may warrant treatment with an alternative regimen; however, there are limited data on their utility. Alternative therapy includes a single-dose of 320 mg oral gemifloxacin with 2 g of oral azithromycin, or a single-dose intramuscular injection with 240 mg gentamicin and 2 g oral dose of azithromycin. Although beta-lactamase production in N. meningitidis is rare, decreased susceptibility to penicillin, mediated by altered penicillin-binding proteins, is emerging. Resistance to sulfonamides is also quite common. The CLSI recommends that susceptibility testing be performed by disk diffusion on Mueller-Hinton agar or using cation-adjusted Mueller-Hinton broth in microdilution. All testing should occur within a biologic safety cabinet to minimize laboratory-acquired infections. 

Prevention The CDC recommends vaccination against N. meningitidis to prevent meningitis in adolescents and young adults. Currently, 11- to 12-year-olds should be vaccinated with a quadrivalent meningococcal conjugate vaccine. The two quadrivalent vaccines in the United States protect against serogroups A, C, W, and Y. The CDC recommends that a booster vaccine be given around age 16. If the initial vaccine is given to an adolescent between 13 and 15 years of age, the booster should be administered at age 18, when the patient enters a period of increased risk. Children with weakened immune systems (human immunodeficiency virus [HIV] positive, asplenia, complement deficiency, or some immunosuppressive drugs) should be given the quadrivalent vaccines earlier; depending on the age and disease, this may start as early as 8 weeks old and is given in multiple doses. There is also a vaccine available for serogroup B N. meningitidis. Historically, this vaccine was recommended only for people 10 years of age and older who had been identified as being at increased risk for exposure to serogroup B (e.g., community outbreak). However, the Advisory Committee on Immunization Practices has revised their position and allows individual clinical decision-making regarding the use of the serogroup B meningococcal vaccine. This is a multistep vaccination, and for each dose, the same manufacturer should be used because they are not interchangeable. Ophthalmia neonatorum occurs when N. gonorrhoeae from an infected mother is transmitted to the newborn during vaginal delivery. Previously, this was a leading cause of neonatal blindness. Prophylactic administration of antimicrobials after birth has dramatically reduced the development of gonococcal conjunctivitis. The CDC recommends a single dose of 0.5% erythromycin ophthalmic ointment into each eye at birth. Silver nitrate is no longer manufactured in the United States, and povidone-iodine has not been adequately studied and is therefore not recommended.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

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Bibliography Abadi FJ, Yakubu DE, Pennington TH: Antimicrobial susceptibility of penicillin-sensitive and penicillin-resistant meningococci, J Antimicrob Chemother 35: 687-190, 1995. Bennett DE, Mulhall RM, Cafferkey MT: PCR-based assay for detection of Neisseria meningitidis capsular serogroups 29E, X, and Z, J Clin Microbiol 42:1764–1765, 2004. Blondeau JM, Ashton FE, Isaacson M, et  al.: Neisseria meningitidis with decreased susceptibility to penicillin in Saskatchewan, Canada, J Clin Microbiol 33:1784–1786, 1995. Bratcher HB, Harrison OB, Maiden MCJ: Genome sequencing and interrogation of genome databases: a guide to Neisseria meningitidis genomics, Methods Mol Biol 1969:51–82, 2019. Carroll KC, Pfaller MA, Landry ML, et  al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Centers for Disease Control and Prevention (CDC): Sexually transmitted disease surveillance 2017: gonorrhea, 2018. Available at: http s://www.cdc.gov/std/stats17/gonorrhea.htm. Centers for Disease Control and Prevention (CDC): Sexually transmitted disease treatment guidelines, 2015. Available at: https://www .cdc.gov/std/tg2015/gonorrhea.htm. Clinical and Laboratory Standards Institute (CLSI): 29th edition, Performance standards for antimicrobial susceptibility testing, M100, Wayne, PA, 2018, CLSI. Committee on Infectious Diseases: 2006 red book: report of the Committee on Infectious Diseases, ed 27, Elk Grove Village, IL, 2006, American Academy of Pediatrics. Dolan Thomas J, Hatcher CP, Satterfield DA, et  al.: sodC-based real-time PCR for detection of Neisseria meningitidis, PloS One 6:e19361, 2011. Guldemir D, Turan M, Bakkaloglu Z, et  al.: Optimization of realtime multiplex polymerase chain reaction for the diagnosis of acute bacterial meningitis and Neisseria meningitidis serogrouping, Mikrobiyol Bul 52:221–232, 2018. Heiddal S, Sverrisson JT, Yngvason FE, et al.: Native valve endocarditis due to Neisseria sicca: case report and review, Clin Infect Dis 16:667, 1993.

Hong E, Bakhalek Y, Taha MK: Identification of Neisseria meningitidis by MALDI-TOF MS may not be reliable, Clin Microbiol Infect 25:717–722, 2019. Kam KM, Wong PW, Cheung MM, Ho NK: Detection of quinolone-resistant Neisseria gonorrhoeae, J Clin Microbiol 34:1462– 1464, 1996. Meyer GA, Shope TR, Waecker Jr NJ, Lanningham FH: Moraxella (Branhamella) catarrhalis bacteremia in children, Clin Pediatr (Phila) 34:146–150, 1995. Peterson ME, Li Y, Shanks H, et  al.: Serogroup-specific meningococcal carriage by age group: a systemic review and meta-analysis, BMJ Open 9(4):e024343, 2019. Rosenstein NE, Perkins BA, Stephens DS, et al.: Meningococcal disease, N Engl J Med 344:1378–1388, 2001. Satterwhite CL, Torrone E, Meites E, et  al.: Sexually transmitted infections among US women and men: prevalence and incidence estimates, 2008, Sex Transm Dis 40:187–193, 2013. Tanaka M, Matsumoto T, Kobayashi I, Uchino U, Kumazawa J: Emergence of in vitro resistance to fluoroquinolones in Neisseria gonorrhoeae isolated in Japan, Antimicrob Agents Chemother 39:2367–2370, 1995. Unemo M, Golparian D, Eyre DW: Antimicrobial resistance in Neisseria gonorrhoeae and treatment of gonorrhea, Methods Mol Biol 1997:37–58, 2019. Vandamme P, Holmes B, Bercovier H, Coenye T: Classification of Centers for Disease Control group eugonic fermenter (EF)-4a and EF-4b as Neisseria animaloris sp. nov. and Neisseria zoodegmatis sp. nov., respectively, Int J Syst Evol Microbiol 56:1801–1805, 2006. Verghese A, Berk SL: Moraxella (Branhamella) catarrhalis, Infect Dis Clin North Am 5:523–538, 1991. Weyant RS, Moss CW, Weaver RE, et  al.: Identification of unusual pathogenic gram-negative aerobic and facultatively anaerobic bacteria, ed 2, Baltimore, 1996, Williams & Wilkins. Woods CR, Smith AL, Wasilauskas BL, Campos J, Givner LB: Invasive disease caused by Neisseria meningitidis relatively resistant to penicillin in North Carolina, J Infect Dis 170:453–456, 1994.

CASE STUDY 39.1 An elderly male with a history of chronic obstructive pulmonary disease (COPD) after 20 years of heavy smoking presents to the emergency department with shortness of breath; severe cough; and profuse, yellow sputum production. Crackles and wheezing can be heard on chest examination. A sputum culture is positive for many intracellular gram-positive, lancet-shaped diplococci. Gram-negative diplococci were observed in the smear. The patient was placed on amoxicillin/clavulanic acid, and other supportive measures.

Questions 1. List the pathogenic agents most often found to be involved in acute infections of patients with COPD. 2. List the tests required to identify Moraxella catarrhalis definitively and rapidly. 3. The culture from this patient grew Streptococcus pneumoniae and M. catarrhalis. What antibiotics could be used to treat both infections? Is susceptibility testing necessary with this isolate or can empiric therapy be used?   

PROCEDURE 39.1

Carbohydrate Utilization Method—Cysteine Trypticase Soy Agar Purpose The carbohydrate utilization method is the traditional method used to identify Neisseria spp. based on carbohydrate utilization in cysteine trypticase soy agar (CTA) with the addition of 1% of a specific carbohydrate (glucose [dextrose], maltose, lactose, or sucrose) and phenol red as a pH indicator. 

Method 1. Using an inoculating loop, prepare a heavy inoculum in saline of a pure isolate from a subculture no more than 24 hours old obtained on nonselective media. 2. Using an inoculating needle, transfer the sample from the prepared tube and stab the top half of the CTA carbohydrate tube. Inoculate one of each of the CTA media: glucose, maltose, lactose, and sucrose. 3. Tightly cap the tubes and incubate at 35°C–37°C in ambient air. Incubate an uninoculated control tube simultaneously. 4. After incubation, examine the tubes within 24–72 hours for a yellow color at the top of the media; this indicates acid production and a positive result for the carbohydrate utilization test. 

Expected Results and Quality Control Positive: Yellow color at the top of the tube only, indicating carbohydrate use. Negative: No color change compared with the control, an uninoculated tube.

Organism Neisseria cinerea Neisseria elongata Neisseria flavescens Neisseria lactamica Neisseria gonorrhoeae Neisseria meningitidis Neisseria mucosa N. subflava biovar flava N. subflava biovar perflava Neisseria subflava biovar subflava Moraxella catarrhalis

Glucose Neg Neg Neg Pos Pos Pos Pos Pos Pos Pos

Maltose Neg Neg Neg Pos Neg Pos Pos Pos Pos Pos

Lactose Neg Neg Neg Pos Neg Neg Neg Neg Neg Neg

Sucrose Neg Neg Neg Neg Neg Neg Pos Neg Pos Pos

Fructose Neg Neg Neg Neg Neg Neg Pos Pos Pos Pos

Neg

Neg

Neg

Neg

Neg

Limitations Avoid incubation in carbon dioxide (CO2), which may alter the pH of the media, resulting in a color change to yellow throughout the entire tube and a false-positive reaction. A yellow color change throughout the tube also may indicate the presence of contaminating organisms. Caution should be used in interpreting the test result, and other confirmatory tests should be performed.   

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Chapter Review 1. Which organism is not a normal inhabitant of the respiratory tract? a.  M. catarrhalis b. N. meningitidis c.  N. mucosa d. N. gonorrhoeae 2. For which of the following are quadrivalent vaccines available? a.  B. pertussis b. M. catarrhalis c.  N. meningitidis d. N. gonorrhoeae 3. Which of the following is a leading sexually transmitted infection? a.  N. gonorrhoeae b. N. lactamica c.  N. sicca d. N. mucosa 4. Identification of N. gonorrhoeae can be accomplished by all of the following except: a. Particle agglutination b. Gram stain c. MALDI-TOF MS d. Amplified assays 5. Which of the following media is not acceptable for primary growth of N. gonorrhoeae? a. Chocolate b. Thayer-Martin c. Blood d. Martin-Lewis 6.  M. catarrhalis grows best under which conditions? a. Anaerobic atmosphere b. 42°C c. Increased CO2 d. 25°C 7.  N. gonorrhoeae can be differentiated from N. meningitidis by: a. Dextrose fermentation b. Maltose fermentation c. Lactose fermentation d. Nitrate reduction 8. All of the following statements about N. gonorrhoeae are true except: a.  Particle agglutination methods are available for identification. b. The sample should be incubated at 36°C for 72 hours in a CO2-enriched environment. c. The sample may be refrigerated if plating is delayed. d. The organism may appear to have up to five different colony types. 9. An organism grows on blood agar at room temperature and nutrient agar at 37°C. The organism has the ability to use carbohydrates and is nitrate positive, DNase positive, and ONPG negative. This organism is: a.  N. gonorrhoeae b. N. meningitidis

c.  N. lactamica d. M. catarrhalis 10. All of the following are advantages of using amplified testing over culturing except: a. It is more sensitive. b. It is suitable for large screening programs. c. It is admissible in medicolegal cases. d. It allows testing for C. trachomatis at the same time. 11.  True or False _____ Men who have sex with men (MSM) are the primary reservoirs for the dissemination of N. gonorrhoeae in the human population. _____ CTA sugars do not require subculture for growth and can be read in 4 hours. _____ Infection with M. catarrhalis may lead to severe disseminated disease. _____ Genital isolates of Neisseria spp. in adults with high-risk behavior must be completely identified. _____ The presence of prolyl-hydroxyl prolyl aminopeptidase alone identifies an organism as N. meningitidis. 12.  Matching: Match the correct term with the appropriate description. _____ N. gonorrhoeae _____ N. meningitidis _____ Thayer-Martin _____ Saprophytic _____ JEMBEC _____ A, B, C, Y, and W135 _____ A, C, Y, and W135

a. nonpathogenic b. CO2-generating culture system c. N. meningitidis groups seen in the United States d. Antigens in a N. meningitidis vaccine e. Leading cause of STIs f. Selective agar for culturing Neisseria spp. g. Cause of fatal bacterial meningitis

13.  Short Answer A synovial fluid sample is submitted for culture during the night shift at approximately 2 a.m. The laboratory is short-staffed and busy testing and supplying blood products after a local nightclub shooting. The synovial fluid sample is placed in the refrigerator until sufficient time is available to plate the sample to the appropriate media. At approximately 6 a.m., the day shift personnel find the specimen in the refrigerator. If the physician suspects an isolate of the organisms discussed in this chapter, how should the laboratory staff proceed? Should the specimen be processed, or should a repeat sample be submitted? Explain your reasoning.

S E C T I ON 1 3    Anaerobic Bacteriology

40

Overview and General Laboratory Considerations OBJECTIVES

>

This chapter provides an overview of the methods used to identify anaerobic microorganisms. The detailed technical procedures discussed are designed for use in conjunction with specifics provided in Chapter 41 to develop a clear understanding of the full process, from specimen collection to identification. However, readers should consider the following general objectives for the information and methods provided. 1. State the specific diagnostic purpose for the test methodology. 2. Briefly describe the test principle associated with the test methodology. 3. Outline limitations and describe a process for troubleshooting or reporting results if a test result is equivocal or indistinguishable. 4. State the appropriate quality-control organisms and results used with each testing procedure. 5. Define and differentiate obligate (strict), moderate, facultative, and aerotolerant anaerobes. 6. List suitable specimens for isolation of anaerobic bacteria and characteristics of these specimens that might suggest the presence of an anaerobic infection. 7. Explain the proper techniques for collecting, transporting, and processing clinical specimens for anaerobic bacteriology. 8. Explain the use of antigen detection methodologies in the diagnosis of anaerobic infections. 9. List the media used for cultivation of anaerobic bacteria. 10. Describe the appropriate incubation conditions for cultivation of anaerobic bacteria. 11. Describe the procedures for the identification of and antibiotic susceptibility testing for anaerobic bacteria.

General Characteristics The organisms described in this chapter and in Chapter 41 are common etiological agents of a variety of clinical conditions. The organisms included in this chapter are predominant in the human microbiome and are often opportunistic pathogens. These organisms do not grow in the presence of oxygen (O2); they are obligate or strict anaerobes (0% O2).

Obligate anaerobes are killed upon brief exposure (less than a few minutes) to atmospheric oxygen. Obligate anaerobes include Prevotella spp., Fusobacterium spp., and Bacteroides spp., which are included in these chapters. These chapters also include some aerotolerant organisms (5% O2), such as Actinomyces spp., Bifidobacterium spp., and Clostridium spp., which are capable of growth in the presence of either reduced or atmospheric oxygen (microaerobic) but grow best under anaerobic conditions. Finally, facultative anaerobes do not require atmospheric oxygen but are capable of growth in oxygen and anaerobic environments. Anaerobic organisms lack superoxide dismutase and catalase, the enzymes required to break down reactive oxygen species produced during respiration or aerobic metabolism. In addition, oxygen has a high affinity for organic compounds containing nitrogen, hydrogen, carbon, and sulfur, which interfere with normal biologic activity. Because they are unable to protect themselves against the action of oxygen, anaerobes require an environment free of oxygen to survive and grow. 

Specimen Collection and Transport The importance of proper collection and transport of specimens for anaerobic culture cannot be overemphasized. Because indigenous anaerobes are often present in large numbers as normal microbiota on mucosal surfaces, even minimal contamination of a specimen can produce misleading results. Box 40.1 shows the specimens acceptable for anaerobic culture; Box 40.2 presents specimens that are likely to be contaminated and therefore are unacceptable for anaerobic culture. In general, material for anaerobic culture is best collected by tissue biopsy or by aspiration using a needle and syringe to prevent contamination with normal microbiota. After collection, the air must be expelled from the device to prevent the reduction or loss of viable anaerobes in the sample. Because of the potential for contamination with normal microbiota, swabs are generally not recommended for the collection of anaerobes. 505

506 PA RT I I I    Bacteriology

• BOX 40.1 Clinical Specimens Suitable for

Anaerobic Culture

Bile Biopsy of endometrial tissue obtained with an endometrial suction curette (Pipelle; Unimar, Wilton, CT) Blood Bone marrow Bronchial washings obtained with a double-lumen plugged catheter Cerebrospinal fluid Culdocentesis aspirate Decubitus ulcer (if obtained from base of the lesion after thorough debridement of the ulcer’s surface) Fluid from normally sterile site (e.g., joint) Material aspirated from abscesses (the best specimens are from loculated or walled-off lesions) Percutaneous (direct) lung aspirate or biopsy Peritoneal (ascitic) fluid Sulfur granules from a draining fistula Suprapubic bladder aspirate Thoracentesis (pleural) fluid Tissue obtained at biopsy or autopsy Transtracheal aspirate Uterine contents (if collected using a protected swab)

• BOX 40.2 Clinical Specimens Unsuitable for

Anaerobic Culture

Bronchial washing or brush (unless collected with a doublelumen plugged catheter) Coughed (expectorated) sputum Feces (except for Clostridioides difficile) Gastric or small-bowel contents (except in blind loop syndrome) Ileostomy or colostomy drainage Nasopharyngeal swab Rectal swab Secretions obtained by nasotracheal or orotracheal suction Swab of superficial (open) skin lesion Throat swab Urethral swab Vaginal or cervical swab Voided or catheterized urine

However, as previously described in Chapter 5, flocked swabs commercially available from numerous manufacturers have an improved design that enhances the recovery of both aerobes and anaerobes but are still prone to contamination and can be used when no other specimen type may be collected. A crucial factor in obtaining valid results with anaerobic cultures is the transport of the specimen; the lethal effect of atmospheric oxygen must be nullified until the specimen can be processed in the laboratory. Recapping a syringe and transporting the needle and syringe to the laboratory

is no longer acceptable because of safety concerns involving needle stick injuries. Even aspirates must be injected into an oxygen-free transport tube or vial. A large variety of transport devices that contain prereduced anaerobic media for the preservation of microorganisms are available commercially. 

Macroscopic Examination of Specimens Upon receipt in the laboratory, specimens should be inspected for characteristics that strongly indicate the presence of anaerobes: (1) foul odor; (2) sulfur granules (associated with Actinomyces spp., Propionibacterium spp., or Eubacterium sp.); and (3) brick red fluorescence under long wavelength ultraviolet (UV) light (associated with pigmented Prevotella or Porphyromonas spp.). 

Direct Detection Methods Gram Staining The Gram stain is an important rapid tool for anaerobic bacteriology. Gram stain morphology from direct specimens should be carefully noted if an anaerobic infection is suspected, because organisms may no longer be viable and additional testing, such as anaerobic cultures, may demonstrate no growth. Not only does a properly performed Gram stain reveal the types and relative numbers of microorganisms and host cells present, it also serves as a quality control measure for the adequacy of anaerobic techniques. The absence of leukocytes does not rule out the presence of a serious anaerobic infection, however, because certain organisms, such as clostridia, produce necrotizing toxins that destroy white blood cells. A positive Gram stain with a negative culture may indicate (1) poor transport methods, (2) excessive exposure to air during specimen processing, (3) failure of the system (jar, pouch, or chamber) to achieve an anaerobic atmosphere, (4) inadequate types of media or old media, or (5) killing of microorganisms by antimicrobial therapy. Standard Gram stain procedures and reagents are used, except that the safranin counterstain is left on for 3 to 5 minutes. Gram-negative anaerobes often stain poorly with safranin, resulting in failure to visualize pathogenic organisms. As an alternative, 0.5% aqueous basic fuchsin can be used as the counterstain to improve identification of gramnegative anaerobes. In addition, some gram-positive anaerobes (e.g., Clostridium spp.) stain pink. Enhanced Gram stain reagents are available that contain different concentrations in the reagents, in addition to a Gram enhancer, which is applied after decolorization to suppress the red color in the background, aiding the differentiation of gram-negative anaerobes. Table 40.1 presents the cellular morphology seen with Gram staining of common anaerobes. 

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507

TABLE   Minimal Criteria Grouping Gram Stain Morphology, Aerotolerance, and Clinical Significance of Anaerobic 40.1  Bacteria

Organism

Gram Staina

Clinical Significance

Actinobaculum spp.

Gram-positive slightly curved rods; some species may branch

Urinary tract infections

Actinomyces spp.

Gram-positive, branching, beaded or banded, thin, filamentous rods

Actinomycosis; orocervicofacial, thoracic, and abdominopelvic forms

Actinotignum spp.

Gram-positive straight or slightly curved rods

Urinary tract infections

Alloscardovia spp.

Gram-positive short irregular shaped rods

Urinary tract infections

Alistipes spp.

Gram-negative rods

Appendicitis, intraabdominal fluids, abscesses, and urine

Anaerococcus spp.

Gram-positive cocci arranged in short chains or tetrads

Wound infections

Anaeroglobus sp.

Gram-negative cocci with cells approximately 0.5–1.1 μm in diameter

Postoperative wounds

Atopobium spp.

Elongated gram-positive cocci or coccobacilli; occur singly, in pairs, or in short chains

Bacterial vaginosis

Bacteroides spp.

Gram-negative, straight rods with rounded ends; occur singly or in pairs; cells may be described as resembling a safety pin (Fig. 41.4)

Bacteremia, ulcers, abscesses, bronchial secretions, bone, intraabdominal infections; body fluids

Bifidobacterium spp.

Gram-positive diphtheroid; coccoid or thin, pointed shape; or larger, highly irregular, curved rods with branching; rods terminate in clubs or thick, bifurcated (forked) ends (“dog bones”)

Predominantly bacterium but may be isolated from a variety of sources

Bilophila wadsworthia

Gram-negative, pale-staining, delicate rods

Intraabdominal infections, abscesses, and bacteremia

Bulleidia spp.

Gram-positive short, straight, or slightly curved; singly or in pairs

Periodontitis and abscesses

Catabacter spp.

Gram-positive coccobacilli or short rods

Bacteremia

Clostridioides difficile

Gram-positive straight rods; may produce chains of up to six cells aligned end to end; spores oval and subterminal

Antibiotic associated disease, diarrhea, and colitis

Clostridium botulinum

Gram-positive, straight rods; occur singly or in pairs; spores usually subterminal and resemble a tennis racket

Food poisoning, botulism; wound botulism and infant botulism, a life-threatening neuromuscular disorder

Clostridium clostridioforme

Gram-positive rod that stains gram negative; long, thin rods; spores usually not seen; elongated football shape with cells often in pairs

Variety of human infections that may be serious and invasive Bacteremia; trauma associated gas gangrene; skin and other soft tissue infections

Clostridium histolyticum

Clostridium novyi

Gram-positive rods with subterminal spores

Skin and other soft tissue infections; cutaneous gas gangrene

Clostridium perfringens

Gram-variable straight rods with blunt ends; occur singly or in pairs; spores seldom seen but if present are large and central to subterminal, oval, and swell cell; large boxcar shapes

Bacteremia; trauma-associated gas gangrene, skin and other soft tissue infections; enteric foodborne disease (food poisoning); enteritis necroticans and necrotizing enterocolitis; anaerobic cellulitis

Clostridium ramosum

Gram-variable straight or curved rods; spores rarely seen but are round and terminal; more slender and longer than C. perfringens

Abscesses, peritonitis, bacteremia, and chronic otitis media in children; bacteremia in adults Continued

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TABLE   Minimal Criteria Grouping Gram Stain Morphology, Aerotolerance, and Clinical Significance of Anaerobic 40.1  Bacteria—cont’d

Organism

Gram Staina

Clinical Significance

Clostridium septicum

Gam positive in young cultures but becomes gram negative with age; stains unevenly; straight or curved rods; occur singly or in pairs; spores subterminal, oval, and swell cells

Bacteremia; trauma-associated gas gangrene; skin and other soft tissue infections; spontaneous nontraumatic gas gangrene

Clostridium sordellii

Gram-positive rods; subterminal spores

Skin and other soft tissue infections; cutaneous gas gangrene; gynecological infections; anaerobic cellulitis

Clostridium tertium

Gram-variable rods; terminal spores

Neutropenic enterocolitis and meningitis in immunocompromised patients

Clostridium tetani

Gram positive, becoming gram negative after 24-h incubation; occur singly or in pairs; spores oval and terminal or subterminal with drumstick or tennis racket appearance

Tetanus associated with puncture wounds

Cryptobacterium spp.

Gram-positive short rods in chains

Oral infections

Curtobacterium spp.

Pleomorphic gram-positive rods, may be branching

Associated with prosthetic infections such as joints and heart valves

Eggerthella spp.

Gram-positive, small, straight rod with rounded ends

Bacteremia

Eggerthii spp.

Gram-positive irregular rods in short chains

Dental abscess with bacteremia; empyema

Eisenbergiella spp.

Gram-positive medium to long, wavy filamentous rods with tapered ends

Bacteremia

Eubacterium spp.

Gram-positive pleomorphic rods or coccobacilli; occur in pairs or short chains; Eubacterium alactolyticum has a seagull-wing shape similar to Campylobacter spp.; Eubacterium nodatum is similar to Actinomyces spp. with beading, filaments, and branching

Oral and other various infections; abscesses, bacteremia, sinusitis, tonsillitis

Filifactor spp.

Gram-positive short, regular bacilli

Oral infections

Finegoldia magna

Gram-positive cocci with cells >0.6 μm in diameter; in pairs and clusters; resemble staphylococci

Infections in various body sites including endocarditis, meningitis, pneumonia, skin, and soft tissue; bone and joint infections, chronic wounds and ulcers, septic arthritis, upper respiratory infections; bacteremia

Fusobacterium spp.

Gram-negative, pale-staining, irregularly stained, highly pleomorphic rods with swollen areas, filaments, and large, bizarre, round bodies

Oral and other various infections; abscesses, bacteremia, sinusitis, tonsillitis

Lactobacillus spp.

Gram-variable pleomorphic rods or coccobacilli; straight, uniform rods have rounded ends; short coccobacilli resemble streptococci

Bacteremia, endocarditis, intraabdominal abscesses; various other infections

Leptotrichia spp.

Gram-negative, large, fusiform rods with one pointed end and one blunt end

Bacteremia in immunocompromised patients; lesions in the oral and gastrointestinal mucosa; endocarditis; bacterial vaginosis

Mobiluncus spp.

Gram-variable, small, thin, curved rods; the two species can be divided based on cell length

Vaginal tract; bacterial vaginosis

Mogibacterium spp.

Gram-positive short rods

Oral infections

Moryella spp.

Gram-positive elongated rods with pointed ends

Abscesses

Olsenella spp.

Short, elliptical gram-positive rods; occur singly, in pairs, or short chains

Dental caries

CHAPTER 40  Overview and General Laboratory Considerations

509

TABLE   Minimal Criteria Grouping Gram Stain Morphology, Aerotolerance, and Clinical Significance of Anaerobic 40.1  Bacteria—cont’d

Organism

Gram Staina

Clinical Significance

Parabacteroides spp.

Gram-negative rods

Intraabdominal infections, bacteremia

Paraeggerthella sp.

Gram-positive, coccobacilli in chains

Bacteremia

Parvimonas micros

Gram-positive cocci with cells 10 mm = sensitive for Km, Van, Col; 9/field

>36/field

>90/field

4+

AFB, Acid-fast bacilli. Modified from Kent PT, Kubica GP. Public Health Mycobacteriology: A Guide for the Level III Laboratory, Washington, DC: Centers for Disease Control and Prevention; 1985; Carroll KC, Pfaller MA. Manual of Clinical Microbiology. 12th ed. Washington, DC: ASM Press; 2019.

with the patient’s CD4 cell count; therefore, interpretations and results should be evaluated with caution. 

Nucleic Acid Detection Nucleic acid probes for the indirect detection and identification of M. tuberculosis complex and the M. avium complex from culture media are available (AccuProbe, Hologic, San Diego, CA). This technique can detect M. avium, M. intracellulare, M. tuberculosis, M. africanum, M. microti, M. pinnipedii, and M. canettii, but it cannot differentiate them at the species level. The assay also lacks enough sensitivity to be used for direct detection of the organisms in clinical samples. 

Genetic Sequencing and Nucleic Acid Amplification PCR-based sequencing for mycobacterial identification consists of PCR amplification of mycobacterial DNA with genus-specific primers and sequencing of the amplicons. The organism is identified by comparison of the nucleotide sequence with reference sequences. The most reliable sequence for identification of mycobacteria to the genus level is the approximately 1500 bp 16S ribosomal ribonucleic acid (rRNA) gene. However, only a 600-bp sequence at the 5′ end is required for identification. The sequence homogeneity in the M. tuberculosis complex prevents the use of this sequence to differentiate these species. This region contains both conserved and variable regions, which makes it an ideal target for identification purposes. Despite the accuracy of PCR-based sequencing to identify mycobacteria, problems remain: the sequences in some databases are not accurate, no present consensus exists as to the quantitative definition of a genus or species based on 16S rRNA gene sequence data, and procedures are not standardized. In addition, the 16S rRNA 5′ region contains two hypervariable regions, A and B. The A region provides the signature sequences for species identification. However,

M. chelonae and M. abscessus both require additional sequencing, because the A and B regions are identical and the 3′ end of the 16S rRNA contains a 4-bp sequence difference. Several other genes have also been used to identify mycobacterial species, including the 23S rRNA, ITS 1, hsp65, rpoB, and gyrB gene. The 23S rRNA sequence is 3100 bp in length, which limits accurate sequencing. ITS 1 is a spacer sequence located between the 16S and 23S rRNA genes. This sequence, which is 200 to 330 bp, is more easily analyzed. The limitation of this sequence is that it is not a genus-specific sequence; therefore results may be affected by contaminating bacteria. The 65-kDa heat shock protein, also referred to as the groEL2 gene, is a 440-bp fragment that can be amplified and analyzed with restriction digestion followed by agarose electrophoresis. The hsp65 is highly conserved but contains a greater variation in polymorphisms than the 16S rRNA, particularly in a 441-bp region referred to as the “Telanti fragment.” This allows for the differentiation of Mycobacterium spp. based on the variation in restriction fragment length polymorphisms (RFLPs). Repetitive sequence–based PCR, Diversilab (bioMérieux, Durham, NC), demonstrates better species discrimination than RFLP. Line probe assays (DNA strip assays) involve PCR amplification coupled with a reverse hybridization step. The target sequence is amplified using biotinylated primers. The amplicon is then hybridized to membrane-immobilized sequence-specific probes for each species. The membrane is developed using an enzyme-mediated reaction and color indicator to analyze the banding pattern. Banding patterns are species-specific based on the immobilized probe map on the membrane. A commercially available system in which the 16S-to-23S rRNA spacer region of mycobacterial species (INNO-LiPA v2 Mycobacteria; Innogenetics, Ghent, Belgium) has been successfully used to directly detect and identify several of the most clinically relevant mycobacterial species in aliquots of positive liquid culture. The assay can be

548 PA RT I I I    Bacteriology

used to identify the MTBC and 13 NTM species. However, the results should be interpreted cautiously because some cross-reactivity has occurred with closely related species. Another commercial system, GenoType Mycobacterium (Hain Lifescience GmbH, Nehren, Germany), which uses a similar format, has additional probes for M. celatum, Mycobacterium malmoense, Mycobacterium peregrinum, M. phlei, and two subgroups of M. fortuitum, in addition to a supplemental kit that allows for the identification of 16 additional mycobacterial species. The rpoB gene encodes the beta-subunit in the organism’s RNA polymerase. Mutations in this gene confer rifampin resistance in M. tuberculosis. Different regions in this gene have been used to identify rapid-growing isolates, but little data are available for the slow-growing species. Finally, the gyrB gene encodes the beta-subunit in the organism’s topo­ isomerase II. Several single nucleotide polymorphisms have been identified in this gene that are useful in distinguishing species in the M. tuberculosis complex. After amplification, identification and differentiation of species require restriction analysis and gel electrophoresis. Additional molecular techniques, such as conventional and real-time PCR, have been used to detect M. tuberculosis directly in clinical specimens. The Xpert MTB/RIF (Cepheid, Sunnyvale, CA) is a real-time PCR, cartridgebased molecular beacon probe assay that also detects the mutations associated with rifampin resistance. The assay can detect the presence or absence of mutations and is unable to recognize silent mutations resulting in incorrect interpretations as rifampin resistance. The CDC recommends followup confirmation should be completed using sequencing technology. The Xpert MTB/Rif Ultra (Cepheid, Sunnyvale, CA), is capable of detecting some of the mutations that would previously go undetected. The Xpert MTB/RIF assay is approved on both smear-positive and smear-negative respiratory specimens only. The Amplified M. tuberculosis Direct Test (AMTD; GenProbe Hologic, San Diego CA) uses rRNA released from the mycobacteria by means of a lysing agent, sonication, and heat. The specific DNA probe is allowed to react with the extracted rRNA to form a stable DNA-RNA hybrid. Any nonhybridized DNA acridinium ester probes are chemically degraded. When an alkaline hydrogen peroxide solution is added to activate the chemiluminescence, only the hybrid bound acridinium ester emits light. The amount of light emitted is directly proportional to the amount of hybridized probe. The light is measured using a chemiluminometer. As previously indicated, the commercially available tests are limited in the number of species they can identify. Some clinical laboratories have developed their own PCR assays to detect M. tuberculosis directly in clinical specimens. Due to the limitation associated with amplified probe assays, sequencing-based methods—including traditional Sanger sequencing, pyrosequencing, next-generation sequencing and whole genome sequencing—are being used to resolve discrepancies associated with detecting drug resistance in Mycobacterium spp. In addition, genome

sequencing is important for epidemiologic students to resolve the transmission of clustered patients and identify false-positive cultures that cause patients to undergo unnecessary treatment. Genotyping and sequencing are essential for tuberculosis control programs. The CDC has established a national tuberculosis genotyping system and a National TB Molecular Surveillance Center that performs whole genome sequencing on all newly diagnosed patients in the United States. Details and updates are available at https://w ww.cdc.gov/tb/topic/laboratory/default.htm. DNA Microarrays

DNA microarrays are also attractive for rapid examination of large numbers of DNA sequences by a single hybridization step. This approach has been used to simultaneously identify mycobacterial species and detect mutations that confer rifampin resistance in mycobacteria. Fluorescentlabeled PCR amplicons generated from bacterial colonies are hybridized to a DNA array containing nucleotide probes. The bound amplicons emit a fluorescent signal that is detected with a scanner. With this approach, 82 unique 16S rRNA sequences allow for the differentiation of 54 mycobacterial species and 51 sequences that contain unique rpoB gene mutations (rifampin resistance).  Matrix-Assisted Laser Desorption Ionization Time-ofFlight Mass Spectrometry

The current approaches for early identification of Mycobacterium spp. in culture include PCR and traditional phenotypic identification schemes. MALDI-TOF MS uses a proteomicbased technique to identify clinical mycobacterial isolates by protein profiling and can be used for accurate and rapid identification of various microorganisms. (See Chapter 7 for more information on MALDI-TOF MS.) Isolates of M. tuberculosis are consistently identified using MALDI-TOF MS; however, discrimination of the other species included in the M. tuberculosis complex is limited. Correct identification of clinically relevant strains of tuberculoid and NTM species has also been demonstrated; however, the capacity to identify all the species is limited to the information available in the current databases. In addition, identification is limited based on successful cultivation in vitro. Moderate growth in culture is required from a specimen to have sufficient sample size for the application to MALDI-TOF MS, whereas sequencing techniques require minimal or scant growth. The application of MALDI-TOF MS to the identification of mycobacterial species requires an extraction process prior to placing the organism on the target plate. This involves chemical and physical disruption of the mycobacterial cell wall. See Fig. 42.4 for the processing of mycobacterial species and extraction required for identification using MALDI-TOF MS. Extraction should be completed in a BSL-3 or BSL-2 using BSL-3 practices. 

Cultivation A combination of different culture media is required to optimize recovery of mycobacteria from culture; at least one solid

CHAPTER 42 Mycobacteria

549

Liquid sample processing method by protein extraction and inactivation for mycobacteria Note: Steps 1–7 must be performed in a biosafety level 3 cabinet.

+

1. Mix the bottle or the tube using a vortex-type mixer for 5 to 10 seconds and immediately aseptically transfer 3.0 mL of medium into the 5 mL conical bottom tube (CBT).

2. Centrifuge CBT for 10 minutes at 3,000 g using a swing bucket centrifuge with a 15 mL adapter to create a pellet.*

4. Add 500 µL of R1 to the 5 mL CBT and resuspend by aspiration / dispensing using a pipette.

5. Transfer the suspension to a tube with glass beads (BEAD). 6. Use a vortex-type mixer with adaptor (at maximum speed) to disrupt the cells for 15 minutes or a bead beater-type homogenizer for 5 minutes.

+

10 mins.

3. Decant medium into a waste container and completely blot CBT dry onto an absorbent pad with protective backing (WIPE). Discard pad after use, avoiding touching the absorbant surface.

7. Remove from the mixer or the bead beatertype homogenizer and incubate the tube at room temperature for 10 minutes to complete the inactivation.

Note: The following steps can be performed out of a biosafety level 3 cabinet. 8. Mix with a vortex-type mixer for 5 to 10 seconds and immediately transfer suspension into an empty 2 mL round-bottomed tube (RBT) using a pipette and avoiding transfer of any glass beads. Discard the pipette tip.

9. Centrifuge sample for 2 minutes at a minimum of 14,000 g to create a pellet.

11. Add 10 µL of R2 to the pellet. Resuspend by aspiration / dispensing using a pipette until the pellet is uniformly dispersed, or directly with a vortex-type mixer. Note: If pellet is not visible, wash sides of tube with R2 to ensure re-suspension.

12. Add 10 µL of R3 and mix using a vortex-type mixer.

+

Note: If the spot is not completely dry before addition of VITEK® MS-CHCA, optimal crystallization of sample may not be achieved and could potentially interfere with VITEK MS results (no identification).

10. Discard all the R1 supernatant using a pipette.

Note: Before the centrifugation steps, note the position of the expected pellet. This could be helpful in case of low intensity pellet.

+

13. Centrifuge for 2 minutes at a minimum of 14,000 g.

16. Add 1 µL of VITEK MS-CHCA matrix to each target slide spot using a new pipette tip after each addition of matrix. Allow matrix to dry.

14. For each organism to be tested, immediately transfer 1 µL of the supernatant onto the target slide spots. 15. Allow each spot to dry completely.

Important: Once the VITEK MS-DS target slide is prepared, it must be tested within 72 hours. Before spectra acquisition, it must be stored at room temperature in its original packaging.

•

Fig. 42.4 Matrix-assisted desorption ionization time-of-flight workflow for extraction and processing Mycobacterium isolates. (Courtesy Biomerieux Inc., Durham, NC.)

medium in addition to a liquid medium should be used. The ideal media combination should be economical and should support the most rapid and abundant growth of mycobacteria, allow for the study of colony morphology and pigment production, and inhibit the growth of contaminants. 

Solid Media Solid media, such as those listed in Box 42.1, are recommended because of the development of characteristic reproducible colonial morphology, good growth from small inocula, and a low rate of contamination. Optimally, at

550 PA RT I I I    Bacteriology

• BOX 42.1 Suggested Media for Cultivation of Mycobacteria From Clinical Specimensa Media

Comments

Solid

Media

Comments

Petragnani medium

Contains twice the concentration of malachite than Löwenstein-Jensen green (an inhibitor of contaminating organisms); improves recovery from heavily contaminated specimens Supplemented with hemin, hemoglobin, or ferric ammonium citrate; increases recovery of Mycobacterium haemophilum

Agar Based—Growth Within 10–12 Days Middlebrook

Middlebrook 7H10 Middlebrook 7H10 selective

Middlebrook 7H11 Middlebrook 7H11 selective

Middlebrook 7H11

Middlebrook 7H11 thin pour plates, 10 × 90 mm (Remel, Lenexa, KS) Middlebrook biplate (7H10/7H11S agar)

Contains 2% glycerol, which enhances the growth of Mycobacterium avium complex (MAC) Supplemented with carbenicillin (for inhibition of pseudomonads), polymyxin B, trimethoprim lactate, and amphotericin B Contains 0.1% enzymatic hydrolysate of casein, which improves recovery of isoniazid-resistant Mycobacterium tuberculosis Supplemented with mycobactin J, which provides for growth of Mycobacterium genavense Enhances visibility of colonies within 11 days

Egg Based—Growth Within 18–24 Days Löwenstein-Jensen (L-J)

L-J Gruft L-J Mycobactosel

L-J with pyruvic acid L-J with glycerol aFor

Commonly used medium; good recovery of M. tuberculosis but poor recovery of many other species; M. genavense fails to grow Supplemented with penicillin and nalidixic acid Supplemented with cycloheximide, lincomycin, and nalidixic acid Enhances recovery of Mycobacterium bovis Enhances recovery of Mycobacterium ulcerans

Heme-supplemented media (egg or agar based)

Liquidb BACTEC 12B medium

Middlebrook 7H9 broth Dubos Tween albumin Septi-Chek AFB

Used in the MGIT960 system; PANTA is added before incubation; 14C-labeled palmitic acid is metabolized to produce 14CO2, which is detected by the instrument

20 mL of Middlebrook 7H9 broth is incubated in 20% CO2; solid phase contains three media: modified L-J, Middlebrook 7H11, and a chocolate agar slab

Media Used in Commercially Supplied Growth and Semiautomated or Fully Automated Systems Mycobacteria Growth Indicator Tube [MGIT] (Becton Dickinson Microbiology Systems, Cockeysville, MD) Versa TREK Culture System (Trek Diagnostic Systems, Cleveland, OH) MB/BacT Alert 3D (bioMérieux, Durham, NC)

MGIT 960 (fully automated system); MGIT is a modified Middlebrook 7H9 broth that incorporates a fluorescence-quenching–based oxygen sensor for detection Modified Middlebrook 7H9 broth

Uses Middlebrook 7H9 broth

optimal recovery of mycobacteria, a minimum combination of liquid medium and solid media is recommended. 80 added to liquid media acts as a surfactant, breaking up clumps of organisms and increasing recovery rates.

bTween

least two solid media (a serum [albumin] agar base medium, [e.g., Middlebrook 7H10] and an egg-potato base medium [e.g., Löwenstein-Jensen, or L-J]) should be used for each specimen (these media are available from commercial

sources). All specimens must be processed appropriately before inoculation. It is imperative to inoculate test organisms to commercially available products for quality control (Evolve Procedure 42.4). An example of interpreting

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551

• BOX 42.2 Example of Interpreting Quality Control Test Results of Decontamination and Concentration

Procedure

SPUTUM SPECIMEN

Unprocessed Quantification of Growth

Processed Quantification of Growth

Sputum Sample 1

104

103

102

3+

2+

50–100 colonies

104 2+

103 1+ or 2+

102 Approximately 10 colonies

Interpretation Media and decontamination procedures acceptable

2

3+

2+

50–100 colonies

1+

0

0

Media acceptable; procedures too toxic

3

2+ or 1+

2+ or 1+

0

1+ or 0

1+ or 0

0

One or more of the media are not supporting growth of acidfast bacilli (AFB) adequately

0, No growth; 1+, scanty, barely discernible countable colonies; 2+, dense, discrete growth, not countable; 3+, confluent, abundant growth

quality-control test results of decontamination and concentration procedures associated with culture is provided in Box 42.2. Cultures are incubated at 35°C in the dark in an atmosphere of 5% to 10% carbon dioxide (CO2) and high humidity. Tube media are incubated in a slanted position with screw caps loose for at least 1 week to allow for the evaporation of excess fluid and the entry of CO2; plated media are either placed in a CO2-permeable plastic bag or wrapped with CO2-permeable tape. If specimens obtained from the skin or superficial lesions are suspected to contain M. marinum or M. ulcerans, an additional set of solid media should be inoculated and incubated at 25°C to 30°C. In addition, a chocolate agar plate (or placement of an X-­factor [hemin] disk on conventional media) and incubation at 25°C to 33°C is needed for recovery of M. haemophilum from these specimens. RGM optimally require incubation at 28°C to 30°C. Cultures are examined weekly for growth. Contaminated cultures are discarded and reported as “contaminated, unable to detect presence of mycobacteria”; additional specimens are also requested. If available, sediment may be recultured after enhanced decontamination or by inoculating the sediment to a more selective medium. Most isolates appear between 3 and 6 weeks; a few isolates appear after 7 or 8 weeks of incubation. When growth appears, the rates of growth, pigmentation, and colonial morphology are recorded. The typical colonial appearance of M. tuberculosis and other mycobacteria is shown in Fig. 42.5. After 8 weeks of incubation, negative

cultures (those showing no growth) are reported, and the cultures are discarded. 

Liquid Media In general, use of a liquid media system reduces the turnaround time for the isolation of AFB to approximately 10 days, compared with 17 days or longer for conventional solid media. Several different systems are available for culturing and detecting the growth of mycobacteria in liquid media. Growth of mycobacteria in liquid media, regardless of the type, requires 5% to 10% CO2; CO2 is either already provided in the culture vials or is added according to the manufacturer’s instructions. When growth is detected in a liquid medium, acid-fast staining of a culture aliquot is performed to confirm the presence of AFB, and the material is subcultured to solid agar. Gram staining can also be performed if contamination is suspected. 

Interpretation Although isolation of MAC organisms indicates infection, the clinician must determine the clinical significance of isolating NTM in most cases; in other words, does the organism represent mere colonization or significant infection? Because these organisms vary greatly in their pathogenic potential, can colonize an individual without causing infection, and are ubiquitous in the environment, interpretation of a positive NTM culture is complicated; therefore the American Thoracic Society has recommended diagnostic criteria for NTM disease to help physicians interpret culture results. 

552 PA RT I I I    Bacteriology

A

B

C

D

E • Fig. 42.5  Typical appearance of some mycobacteria on solid agar media. (A) Mycobacterium tuberculo-

sis colonies on Löwenstein-Jensen agar after 8 weeks of incubation. (B) A different colonial morphology is seen on culture of one strain of Mycobacterium avium complex. (C) Mycobacterium kansasii colonies exposed to light. (D) Scotochromogen Mycobacterium gordonae showing yellow colonies. (E) Smooth, multilobate colonies of Mycobacterium fortuitum on Löwenstein-Jensen medium.

Approach to Identification Regardless of the identification methods used, the first test always performed on organisms growing on solid or liquid mycobacterial media is acid-fast staining to confirm that the organisms are indeed mycobacteria. Identification of species other than MAC and the more commonly isolated NTM (MAC, M. avium, M. intracellulare, M. gordonae, and M. kansasii) has become challenging for routine clinical microbiology laboratories, particularly considering the ever-increasing number of new mycobacterial species.

Traditional methods (i.e., phenotypic methods) for identifying mycobacteria, particularly the NTM, are based on growth parameters, biochemical characteristics, and analysis of cell wall lipids, all of which are slow, cumbersome, and often inconclusive procedures. Over the past decade, the rate of non–AIDS-associated infections have been increasing, and many of the newly identified NTM species have been associated with various diseases. As a result, identification of species is vital to selecting effective antimicrobial therapy and to deciding whether to perform susceptibility

CHAPTER 42 Mycobacteria

testing on accurately speciated NTM. Newer species have been identified using nucleic acid sequencing with limited published phenotypic characteristics. Because of these issues and limitations with conventional phenotypic methods for identification, molecular and genetic investigations are becoming indispensable to identifying the NTM accurately. Therefore, for timely and accurate identification of mycobacteria, molecular approaches in conjunction with some phenotypic characteristics should be used. Regardless of whether molecular or phenotypic methods are used, when growth is detected, broth subcultures of colonies growing in liquid media or on solid media (several colonies inoculated to Middlebrook 7H9 broth [5 mL] and incubated at 35°C for 5 to 7 days with daily agitation to enhance growth) are then used to determine pigmentation and growth rate and to inoculate all test media for biochemical tests if performed. Additional cultures may be inoculated and then incubated at different temperatures when more definitive identification is needed.

Conventional Phenotypic Tests Growth Characteristics

Preliminary identification of mycobacterial isolates depends on the organisms’ rate of growth, colonial morphology (Fig. 42.5), colonial texture, pigmentation, and, in some instances, the permissive incubation temperatures of mycobacteria. Despite the limitations of phenotypic tests, the mycobacterial growth characteristics are helpful for determining a preliminary identification (e.g., an isolate appears to represent RGM). To perform identification procedures, quality-control organisms should be tested along with unknowns (Table 42.8). The commonly used quality-control organisms can be maintained in broth at room temperature and transferred monthly. In this way they are always available for inoculation to test media along with suspensions of the unknown mycobacteria being tested. Growth Rate. The rate of growth is an important criterion for determining the initial category of an isolate. Rapid growers usually produce colonies within 3 to 4 days after subculture. However, even a rapid grower may take longer than 7 days to initially produce colonies because of inhibition by a harsh decontaminating procedure; therefore the growth rate (and pigment production) must be determined by subculture (Evolve Procedure 42.5). The dilution of the organism used to assess the growth rate is critical. Even slow-growing mycobacteria appear to produce colonies in less than 7 days if the inoculum is too heavy. One organism particularly likely to exhibit false-positive rapid growth is Mycobacterium flavescens. It therefore serves as an excellent quality-control organism for this procedure.  Pigment Production. As previously discussed, mycobacteria can be categorized into three groups based on pigment production. Evolve Procedure 42.5 describes how to determine pigment production. To achieve optimum photochromogenicity, colonies should be young, actively metabolizing, isolated, and well aerated. Although some species (e.g., M. kansasii) turn yellow after a few hours of

553

light exposure, others (e.g., Mycobacterium simiae) may require prolonged exposure to light. Scotochromogens produce pigmented colonies even in the absence of light, and colonies often become darker with prolonged exposure to light (Fig. 42.6). One member of this group, Mycobacterium szulgai, is peculiar in that it is a scotochromogen at 35°C and nonpigmented when grown at 25°C to 30°C. For this reason, all pigmented colonies should be subcultured to test for photoactivated pigment at both 35°C and 25°C to 30°C. Nonchromogens are not affected by light.  Biochemical Testing

Once categorized into a preliminary subgroup based on its growth characteristics, an organism must be definitively identified to species or complex level. Although conventional biochemical tests can be used for this purpose, molecular methods have replaced biochemical tests for identifying mycobacterial species because of the previously discussed limitations of phenotypic testing. Although key biochemical tests are still discussed in this edition, the reader must be aware that this approach to identification will ultimately be replaced by molecular methods or at the very least be utilized to resolve unusual cases in the absence of availability of whole genome sequencing and/or database limitations. Table 42.9 summarizes distinctive properties of the more commonly cultivable mycobacteria isolated from clinical specimens; key biochemical tests for each of the major mycobacterial groupings, including M. tuberculosis complex, are listed in Table 42.10. The following sections address key biochemical tests. Niacin. Niacin (nicotinic acid) plays an important role in the oxidation-reduction reactions that occur during mycobacterial metabolism. Although all species produce nicotinic acid, M. tuberculosis accumulates the largest amount. (M. simiae and some strains of M. chelonae also produce niacin.) Niacin therefore accumulates in the medium in which these organisms are growing. A positive niacin test (Evolve Procedure 42.6) is preliminary evidence that an organism that exhibits a buff-colored, slow-growing, rough colony may be M. tuberculosis (Fig. 42.7). However, this test is not sufficient to confirm identification. If sufficient growth is present on an initial L-J slant (the egg-base medium enhances accumulation of free niacin), a niacin test can be performed immediately. If growth on the initial culture is scant, the subculture used for growth rate determination can be used. If this culture yields rare colonies, the colonies should be spread around with a sterile cotton swab (after the growth rate has been determined) to distribute the inoculum over the entire slant. The slant then is incubated until light growth over the surface of the medium is visible. For reliable results, the niacin test should be performed from cultures on L-J medium that are at least 3 weeks old and show at least 50 colonies; otherwise, enough detectable niacin might not have been produced.  Nitrate Reduction. A nitrate reduction test is valuable for identifying M. tuberculosis, M. kansasii, M. szulgai, and

Control Organisms

Biochemical Test Tables Are Difficult to Review in This Format

Result

Incubation Conditions

Positive

Negative

Positive

Negative

Medium Used

Duration

Niacin

Mycobacterium tuberculosis

Mycobacterium intracellulare

Yellow

No color change

0.5 mL DH2O

15–30 min

Room temperature

Nitrate

Mycobacterium tuberculosis

Mycobacterium intracellulare

Pink or red

No color change

0.3 mL DH2O

2h

37°C bath

Urease

Mycobacterium fortuitum

Mycobacterium avium

Pink or red

No color change

Urea broth for AFB

1, 3, and 5 days

37°C incubator (without CO2)

68°C Catalase

Mycobacterium fortuitum or Mycobacterium gordonae

Mycobacterium tuberculosis

Bubbles

No bubbles

0.5 mL phosphate buffer (pH, 7.0)

20 min

68°C bath

SQ Catalase

Mycobacterium kansasii or M. gordonae

Mycobacterium avium

>45 mm

200 species)

Ureaplasma (7 species)

*Mollicutes that have been detected in humans.

• Fig. 44.1  Taxonomy of the class Mollicutes.

of all free-living bacteria. Mollicutes are descended from low G+C gram-positive bacteria (i.e., the Firmicutes), and their closest living relatives include bacilli, streptococci, and lactobacteria. 

Epidemiology and Pathogenesis Mycoplasmas are part of the human microbiota of the oropharynx and upper respiratory tract and can be part of the microbiota of the genitourinary tract. Besides those that are considered commensals, considerable evidence indicates the pathogenicity of some mycoplasmas; for others, a role in human disease is unclear.

Epidemiology The mycoplasmas usually considered as commensals are listed in Table 44.1, along with their respective sites of colonization. These organisms are transmitted by direct sexual contact, by transplanted tissue from donor to recipient, or from mother to fetus during childbirth or in utero. Mycoplasma pneumoniae are transmitted by respiratory secretions. One species of Acholeplasma (organisms that are widely disseminated in animals), Acholeplasma laidlawii, has been isolated from the oral cavity of humans; however, the significance of these mycoplasmas and their colonization of humans remains uncertain. Of the other mycoplasmas isolated from humans, the possible role that Mycoplasma pirum, Mycoplasma amphoriforme, Mycoplasma fermentans, and Mycoplasma penetrans might play in human disease is uncertain at this time. M. pirum, M. fermentans, and M. penetrans have been isolated from patients infected with human immunodeficiency virus (HIV); however, no clear link exists between acquisition or shedding of HIV and infections with these mycoplasmas. The potential for M. penetrans to play a role in the transition from HIV-positive patients

to clinical acquired immune deficiency syndrome (AIDS) has been examined but no connection has been fully demonstrated. In contrast, the ability of M. genitalium to facilitate both the shedding and the acquisition of HIV following exposure has been determined. M. fermentans has been isolated from specimens such as bronchoalveolar lavage, bone marrow, peripheral blood, and the throats of children with pneumonia. M. fermentans has also been associated with infection in children and immunocompromised individuals. M. amphoriforme has been detected in the lower respiratory tract in patients with chronic respiratory disease and antibody deficiencies, and isolation from healthy asymptomatic carriers has never been reported. However, until recent advances in molecular diagnostic offering for M. pneumoniae, the serologic cross reactivity between M. pneumoniae and M. amphoriforme may have masked primary respiratory tract infections with M. amphoriforme in previously healthy patients by miscategorizing them as M. pneumoniae infections. Prospective studies evaluating culture-positive, M. pneumoniae DNA-negative respiratory specimens can be evaluated to determine the role of M. amphoriforme as a primary or opportunistic pathogen. The presence of various Mycoplasma spp. infections in immunocompromised patients has been demonstrated by genital or respiratory tract colonization associated with medical procedures such as renal transplantation or genitourinary manipulations or following trauma resulting in wound infections. Finally, the remaining species that have been isolated from humans—M. pneumoniae, M. genitalium, M. phocicerebrale, Ureaplasma urealyticum, Ureaplasma parvum, and Mycoplasma hominis—have well-established roles in human infections. M. genitalium, U. urealyticum, U. parvum, and M. hominis have been isolated from the genitourinary tract of humans, and M. pneumoniae has been isolated from the respiratory tract. M. genitalium accounts for approximately 15% to 20% of nongonococcal urethritis (NGU), which

578 PA RT I I I    Bacteriology

TABLE 44.1    Mycoplasmas Isolated From Humans

Organism

Source of Isolation

Clinical Presentations

Mycoplasma amphoriforme

Respiratory tract

• C  hronic bronchopneumonia and relapsing airway infection in immunocompromised patients • Primary atypical pneumonia or secondary pneumonia (proposed/ suspected)

Mycoplasma buccale

Oropharynx

• Oral microbiome

Mycoplasma orale

Oropharynx

• O  ral microbiome • Opportunistic pneumonia, infectious synovitis, osteomyelitis, or abscess in immunocompromised patients

Mycoplasma salivarium

Oropharynx, gingiva

• O  ral microbiome • Opportunistic arthritis, submasseteric abscess,a gingivitis, or periodontitis in immunocompromised patients

Mycoplasma hominis

Genital tract, joints, respiratory tract (neonates), central nervous system (neonates)

• • • • • •

• • • •

 aginal microbiome V Bacterial vaginosis Pelvic inflammatory disease Pyelonephritis Chorioamnionitis Bronchopulmonary dysplasia (neonates) Meningitis/meningoencephalitis (neonates) Septic arthritis Abscess (pelvic, brain, aortal) Meningitisa Osteomyelitisa

• • • • • • • • • •

 rimary atypical pneumonia P Bronchitis Pharyngitis Meningoencephalitis Pericarditis Arthritis Hemolytic anemia Nephritis Bell palsy Stevens-Johnson syndrome

•

Mycoplasma pneumoniae

Mycoplasma faucium

Oropharynx

• O  ral microbiome • Cerebral abscessa

Mycoplasma fermentans

Oropharynx, peripheral blood, respiratory tract, bone marrow, urine, genital tract

• • • •

Mycoplasma lipophilum

Respiratory tract

• Pneumoniaa

Mycoplasma penetrans

Urine, genital tract, blood, respiratory tract

• HIV disease progression (proposed) • Idiopathic nongonococcal urethritis

Mycoplasma phocicerebrale

Cutaneous lesions

• “ Seal finger” (ulcerative keratitis secondary to seal bite)

Mycoplasma pirum

Rectum, peripheral blood, urine

• Gastrointestinal microbiome

 ipshütz ulcera L Infectious rheumatoid arthritis Malignant transformation (proposed) Opportunistic pneumonia (proposed)

CHAPTER 44  Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma

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TABLE 44.1    Mycoplasmas Isolated From Humans—cont’d

Organism

Source of Isolation

Clinical Presentations

Mycoplasma genitalium

Genital tract, urine

• • • • • • •

Mycoplasma primatum

Genital tract, oropharynx

• O  ral microbiome • Opportunistic keratitisa

Mycoplasma spermatophilum

Genital tract, semen

• Impaired fertility (proposed)

Acholeplasma laidlawii

Cutaneous burns

• Complex burn infectionsa,b

Ureaplasma parvum

Genital tract, urine, semen, blood, neonatal respiratory tract, neonatal central nervous system

• • • •

Ureaplasma urealyticum

Genital tract, urine, semen, blood, neonatal respiratory tract, neonatal central nervous system

• U  rogenital microbiome • Chorioamnionitis • Fatal hyperammonemia syndrome in immunocompromised patients • Preterm labor • Bronchopulmonary dysplasia (neonates) • Meningitis/meningoencephalitis (neonates) • Nongonococcal urethritis

 ongonococcal urethritis N Prostatitis Pelvic inflammatory disease Infertility Cervicitis Sexually acquired reactive arthritis Enhanced HIV transmission

 rogenital microbiome U Chorioamnionitis Preterm labor Bronchopulmonary dysplasia (neonates) • Meningitis/meningoencephalitis (neonates) • Nongonococcal urethritis

HIV, Human immunodeficiency virus. aThis presentation has been reported less than 5 times. bThis presentation is described in a single report, and all Acholeplasma-positive lesions were coinfected with Gram-positive bacilli. Evidence for an association between A. laidlawii and burn infections or carriage in humans is minimal.

is urethritis not attributable to Neisseria gonorrhoeae, the most common cause of urethritis in males. M. genitalium is not associated with the presence of other mycoplasmas or ureaplasmas in the urogenital tract. In females, this organism can cause cervicitis, pelvic inflammatory disease (PID), and postinfectious tubal factor infertility. Both Ureaplasma and Mycoplasma species have been isolated from the internal organs of stillborn, premature, and spontaneously aborted fetuses. Although M. genitalium and, to a lesser extent, M. hominis are associated but not yet definitely implicated in pregnancy-related complications, U. urealyticum and U. parvum are recognized as causes of chorioamnionitis, preterm labor, and premature rupture of the membranes both in clinical studies and prospective studies using animal (sheep) models. In addition, delivery following invasion of the chorioamnion and amniotic fluid by ureaplasmas is strongly associated with risk for bronchopulmonary dysplasia and meningitis/meningoencephalitis in preterm infants. Recently, Ureaplasma sp. and M. hominis have been

identified as a cause of hyperammonemia in immunocompromised patients. Risks associated with prenatal ureaplasmosis are highly strain dependent. Infants are commonly colonized with benign strains of U. urealyticum and M. hominis. Once an individual reaches puberty, colonization with these organisms occurs primarily as a result of sexual contact. M. hominis is associated with bacterial vaginosis (BV) and should be considered in clinical BV when other agents have been ruled out. The organism also has a commensal relationship with the protozoal parasite Trichomonas vaginalis, wherein it invades T. vaginalis cells and lives intracellularly. This relationship may play a role in elevated pathogenicity for T. vaginalis in the form of enhanced cytopathology and metronidazole resistance. The role of T. vaginalis in the improved survival and enhanced transmission for M. hominis has been confirmed during antibiotic treatment. M. hominis is rarely associated with infection at distal body sites, most notably cerebral abscess.

580 PA RT I I I    Bacteriology

M. pneumoniae is a cause of community-acquired atypical pneumonia, often referred to as walking pneumonia (Chapter 68). The organism causes infections worldwide, with an estimated 2 million cases per year in the United States. M. pneumoniae infection may also result in bronchitis or pharyngitis. M. pneumoniae may be transmitted person to person by respiratory secretions or indirectly via inanimate objects contaminated with respiratory secretions (fomites). Infections can occur singly or as outbreaks in closed populations such as families and military recruit camps. Pneumonia caused by M. pneumoniae may present as asymptomatic to mild disease, with early nonspecific symptoms including malaise, fever, headache, sore throat, earache, and nonproductive cough. This differs significantly from the classic symptoms associated with pneumonia associated with a Streptococcus pneumoniae infection (Chapters 14 and 68). M. pneumoniae strongly attaches to the mucosal cells and may reside intracellularly within host cells, resulting in a chronic persistent infection that may last for months to years. The infections do not follow seasonal patterns as seen with influenza and other respiratory pathogens. Besides respiratory infection, M. pneumoniae can cause extrapulmonary manifestations such as pericarditis, hemolytic anemia, arthritis, nephritis, Bell palsy, and meningoencephalitis. Finally, M. phocicerebrale, an oropharyngeal commensal and occasional cause of wound infections of seals, is the only confirmed mycoplasmal zoonosis described to date. This organism is associated with ulcerative keratitis secondary to marine mammal contact colloquially known as “seal finger.” 

Pathogenesis In general, mycoplasmas colonize mucosal surfaces of the respiratory and urogenital tracts. Except for those mycoplasmas noted, most rarely produce invasive disease except in immunocompromised hosts or infections associated with medical device implants. Of the mycoplasmas that are established as causes of human infections, these agents predominantly reside extracellularly, attaching with great affinity to the surfaces of ciliated and nonciliated epithelial cells. Recently M. fermentans, M. penetrans, M. genitalium, M. hominis, and M. pneumoniae have been identified intracellularly. Intracellular invasion in bacterial infections is generally considered a means for immune evasion and may contribute to the persistent nature of infections, capacity for recrudescence after cessation of treatment, and difficulties in cultivation or isolation of Mycoplasma spp. M. pneumoniae has a complex and specialized attachment organelle that includes a P1 adhesin protein that primarily interacts with host cells. Similarly, M. genitalium uses a tip organelle to attach to host cells using the MgPa adhesion, which is homologous to the P1 adhesin of M. pneumoniae. With respect to the mycoplasmas that are clearly able to cause disease, many of the disease processes include a strong competent host immune response (inflammation), which leads

to long-term inflammatory syndromes because of the chronic nature of these infections. In addition to adherence properties and host immune responses to infection, the ability to directly cause cell death may also contribute to their pathogenicity. M. pneumoniae produces a potent ADP-ribosylating toxin (“community-acquired respiratory distress syndrome” [CARDS] toxin) that is strongly associated with disease capacity. Of interest, the mycoplasmas associated with HIV patients (M. fermentans, M. penetrans, and M. pirum) are all capable of invading human cells and modulating the immune system. Based on these findings, some investigators have proposed that these mycoplasmas might play a role in certain disease processes in these patients. 

Spectrum of Disease The clinical manifestations of mycoplasmosis in humans are summarized in Table 44.1. 

Laboratory Diagnosis The laboratory diagnosis of mycoplasma infections can be extremely challenging because of complex and timeconsuming culture requirements; however, the recent availability of rapid, molecular diagnostic tests represents a major step forward in the diagnosis of mycoplasmosis and ureaplasmosis. Accurate, rapid diagnosis for M. pneumoniae and M. genitalium is highly desired because penicillin and other beta-lactam agents are ineffective treatments. The laboratory diagnosis of the cell wall deficient organisms implicated in human diseases (i.e., M. pneumoniae, U. urealyticum, U. parvum, M. hominis, and M. genitalium) is addressed.

Specimen Collection, Transport, and Processing Consistent with the diverse spectrum of diseases, various specimens are appropriate for the diagnosis of mycoplasma infections by culture or other means of detection. Acceptable specimens include body fluids (e.g., blood, joint fluid, amniotic fluid, urine, prostatic secretions, semen, pleural secretions, sputum, and bronchoalveolar lavage specimens); tissues; wound aspirates; and swabs of wounds, the throat, nasopharynx, urethra, cervix, or vagina. Blood for culture of genital mycoplasmas should be collected without anticoagulants and immediately inoculated into an appropriate broth culture medium. Mycoplasmas are inhibited by sodium polyethyl sulfonate (SPS), the anticoagulant typically found in commercial blood culture media. This may be overcome by the addition of 1% wt/vol of gelatin. However, commercial blood culture media and automated instruments are not adequate for the detection of Mycoplasma spp. Swab specimens should be obtained without the application of any disinfectants, analgesics, or lubricant. Dacron or polyester swabs on aluminum or plastic shafts should be

CHAPTER 44  Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma

used. Care must be taken when collecting urine samples to prevent contamination with lubricants or antiseptics used during gynecologic examination. Because mycoplasmas have no cell wall, they are highly susceptible to drying; therefore, transport media is necessary, particularly when specimens are collected on swabs. Liquid specimens such as body fluids do not require transport media if inoculated to appropriate media within 1 hour of collection. Tissues should be kept moist; if a delay in processing is anticipated, they should also be placed in transport media. Specific media for the isolation of Mycoplasma spp. include those containing 10% heat-inactivated calf serum containing 0.2 M sucrose in a 0.02 M phosphate buffer, pH 7.2, such as SP4 glucose broth, or Shepard 10B broth. Additional commercial media available for cultivation of these organisms include Stuart medium, trypticase soy broth supplemented with 0.5% bovine serum albumin, Mycotrans (Irvine Scientific, Irvine, CA), and A3B broth (Remel, Inc., Lenexa, KS). Excessive delays in processing can result in decreased viability and recovery of organisms from clinical specimens. If the storage time is expected to exceed 24 hours before cultivation, the samples should be placed in transport media and frozen at −80°C. Frozen samples should be thawed in a hot water bath at 37°C. Transport and storage conditions of various types of specimens are summarized in Table 44.2. 

Direct Detection Methods No direct methods for identifying M. pneumoniae, Ureaplasma spp., or other Mycoplasma spp. in clinical samples are recommended, although some methods have been described, such as immunoblotting and indirect immunofluorescence. Direct detection by Gram staining may rule out the presence of other infectious organisms, but does not informatively stain cell wall–deficient mycoplasmas and ureaplasmas. Acridine orange or a fluorochrome stain may be useful to visualize organisms. However, these are nonspecific stains that will stain nucleic acids in bacteria as well as human cells.

Nucleic Acid Detection Several amplification methods, such as polymerase chain reaction (PCR), have been developed for the detection of the clinically relevant Mycoplasma and Ureaplasma species, including US Food and Drug Administration (FDA)cleared or approved tests to detect M. pneumoniae and M. genitalium. Various targets including 16S ribosomal ribonucleic acid (rRNA) sequences, insertion sequences, and organism-specific genes have been used in the development of these assays. As a result of the fast turnaround time, specificity, and lack of need to cultivate fastidious organisms, PCR amplification for the diagnosis of these organisms is currently the “gold standard.” The Illumigene (Meridian Biosciences, Inc., Cincinnati, OH) ab FDA cleared assay, is a single-target isothermal loop-mediated PCR test that may be used for detection. When considering the use of

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molecular amplification methods for the detection of infectious diseases, it is important to note that, although an organism is detectable, the patient’s signs and symptoms must be correlated with the identified agent. It is possible to detect an organism by one method but not another— in other words, a patient may be PCR positive but culture negative or serologically negative for a Mycoplasma based on the patient’s response to infection and current state of disease manifestation. Chapter 8 provides a more detailed description of the advantages, limitations, and methods used in the development of amplification assays. Multiplex real-time PCR assays that detect M. pneumoniae as well as other atypical respiratory tract pathogens such as Chlamydia pneumoniae and Legionella pneumophila are widely available in routine clinical laboratories. M. genitalium has been detected directly in urine and urethral swabs in males, using PCR methods targeting several unique genomic loci. In females, similar methods are used for detection from vaginal or cervical swabs. After successful implementation of commercial PCR-based diagnostics in Europe and Australia, a similar test for M. genitalium is also FDA cleared and available for use in the United States (Aptiva M. genitalium Assay by Hologic [Marlborough, MA]). Because macrolide resistance is increasing in M. genitalium at an alarming rate, the Australian company SpeeDx (Eveleigh, NSW, Australia) has developed ResistancePlus MG that detects the organism and the five distinct genotypes associated with macrolide resistance. 

Cultivation In general, the medium for mycoplasma isolation contains a beef or soybean protein with serum, fresh yeast extract, and other specific growth factors. As a result of the slow growth of these organisms, the medium must be highly selective to prevent overgrowth of faster-growing organisms that may be present in a clinical sample. Culture media and incubation conditions for these organisms are summarized in Table 44.3. Culture methods for M. pneumoniae, U. urealyticum, and M. hominis are provided in Evolve Procedures 44.1, 44.2, and 44.3, respectively. The quality control of the growth media with a fastidious isolate is of great importance. For the most part, the different metabolic activity of the mycoplasmas for different substrates is used to detect their growth. Glucose (dextrose) is incorporated into media selective for M. pneumoniae, because this mycoplasma ferments glucose to lactic acid; the resulting pH change is then detected by a color change in a dye indicator. Similarly, urea or arginine can be incorporated into media to detect U. urealyticum, U. parvum, and M. hominis, respectively (Table 44.4). If a color change is observed (i.e., a change in pH detected by a chemical indicator added to the culture medium), a 0.1- to 0.2-mL aliquot is immediately subcultured to fresh broth and/or agar medium. In some clinical situations, it may be necessary to provide quantitative information regarding the burden of

582 PA RT I I I    Bacteriology

TABLE 44.2    Transport and Storage Conditions for Mycoplasma and Related Organisms

Specimen Type

Transport Conditions

Transport Media (Examples)a

Storage

Processing hc

Body fluid or liquid specimensb

Within 1 h of collection on ice or at 4°C

Not required

4°C up to 24

Concentrate by high-speed centrifugation and dilute (1:10–1:1000) in broth culture media to remove inhibitory substances and contaminating bacteria; urine should be filtered through a 0.45-μm pore size filter

Swabs

Place immediately into transport media

0.5% albumin in trypticase soy broth modified Stuart 2SP (sugar-phosphate medium with 10% heat-inactivated fetal calf serum) Shepard 10B broth for ureaplasmas SP4 broth for other mycoplasmas and M. pneumoniaed Mycoplasma transport medium (trypticase phosphate broth, 10% bovine serum albumin, 100,000 U of penicillin/milliliter and universal transport media [Copan, Murrieta, CA])

4°C up to 24 hc

None

Tissue

Within 1 h of collection on ice or at 4°C

Not required as long as prevented from drying out

4°C up to 24 hc

Mince (not grind) and dilute (1:10 and 1:100) in transport media

aNot

a complete list. A variety of commercial media is available. blood (see text). cCan be stored indefinitely at −80°C if diluted in transport media after centrifugation. dSP4 broth: sucrose phosphate buffer, 20% horse serum, Mycoplasma base, and neutral red. bExcept

TABLE 44.3    Cultivation of Mycoplasma pneumoniae, Ureaplasma spp., and Mycoplasma hominis

Organism

Media (Examples)

Incubation Conditions

M. pneumoniae

Biphasic SP4 (pH 7.4) Triphasic system (Mycotrim RS, Irvine Scientific, Irvine, CA) PPLOa broth or agar with yeast extract and horse serum Modified New York City medium

Broths: 37°C, ambient air for up to 4 weeks Agars: 37°C, ambient air supplemented with 5%–10% CO2 or anaerobically in 95% N2 plus 5% CO2 All cultures should be retained for 4 weeks before reporting as negative

U. urealyticum/U. parvumb/M. hominisc

A7 or A8 agar medium (Remel, Lenexa, KS); penicillin should be included to minimize bacterial overgrowtha New York City medium Modified New York City medium SP4 glucose broth with arginined SP4 glucose broth with ureae Triphasic system (Mycotrim GU, Irvine Scientific) Shepard 10B broth (or Ureaplasma 10C broth)e

Broths: 37°C, ambient air for up to 7 days Agars: 37°C in 5%–10% CO2 or anaerobically in 95% N2 plus 5% CO2 for 2–5 days Genital cultures should be retained for 7 days before reporting as negative

PPLO, pleuropneumonia-like organisms. aCommercially available. bUses urea and requires acidic medium. cConverts arginine to ornithine and grows over a broad pH range. dFor M. hominis isolation. eFor U. urealyticum isolation.

CHAPTER 44  Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma

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TABLE   Basic Biochemical Differentiation of the 44.4  Major Mycoplasma and Ureaplasma spp.

Glucose Metabolism

Arginine Metabolism

Urease

M. fermentans

Positive

Positive

Negative

M. genitalium

Positive

Negative

Negative

M. hominis

Negative

Positive

Negative

M. pneumoniae

Positive

Negative

Negative

U. parvum

Negative

Negative

Positive

U. urealyticum

Negative

Negative

Positive

Organism

•

Fig. 44.2  Colonies of Mycoplasma pneumoniae visualized under 100× magnification. Note the variation in the size of the colonies (arrows). (Courtesy Clinical Microbiology Laboratory, SUNY Upstate Medical University, Syracuse, NY.)

mycoplasmas in a clinical specimen. For example, quantitation of specimens taken at different stages during urination or after prostatic massage can help to determine the location of mycoplasmal infection in the genitourinary tract.

Approach to Identification On agar, M. pneumoniae will appear as spherical, grainy, yellowish forms that are embedded in the agar, with a thin outer layer similar to those shown in Fig. 44.2. The agar surface is examined under 20× to 60× magnification using a stereomicroscope daily for Ureaplasma spp., at 24 to 72 hours for M. hominis, and every 3 to 5 days for M. pneumoniae and other slow-growing species. Because only M. pneumoniae and one serovar of U. urealyticum hemadsorb, M. pneumoniae is definitively identified by overlaying suspicious colonies with 0.5% guinea pig erythrocytes in phosphate-buffered saline. After 20 to 30 minutes at room temperature, colonies are observed for adherence of red blood cells.

B

A

• Fig. 44.3  Isolation of Mycoplasma hominis and Ureaplasma urealyti-

cum (100× magnification). Note the “fried egg” appearance of the large M. hominis colony (arrow A) and the relatively small size of the U. urealyticum colony (arrow B). (Courtesy Clinical Microbiology Laboratory, SUNY Upstate Medical University, Syracuse, NY.)

Cultures for the genital mycoplasmas are handled in a similar fashion, including culture examination and the requirement for subculturing. Colonies may be definitively identified on A8 agar (Hardy Diagnostics, Santa Maria CA.) as U. urealyticum by urease production in the presence of a calcium chloride indicator. U. urealyticum colonies (15 to 60 μm in diameter) will appear as dark brownish clumps. Colonies that are typical in appearance for U. urealyticum are shown in Fig. 44.3. M. hominis colonies are large (approximately 20 to 300 μm in diameter) and are urease negative (Fig. 44.3), with a characteristic “fried egg” appearance (Fig. 44.4). Similar fried egg colonies are produced by M. genitalium but typically take several weeks to develop and thus are not useful for diagnostic purposes. On conventional blood agar, strains of M. hominis, but not of U. urealyticum, produce nonhemolytic, pinpoint colonies that do not Gram stain. These colonies can be stained with the Dienes or acridine orange stains. Numerous transport and growth media systems for the detection, quantitation, identification, and antimicrobial susceptibility testing of the genital mycoplasmas are commercially available in the United States and Europe. 

Serodiagnosis Laboratory diagnosis of M. pneumoniae is usually made serologically. Nonspecific production of cold agglutinins occurs in approximately half of patients with atypical pneumonia caused by this organism. Antibodies to M. pneumoniae are typically detectable after approximately 1 week of illness, peaking between 3 to 6 weeks, followed by a gradual decline. The antibody response to M. pneumoniae varies greatly from patient to patient. Some patients fail to produce a detectable immunoglobulin M (IgM) level, whereas in others the IgM level will persist

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Petri dish Agar medium

Surface view “fried egg” appearance

Agar, side view Central zone embeds into the agar below the surface

•

Fig. 44.4  Colonial growth characteristics of Mycoplasma in agar medium.

for months. The variability associated with the antibody response necessitates the comparison of paired sera for proper diagnosis. In addition, cold agglutinins form in association with M. pneumoniae infection. The most widely used serologic tests are enzyme-linked immunosorbent assay (ELISA) tests, although indirect fluorescent antibody tests have been used with some success. IgM-specific tests such as the Immuno Card (Meridian Diagnostics, Cincinnati, OH) are commercially available, and a single positive result in children, adolescents, and young adults may be considered diagnostic in some cases. In addition, commercially available, membranebased assay that simultaneously detects IgM and IgG against M. pneumoniae (Remel EIA, Lenexa, KS) has demonstrated good sensitivity and specificity compared with other tests. Several additional commercial assays are available that include enzyme immunoassay (EIA) microtiter assays. Although serologic tests such as indirect hemagglutination and metabolism inhibition for genital mycoplasmas are available, they are rarely used. Because of the antigenic complexity of the mycoplasmas, and serologic cross reactivity among the outer membrane antigens, the development of a specific and useful serologic assay is a challenge. 

Susceptibility Testing and Therapy Although agar and broth dilution methods may be used to determine antibiotic susceptibilities, the complex growth requirements of mycoplasmas have restricted their performance to a few laboratories. The Human Mycoplasma Susceptibility Testing Subcommittee of the Clinical and Laboratory Standards Institute has formulated agar and

broth dilution methods. Most mycoplasmal infections are treated empirically. Most M. pneumoniae infections are self-limited and usually do not require treatment, but antibiotic treatment can markedly reduce or shorten the disease process. Because of the lack of a cell wall, M. pneumoniae and the other Mollicutes are innately resistant to all beta-lactam antibiotics, because this class of drugs interferes with cell wall synthesis. In addition, they are resistant to sulfonamides, trimethoprim, and rifampin. Susceptibility patterns vary by species to macrolides and lincosamides. M. pneumoniae and M. genitalium have historically been susceptible to the macrolides, tetracycline, ketolides, and fluoroquinolones. Although macrolides are still an appropriate first-line therapy for M. pneumoniae, they are rapidly becoming ineffective against predominant strains of M. genitalium. Singledose macrolides (i.e., azithromycin) are beneficial for treating gonococcal and nongonococcal urethritis in sexually transmitted disease clinics, because the success of extended dosing regimens relies on patient compliance, which is often unreliable in high-risk clinical settings. Although this was initially a successful intervention against M. genitalium infection, the organism’s facultative intracellular location affords it protection against macrolides during high-dose treatment, resulting in incomplete clearance, disease recrudescence, and increasing rates of macrolide resistance. Given the factors that single-dose macrolide treatment successfully addressed, it is predictable that outpatient treatment of M. genitalium will pose a unique challenge. Unfortunately, the susceptibility of M. hominis and U. urealyticum to various agents is not as predictable. For the most part, the tetracyclines are the drugs of choice for these agents, although resistance has been reported. Multidrug-resistant mycoplasmas and ureaplasmas have been identified in genital and extragenital infections in immunocompromised patients. Treatment and clearance of these infections is extremely difficult and limited by the bacteriostatic concentrations of antimicrobials, as well as the slow growth and immune modulation associated with infections with these agents. 

Prevention No vaccines are available for the human mycoplasmas. Early studies both in human subjects and animal models have proven either ineffective or deleterious upon challenge, likely due to the inflammatory, immunomodulatory nature of the organisms. Urogenital transmission of mycoplasmas and ureaplasmas can be prevented through barrier protection, such as with the use of male condoms.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

CHAPTER 44  Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma

Bibliography Ainsworth JG, Katseni V, Hourshid S, et al.: Mycoplasma fermentans and HIV-associated nephropathy, J Infect 29:323–326, 1994. Bauer FA, Wear DJ, Angritt P, Lo SC: Mycoplasma fermentans (incognitus strain) infection in the kidneys of patients with acquired immunodeficiency syndrome and associated nephropathy: a light microscopic, immunohistochemical, and ultrastructural study, Hum Pathol 22:63–69, 1991. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Bharat A, Cunningham SA, Scott Budinger GR, et al.: Disseminated Ureaplasma infection as a cause of fatal hyperammonemia in humans, Sci Transl Med 7(284):284re3, 2015. https://doi. org/10.1126/scitranslmed.aaa8419. Blanchard A: Mycoplasmas and HIV infection, a possible interaction through immune activation, Wien Klin Wochenschr 109(14– 15):590–593, 1997. Browning GF, Whithear KG, Geary SJ: Vaccines to control mycoplasmosis. In Mycoplasmas: molecular biology, pathogenicity, and strategies for control, Norfolk, NR, United Kingdom, 2005, Horizon Bioscience. Caroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Dessì D, Delogu G, Emonte E, Catania MR, Fiori PL, Rappelli P: Long-term survival and intracellular replication of Mycoplasma hominis in Trichomonas vaginalis cells: potential role of the protozoon in transmitting bacterial infection, Infect Immun 73(2):1180– 1186, 2005. Gaydos CA: Mycoplasma genitalium: accurate diagnosis is necessary for adequate treatment, J Infect Dis 216(Suppl 2):S406–S411, 2017. Goldenberg RL, Thompson C: The infectious origins of stillbirth, Am J Obstet Gynecol 189:861–873, 2003. Loens K, Ursi D, Goossens H, Ieven M: Molecular diagnosis of Mycoplasma pneumoniae respiratory tract infections, J Clin Microbiol 4:4915–4923, 2003.

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May M, Balish MF, Blanchard A: The order Mycoplasmatales in the prokaryotes, New York, NY, 2014, Springer Inc. Mena L, Wang X, Mroczkowski TF, Martin DH: Mycoplasma genitalium infections in asymptomatic men and men with urethritis attending a sexually transmitted diseases clinic in New Orleans, Clin Infect Dis 35:1167–1173, 2001. Montagnier L, Blanchard A: Mycoplasmas as cofactors in infection due to the human immunodeficiency virus, Clin Infect Dis 17(Suppl 1):S309–S315, 1993. Razin S, Yogev D, Naot Y: Molecular biology and pathogenicity of mycoplasmas, Microbiol Mol Biol Rev 62:1094–1156, 1998. Rittenschober-Böhm J, Waldhoer T, Schulz SM, et  al.: Vaginal Ureaplasma parvum serovars and spontaneous preterm birth, Am J Obstet Gynecol 220(6):594.e1–594.e9, 2019. https://doi. org/10.1016/j.ajog.2019.01.237. Sanchez PJ: Perinatal transmission of Ureaplasma urealyticum: current concepts based on review of the literature, Clin Infect Dis 17(Suppl 1):S107–S111, 1993. Silwedel C, Speer CP, Glaser K: Ureaplasma-associated prenatal, perinatal, and neonatal morbidities, Expert Rev Clin Immunol 13(11):1073–1087, 2017. Totten PA, Schwartz MA, Sjöström KE, et  al.: Association of Mycoplasma genitalium with nongonococcal urethritis in heterosexual men, J Infect Dis 183:269–276, 2001. Waites KB, Bebear CM, Robertson JA, et  al.: Cumitech 34, laboratory diagnosis of mycoplasmal infections, Washington, DC, 2001, American Society for Microbiology. Waites KB, Talkington DF: Mycoplasma pneumoniae and its role as a human pathogen, Clin Microbiol Rev 17:697–728, 2004. Wang PJ, Xie CB: Mycoplasma hominis symbiosis and Trichomonas vaginalis metronidazole resistance, Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi 30(3):210–213, 2012.

Chapter Review 1. Why are Mycoplasmataceae difficult to cultivate in the clinical laboratory? a. They lack cell walls. b. They require serum components including sterols in the growth medium. c. They are extremely sensitive to pH changes in the environment. d. All of the above 2. Mycoplasmas generally may be transmitted by all of the following except: a. Direct sexual contact b. Vertical transmission from mother to fetus c. Oral sexual practices d. A contaminated toothbrush 3. All of the following are considered commensal organisms capable of causing opportunistic infections except: a. U. urealyticum b. M. fermentans c. M. genitalium d. M. hominis e. M. faucium f. U. parvum 4.  Which of the following would be the most reliable serologic diagnosis for M. pneumoniae, considering the pathogenesis and the immunologic response and based on the descriptions provided? a. A single acute immunoglobulin M (IgM) titer b. A single elevated acute IgM titer c. A negative IgM titer followed by two successive IgG titers demonstrating an increase in titer d. A classic paired sera with a definitive fourfold increase in titer 5. A genital swab was submitted for culture. U. urealyticum was isolated. The suspected biochemical profile to confirm identification would be: a. glucose +, arginine +, urease − b. glucose −, arginine +, urease + c. glucose −, arginine −, urease + d. glucose +, arginine −, urease − 6. A 21-year-old male presented to his family physician complaining of flulike symptoms, including a lowgrade fever and body aches. A complete blood count was drawn, and the patient exhibited no elevation in the white blood cell count. Serology was negative for influenza. Chest x-ray indicated lobar consolidation representative of a respiratory infection. Based on the laboratory results and patient presentation, what is the most likely cause? a. Influenza A b. Klebsiella pneumoniae c. Streptococcus pneumoniae d. Mycoplasma pneumoniae e. Pseudomonas aeruginosa

7. A patient presented to his doctor around 1 p.m., complaining of frequent urination and abdominal pain. A urine sample was collected by the patient at 8 a.m. and submitted for culture. The routine urinalysis indicated a positive leukocyte esterase test. All other tests were normal. Polymerase chain reaction (PCR) was negative for Ureaplasma spp. After 24 hours of incubation, routine urine culture demonstrated the growth of three different organisms with no significant pathogens noted. What is the most likely cause for the conflicting laboratory results? a. The original urine sample was not inoculated onto proper growth media. b. The PCR test was likely contaminated. c. The sample delay in processing of the urine affected the viability of the organism. d. Both A and C e. All of the above 8. True or False _____ All Mycoplasma spp. are capable of immune evasion through intracellular growth cycles. _____ Sodium polyanethol sulfonate (SPS) in commercial blood culture media inhibits the growth of Mycoplasma spp. _____ Specimens collected for the isolation of Mycoplasma or Ureaplasma should be placed in transport media and incubated at 37°C until processed. _____ Current Clinical and Laboratory Standards Institute (CLSI) standards recommend broth or agar dilution methods for susceptibility testing for Mycoplasma and Ureaplasma species. 9. Matching: Match each term with the appropriate description. ____ M. genitalium ____ M. fermentans ____ U. urealyticum ____ M. pneumoniae ____ M. hominis

a.  walking pneumonia b. crosses placenta after

colonization of the urogenital tract c.  nongonococcal urethritis, cervicitis, and pelvic inflammatory disease (PID) d. arthritis associated with agammaglobulinemia e.  infections in immunocompromised patients

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PROCEDURE 44.1

Isolation of Mycoplasma pneumoniae Principle Mycoplasma pneumoniae is primarily a respiratory tract pathogen that is extremely fastidious. Because this bacterial agent lacks a cell wall and has strict nutritional requirements, the recovery of this organism from clinical specimens requires special conditions in terms of both culture media and incubation conditions. 

Method 1. Prepare biphasic SP4 culture media as follows: A. Combine the ingredients for the media base. Mycoplasma broth base (BD), 3.5 g Tryptone (Difco), 10 g Bacto-Peptone (Difco), 5 g (50% aqueous solution, filter sterilized), 10 mL Distilled water, 615 mL B. Stir the solids into boiling water to dissolve them and adjust the pH to 7.4 to 7.6. Autoclave according to the manufacturer’s instructions. Cool to 56°C before adding the following supplements for each 625 mL of base to make a final volume of 1 L: CMRL 1066 tissue culture medium with glutamine, 10× (GIBCO), 50 mL Aqueous yeast extract (prepared as described in New York City Medium), 35 mL Yeastolate (Difco; 2% solution), 100 mL Fetal bovine serum (heat inactivated at 56°C for 30 min), 170 mL Penicillin G sodium, 1000 IU/mL Amphotericin B, 0.5 g Polymyxin B, 500,000 U C. To prepare the agar necessary for making biphasic media, add 8.5 g Noble agar (Difco) to the basal medium ingredients before adding the supplements. D. Dispense 1 mL of SP4 agar aseptically into the bottom of sterile 4-mL screw-capped vials. Allow the agar to set and dispense 2 mL of SP4 broth above the agar layer in each vial. Seal caps tightly and store at −20°C.

2. Place 0.1 to 0.2 mL of liquid specimen or dip and twirl a specimen received on a swab in a vial of biphasic SP4 culture medium. After expressing as much fluid as possible from the swab, remove the swab to prevent contamination. 3. Seal the vial tightly and incubate in air at 35°C for up to 3 weeks. 4. Inspect the vial daily. During the first 5 days, a change in pH, indicated by a color shift from orange to yellow or violet, or increased turbidity is a sign that the culture is contaminated and should be discarded. 5. If either a slight acid pH shift (yellow color) with no increase in turbidity or no change occurs after 7 days’ incubation, subculture several drops of the broth culture to agar. Continue to incubate the original broth. 6. If broth that exhibited no changes at 7 days shows a slight acid pH shift at any time, subculture to agar as noted earlier. Broths that show no change at 3 weeks are subcultured to agar. 7. Incubate the agar plates in a very moist atmosphere with 5% to 10% CO2 at 35°C for 7 days. 8. Observe the agar surface under 20× to 60× magnification using a stereomicroscope after 5 days for colonies, which appear as spherical, grainy, yellowish forms, embedded in the agar, with a thin outer layer (Fig. 44.2). 9. Definitive identification of M. pneumoniae is accomplished by overlaying agar plates showing suspicious colonies with 5% sheep or guinea pig erythrocytes in 1% agar prepared in physiologic saline (0.85% NaCl) instead of water. The 1% agar is melted and cooled to 50°C, the blood cells are added, and a thin layer is poured over the original agar surface. 10. Reincubate the plate for 24 h, and observe for beta hemolysis around colonies of M. pneumoniae caused by production of hydrogen peroxide. Additional incubation at room temperature overnight enhances the hemolysis. No other species of Mycoplasma produces this reaction.   

CHAPTER 44  Cell Wall–Deficient Bacteria: Mycoplasma and Ureaplasma

PROCEDURE 44.2 ISOLATION OF UREAPLASMA UREALYTICUM AND UREAPLASMA PARVUM

PROCEDURE 44.3 ISOLATION OF MYCOPLASMA HOMINIS

Principle

Principle Similar to other mycoplasmas, Ureaplasma spp. require special culture conditions to facilitate recovery of this bacterial agent from clinical specimens. 

Similar to the other mycoplasmas, the successful recovery of Mycoplasma hominis by culture from clinical specimens requires special conditions. 

Method A

Method 1. Inoculate one Ureaplasma agar plate and one Ureaplasma broth each with a 0.1 mL specimen from transport medium. 2. Incubate broth in tightly sealed test tubes for 5 days. Observe twice daily for a color change in the broth to red, with no increase in turbidity. If color change occurs, immediately transfer one loopful to a Ureaplasma agar plate and streak for isolated colonies. 3. Agar plates are incubated in a candle jar or, optimally, in an anaerobic environment at 35°C. Colonies appear on agar within 48 h. Plates are inspected in the same way as described for Mycoplasma pneumoniae (Procedure 44.1). Ureaplasma colonies appear as small, granular, yellowish spheres. 4. To definitively identify colonies on Ureaplasma agar after 48 hours’ incubation, pour a solution of 1% urea and 0.8% MnCl2 in distilled water over the agar surface. Ureaplasma urealyticum stains dark brown because of production of urease (Fig. 44.3).   

1. Inoculate one M. hominis agar plate and two M. hominis broth tubes, one broth containing phenol red indicator and one without the possibly inhibitory phenol red, each with a 0.1-mL specimen from transport media. 2. Incubate broths in tightly sealed test tubes for 5 days. If the phenol red–containing broth changes color to red or violet, both broths are subcultured to M. hominis agar. After 48 h of incubation, transfer 0.1 mL or a loopful of broth from tubes that exhibited no change or only a slight increase in turbidity to M. hominis agar and streak for isolated colonies. 3. M  . hominis agar plates are incubated in the same manner as Ureaplasma cultures. Plates should be observed daily for up to 5 days for colonies. 

Method B (Alternative Method) 1. Inoculate specimen onto a prereduced colistin-nalidixic acid (CNA) sheep blood agar or anaerobic blood plate and incubate anaerobically for 48–72 h. 2. Examine for pinpoint colonies that show no bacteria on Gram stain. 3. Streak suspicious colonies to M. hominis agar and incubate as in step 3.   

CASE STUDY 44.1 A 29-year-old previously healthy female presented with a productive cough, fever of 102°F, and severe headache. She had cervical adenopathy (swollen glands), although she had a nonerythematous throat with no exudate. Chest examination showed crackles bilaterally at the lung base with decreased breath sounds diffusely. This finding was confirmed by chest film that showed bilateral multifocal areas of patchy consolidation. Her neck was not stiff, but because of the severity of the headache, she was admitted to the neurologic service. Spinal fluid was obtained and was negative for bacteria upon Gram staining, Cryptococcus spp. negative using an India ink preparation and cryptococcal antigen, and negative for the presence of acid-fast organisms such as Mycobacterium spp. No pathogens were isolated from blood or sputum cultures. The patient did not improve on ceftriaxone. On day 3, she was prescribed ciprofloxacin. The patient gradually improved, although the headache, photophobia, and cough continued for some time.

Questions 1. What is the etiologic agent of disease in this case? 2. Can you explain why the bacterial cultures were negative? 3. Why is ciprofloxacin an effective therapy for Mycoplasma pneumoniae but ceftriaxone is not?   

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The Spirochetes OBJECTIVES 1. Describe the bacterial agents discussed in this chapter in terms of morphology, taxonomy, and growth conditions. 2. Identify the four stages of syphilis (i.e., primary, secondary, latent, and tertiary), according to clinical symptoms, antibody production, transmission, infectivity, and treatment. 3. Explain congenital syphilis, including transmission and clinical manifestations. 4. Define reagin, cardiolipin, and biologic false positive. 5. Differentiate reagin and treponemal antibodies, including specificity and association with disease. 6. Identify the various serologic methods that use specific treponemal or nonspecific nontreponemal antigens. 7. Describe the basic principles for the RPR, VDRL, FTA-ABS, TP-PA, PaGIA, MHA-TP, EIA, CIA, and MBIA assays. 8. Compare Borrelia spp. to the other spirochetes discussed in this chapter, including morphology and growth conditions. 9. Describe the pathogenesis for and diagnosis of relapsing fever and Lyme disease, including the routes of transmission, vector, and disease presentation. 10. Explain the methodology and clinical significance for using a two-step diagnostic procedure for Borrelia spp. infections. 11. Compare and contrast the standard two-tier testing algorithm (STTT) with the modified two-tier testing algorithm (MTTT) for Lyme disease. 12. Describe the pathogenesis associated with leptospirosis, including the two major stages of the disease and the recommended clinical specimens. 13. Describe Brachyspira spp., including potential patho­ genesis, appropriate specimen, transmission, and clinical ­significance. 14. Correlate patient signs and symptoms with laboratory data to identify the most likely etiologic agent.

GENERA AND SPECIES TO BE CONSIDERED Treponema pallidum subsp. pallidum Treponema pallidum subsp. pertenue Treponema pallidum subsp. endemicum Treponema carateum Treponema denticola

Lyme Borreliosis Borrelia afzelii Borrelia bavariensis

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Borrelia bissettiae Borrelia burgdorferi sensu stricto Borrelia garinii Borrelia lusitaniae Borrelia mayonii Borrelia spielmanii Borrelia valaisiana 

Relapsing Fever Borrelia caucasica Borrelia crocidurae Borrelia duttonii Borrelia hermsii Borrelia hispanica Borrelia mazzottii Borrelia miyamotoi (hard tick-borne relapsing fever) Borrelia parkeri Borrelia persica Borrelia recurrentis Borrelia turicatae Borrelia venezuelensis Brachyspira aalborgi Brachyspira hominis (provisionally named) Brachyspira pilosicoli

Pathogenic Species Leptospira alexanderi Leptospira alstonii Leptospira borgpetersenii Leptospira interrogans Leptospira kirschneri Leptospira mayottensis Leptospira noguchii Leptospira santarosai Leptospira weilii 

Intermediate Species Leptospira broomii Leptospira fainei Leptospira inadai Leptospira licerasiae Leptospira venezuelensis Leptospira wolffii 

Saprophytic Species Leptospira biflexa Leptospira idonii Leptospira meyeri Leptospira terpstrae Leptospira vanthielii Leptospira wolbachii Leptospira yanagawae

CHAPTER 45  The Spirochetes

This chapter discusses the bacteria that belong to the phylum Spirochaetes. The phylum includes three orders, which include four medically relevant genera: the order Brachyspirales (Brachyspira spp.), the order Leptospirales (Leptospira spp.), and the order Spirochaetales. Within the order Spirochaetales, there are two families: Borreliaceae (Borrelia spp.) and Spirochaetaceae (Treponema spp.). The spirochetes are all long, slender, helically curved organisms, with the unusual morphologic features of axial fibrils and an outer sheath. These fibrils, or axial filaments, are flagella-like organelles that wrap around the bacteria’s cell walls, are enclosed within the outer sheath, and facilitate motility of the organisms. The fibrils are attached in the cell wall by platelike structures, called insertion disks, located near the ends of the cells. The protoplasmic cylinder gyrates around the fibrils, causing bacterial movement to appear as a corkscrewlike winding. Differentiation of genera (Treponema) within the family Spirochaetaceae is based on the number of axial fibrils (endoflagella), the number of insertion disks present (Table 45.1), and biochemical and metabolic features. The spirochetes also fall into genera based loosely on their morphology (Fig. 45.1): Treponema appear as slender with tight coils; Borrelia (family Borreliaceae) are somewhat thicker with fewer and looser coils; and Leptospira (family Leptospiraceae) resemble Borrelia, except for their hooked ends. Brachyspira (family Brachyspiraceae) are comma-shaped or helical, with tapered ends with four flagella at each end.

Treponema General Characteristics The major pathogens in the genus Treponema—Treponema pallidum subsp. pallidum, Treponema pallidum subsp. pertenue, Treponema pallidum subsp. endemicum, and Treponema carateum—infect humans and have not been cultivated for more than one passage in vitro. Most species stain poorly with Gram staining or Giemsa methods and are best observed with the use of dark-field or phase-contrast microscopy. The organisms are microaerophilic. Other treponemes such as Treponema vincentii, T. denticola, T. refringens, T. socranskii, T. parvum, T. pectinovorum, T. putidum, T. lecithinolyticum, T. amylovorum, T. medium, and T. maltophilum are normal microbiota of the oral cavity or the human genital tract. These organisms are cultivable anaerobically on artificial media. Acute necrotizing ulcerative gingivitis, also known as Vincent disease, is a

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destructive lesion of the gums. Methylene blue–stained material from the lesions of patients with Vincent disease shows certain morphologic types of bacteria. Observed morphologies include spirochetes and fusiform; oral spirochetes, particularly unusually large ones, may be important in this disease, along with other anaerobes. 

Epidemiology and Pathogenesis Key features of the epidemiology of diseases caused by the pathogenic treponemes are summarized in Table 45.2. In general, these organisms enter the host by either penetrating intact mucous membranes (as is the case for T. pallidum subsp. pallidum—hereafter referred to as T. pallidum) or entering through breaks in the skin. T. pallidum is transmitted by sexual contact and vertically from mother to the unborn fetus. Neonates can also become infected from contact with contaminated lesions or infected maternal blood during the birthing process. Although rare in the United States, the transmission of T. pallidum in blood products still occurs in underdeveloped countries that do not have routine donor screening and processing facilities. After T. pallidum penetrates the host, the organism subsequently invades the bloodstream and spreads to other body sites. The mechanisms associated with disease pathology to the host are unclear. Molecular studies have not identified a T.  ­pallidum protein that binds to host fibronectin and ­laminin receptors on various human cells. Binding to different host cells is apparent in cells that produce fibronectin. The organism has an inherent ability to cross the endothelial, blood-brain, and placental barriers. T. pallidum has been shown to activate platelets to aid in movement and dissemination throughout the host. As a result, T. pallidum has a remarkable tropism (attraction) to arterioles; infection ultimately leads to endarteritis (inflammation of the lining of arteries) and subsequent progressive tissue destruction.  Borrelia and Brachyspira Leptospira

Treponema

• Fig. 45.1  Species designation of spirochetes based on morphology.

TABLE 45.1    Spirochetes Pathogenic for Humans

Genus

Axial Filaments

Insertion Disks

Treponema

6–10

1

Borrelia

30–40

2

Leptospira

2

3–5

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TABLE 45.2    Epidemiology and Spectrum of Disease of the Treponemes Pathogenic for Humans

Agent

Transmission

Treponema pallidum subsp. pallidum

Sexual contact or congenital (mother to fetus).

Treponema pallidum subsp. pertenue

Traumatized skin contact with an infected lesion (person-to-­ person contact).

Geographic Location

Disease

Clinical Manifestationsa

Worldwide

Venereal syphilisb

Refer to text in this chapter

All ages

Humid, warm climates: Africa, South and Central America, Pacific Islands

Yaws

Skin—papules,b ­nodules, ulcers

Children

Age Group

Primary lesion (mother yaw), disseminated lesions (frambesia) May progress to a latent stage and late infection involving destructive lesions to bone and cartilage

Treponema pallidum subsp. endemicum

Mouth to mouth by utensils (personto-person contact).

Arid, warm climates: North Africa, Southeast Asia, Middle East

Endemic nonvenereal syphilis

Skin/mucous membrane patches, papules, macules, ulcers, scarsb

Children or adults; rarely congenital

May progress to disseminated oropharyngeal with generalized lymphadenopathy May demonstrate a latent stage, and late syphilis destructive to skin, bone, and cartilage Treponema carateum

Traumatized skin contact with an infected lesion (person-toperson contact).

Semiarid, warm climates: Central and South America, Mexico

Pinta

Skin papules, macules. Hyperkeratotic pigmented may lead to disseminated skin lesions and lymphadenopathy; late stage may result in pigmentary changes in skin (hyper- or hypopigmentation)

All ages but primarily children and adolescents

aAll bIf

diseases have a relapsing clinical course and prominent cutaneous manifestations. untreated, organisms can disseminate to other parts of the body, such as bone.

Spectrum of Disease In recent years, there has been a resurgence of primary and secondary syphilis cases in the United States, with a 10.5% increase in the number of reported cases between 2016 and 2017. T. pallidum causes venereal (transmitted through sexual contact) syphilis. The clinical presentation of venereal syphilis is varied and complex, often mimicking many other diseases. The disease is divided into stages: incubating, primary, secondary, early nonprimary nonsecondary syphilis, unknown duration or late syphilis, and tertiary.

Primary syphilis is characterized by the appearance of one or more hard chancres (painless ulcers) that develop(s) at the site of inoculation, most commonly the genitalia. The primary stage is extremely infectious because the lesion contains a large number of organisms. Disease at this stage may be undetectable in the vagina or rectum. Within 3 to 6 weeks, the chancre heals spontaneously (without treatment). Dissemination of the organism occurs during this primary stage; once the organism has reached a sufficient number (usually within 4 to 10 weeks), clinical manifestations of secondary syphilis become apparent.

CHAPTER 45  The Spirochetes

During secondary syphilis, the patient usually seeks medical attention. Systemic symptoms such as fever, weight loss, malaise, and loss of appetite are present in about half of patients. Common symptoms are flulike (fever, sore throat, and lymphadenopathy). The skin is the organ most commonly affected in secondary syphilis, with patients having a widespread rash (generally on the face, scalp, palms of hands, and soles of the feet) and generalized lymphadenopathy. Patchy hair loss may occur, such as a loss of eyebrows (“moth-eaten” area). This stage is a highly infectious state, again because large numbers of spirochetes are present. Aseptic meningitis may also occur. This stage may be mild and go unnoticed by the patient, or symptoms may disappear without treatment. This is when the disease becomes subclinical (asymptomatic) but not necessarily dormant (inactive). During this period, diagnosis can be made using serologic methods. Relapses are common during early, nonprimary, nonsecondary (≤1 year) syphilis. Unknown duration or late latent syphilis (≥1 year) is usually asymptomatic and noninfectious. Many untreated cases develop into tertiary syphilis. Tertiary syphilis is the tissue-destructive phase that appears 10 to 25 years after the initial infection in up to 35% of untreated patients. Complications of syphilis at this stage include central nervous system disease (neurosyphilis) and cardiovascular abnormalities (cardiovascular syphilis), such as aortic valve insufficiency associated with the presence of cardiovascular lesions, eye disease (ocular syphilis), and granuloma-like lesions (gummas) that are soft, painless, and noninfectious, and found on the skin or in the bones or visceral organs. Neurosyphilis has been categorized into five major clinical presentations: asymptomatic, meningeal, meningovascular, parenchymatous, and gummatous. As seen with primary and secondary syphilis, there has also been an increase in the number of reported congenital syphilis cases in recent years; cases of congenital syphilis have increased every year in the United States since 2013. Congenital syphilis is transmitted from a mother to an unborn fetus during any stage of infection but is most often associated with early syphilis. The unborn fetus may develop an asymptomatic infection or symptomatic infection. Clinical signs known as Hutchinson’s triad (deafness, blindness, notched peg-shaped teeth) may occur. Additionally, poor bone formation may result, such as “saber shin” bowing of the tibia and the “bulldog” appearance of a deformed maxilla. Finally, neurosyphilis or neonatal death can occur. The additional pathogenic treponemes are major health concerns in developing countries. Although morphologically and antigenically similar, these agents differ epidemiologically and with respect to their clinical presentation from T. pallidum. The diseases caused by these treponemes are summarized in Table 45.2. 

Laboratory Diagnosis Specimen Collection Samples collected from ulcers and lesions should not be contaminated with blood, microorganisms, or tissue debris.

589

The site should be cleansed with sterile gauze moistened with saline. The sample should be placed on a clean glass slide and cover slipped. Polymerase chain reaction (PCR) samples should be collected on a sterile Dacron or cotton swab and placed in a cryotube containing nucleic acid transport medium or universal transport medium. Tissue or needle aspirates of lymph nodes should be placed in 10% buffered formalin at room temperature. To test for congenital syphilis, a small section of the umbilical cord is collected and fixed in 10% buffered formalin at room temperature until processed. A 3 to 4 cm section of umbilical cord distal from the placenta for the detection of congenital syphilis should be collected immediately following delivery and processed as a tissue. Serum is the specimen of choice for serology; however, plasma may be used in some assays. Plasma should be tested within 24 hours to avoid false-positive results. Capillary draws of whole blood, serum, or plasma may be used for rapid syphilis tests. Maternal serum may also be used to screen for congenital syphilis. Infants’ serum should be used for immunoglobulin (Ig)M specific tests because cord blood specimens may be contaminated with maternal blood. Serum, plasma, and cerebrospinal fluid (CSF) should be stored at 4°C if testing is delayed more than 4 hours and at –20°C if delayed more than 5 days. Samples collected for PCR, such as unfixed tissue, ulcer exudate, mucosal and skin lesions, CSF or amniotic fluid, and whole blood in EDTA, should be stored at −80°C if testing is delayed. 

Direct Detection Treponemes can be detected in material taken from skin lesions by dark-field examination or fluorescent antibody staining and microscopic examination. Material for microscopic examination is collected from suspicious lesions. The area around the lesion must first be cleansed with a sterile gauze pad moistened in saline. The surface of the ulcer is then abraded until some blood is expressed. After blotting the lesion until there is no further bleeding, the area is squeezed until serous fluid is expressed. Dark-field microscopy should be completed within 20 minutes of collection in order to identify motile treponemes. For other microscopic techniques, the surface of a clean glass slide is touched to the exudate, allowed to air dry, and transported in a dust-free container for fluorescent antibody staining. A T. pallidum fluorescein-labeled antibody is commercially available for staining. For dark-field examination, the expressed fluid is aspirated using a sterile pipette, dropped onto a clean glass slide, and cover slipped. The slide containing material for dark-field examination must be transported to the laboratory immediately. Because positive lesions may be teeming with viable spirochetes that are highly infectious, all supplies and patient specimens must be handled with extreme caution and carefully discarded as required for contaminated materials. Gloves should always be worn. Material for dark-field examination is examined under 400× high-dry magnification for the presence of motile spirochetes. Treponemes are long (8 to 10 μm, slightly larger

590 PA RT I I I    Bacteriology

than a red blood cell) and consist of 8 to 14 tightly coiled, even spirals (Fig. 45.2). Once seen, characteristic forms should be verified by examination under oil immersion magnification (1000×). Although the dark-field examination depends greatly on technical expertise and the numbers of organisms in the lesion, it can be highly specific when performed on genital lesions. Dark-field methods cannot be used to evaluate oral or rectal lesions. Lesion exudates or tissue samples may be used for direct fluorescent antibody detection for T. pallidum (DFA-TP). DFA-TP visualizes specimens on slides with fluorescein isothiocyanate (FITC) labeled antibodies. Polyclonal and monoclonal antibodies may be used; however, the US Food and Drug Administration (FDA) has not approved this test. Dark-field microscopy and fluorescent antibody methods are insensitive. Dark-field microscopy is no longer performed in most clinical laboratories. 

Nucleic Acid Detection Although nucleic acid–based assays are not currently available within many clinical laboratories, several methods have been developed using PCR for the detection of T. pallidum. These methods are primarily useful in the identification of organisms within exudate or lesions, and are sensitive and specific when used to analyze genital lesions. Commercially available extraction kits can be used, such as the QIAamp DNA mini kit (Qiagen, Inc., Valencia, CA). Although there are currently no FDA-approved nucleic acid tests available for the detection of T. pallidum, the Center for Disease Control Laboratory Reference and Research Branch, Division of Sexually Transmitted Disease Prevention, has developed a multiplex PCR assay. The TaqMan-based assay simultaneously detects T. pallidum, Haemophilus ducreyi, and herpes simplex viruses (HSV1 and HSV2). In addition, laboratory-developed tests have been shown to detect atypical cases of syphilis in tonsillar, vertebral, and ocular syphilis. However, a major limitation of PCR is that the sensitivity of the test begins to decrease during secondary syphilis. 

Serodiagnosis Serologic tests for treponematosis measure the presence of two types of antibodies: treponemal and nontreponemal. Treponemal antibodies are produced against antigens of the organisms themselves, whereas nontreponemal antibodies, often referred to as reagin antibodies, are produced in infected patients against components of mammalian cells. Reaginic antibodies, although almost always produced in patients with syphilis, are also produced in patients with other infectious diseases such as leprosy, tuberculosis, chancroid, leptospirosis, malaria, rickettsial disease, ­trypanosomiasis, lymphogranuloma venereum (LGV), measles, chickenpox, hepatitis, and infectious mononucleosis; noninfectious conditions such as drug addiction; autoimmune disorders, including rheumatoid disease and systemic lupus erythematosus; and in conjunction with increasing age, pregnancy, and recent immunization. Nontreponemal serologic tests are useful in monitoring treatment, whereas treponemal tests are not, as titers tend to remain elevated, even after successful treatment. Serologic testing may be negative during the early course of infection. 

Rapid Syphilis Tests Rapid point-of-care immunochromatographic strip assays have been developed that detect treponemal antibodies from whole blood, serum, or plasma. These rapid tests are capable of detecting IgM and IgG antibodies with antigens bound to a solid phase membrane. The tests provide results within 20 to 25 minutes and are useful for screening in the clinical setting. Although the rapid tests demonstrate similar sensitivities and specificities in comparison to laboratorybased methods, follow-up nontreponemal antibody titers are required for further evaluation. The Chembio DPP Syphilis Screen & Confirm Assay (Chembio Diagnostic Systems, Inc., Medford, NY) utilizes an immunochromatographic test strip and is able to detect both treponemal and nontreponemal antibodies by utilizing two test lines on the same strip. There are other rapid

• Fig. 45.2  Appearance of Treponema pallidum in dark-field preparation.

CHAPTER 45  The Spirochetes

point-of-care immunochromatographic strip assays that qualitatively detect antibodies to both T. pallidum and HIV1/2 (SD Bioline HIV/Syphilis Duo, Abbott, Lake ­Forest, IL; Chembio DPP HIV-Syphilis Assay, Chembio Diagnostic Systems, Inc., Medford, NY). 

Nontreponemal Antibody Tests The three nontreponemal serologic tests are the Venereal Disease Research Laboratory (VDRL), the TRUST assay, and the rapid plasma reagin (RPR) tests, which measure IgM and IgG antibodies. Each of these tests is a flocculation (or agglutination) test, in which soluble antigen particles are coalesced to form larger particles that are visible as clumps when they are aggregated in the presence of an antibody. The VDRL is used as a quantitative test and may be performed on serum or CSF in suspected cases of neurosyphilis. The RPR uses charcoal particles to detect antibodies in serum or plasma. There are numerous RPR kits commercially available that vary in methodology and procedures. Automated RPR tests use latex agglutination or immunoassay methods to increase the ability to screen multiple samples simultaneously. The TRUST assay is similar to the RPR but utilizes toluidine red in place of charcoal particles and is considered a macroflocculation. See Evolve Procedures 45.1 and 45.2 for details and limitations for the VDRL and RPR. Nontreponemal serologic tests for syphilis can be used to determine antibody quantitative titers, which are useful to follow the patient’s response to therapy. The relative sensitivity of each test is shown in Table 45.3 to confirm that a positive nontreponemal test result is from syphilis rather than from one of the other infections or biologic false-positive conditions noted previously. For syphilis, traditional diagnosis is useful in active infections. However,

early or treated infections may be incorrectly diagnosed. In addition, primary testing using RPR or VDRL may result in a high rate of false positives. The CDC has recommended a reverse algorithm to detect early primary or treated infections that may be missed using traditional nonspecific screening methods. Reverse testing suggests the use of specific antibody testing for syphilis, using EIA for IgM and IgG or a similar technique. T. pallidum antibodies persist for many years after infection. Specific tests may then be followed by nonspecific screening tests, which become less reactive over time. However, reverse testing is not currently widely accepted, and more data are needed to resolve clinical diagnostic discrepancies (Fig. 45.3). 

Treponemal Serologic Tests Specific treponemal serologic tests are typically positive within 1 to 2 weeks after the appearance of the primary lesion and include automated enzyme immunoassays (EIAs) and agglutination tests, such as the T. pallidum particle agglutination (TP-PA) test, the microhemagglutination assay (MHA-TP), T. pallidum indirect hemagglutination (TPHA), particle gel immunoassay (PaGIA), the fluorescent treponemal antibody absorption (FTA-ABS) test, chemiluminescence immunoassays (CIAs), and microbead immunoassays (MBIA). Once positive, their usefulness is limited because these tests tend to yield positive results throughout the patient’s life. The specificity of these tests is around 99%. The FTA-ABS test is performed by overlaying whole treponemes fixed to a slide with serum from patients suspected of having syphilis. This test is typically performed after a positive VDRL or RPR screening test. The patient’s serum is first absorbed with non–T. pallidum treponemal antigens (sorbent) to reduce nonspecific cross-reactivity.

TABLE 45.3    Sensitivity of Commonly Used Serologic Tests for Syphilis

Stage Method

Primary

Secondary

Late

Venereal Disease Research Laboratory 78% (reaginic) test (VDRL)

100%

96%

Rapid plasma-reagin (RPR) card test and automated reagin test (ART)

100%

98%

Nontreponemal (Reaginic Tests)—Screening

86%

591

Specific Treponemal Tests—Confirmatory FTA-ABS

78%

93%

93%

TP-PA

95%

100%

87%

Trep-Sure EIA

95%

100%

99%

Centaur CIA

95%

100%

94%

Liason CIA

96%

100%

93%

Bioplex MBIA

96%

100%

94%

FTA-ABS, Fluorescent treponemal antibody absorption; TP-PA, Treponema pallidum particle agglutination; EIA, enzyme immunoassay; CIA, chemiluminescence immunoassay; MBIA, microbead immunoassay.

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Fluorescein-conjugated antihuman antibody reagent is applied as a marker for specific antitreponemal ­antibodies in the patient’s serum. This test should not be used as a ­primary screening procedure. It has been demonstrated that FTA-ABS is less sensitive than TP-PA and other immunoassays (CIA, EIA, MBIA) at all stages of syphilis, especially primary syphilis. TP-PA (Fujirebio America, Fairfield, NJ) tests use gelatin particles sensitized with T. pallidum subsp. pallidum antigens. Serum samples are diluted in a microtiter plate, and sensitized gelatin particles are added. The presence of specific antibodies causes the gelatin particles to agglutinate and form a flat mat across the bottom of the microdilution well in which the test is performed. The MHA-TP is a passive hemagglutination assay of sensitized erythrocytes that are tested against the patient’s serum. Agglutination indicates the presence of IgG or IgM antitreponemal antibodies in the patient’s serum. TPHA is an indirect hemagglutination assay that uses sensitized red blood cells, which aggregate when exposed to positive patient serum. This test is similar to the MHA-TP. The PaGIA test, which uses gel immunoassay technology, is an established method in blood group serology. The assay contains recombinant antigens for the detection of T. pallidum antibodies in the patient’s serum or plasma. The results are available in approximately 15 minutes. Several EIAs are available that use direct, antibody class capture, indirect sandwich or competitive assay methodology. EIAs use recombinant antigens to detect IgM, IgG, or both. Trep-Sure EIA (Phoenix Biotech, Mississauga, Ontario, Canada) is a qualitative EIA that measures IgG and IgM treponemal antibodies. It utilizes microplates that are coated with

treponemal antigens. Patient antibodies bind to these antigens, and horseradish peroxidase conjugated to treponemal antigens is added, which binds to the antigen-antibody complexes. A chromogenic reaction occurs when the substrate for horseradish peroxidase, tetramethylbenzidine is added. The reaction is detected with a spectrophotometer at 450 nm. Several automated multiplex flow or MBIA systems exist that use bead-capture technology. These assays use a capture antibody attached to a suspension of small micropolystyrene beads. The beads are tagged with fluorophores of differing intensity, giving each a unique fingerprint. The sandwich immunoassay uses a flow cytometry dual-laser ­system for detection. There are currently three Luminex commercial platforms that use this technology: Abbott Architect (Abbott Laboratories, Abbott Park, IL), Bio-Rad Bioplex (Bio-Rad Laboratories, Hercules, CA), and Zeus AtheNA (Zeus Scientific, Branchburg, NJ). CIA also utilize bead technology and are automated. This type of assay utilizes a luminescent molecule as the antigen conjugate. The Liaison Treponema screen (DiaSorin, Stillwater, MN) uses magnetic beads to capture patient antibodies with an isoluminol-antigen conjugate. Positive samples are detected using a flash-chemiluminescent signal. The ADVIA Centaur syphilis assay (Siemens Healthcare Diagnostics, Inc., Newark, DE) is a direct sandwich immunoassay utilizing acridinium ester-labeled T. pallidum recombinant antigens, which bind to patient antibodies present in the specimen. Streptavidin-coated magnetic latex particles with biotinylated T. pallidum antigens bind to the antigen-antibody complexes. A light signal is produced during the reaction. 

Traditional testing

Reverse testing

Quantitative RPR + –

TP-PA or other specific treponemal +

Syphilis

test

EIA

No further testing

–

Differential diagnosis; syphilis unlikely

+

–

Quantitative RPR

No further testing

+

–

Syphilis

TP-PA or other specific tests –

+

Syphilis

•

Differential diagnosis; syphilis unlikely

Fig. 45.3  Traditional testing versus reverse testing.  EIA, Enzyme immunoassays; RPR, rapid plasma reagin; TP-PA, T. pallidum particle agglutination.

CHAPTER 45  The Spirochetes

Antimicrobial Susceptibility Testing and Therapy Because the treponemes cannot be cultivated, susceptibility testing is not performed. For all treponemal infections, penicillin G is the drug of choice. Ceftriaxone is also highly effective in most cases of syphilis, other than early syphilis. Tetracycline or doxycycline is often the treatment of choice when patients are allergic to penicillin. Treatment varies depending on the stage of disease and the host (e.g., children or adults, HIV-infected, or infected with congenital syphilis). 

Prevention No vaccines are available for the treponematoses. Prevention begins with early and appropriate treatment, thereby preventing person-to-person spread. 

Borrelia General Characteristics Borreliosis is considered a relapsing fever transmitted by a human-specific body louse or the soft-body tick of the genus Ornithodoros. Organisms belonging to the genus

Borrelia are helical shaped and composed of 3 to 10 loose coils (Fig. 45.1) without hooked ends. They contain endoflagella located beneath the outer membrane that enables the organism to actively demonstrate a characteristic corkscrew motility. The cells contain a protoplasmic cylinder that is composed of a peptidoglycan layer and an inner membrane. In contrast to the treponemes, Borrelia spp. stain well with a Giemsa stain. Species that have been grown in  vitro are microaerophilic or anaerobic. 

Epidemiology and Pathogenesis Although pathogens for mammals and birds, Borrelia are the causative agents of tick-borne and louse-borne relapsing fever and tick-borne Lyme disease in humans (Table 45.4).

Relapsing Fever Human relapsing fever is caused by more than 20 species of Borrelia and is transmitted to humans by the bite of a louse or tick. Borrelia recurrentis is responsible for louse-borne or epidemic relapsing fever. This spirochete is transmitted from the louse Pediculus humanus subsp. humanus, and disease is found worldwide; humans are the only reservoir for B. recurrentis. All other borreliae that cause disease in the United States are transmitted via tick bites and are named after the species of tick, usually of

TABLE 45.4    Epidemiology and Spectrum of Disease of Borrelia spp. Pathogenic for Humans

Primary Arthropod Vector

Geographic Location

Borrelia burgdorferi sensu lato (B. burgdorferi sensu stricto, B. mayonii, B. afzelii,a B. garinii,b B. valaisiana, B. lusitaniae, B. bavariensis, B. spielmanii, B. finlandensis, B. bissettiae, B. carolinensis)

Ixodes scapularis and I. pacificus in the United States, I. ricinus in Europe, and I. persulcatus in Asia

Northeast, MidAtlantic, Upper Midwest, West Coast United States, Europe, and Asia

Refer to text in this chapter

Tick-borne relapsing fever (TBRF)

Borrelia hermsii, B. turcatae,c B. parkeri, B. mazzottii, B. caucasica, B. crocidurae,c B. duttonii,c B. hispanica,c B. persica, B. venezuelensis

Ornithodoros spp.

North America, South America, Europe, Asia, Africa

Refer to text in this chapter

Louse-borne relapsing fever (LBRF)

Borrelia recurrentis

Pediculus humanus subsp. humanus

Worldwide

Refer to text in this chapter

Hard tick-borne relapsing fever (HTBRF) or Borrelia miyamotoi disease (BMD)

Borrelia miyamotoi

Ixodes scapularis and I. pacificus in the United States, I. ricinus in Europe, and I. persulcatus in Asia

Northeast, MidAtlantic, Upper Midwest, West Coast United States, Europe, and Asia

Fever, chills, headache, myalgia, arthralgia, and fatigue. Neurologic involvement in immunocompromised. Skin lesions are rare.

Disease

Agent

Lyme disease (Lyme borreliosis)

aAcrodermatitis

chronica atrophicans. is strongly associated. cNeurological involvement is common. bNeuroborreliosis

593

Clinical Manifestations

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the genus Ornithodoros (soft tick), from which they are recovered. Common species in the United States include Borrelia hermsii, Borrelia turicatae, Borrelia parkeri, and Borrelia mazzottii. Depending on the organisms and the disease, their reservoir is either humans or rodents, in most cases. Although their pathogenic mechanisms are unclear, these spirochetes exhibit antigenic variability that may account for the cyclic fever patterns associated with this disease. 

Lyme Disease Although there are currently more than 20 different Borrelia genospecies within the B. burgdorferi sensu lato complex, only Borrelia burgdorferi sensu stricto (strict sense of B. burgdorferi), as well as Borrelia garinii, Borrelia afzelii, Borrelia spielmanii, Borrelia lusitaniae, and Borrelia valaisiana, and the recently discovered Borrelia mayonii, are agents of Lyme disease and are transmitted by the bite of Ixodes ticks. B. burgdorferi and B. mayonii (less commonly) cause Lyme disease in North America. Currently, B. mayonii has been detected in ticks in the Upper Midwest. Lyme disease is the most common vector-borne disease in North America and Europe and is an emerging problem in northern Asia. Hard ticks, belonging primarily to the genus Ixodes, act as vectors in the United States, including Ixodes pacificus in California and Ixodes scapularis in other areas. The ticks’ natural hosts are deer and rodents. However, the adult ticks feed on a variety of mammals, including raccoons, domestic and wild carnivores, and birds. The ticks will attach to pets as well as to humans; all stages of ticks—larva, nymph, and adult—can harbor the spirochete and transmit disease. The nymphal form of the tick is most likely to transmit disease because it is active in the spring and summer, when people are dressed lightly and participate in outdoor activities in wooded areas. At this stage, the tick is the size of a pinhead and the initial tick bite may be overlooked. Ticks generally require a period of attachment of at least 24 hours before they transmit disease; however, other common diseases associated with Ixodes ticks, such as anaplasmosis, may be transmitted earlier on in the attachment period. Endemic areas of disease have been identified in many states, including Massachusetts, Connecticut, Maryland, Minnesota, Oregon, and California, as well as in Europe, Russia, Japan, and Australia. Direct invasion of tissues by the organism is responsible for the clinical manifestations. However, IgM antibodies are produced continually months to years after initial infection as the spirochete changes its antigens. B. burgdorferi’s potential ability to induce an autoimmune process in the host because of cross-reactive antigens may contribute to the pathology associated with Lyme disease. Moreover, by virtue of its ability to vary its surface antigens (e.g., outer surface protein [Osp] A to G), as well as avoid complement, B. burgdorferi is able to avoid the human host response. The pathologic findings associated with Lyme ­disease are also believed to result from the release of host cytokines initiated by the presence of the organism. 

Spectrum of Disease Relapsing Fever Between 2 and 15 days after infection, patients have an abrupt onset of fever, headache, and myalgia that lasts for 4 to 10 days. Physical findings often include petechiae, diffuse abdominal tenderness, and conjunctival effusion. As the host produces specific antibodies in response to the agent, organisms disappear from the bloodstream, becoming sequestered (hidden) in different organs during the afebrile period. Subsequently, organisms reemerge with newly modified antigens and multiply, resulting in another febrile period. Subsequent relapses are usually milder and of shorter duration. In general, more relapses are associated with cases of untreated tick-borne relapsing fever, but louseborne relapsing fevers tend to be more severe. B. miyamotoi can manifest as meningoencephalitis in immunocompromised patients. Treatment of relapsing fever with antibiotics may result in the formation of the Jarisch-Herxheimer reaction. This reaction is associated with the clearance of the organisms from the bloodstream and the release of cytokines within hours of antibiotic treatment. The patient experiences tachycardia, chills, rigors, hypotension, fever, and diaphoresis. Death may be associated with the reaction. An acute respiratory distress syndrome has also been recognized in cases associated with tick-borne relapsing fever. 

Lyme Disease Lyme disease is characterized by three stages, not all of which occur in any given patient. The first stage, early localized, is characterized by erythema migrans (EM), a red, ring-shaped skin lesion sometimes with a central clearing (“bull’s-eye” appearance) that first appears at the site of the tick bite but may develop at distant sites as well (Fig. 45.4). Patients may experience headache, fever, muscle and joint pain, and malaise during this stage. The second stage, early disseminated, beginning weeks to months after infection, may include arthritis, but the most important features are neurologic disorders (i.e., meningitis, neurologic deficits) and carditis. This is a result of the hematogenous spread of

•

Fig. 45.4  Appearance of the classic erythema migrans lesion of acute Lyme disease.

CHAPTER 45  The Spirochetes

spirochetes to organs and tissues. In addition, neurologic symptoms and infection may occur in the meninges, spinal cord, peripheral nerves, and brain (neuroborreliosis). The third stage, late disseminated, is usually characterized by chronic arthritis or acrodermatitis chronica atrophicans (ACA), a diffuse skin rash, and may continue for years. There is an association between Borrelia species and distinct clinical manifestations. For example, B. garinii has been associated with up to 72% of European cases of neuroborreliosis. Infection with B. mayonii has been associated with nausea and vomiting, as well as the presence of a diffuse skin rash and higher levels of spirochetemia, distinguishing it from B. burgdorferi. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing Peripheral blood in EDTA is the specimen of choice for direct detection of borreliae that cause relapsing fever. B.  burgdorferi can be visualized, cultured, and identified using PCR; CSF samples may be used for nucleic acid detection in the diagnosis of B. miyamotoi–associated meningoencephalitis or in patients with neurological symptoms associated with B. burgdorferi. Serum should be collected for serology during acute and convalescent phases, separated by at least 2 weeks. Specimens submitted for stain or culture include blood, biopsy specimens, and body fluids, including joint and CSFs. Body fluids should be transported without any preservatives. Tissue biopsy specimens should be placed in sterile saline to prevent drying. Skin biopsies of the EM lesion can be used for culture and nucleic acid detection. Whole blood in EDTA can also be used for culture or nucleic acid detection during early disseminated infection. Synovial fluid and tissue may also be tested using nucleic acid tests to monitor treatment in Lyme arthritis. 

Direct Detection Methods Relapsing Fever

Clinical laboratories rely on direct observation of the organism in peripheral blood from patients for diagnosis. Organisms can be found in 70% of cases when blood specimens from febrile patients are examined. The organisms can be seen directly in wet preparations of peripheral blood (mixed with equal parts of sterile, nonbacteriostatic saline) under dark- or bright-field illumination, in which the spirochetes move rapidly, often pushing the red blood cells around. The organisms may be visualized by staining thick and thin films with Wright or Giemsa stains or the quantitative buffy coat (QBC) method, using procedures similar to those used to detect malaria. B. miyamotoi may be detected microscopically in CSF in patients experiencing meningoencephalitis. B. burgdorferi may be visualized in tissue sections stained with Warthin-Starry silver stain. In general, the number of spirochetes in the blood of patients with Lyme borreliosis is below the lower limits of microscopic detection, although B. mayonii may be seen on a blood smear due to high spirochetemia. 

595

Nucleic Acid Detection PCR has become important in diagnosing Lyme disease. PCR has detected B. burgdorferi DNA in clinical specimens from patients with early and late clinical manifestations; optimal specimens include synovial tissue, CSF, synovial fluid, and skin biopsies from patients with EM. PCR amplification of B. burgdorferi nucleic acid sequences has been successful from urine but is not recommended. Laboratories have used a variety of nucleic acid–based methods to increase sensitivity and specificity and decrease turnaround time for diagnosing Lyme borreliosis. PCR has confirmed EM with an overall sensitivity and specificity of 68% and 100%, respectively. The ability to detect spirochetes in blood or plasma by PCR is dependent on the stage of illness (from 40% of patients with secondary EM to only 9.5% of patients with primary EM); PCR also does relatively well in detecting B. burgdorferi sensu lato in synovial fluids. In contrast, variable results using PCR occur in CSF specimens obtained from patients with peripheral or central nervous system involvement with Lyme borreliosis. PCR testing can be used to diagnose Lyme borreliosis caused by B. mayonii. PCR has also been used for the detection of organisms within the relapsing fever Borrelia group. However, 16SrRNA gene sequences do not provide good discrimination between species. Other sequences have been used for speciation, including the chromosomally encoded flagellin sequence (flaB), the 16-23S ribosomal RNA intergenic spacer gene, and the glycerophosphodiester (glpQ) gene unique to relapsing fever Borrelia species. 

Serodiagnosis Relapsing Fever

Serologic tests for relapsing fever have not demonstrated reproducible or reliable data for diagnosis because of the many antigenic shifts Borrelia organisms undergo during the course of disease. Protein heterogeneity in strains of different species is quite variable. For example, the OspC protein has 21 major recognized antigenic types. In addition, patients may exhibit increased titers to Proteus OX K antigens (up to 1:80), but other cross-reacting antibodies are rare. EIA using recombinant GlpQ antigen has been successful in the relapsing fever Borrelia group. Patients infected with relapsing fever Borrelia can demonstrate reactivity in B. burgdorferi EIA antibody assays, including the C6 antibody assay. References laboratories and the CDC perform Western blot testing for relapsing fever. Infection is confirmed by Western blot when the organisms GlpQ and 22-kDa antigens are positive with IgG and IgM.  Lyme Disease

Despite its inadequacies, serology continues to be the standard for the diagnosis of Lyme disease. B. burgdorferi has numerous immunogenic lipids, proteins, lipoproteins, and carbohydrate antigens on its surface and outer membrane. The earliest antibody response and development of IgM are in response to the OspC membrane protein, the flagellar

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antigens (FlaA and FlaB), or the fibronectin-binding protein (BBK32). The IgM levels peak within several weeks but may be detectable for several months. The IgG response develops slowly during the first several weeks of disease and increases with antibody responses to Osp17 (decorin-binding protein) and additional proteins, including p39 (BmpA) and p58. The late-stage infection demonstrates IgG antibodies to numerous antigens. Numerous serologic tests are commercially available. These include indirect immunofluorescence assays (IFA), which are being used less frequently, EIA, CIA, Immunoblots, and Western blots. The CDC recommends using validated and FDA approved IFA, CIA, or EIA as a first step, followed by an immunoblot, when performing serologic testing using the STTT. Measuring antibody by EIA or CIA are the most common screening methods used because they are quick, reproducible, and relatively inexpensive. However, false-positive rates are high, mainly as a result of crossreactivity. In addition, there is a lack of standardization between assays regarding the antigenic composition and antibodies detected from different manufacturers. The specificity of IFA may be improved by the adsorption of serum with the Reiter Treponema, Treponema phagedenis sonicate, or egg yolk (IFA-ABS). Patients with syphilis, HIV infection, leptospirosis, mononucleosis, parvovirus infection,

rheumatoid arthritis, and other autoimmune diseases commonly show positive results. Capture EIAs have been developed to prevent false-positive reactions with rheumatoid factor. In addition, this may be overcome by pretreatment of the patient’s sera with anti-IgG. For the United States, the CDC recommends a two-step approach to the serologic diagnosis of Lyme disease. In the STTT, the first step is a sensitive screening test such as an EIA, IFA, or CIA; if this test is positive for IgM or IgG or equivocal, the result must be confirmed by separate IgM and IgG immunoblotting (Fig. 45.5). Western blots should not be performed without a previously positive CIA, IFA, or EIA result, as this decreases the specificity of the test and can lead to false-positive results. IgM immunoblot requires reactivity in at least two of the three antigens tested: 23 kDaOspC, 39, and 41 kDa antigens. An IgM immunoblot requires reactivity in at least 5 of 10 antigens included in the assay: 18, 23, 28, 30, 39, 41, 45, 58, 66, and 93 kDa. IgM should only be used during the first 30 days of the infection, whereas IgG may be used at any stage during the disease process. As with B. burgdorferi, Lyme borreliosis caused by B. mayonii can also be detected with two-tier serologic testing. Recently, the CDC changed its recommendations for Lyme disease testing to include an MTTT. In this algorithm,

Screening assay (EIA IgM and IgG)

Positive or slightly positive

Negative

IgG and IgM immunoblot Report as negative. No further testing required.

Positive

Negative

Report as positive

Report as negative*

• Fig. 45.5  Two-step serodiagnostic procedure. *Note: If neuroborreliosis is suspected, a paired sera and cere-

brospinal fluid (CSF) specimen is recommended for testing. Any disease of short duration. For Lyme disease, it is recommended that a follow-up serologic test be performed at a later date. EIA, Enzyme immunoassays.

CHAPTER 45  The Spirochetes

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two EIAs are used: one as the primary screening test and the other as the second confirmatory test, which is performed if the first EIA is positive or equivocal. The FDA cleared ZEUS Scientific’s EIAs (ZEUS ELISA) for use in the MTTT. The first-tier test, ZEUS ELISA Borrelia VlsE1/pepC10 IgG/ IgM Test System, measures IgM and IgG antibodies to the Borrelia burgdorferi antigens VlsE1 and pepC10. Positive or equivocal results are followed by the second-tier test, either the ZEUS ELISA Borrelia burgdorferi IgG/IgM Test ­System, ZEUS ELISA Borrelia burgdorferi IgM Test System, or ZEUS ELISA Borrelia burgdorferi IgG Test System, which measure antibodies to Borrelia burgdorferi whole-cell ­antigen. All of the assays are horseradish peroxidase based. The MTTT allows for a greater ease of performance and interpretation as compared with the STTT by eliminating the need to perform a Western blot and decreases turnaround time. In certain clinical situations, results of serologic tests must be interpreted with caution. For example, patients with Lyme arthritis commonly remain antibody-positive despite treatment but do not necessarily have a persistent infection. Conversely, patients with a localized EM may be seronegative. Because of these limitations and others, the FDA and the American College of Physicians have published guidelines regarding the use of laboratory tests for Lyme disease diagnosis. Of paramount importance is the clinician’s determination before ordering serologic tests of the pretest probability of Lyme disease based on clinical symptoms and the incidence of Lyme disease in the population represented by the patient. 

Antimicrobial Susceptibility Testing and Therapy

Cultivation

Brachyspira spp. are comma or helical shaped, with tapered ends containing four flagella. Brachyspira aalborgi requires anaerobic incubation and has not been isolated from animals, whereas Brachyspira pilosicoli colonizes the intestine of a variety of animal species. The organisms reside in the brush border within the intestine and appear as a basophilic fringe, referred to as a “false brush border,” upon histologic staining with hematoxylin and eosin. 

Although the organisms that cause relapsing fever can be cultured in nutritionally rich media under microaerobic conditions, the procedures are cumbersome and unreliable and are used primarily as research tools. Similarly, the culture of B. burgdorferi may be attempted, although the yield is low. The best specimens for culture in untreated patients include the peripheral area of the EM ring lesion or synovial tissue. CSF and blood or plasma (greater than 9 mL), in general, are of low diagnostic yield by culture. This seems to correlate with the duration of the neurologic disease—in other words, positive results decrease as the duration of the disease increases. To cultivate the organism, the plasma, spinal fluid sediment, or macerated tissue biopsy is inoculated into a tube of modified Kelly’s medium (BSK II, BSK-H, or Preac-Mursic) and incubated at 30°C to 34°C for up to 12 weeks under microaerophilic conditions. Likewise, B. mayonii can be cultivated under these conditions in BSK II medium. Blind subcultures (0.1 mL) are performed weekly from the lower portion of the broth to fresh media, and the cultures are examined by dark-field microscopy or by fluorescence microscopy after staining with acridine orange for the presence of spirochetes. Because of the long incubation time and low sensitivity associated with cultivation, cultivation is often confined to reference or research laboratories. 

Currently there are no standardized methods, and borreliae are difficult to culture; therefore, antimicrobial susceptibility testing is not routinely performed. Several antibiotics, including tetracycline, are effective in treating relapsing fever, although it is associated with a higher rate of Jarisch-Herxheimer reaction, which can be life threatening. Doxycycline, amoxicillin, or cefuroxime and parenteral cephalosporins are drugs of choice during the first stage of Lyme disease. Broad-spectrum cephalosporins, particularly ceftriaxone or cefotaxime, have been used successfully with patients who either fail initial treatment or present in later stages of the disease, such as neuroborreliosis. Oral regimens are typically successful; however, patients with atrioventricular block may require ­intravenous (IV) therapy. Symptomatic treatment failures have been reported. 

Prevention Avoiding tick-infested areas; wearing protective clothing; checking the clothing, body, and pets for ticks; and removing them promptly also assist in the prevention of infection. There are no vaccines against infections caused by other ­Borrelia spp. 

Brachyspira General Characteristics

Epidemiology and Pathogenesis B. aalborgi is most likely transmitted via fecal-oral contamination. B. pilosicoli infection results from ingesting water contaminated with feces from infected animals. Brachyspira spp. cause intestinal spirochetosis, which may manifest as chronic diarrhea. The organisms have been associated with crypt abscesses, ulceration, necrosis, and multiple organ failure. Pathogenic mechanisms are not well understood. ­Clinical significance must be carefully correlated with patient signs and symptoms. 

Laboratory Diagnosis Specimen Collection and Direct Detection Fresh stool or rectal swabs may be collected and examined by dark-field microscopy. In addition, tissue biopsy specimens may be submitted for culture, PCR and histologic

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examination using periodic acid-Schiff (PAS) or hematoxylin and eosin staining. PCR amplification methods have been developed but are not available within the clinical laboratory. 

Cultivation Brachyspira spp. can be grown in the laboratory on brainheart infusion (BHI) or tryptic soy agar containing 10% fetal bovine blood, 400 μg/mL of spectinomycin, and 5 μg/ mL polymyxin in anaerobic conditions at 37°C. Increased growth can be seen at 41°C. Confluent growth patterns typically occur. B. aalborgi is weakly beta-hemolytic on BHI containing 10% fetal bovine blood. 

Approach to Identification B. aalborgi can be differentiated from B. pilosicoli by a strong positive hippurate hydrolysis reaction and a weak indole reaction. B. pilosicoli is indole-negative and has a weak hippurate hydrolysis reaction. Matrix-assisted laser desorption ionization and time-of-flight mass spectrometry have been successfully used for the identification of Brachyspira species. 

Antimicrobial Susceptibility Testing and Therapy Isolates of B. pilosicoli have demonstrated susceptibility to Augmentin (amoxicillin-clavulanic acid), ceftriaxone, chloramphenicol, meropenem, tetracycline, and metronidazole. 

Leptospira General Characteristics The leptospires include both free-living and parasitic forms. The organisms are spiral-shaped, right-handed helices with hooked ends. The organisms contain two axial filaments and exhibit either a spinning motility or a rapid back-andforth movement. Leptospira spp. are typically classified into three major groups, with Leptospira interrogans sensu stricto being the main species associated with human leptospirosis; in France, this organism is responsible for about 60% of human cases. Molecular classification using 16S ribosomal ribonucleic acid (rRNA) sequencing currently separates the genus into three distinct groups of pathogens, intermediate pathogens, and environmental saprophytes. The pathogens include more than 200 serologically defined types that were formerly designated as species and are now referred to as serovars, or serotypes, of L. interrogans sensu stricto and more than 60 serovars of L. biflexa sensu lato. Each serovar is usually associated with a particular animal host, and therefore serovar identification is important for epidemiology studies and prevention strategies. The genotypic classification scheme now includes approximately 23 genomospecies, which incorporates all current serovars of Leptospira. Serovars cross species lines as a result of the horizontal transfer of genetic elements, making it difficult to fully classify species phenotypically. 

Epidemiology and Pathogenesis Leptospirosis, a zoonosis, has a worldwide distribution but is most common in developing countries and warm climates via contact with infected animals or contaminated water. L. interrogans can infect most mammals throughout the world, as well as reptiles, amphibians, fish, birds, and invertebrates. The organism is maintained in nature by virtue of persistent colonization of renal tubules of carrier animals. Humans become infected through direct or indirect contact with the urine of infected animals. Humans and animals can be classified as a maintenance (infection is endemic) host or accidental host for leptospires. Leptospires enter the human host through breaks in the skin, mucous membranes, or conjunctivae. Infection can be acquired in home and recreational settings (e.g., swimming, hunting, canoeing) or in people who work in certain occupational settings (e.g., farmers, ranchers, abattoir workers, trappers, veterinarians). Pathogenic leptospires rapidly invade the bloodstream after entry, and spread throughout all sites in the body, such as the central nervous system and kidneys. Virulent strains show chemotaxis toward hemoglobin as well, as the ability to migrate through host tissues. A number of potential virulence factors that might facilitate this process are shown in Box 45.1. How L. interrogans causes disease is not completely understood, but it appears that the presence of endotoxin and other toxins may play a role in activation of the hemostasis pathways as an autoimmune response in the human host. 

Spectrum of Disease Symptoms begin abruptly 2 to 30 days after infection and include fever, headache, and myalgia. The most common clinical syndrome is anicteric leptospirosis, which is a selflimiting illness consisting of a septicemic stage, with high fever and severe headache that lasts 3 to 7 days, followed by the immune stage. Symptoms associated with the immune stage (onset coincides with the appearance of IgM) are varied but are generally milder than those associated with the septicemic stage. The hallmark of the immune stage is aseptic meningitis. Weil disease, or icteric leptospirosis, is generally the most severe illness, with symptoms caused by liver, kidney, or vascular dysfunction with lethal pulmonary hemorrhage; death can occur in up to 10% of cases.

• BOX 45.1 Potential Virulence Factors of

Leptospira

Hemolysins Sphingomyelinases C and H Proteolytic enzymes (thermolysin, collagenase) Catalase Cobalamin biosynthesis Sialic acids Fibronectin-binding protein for adhesion and invasion Lipopolysaccharide and outer membrane proteins

CHAPTER 45  The Spirochetes

Unfortunately, the clinical presentations of leptospirosis mimic those of many other diseases. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing During the first 10 days of illness, leptospires are present in the blood, CSF, and peritoneal dialysate. Urine specimens can be collected beginning in the second week of illness and up to 30 days after the onset of symptoms. Specimens may be collected in citrate, heparin, or oxalate anticoagulants. There are no other special requirements for specimen collection, transport, or processing. Citrate or ethylenediaminetetraacetic acid (EDTA) is the preferred anticoagulant for nucleic acid–based testing. Urine specimens should not be placed in preservatives and should be processed within 1 hour for optimal results. Urine specimens collected for nucleic acid testing can be placed in several commercial products and transported over long periods. Specimens should be transported at room temperature and inoculated for culture within 24 hours. 

Direct Detection Blood, CSF, and urine may be examined directly by darkfield microscopy. The detection of motile leptospires in these specimens is optimized after centrifuging at 1500× g for 30 minutes; sodium oxalate or heparin-treated blood is initially spun at 500× g for 15 minutes to remove blood cells. Other techniques, such as fluorescent antibody staining and hybridization techniques using leptospira-specific DNA probes, have also detected leptospires in clinical specimens. 

Nucleic Acid Detection Conventional and real-time PCR assays have been used to detect leptospires in blood, plasma, serum, urine, aqueous humor, CSF, autopsy tissue, and environmental samples. Nucleic acid detection is at least comparable to culture and useful in confirming diagnosis during the acute stages of the infection. 

Cultivation Albeit insensitive, the definitive method for laboratory diagnosis of leptospirosis is to culture the organisms from blood, CSF, or urine. A few drops of heparinized or sodium oxalate– anticoagulated blood are inoculated into tubes of semisolid media enriched with rabbit serum (Fletcher’s or Stuart’s) or bovine serum albumin. Urine should be inoculated soon after collection, because acidity (diluted out in the broth medium) may harm the spirochetes. One or two drops of undiluted urine and a 1:10 dilution of urine are added to 5 mL of medium. The addition of 200 μg/mL of 5-fluorouracil (an anticancer drug) may prevent contamination by other bacteria without harming the leptospires. Commercial media such as the Ellinghausen-McCullough-Johnson-Harris (EMJH) or Fletcher’s medium (Difco EMJH or Difco Fletcher’s medium; BD Diagnostic Systems, Sparks, MD) are available, which contain 5-fluorouracil for use at the patient’s bedside. Tissue specimens, especially from the liver

599

and kidney, may be aseptically macerated and inoculated in dilutions of 1:1, 1:10, and 1:100 as for urine cultures. All cultures are incubated at room temperature or 30°C in the dark for up to 6 to 8 weeks. Because organisms grow below the surface, material collected from a few centimeters below the surface of broth cultures should be examined weekly for the presence of growth, using a direct wet preparation under dark-field illumination. Leptospires exhibit corkscrewlike motility. 

Approach to Identification Based on the number of coils and hooked ends, leptospires can be distinguished from other spirochetes. Physiologically, the saprophytes can be differentiated from pathogens by their ability to grow at 10°C and lower, or at least 5°C lower than the growth temperature of pathogenic leptospires. Leptospires may also be visualized using dark-field microscopy or immunofluorescence. 

Serodiagnosis Serodiagnosis of leptospirosis requires a fourfold or greater rise in titer of agglutinating antibodies. The microscopic agglutination test (MAT) using live cells is the standard serologic procedure. Serologic diagnosis of leptospirosis is performed using pools of bacterial antigens containing many serotypes in each pool. Positive results are indicated by the presence of agglutination using dark-field microscopy. However, a macroscopic agglutination procedure is more readily accessible to routine clinical laboratories. Reagents are available commercially. Indirect hemagglutination and an ELISA test for IgM antibody are also available; IgM-detection assays are primarily used because IgM antibodies become detectable during the first week of illness. Rapid immunochromatographic assays are now available for the detection of IgM and/or IgG antibodies to Leptospira interrogans (Abbott, Chicago, IL). 

Molecular Typing Methods Several nucleic acid–based amplification methods have been developed. However, PCR methodologies are not useful for the differentiation of serovars and therefore are of limited use in epidemiologic studies. Highly complex and laborintensive techniques such as pulsed-field gel electrophoresis (PFGE) and restriction-fragment-length polymorphism (RFLP) and ribotyping are more useful for the identification of serovars. Nucleic acid sequencing has been used to successfully identify Leptospira directly in clinical specimens when the bacterial load was high. 

Antimicrobial Susceptibility Testing and Therapy Treatment of leptospirosis is supportive management and the use of appropriate antibiotics. Ceftriaxone, penicillin, amoxicillin, doxycycline, and tetracycline are recommended for the treatment of leptospirosis. Standardized procedures for antibiotic susceptibility are limited by the slow growth of the organisms and the need for serum during cultivation. 

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Prevention General preventive measures include the vaccination of domestic livestock and pet dogs. In addition, protective clothing, rodent control measures, and preventing recreational exposures, such as avoiding freshwater ponds, are indicated in preventing leptospirosis.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Aguero-Rosenfeld ME, Wang G, Schwartz I, Wormser GP: Diagnosis of Lyme borreliosis, Clin Microbiol Rev 18:484–509, 2005. Caroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Centers for Disease Control and Prevention: Discordant results from reverse sequence syphilis screening—five laboratories, United States, 2006-2010, MMWR Morb Mortal Wkly Rep 60:133–137, 2011. Centers for Disease Control and Prevention: Sexually transmitted disease surveillance 2017, Atlanta, 2018, U.S. Department of Health and Human Services. Church B, Wall E, Webb JR, Cameron CE: Interaction of Treponema pallidum, the syphilis spirochete, with human platelets, PloS One 14(1):e0210902, 2019. Cutler S, Vayssier-Taussat M, Estrada-Peña A, Potkonjak A, Mihalca AD, Zeller H: A new Borrelia on the block: Borrelia miyamotoi – a human health risk? Euro Surveill 24(18), 2019. https://doi. org/10.2807/1560-7917.ES.2019.24.18.1800170. Djokic V, Giacani L, Parveen N: Analysis of host cell binding specificity mediated by the Tp0136 adhesin of the syphilis agent T. pallidum subsp. pallidum, PLoS Negl Trop Dis 13(5):e0007401, 2019. Erlandson KM, Klingler ET: Intestinal spirochetosis: epidemiology, microbiology and clinical significance, Clin Microbiol Newsl 27:91, 2005. Fakile YF, Jost H, Hoover KW, et  al.: Correlation of treponemal immunoassay signal strength values with reactivity of confirmatory treponemal testing, J Clin Microbiol 56(1):e01165-e01117, 26, 2017. https://doi.org/10.1128/JCM.01165-17. Food and Drug Administration: FDA clears new indications for existing Lyme disease tests that may help streamline diagnoses. [News release], Silver Spring, MD, 2019, US Department of Health and Human Services, Food and Drug Administration, Available at: https://www.fda.gov/news-events/press-announcements/fdaclears-new-indications-existing-lyme-disease-tests-may-helpstreamline-diagnosesexternal. Fouts DE, Matthias MA, Adhikarla H, et al.: What makes a bacterial species pathogenic? comparative genomic analysis of the genus Leptospira, PLoS Negl Trop Dis 10(2):e0004403, 2016. https://doi. org/10.1371/journal.pntd.0004403. Hampson DJ: The spirochete Brachyspirapilosicoli, enteric pathogen of animals and humans, Clin Microbiol Rev 31(1):e00087-17, 2017. https://doi.org/10.1128/CMR.00087-17. Johnson TL, Graham CB, Maes SE, et al.: Prevalence and distribution of seven human pathogens in host-seeking Ixodes scapularis (Acari: Ixodidae) nymphs in Minnesota, USA, Ticks Tick Borne Dis 9(6): 1499–1507, 2018. https://doi.org/10.1016/j.ttbdis.2018.07.009.

Li SJ, Zhang CC, Li XW, et al.: Molecular typing of Leptospira interrogans strains isolated from Rattus tanezumi in Guizhou province, southwest of China, Biomed Environ Sci 25:542–548, 2012. Malincarne L, Schiaroli E, Ciervo A, et al.: Meningitis with cranial polyneuritis and cavernous sinus thrombosis by Borrelia crocidurae: first autochthonous case in Europe, Int J Infect Dis 82:30–32, 2019. https://doi.org/10.1016/j.ijid.2019.02.028. Mead P, Petersen J, Hinckley A: Updated CDC recommendation for serologic diagnosis of Lyme disease, MMWR Morb Mortal Wkly Rep 68:703, 2019. https://doi.org/10.15585/mmwr.mm6832a4. Package Insert: ZEUS ELISA Borrelia VlsE1/pepC10 IgG/IgM test System, Branchburg, NJ 08876, USA, ZEUS Scientific, Inc. Package Insert: ZEUS ELISA Borrelia burgdorferi IgG/IgM test System, Branchburg, NJ 08876, USA, ZEUS Scientific, Inc. Park IU, Fakile YF, Chow JM, et al.: Performance of treponemal tests for the diagnosis of syphilis, Clin Infect Dis 68(6):913–918, 2019. https://doi.org/10.1093/cid/ciy558. Peeling RW, Mabey D, Kamb ML, Chen XS, Radolf JD, Benzaken AS: Syphilis, Nat Rev Dis Primers 3:17073, 2017. https://doi. org/10.1038/nrdp.2017.73. Phillips ND, La T, Hampson DJ: Brachyspira catarrhinii sp. nov., an anaerobic intestinal spirochaete isolated from Vervet monkeys may have been misidentified as Brachyspira aalborgi in previous studies, Anaerobe 59:8–13, 2019. https://doi.org/10.1016/j. anaerobe.2019.05.004. Pritt BS, Respicio-Kingry LB, Sloan LM, et al.: Borrelia mayonii sp. nov., a member of the Borrelia burgdorferi sensu lato complex, detected in patients and ticks in the upper Midwestern United States, Int J Syst Evol Microbiol 66(11):4878–4880, 2016. Ratnam S: The laboratory diagnosis of syphilis, Can J Infect Dis Med Microbiol 16(1):45–51, 2005. Shen AK, Mead PS, Beard CB: The Lyme disease vaccine—a public health perspective, Clin Infect Dis 52(Suppl 3):s247–s252, 2011. Available at: https://doi.org/10.1093/cid/ciq115. Strnad M, Hönig V, Růžek D, Grubhoffer L, Rego ROM: Europe-wide meta-analysis of Borrelia burgdorferi sensu lato prevalence in questing Ixodes ricinus ticks, Appl Environ Microbiol 83(15):e00609– e00617, 2017. https://doi.org/10.1128/AEM.00609-17. Spirochaetes. National Center for Biotechnology Information (NCBI) taxonomy database. Accessed May 15,2019. Available at: https:// www.ncbi.nlm.nih.gov/Taxonomy/Browser/wwwtax.cgi?mode=U ndef&id=203691&lvl=6&lin. Talagrand-Reboul E, Boyer PH, Bergström S, Vial L, Boulanger N: Relapsing fevers: neglected tick-borne diseases, Front Cell Infect Microbiol 8:98, 2018. https://doi.org/10.3389/ fcimb.2018.00098. Wang Y, Li S, Wang Z, Zhang L, Cai Y, Liu Q: Prevalence and identification of Borrelia burgdorferi sensu lato genospecies in ticks from Northeastern China, Vector Borne Zoonotic Dis 19(5):309–315, 2019. https://doi.org/10.1089/vbz.2018.2316. Wilske B: Diagnosis of Lyme borreliosis in Europe, Vector Borne Zoonotic Dis 3:215–227, 2003. You M, Mo S, Leung WK, Watt RM: Comparative analysis of oral treponemes associated with periodontal health and disease, BMC Infect Dis 13:174, 2013. https://doi.org/10.1186/ 1471-2334-13-174. Zhou C, Zhang X, Zhang W, Duan J, Zhao F: PCR detection for syphilis diagnosis: status and prospects, J Clin Lab Anal 33(5):e22890, 2019. https://doi.org/10.1002/jcla.22890.

Chapter Review 1. Primary syphilis is characterized by what clinical sign: a. Chancre b. Rash c. Lymphadenopathy d. Gummas 2. Rapid point-of-care tests for syphilis utilize which of the following methods: a. Horseradish peroxidase–conjugated antigens b. Fluorescent-tagged antihuman antibodies c. Chemiluminescent molecules d. Antigens bound to a solid-phase membrane 3. The reverse testing algorithm for syphilis includes: a. Testing first for nonspecific antibodies and, if positive, testing for treponemal specific antibodies b. Testing first for specific treponemal antibodies and, if positive, testing for nonspecific antibodies c. Testing for nonspecific antibodies and, if positive, no further testing is required d. Testing for specific treponemal antibodies and, if positive, no further testing is required 4.  In addition to syphilis, reaginic antibodies can also be produced in patients with which of the following conditions? a. Leptospirosis b. Measles c. Autoimmune disorders d. All of the above 5. Which spirochete resides in the brush border of the intestine? a. Treponema pallidum b. Brachyspira pilosicoli c. Borrelia burgdorferi d. Leptospira interrogans 6. The standard serologic procedure for the diagnosis of leptospirosis is: a. PaGIA b. MBIA c. MA d. EIA followed by Western blot

7. A patient presents with a red skin lesion that has central clearing and flulike symptoms. She reports spending a lot of time outdoors hiking in wooded areas. What laboratory test should be ordered? a. Tissue biopsy of the skin lesion to stain and visualize spirochetes b. Whole blood specimen for culture of spirochetes c. Urine specimen for PCR d. Serum specimen for serologic testing 8. True or False _____ Treponemal serologic tests are useful in monitoring treatment for syphilis. _____ Infection with Borrelia mayonii can be detected using the two-tiered serologic test algorithm for Lyme disease. 9. Matching: Match each term with the appropriate definition. _____ Borrelia burgdorferi _____ Brachyspira spp. _____ MBIA _____ TP-PA _____ Leptospirosis _____ Prozone reaction _____ Dark-field microscopy _____ VDRL _____ Hutchinson’s triad

a. Renal involvement b. Erythema migrans c. Direct detection method d. Antigen-coated beads e. Congenital syphilis f. Intestinal spirochetosis g. Nontreponemal test h. False negative i. Agglutination of gelatin particles sensitized with T. pallium antigens

600.e1

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CASE STUDY 45.1 A 43-year-old female was referred to the infectious disease clinic for recurrent symptoms beginning about 4 months after she noticed a painful, swollen spot on her leg after removal of a tick. She had been treated with doxycycline for 14 days for presumed Lyme disease, documented by a positive IFA titer and presentation with migratory arthralgias, which were worse in the small joints of the hands. She also complained of fatigue, poor mentation, and occasional headaches. Because her symptoms recurred 2 months after treatment, she was unable to continue employment. Six months after a course of amoxicillin and later two courses of ceftriaxone, she again became symptomatic. A Western blot and PCR of her serum were ordered. The Western blot result was equivocal, but the PCR was positive for the agent of Lyme disease.

Questions 1. How did this patient acquire Lyme disease? 2. Why did the physician order further testing to diagnose Lyme disease in this patient? 3. The patient did not respond to therapy for B. burgdorferi. Can you explain why this can happen?   

PROCEDURE 45.1

Rapid Plasma Reagin Test Purpose The rapid plasma reagin (RPR) test is a presumptive serologic screening test for syphilis. 

5. Place the test card on a mechanical rotator or shaker at 80–100 rpm for 8 min. 6. Read the results before drying. 

Principle

Expected Results

The test is based on the presence of reagin, a nonspecific antilipid antibody found in the serum of patients infected with Treponema pallidum. Syphilis infection results in the breakdown of human tissue, releasing fatty substances that combine with T. pallidum protein to form an antigen, resulting in the formation of nonspecific and specific T. pallidum antibodies. The RPR antigen consists of cardiolipin, lecithin, and cholesterol bound to charcoal particles. The nonspecific antibody reacts with the test antigen, resulting in flocculation. The flocculation is visible on a white test card by incorporating charcoal particles in the antigen preparation. 

Specimen Patient serum or plasma is tested at room temperature, 20°C–25°C. 

Method 1. Place a 50 μL sample of patient serum or plasma and positive and negative controls into three separate circles on the test card. 2. Mix the RPR-carbon reagent gently before applying to each circle. 3. Add exactly one drop (20 μL) of reagent to each sample. 4. Mix the specimen and reagent with a stirrer, carefully spreading each sample over the entire surface of the circle. Use a clean stirrer for each sample.

Reactive: samples that demonstrate small to large clumps; a weak positive may show fine granulation or partial clumping. Nonreactive: samples that demonstrate no clumping or very slight roughness. 

Limitations 1. Slide flocculation tests are affected by room temperature. All test reagents and specimens should be warmed to room temperature before testing. 2. Failure to read results within 8 min may lead to drying and the appearance of false-positive results. 3. Patient samples that are hemolyzed, lipemic, or contain a high concentration of bilirubin may interfere with test results. 4. The RPR test cannot be used to test CSF specimens. 5. Biologic false positives occur as a result of other physiologic or pathologic conditions. 6. False negatives may be seen in specimens with high titers (prozone reaction). 

Quality Control Positive controls should demonstrate readily visible flocculation. Negative controls will appear uniformly turbid.   

CHAPTER 45  The Spirochetes 600.e3

PROCEDURE 45.2

Venereal Disease Research Laboratory Test Purpose The Venereal Disease Research Laboratory (VDRL) test is a slide flocculation test used for screening patients for infection with Treponema pallidum. 

Principle Patients infected with T. pallidum produce nonspecific antibodies capable of reacting with the cardiolipin test antigen. Cardiolipin is a lipid antigen extracted from beef heart that contains cardiolipin, lecithin, and cholesterol. The VDRL test is typically positive within 1–2 weeks after the appearance of the primary lesion. The test becomes reactive in late phase primary syphilis and highly reactive in secondary syphilis. The results will slowly decrease and become less reactive in late or tertiary syphilis. The VDRL test is also useful in the diagnosis of congenital syphilis and neurosyphilis. Maternal antibodies are capable of crossing the placenta; a positive VDRL immediately after birth may be solely a result of the presence of maternal antibodies. A quantitative titer is thus required at birth, followed by a second titer approximately 1 month after birth. No increase in titer will assist in ruling out the possibility of congenital syphilis. 

Specimen Patient serum or cerebrospinal fluid (CSF) samples. The serum must be inactivated before testing. The sample is heated at 56°C for 30 min. This inactivates serum proteins, such as complement within the patient specimen. CSF samples should not be heated before testing. 

Method 1. Place 50 μL of the patient sample, a positive reactive, and a negative nonreactive control into separate circles on the test card. 2. Place 20 μL of reagent next to the sample and stir each, making sure to completely cover the circle on the test card. Use a separate stirrer for each sample to prevent contamination. 3. Place the test card on a rotator or shaker at 180 rpm for 4 min. 

Expected Results Strongly reactive results will demonstrate large visible clumps. Weakly reactive results will appear as small clumps. Nonreactive results will appear as a diffuse, evenly turbid, reaction. 

Limitations 1. Results should be read immediately to prevent drying of the sample and the appearance of clumping caused by evaporation. 2. Biologic false positives occur as a result of other physiologic or pathologic conditions. Various conditions including pregnancy, menstruation, vaccination, trauma, and a variety of infectious diseases and immunologic disorders may cause biologic false positives. 3. False negatives may be seen in specimens with high titers (prozone reaction). 

Quality Control A reactive and nonreactive control should be included in each assay, demonstrating results as indicated in expected outcomes.   

PA RT IV   Parasitology

46

Overview of the Methods and Strategies in Parasitology OBJECTIVES This chapter provides an overview of the general epidemiology, pathogenesis, spectrum of disease, and approach to the identification of parasites. The detailed technical procedures should be used in conjunction with additional organism-specific chapters in this section to provide a clear description of the process, from specimen collection to identification. However, students should consider the following general objectives for the methods provided: 1. State the specific diagnostic purpose for each test methodology as well as its advantages and disadvantages. 2. Briefly describe the principle associated with the test method. 3. Determine specimen acceptability for parasite identification, including collection method, collection time/receipt time, number or quantity of specimen, and presence of interfering and contaminating substances. 4. Select appropriate preservatives for parasite specimens and explain the chemical principle and rationale for the preservative, including polyvinyl alcohol (PVA), universal fixative, formalin, and sodium acetate acetic acid formalin (SAF). 5. Select the appropriate method of detection and identification of parasites based on the type of specimen. 6. Discuss the effectiveness of antibody serology, antigen detection, molecular methods, and the traditional processing of ova and parasites (O&Ps) for the diagnosis of various parasite infections.

P

arasitic diseases are often associated with the earth’s tropical zones; however, many parasitic organisms that infect humans are worldwide in distribution and occur with some frequency in the temperate zones. In addition, people who travel to and from areas where parasites are endemic may contract one of these infections and then seek medical attention in their home country, far distant from the expected geographic area. An increase in the number

of compromised patients, particularly those who are immunodeficient or immunosuppressed, has also led to increased interest in the field of parasitology. These individuals are greatly at risk for certain parasitic infections. Parasites that infect humans are classified into six major divisions: • Protozoa (amebae, flagellates, ciliates, sporozoans, coccidia, microsporidia) • Nematoda, or roundworms • Platyhelminthes, or flatworms (cestodes, trematodes) • Pentastomids, or tongue worms • Acanthocephala, or thorny-headed worms • Arthropoda (e.g., insects, spiders, mites, ticks) The identification of parasitic organisms traditionally depends on the observation of characteristic morphologic criteria; accurate depiction of these criteria, in turn, depends on correct specimen collection, processing, and adequate fixation. Improperly submitted specimens may result in failure to detect the organisms or their misidentification. Tables 46.1 to 46.3 present information on the various groups of parasites, those that may be recovered from various body sites, the most frequently used specimen collection approaches, and appropriate processing methods.

Epidemiology Parasites are usually restricted to specialized environments inside and outside of their hosts. The host specificity of any individual parasite greatly influences factors associated with transmission and control. A disease of wild or domestic animals, zoonotic disease, also occurs in humans. Animals that are potential sources of a parasitic infection for humans are reservoir hosts. Some parasitic organisms are free living during specific stages of their life cycles and do not depend on human or other living hosts for survival. In some cases, humans become an accidental or unintended host. When humans are the only host for a parasite or a stage of its development, control and prevention options are relatively easy to define. However, if an infection is zoonotic or has an 601

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TABLE 46.1    Description of the More Common Groups of Human Parasites

Parasite Group

Description

Protozoa, Intestinal Amebae

Single-celled organisms; pseudopodia (motility), trophozoite, and cyst stages in the life cycle. Exceptions: Some have no identified cyst. Fecal-oral transmission of the infective cyst. Entamoeba histolytica causes amebiasis and is the most significant organism in this group.

Flagellates

Protozoa with characteristic flagella. Fecal-oral transmission. Exceptions: Trichomonas has a trophozoite and no cyst stage. Reproduction by longitudinal binary fission. Examples: Giardia duodenalis and Dientamoeba fragilis.

Ciliates

Single-celled protozoa; cilia (motility), which beat in a coordinated, rhythmic pattern, moving the trophozoite in a spiral path. Trophozoite and cyst stages in the life cycle; both stages show a large macronucleus and a micronucleus. Fecal-oral transmission. Neobalantidium coli is the single human pathogen in the group.

Coccidia

Protozoa; asexual and sexual life cycles. Fecal-oral transmission via contaminated food and/or water. Infective stage (oocyst) containing sporocysts and/or sporozoites. Examples: Cryptosporidium spp., Cyclospora cayetanensis, Cystoisospora belli, and Sarcocystis spp.

Microsporidia

Small (1–2.5 μm) intestinal protozoa. Transmission by ingestion, inhalation, or direct inoculation of spores. Nine genera cause disease in humans; the two most important are Encephalitozoon and Enterocytozoon.

Protozoa, Other Sites Amebae

Pathogenic free-living organisms associated with warm freshwater environments. Except for Entamoeba gingivalis (found in the mouth), they have been isolated from the central nervous system (CNS), eye, and other body sites. Examples: Naegleria fowleri—acute CNS infection and death. Chronic CNS disease (Acanthamoeba spp., Balamuthia mandrillaris), and Acanthamoeba spp. can also cause keratitis.

Flagellates

Have flagella (long, proteinaceous organelles used for motility). Sexual transmission. Examples: Trichomonas vaginalis is located in the genitourinary system. Trichomonas tenax can be identified in the mouth and is considered nonpathogenic.

Coccidia

Obligate intracellular, spore forming. Transmission is typically fecal-oral through ingestion of contaminated materials or food. Examples: Cryptosporidium spp. and Toxoplasma gondii.

Microsporidia

Small (1–2.5 μm) spore-forming protozoa. Transmission is typically by ingestion of spores. Life cycles vary considerably; some have an asexual life cycle, whereas others are complex and have both asexual and sexual life cycles and multiple hosts. Examples: Encephalitozoon, Pleistophora, Trachipleistophora, and Anncaliia spp.

Protozoa, Blood and Tissue Malaria, babesiosis

Arthropod vector–borne protozoa. Transmission via insect bite. Examples: Plasmodium spp. includes parasites that undergo exoerythrocytic and pigment-producing erythrocytic schizogony in vertebrates and a sexual stage followed by sporogony in mosquitoes. Babesia spp. are tick-borne and can cause severe disease in patients who have been splenectomized or otherwise immunologically compromised.

Flagellates (leishmaniae)

Trypanosomatid protozoa; two morphologic forms—promastigotes (anterior flagellum) in the insect host and amastigote (no flagella) in the vertebrate host. Transmission is through an insect vector. Recovery and identification of the organisms are related to body site. Recovery of leishmanial amastigotes is limited to the site of the lesion in infections other than those caused by the Leishmania donovani complex (visceral leishmaniasis).

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

TABLE 46.1    Description of the More Common Groups of Human Parasites—cont’d

Parasite Group

Description

Flagellates (trypanosomes)

Trypanosomatid protozoa; morphologic forms are identified based on the position, length, and attachment site of the flagella. At some time in their life cycle, these protozoa have the trypomastigote form with the typical undulating membrane and free flagellum at the anterior end. Transmission is typically through an insect vector. Some organisms cause African sleeping sickness (e.g., Trypanosoma brucei gambiense, T. brucei rhodesiense). The etiologic agent of American trypanosomiasis is Trypanosoma cruzi, which has amastigote and trypomastigote stages in the mammalian host and an epimastigote form in the arthropod host.

Nematodes, intestinal

Helminthic parasites; roundworms. Nematodes have separate sexes, are elongate-cylindrical and bilaterally symmetric with a triradiate symmetry at the anterior end. Nematodes have an outer cuticle layer, no circular muscles, and a pseudocele that contains all systems (digestive, excretory, nervous, and reproductive). Transmission is by ingestion of eggs or by skin penetration of larval forms from the soil. Examples: Ascaris, Enterobius, Trichuris, and Strongyloides spp. and hookworm.

Nematodes, tissue

Helminthic parasites; roundworms. Many of these organisms are rarely seen in the United States; however, some are important and are found worldwide. Diagnosis may be difficult if the only specimens are biopsy and/or autopsy, and interpretation must be based on examination of histologic preparations. Examples: Trichinella spp., visceral larva migrans (VLM), ocular larva migrans (OLM), cutaneous larva migrans (CLM).

Nematodes, filarial

Helminthic parasites; roundworms. Transmission is via arthropods. Adult worms tend to live in the tissues or lymphatics of the vertebrate host. The diagnosis is made based on recovery and identification of the larval worms (microfilariae) in the blood, other body fluids, or skin. Examples: Wuchereria, Brugia, Loa, and Onchocerca spp.

Cestodes, intestinal

Helminthic parasites; tapeworms. Adult tapeworm consists of a chain of egg-producing units called proglottids, which develop from the neck region of the attachment organ (scolex). Food is absorbed through the worm’s integument. The intermediate host contains the larval forms acquired through ingestion of the adult tapeworm eggs. Transmission is through the ingestion of larval forms in poorly cooked or raw meat or freshwater fish. Examples: Dipylidium caninum (infection is acquired by accidental ingestion of dog fleas). Hymenolepis nana and Hymenolepis diminuta are transmitted via ingestion of certain arthropods (fleas, beetles). In addition, H. nana can be transmitted through egg ingestion (life cycle can bypass the intermediate beetle host). Humans can serve as both the intermediate and definitive hosts in H. nana and Taenia solium infections.

Cestodes, tissue

Tissue tapeworms. Transmission is through ingestion of certain tapeworm eggs or accidental contact with certain larval forms, leading to tissue infection. Humans serve as the accidental intermediate host. Examples: T. solium, Echinococcus granulosus, and several other species.

Trematodes, intestinal

Flatworms that are exclusively parasitic. Except for the schistosomes (blood flukes), flukes are hermaphroditic. They may be flattened; most have oral and ventral suckers. Transmission: Intestinal trematodes require a freshwater snail to serve as an intermediate host; these infections are food-borne (freshwater fish, mollusks, or plants). Example: Fasciolopsis buski, the giant intestinal fluke.

Trematodes, liver, lung

Transmission: Liver and lung trematodes require a freshwater snail to serve as an intermediate host; these infections are food-borne (freshwater fish, crayfish or crabs, or plants). Examples: Public health concerns include cholangiocarcinoma associated with Clonorchis and Opisthorchis infections, severe liver disease associated with Fasciola infections, and misdiagnosis of tuberculosis in individuals infected with Paragonimus spp.

Trematodes, blood

Schistosomes; sexes are separate. Males are characterized by an infolded body that forms the gynecophoral canal in which the female worm is held during copulation and oviposition. Transmission: Infection is acquired by skin penetration by the cercarial forms that are released from freshwater snails. The adult worms reside in the blood vessels over the small intestine, large intestine, or bladder. Examples: Schistosoma mansoni, Schistosoma haematobium, and Schistosoma japonicum.

603

TABLE   Body Sites and Parasite Recovery (Trophozoites, Cysts, Oocysts, Spores, Adults, Larvae, Eggs, 46.2  Amastigotes, Trypomastigotes)

Site

Parasites

Blood Red cells

Plasmodium spp. Babesia spp.

White cells

Leishmania spp. Toxoplasma gondii

Whole blood/plasma

Trypanosoma spp. Microfilariae

Bone marrow

Leishmania spp. Trypanosoma cruzi Plasmodium spp.

Site

Parasites

Liver, spleen

Capillaria hepatica Clonorchis/Opisthorchis Echinococcus spp. Entamoeba histolytica Fasciola hepatica Leishmania donovani Toxocara spp. Microsporidia

Lung

Cryptosporidium spp.a Dirofilaria immitis Echinococcus spp. Hookworm larvae Paragonimus spp. Microsporidia

Muscle

Gnathostoma spinigerum Taenia solium (cysticerci) Taenia/Multiceps spp. Trichinella spp. Onchocerca volvulus (nodules) Spirometra/Diphyllobothrium spp. Trypanosoma cruzi Microsporidia

Skin

Ancylostoma spp. Dracunculus medinensis Gnathostoma spinigerum Leishmania spp. Onchocerca spp. Microfilariae Taenia spp. Demodex spp. Sarcoptes scabiei

Urogenital system

Trichomonas vaginalis Schistosoma spp. Baylisascaris procyonis Microsporidia Microfilariae

Eye

Acanthamoeba spp. Dirofilaria spp. Toxoplasma gondii Toxocara spp. Loa loa Microsporidia

Central Nervous System Cutaneous ulcers

Taenia solium (cysticerci) Echinococcus spp. Naegleria fowleri Acanthamoeba spp. Balamuthia mandrillaris Sappinia diploidea Toxoplasma gondii Microsporidia Trypanosoma spp. Leishmania spp.

Intestinal tract

Entamoeba bangladeshi Entamoeba histolytica Entamoeba dispar Entamoeba coli Entamoeba hartmanni Entamoeba polecki Endolimax nana Iodamoeba bütschlii Blastocystis hominis Giardia duodenalis Chilomastix mesnili Dientamoeba fragilis Pentatrichomonas hominis Neobalantidium coli Cryptosporidium spp. Cyclospora cayetanensis Cystoisospora belli Sarcocystis spp. Microsporidia Ascaris lumbricoides Anisakis spp. Enterobius vermicularis Hookworm Strongyloides stercoralis Angiostrongylus spp. Trichostrongylus spp. Trichuris trichiura Hymenolepis nana Hymenolepis diminuta Taenia saginata Taenia solium Diphyllobothrium latum Clonorchis sinensis (Opisthorchis) Paragonimus spp. Schistosoma spp. Fasciolopsis buski Fasciola hepatica Metagonimus yokogawai Heterophyes heterophyes

Note: This table does not include every possible parasite that can be identified in each body site; the most likely organisms have been listed. aDisseminated in severely immunosuppressed individuals.

TABLE 46.3    Specimens and Body Site: Specimen Options, Collection and Transport Methods, and Processing

Specimens and/or Body Site

Specimen Options

Collection and Transport Methods

Fresh stool

Comments

½ pint waxed container; 30 min if liquid, 60 min if semi formed, 24 h if formed; delivery to laboratory

Direct wet smear (not on formed specimen), concentration, permanent stained smear

Preserved stoola

5% or 10% formalin, MIF, SAF, Schaudinn’s, PVA, modified PVA, single vial systems, universal fixative

Concentration, permanent stained smear Depending on specimen (fresh or preserved) and patient’s clinical history, immunoassays may also be performed.

Stool specimens containing barium are unacceptable; intestinal protozoa may be undetectable for 5–10 days after barium use. Certain substances and medications also impede detection of intestinal protozoa: mineral oil, bismuth, antibiotics, antimalarial agents, and nonabsorbable antidiarrheal preparations. After administration of any of these compounds, parasitic organisms may not be recovered for a week to several weeks. Specimen collection should be delayed after barium or antibiotics are administered for 5–10 days or at least 2 weeks, respectively.

Stool for culture of nematodes

Fresh stool, entire stool specimen

½ pint waxed container; immediate delivery to laboratory

Filter paper strip, Petri dish, and agar plate, charcoal cultures are all available.

Fresh stool (do not refrigerate) is required for these procedures.

Stool for recovery of tapeworm scolex

Preserved stool, entire stool specimen

5% or 10% formalin (10% recommended)

The stool is filtered with a series of mesh screens and examined for the very small tapeworm scolex (proof of therapy efficacy) and/or proglottids.

Adult worms, worm segments

Saline, 70% alcohol

After treatment for tapeworm removal, the patient should be instructed to take a saline cathartic and to collect all stool material passed for the next 24 h. The stool should be immediately placed in 10% formalin and thoroughly broken up and mixed with the preservative (1-gallon [3.8-L] plastic jars are recommended, half-full of 10% formalin).

Cellophane tape preparation for pinworms

Surface sample from perianal skin; anal impression smear

Cellophane (Scotch) tape preparation or commercial sampling paddle or swab

Tape is lifted from a slide, a drop of xylene substitute is added, the tape is replaced, and the specimen is ready for examination under the microscope.

Specimens should be collected late at night after the person has been asleep for several hours or first thing in the morning before going to the bathroom or taking a shower. At least 4–6 consecutive negative tapes are required to rule out the infection.

Sigmoid colon

Sigmoidoscopy material, prepared as smears

Fresh or PVA or Schaudinn’s smears; specimen is taken with a spatula rather than cotton-tipped swabs; transported as smears in preservative

Direct wet smear, permanent stained smears

Material from the mucosal surface should be aspirated or scraped; it should not be collected with cotton-tipped swabs. At least 6 representative areas of the mucosa should be sampled and examined (6 samples, 6 slides). A parasitology specimen tray (containing Schaudinn’s fixative, PVA, and 5% or 10% formalin) should be provided, or a trained technologist should be available at the time of sigmoidoscopy to prepare the slides. Examination of sigmoidoscopy specimens does not take the place of routine O&Ps examinations. If the amount of material is limited, use of a fixative containing PVA is highly recommended.

Stool for ova and parasites (O&Ps) examination

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

Specimen Processing

Continued

605

Specimens and/or Body Site

Specimen Options

Collection and Transport Methods

Duodenum

Duodenal contents

Entero-Test capsule (string)

Urogenital tract

Specimen Processing

Comments

Entero-Test or aspirates; string in Petri dish or tube; immediate transport to laboratory

The specimen may be centrifuged (10 min at 500×) and should be examined immediately as a wet mount for motile organisms. Iodine also can be used. Direct wet smear of mucus; permanent stained smears can also be prepared.

A fresh specimen is required; the amount may vary from ＜0.5 mL to several milliliters of fluid. If the specimen cannot be completely examined within 2 h after collection, any remaining material should be preserved in 5%–10% formalin.

Duodenal contents

Entero-Test (string test) in Petri dish (fresh) or preserved in PVA

Bile-stained mucus clinging to the yarn should be scraped off (mucus can also be removed by pulling the yarn between the thumb and finger) and collected in a small Petri dish; disposable gloves are recommended. Usually 4 or 5 drops of material are collected. The specimen should be examined immediately as a wet mount for motile organisms (iodine may be added later to facilitate identification of any organisms present). The pH of the terminal end of the yarn should be checked to ensure adequate passage into the duodenum (a very low pH means that it never left the stomach). The terminal end of the yarn should be yellow-green, indicating that it was in the duodenum (the bile duct drains into the intestine at this point). Permanent stained smears can also be prepared.

If the specimen cannot be completely examined within 1 h after removal of the yarn, the material should be preserved in 5%–10% formalin, or PVA-mucus smears should be prepared.

Vaginal discharge Urethral discharge Prostatic secretions

Saline swab, transport swab (no charcoal), culture medium, plastic envelope culture, air-dried smear for FA

Direct wet smear; fluorescence; urine must be centrifuged before examination.

Fresh specimens are required; an air-dried smear may be an option for fluorescence. Do not refrigerate swabs and/or culture containers at any time, because motility and/or ability to grow will probably be lost.

Urine

Single unpreserved specimen, 24-h unpreserved specimen, early morning Nucleic acid–based testing media according to manufacturer’s instructions

Examination of urinary sediment may be indicated in certain filarial infections. Administration of the drug diethylcarbamazine (Hetrazan) has been reported to enhance the recovery of microfilariae from the urine. The triple-concentration technique is recommended for the recovery of microfilariae. The membrane filtration technique can be used with urine for the recovery of microfilariae. A membrane filter technique for the recovery of Schistosoma haematobium eggs has also been useful.

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TABLE 46.3    Specimens and Body Site: Specimen Options, Collection and Transport Methods, and Processing—cont’d

TABLE 46.3    Specimens and Body Site: Specimen Options, Collection and Transport Methods, and Processing—cont’d

Specimen Options

Collection and Transport Methods

Sputum

Sputum

True sputum (not saliva)

Induced sputum

No preservative (10% formalin if time delay)

Bronchoalveolar lavage (BAL)

Sterile; immediate delivery to laboratory

Bone marrow

Sterile; immediate delivery to laboratory

Cutaneous ulcers

Sterile plus air-dried smears

Liver, spleen

Sterile, collected in 4 separate aliquots (liver)

Aspirates

Specimen Processing

Comments

Direct wet smear; permanent stained smears; True sputum is required; all specimens, especially induced fluorescence also available (Calcofluor for specimens and BAL, should be delivered immediately to the microsporidia). Sputum is usually examined laboratory (do not refrigerate). as a wet mount (saline or iodine), using low and high dry power (10× and 400×). The specimen is not concentrated before preparation of the wet mount. If the sputum is thick, an equal amount of 3% sodium hydroxide (NaOH) (or undiluted chlorine bleach) can be added; the specimen is thoroughly mixed and then centrifuged. NaOH should not be used if the examiner is looking for Entamoeba spp. or Trichomonas tenax. After centrifugation, the supernatant is discarded, and the sediment can be examined as a wet mount with saline or iodine. If examination is delayed, the sputum should be fixed in 5% or 10% formalin to preserve helminth eggs or larvae or in PVA. Permanent stained smears; cultures can also be set (specifically designed for the recovery of blood parasites).

All aspirates for culture must be collected using sterile conditions and containers; this is mandatory for culture isolation of leishmania and trypanosomes.

Direct wet smear, permanent stained smears; culture for free-living amebae (Naegleria, Acanthamoeba spp.).

All specimens must be transported immediately to the laboratory (STAT procedure).

Lung Transbronchial aspirate

Air-dried smears

TracheobronAir-dried smears chial aspirate Central nervous system

Cerebrospinal fluid (CSF)

Sterile

Continued

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

Specimens and/or Body Site

607

Specimens and/or Body Site

Specimen Options

Collection and Transport Methods

Biopsy

Intestinal tract

Routine histology

Cutaneous ulcers Eye

Specimen Processing

Comments

Direct wet smears, permanent stained smears; The more material that is collected and tested, the more likely specimens to histology for routine processing. the organism is to be isolated and subsequently identified. Sterile, nonsterile to histopaSterile collection is required for all specimens that will be thology (formalin acceptable) cultured; bacterial and/or fungal contamination prevents Sterile (in saline), nonsterile to isolation of parasites in culture. histopathology

Scrapings

Sterile (in saline)

Cornea (scrapings)

Collected by physician, placed directly on microscope slide

Liver, spleen

Sterile, nonsterile to histopathology

Fixed using methyl alcohol and stained using Calcofluor white.

Helpful in diagnosis of Acanthamoeba keratitis.

Lung

Blood

Brush biopsy

Air-dried smears

Open lung biopsy

Air-dried smears

Muscle

Fresh, squash preparation, nonsterile to histopathology

Skin biopsy

Nonsterile to histopathology (formalin acceptable)

Scrapings

Sterile (in saline), nonsterile to histopathology

Skin snip

Aseptic, smear or vial No preservative

Smears of whole blood

Fresh (first choice) Thick and thin films; immediate delivery to laboratory

Thick and thin films, specialized concentrations and/or screening methods

Examination of blood films (particularly for malaria) is considered a STAT procedure; immediate delivery to the laboratory is mandatory.

Anticoagulated blood

Anticoagulant (second choice) EDTA (first choice) Heparin (second choice)

Thick and thin films, specialized concentrations and/or rapid methods Quantitative Buffy Coat (QBC) Microhematocrit Centrifugation Method (Becton Dickinson, Tropical Disease Diagnostics, Sparks, MD)

Delivery to the laboratory within 30 min or less. If delayed, typical parasite morphology may not be seen in blood collected using anticoagulants. Knott concentration procedure: Used primarily to detect microfilariae in the blood, especially when a light infection is suspected. The disadvantage of the procedure is that the microfilariae are killed by the formalin and are no longer motile. Membrane filtration technique: This technique, using Nuclepore filters, has proved highly efficient in demonstrating filarial infections when microfilaria are of low density. It has also been successfully used in field surveys.

EDTA, Ethylenediaminetetraacetic acid; FA, fluorescent antibody; MIF, merthiolate-iodine-formalin; PVA, polyvinyl alcohol; SAF, sodium acetate–acetic acid–formalin. aA number of new stool fixatives are available; some use a zinc sulfate base rather than mercuric chloride. Some collection vials can be used as a single-vial system; both the concentration and permanent stained smear can be performed from the preserved stool. However, not all single-vial systems (proprietary formulas) provide material that can be used for fecal immunoassay procedures. A universal fixative is now available (TOTAL-FIX) that contains no formalin, mercury, or PVA. Modified from Garcia LS. Diagnostic Medical Parasitology. 6th ed. Washington, DC: ASM Press; 2016.

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TABLE 46.3    Specimens and Body Site: Specimen Options, Collection and Transport Methods, and Processing—cont’d

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

environmental phase, these measures can be complex, especially if more than one reservoir species or environmental resource is involved. A human parasitic infection can be initiated through a variety of mechanisms and pathways depending on the species of microorganism. Many parasites of the intestinal tract are transmitted through ingestion of the infective form of the parasite in contaminated food or water (e.g., Giardia duodenalis, Cryptosporidium spp., Ascaris lumbricoides). Other parasites can be transmitted from host to host through sexual means (venereal transmission) (e.g., Trichomonas vaginalis), through skin penetration of infective larvae (e.g., Strongyloides stercoralis, hookworm), or through the bites of various arthropods (e.g., Plasmodium, Trypanosoma, Leishmania) (Table 46.4). 

Pathogenesis and Spectrum of Disease Although a number of parasites can cause serious and lifethreatening disease, particularly in compromised patients, many organisms reach a “status quo” with the host and cause no significant damage. Obvious disease symptoms may not be the ultimate outcome of infection. Depending on the parasite, one or multiple body sites may be infected. Some parasitic infections can result in few or no symptoms, whereas infection with other parasites can result in devastating permanent damage and eventual death of the host. Some parasites multiply in the human body, whereas others mature but do not increase in number inside the body. These life-cycle differences play important roles in pathogenicity and disease outcome. It is also important to remember that a patient who is debilitated or immunocompromised (including the very young and the very old) may react differently to a parasitic infection than a healthy adult. Certainly, it is not to the parasite’s advantage to damage the host to the extent that severe illness or death results; this makes survival of the parasite difficult at best. When this occurs, long-term survival of the parasite may depend on rapid and efficient transmission from host to host or the parasite’s ability to survive within the environment without a live host. It is important to understand the life cycle of parasites in terms of potential prevention and control as well as mechanisms of infectivity and disease outcome (Table 46.5). Table 46.6 lists mechanisms of pathogenesis and the spectrum of parasitic diseases. Specific guidelines for physician requests are presented in Table 46.7. 

609

collection fixatives, clinical specimens, diagnostic tests, the elements of a positive finding, and comments are presented in Tables 46.9 and 46.10.

Specimen Collection and Transport Depending on its stage of development in the clinical specimen (adult, larvae, eggs, trophozoites, cysts, oocysts, spores), a particular parasite may not be able to survive outside the host. For this reason, clinical specimens should be transported to the laboratory immediately to increase the likelihood of finding intact organisms. Because a lag time often occurs between collection of the specimen and its arrival in the laboratory, most facilities routinely use preservatives for collection and transport (Fig. 46.1). This approach ensures that any parasites present maintain their morphology and can be identified after processing. Correct processing depends on the use of appropriate fixatives, immediate fixation upon collection of the specimen, and adequate mixing between the fixative and specimen (Table 46.9). It is mandatory that specimen collection guidelines be available in plain and direct language for health care personnel or patients performing specimen collection and that all clients recognize the importance of following such guidelines. In areas with non-native English speakers, alternate language versions should be readily available to provide clear directions for specimen collection. Specimen rejection criteria must be included as a part of the guidelines; guidelines must be followed and enforced to limit the possibility of reporting misleading or incorrect results. Detailed specimen descriptions and body sites, in addition to collection and transport information, are included in Table 46.3.

Laboratory Diagnosis As with all laboratory testing, the ability to detect and correctly identify human parasites is directly linked to the quality of the clinical specimen, submission of the appropriate specimen or specimens, appropriate processing and handling before analysis, relevant diagnostic test orders, and the experience and training of laboratory personnel (Table 46.8). A summary of human parasites and applicable

• Fig. 46.1  Stool collection vial.  Most laboratories recommend immediate placement of stool samples into a preservative such as this to preserve morphology.

TABLE 46.4    Epidemiology of the More Common Groups of Human Parasites

Parasite Group Habitat (Reservoir)

Mode of Transmission

Prevention

Protozoa, Intestinal Amebae

Single-celled organisms generally found in humans. Although certain animals harbor some of these organisms, they are not considered important reservoir hosts.

Humans acquire infections by ingesting food and water contaminated with fecal material containing the resistant infective cyst stage of the protozoa. Various sexual practices have been documented in transmission.

Preventive measures include increased attention to personal hygiene, sanitation measures, and elimination of sexual activities that may involve fecal-oral contact.

Flagellates

The flagellates are generally found in humans. Although certain animals harbor some of these organisms, they are not considered important reservoir hosts; one exception may be animals such as the beaver that harbor Giardia duodenalis. Contaminated water supplies are also a source.

Humans acquire infections by ingesting food and water contaminated with fecal material containing the resistant infective cyst stage of the protozoa; the trophozoite forms may be transmitted from person to person in helminth eggs.

Preventive measures include increased attention to personal hygiene and sanitation measures, elimination of sexual activities that may involve fecal-oral contact, adequate water treatment (including filtration), and awareness of environmental sources of infection.

Ciliates

Neobalantidium coli is generally found in humans and pigs. In some areas of the world, pigs are considered important reservoir hosts.

Humans acquire infections by ingesting food and water contaminated with fecal material containing the resistant infective cyst stage of the protozoa.

Preventive measures include increased attention to personal hygiene and sanitation measures as well as the elimination of sexual activities that may involve fecal-oral contact.

Coccidia

Coccidia are found in humans. In some cases (e.g., cryptosporidiosis) animal reservoirs (cattle) can serve as important hosts. The muscle of various animals may contain sarcocysts that are infective for humans through the consumption of raw or poorly cooked meat. Numerous water-borne outbreaks with Cryptosporidium spp. have been reported throughout the world. Coccidian oocysts are extremely resistant to environmental conditions, particularly if kept moist.

These protozoa are acquired through ingestion of various meats or by fecal-oral transmission through contaminated food and/or water. The infective forms are called oocysts (Cryptosporidium spp., Cystoisospora belli, Cyclospora cayetanensis) or sarcocysts (Sarcocystis spp.), which are contained in infected meat. Cryptosporidia have also been implicated in nosocomial infections.

Preventive measures include increased attention to personal hygiene and sanitation measures and elimination of sexual activities that may involve fecal-oral contact. Adequate water treatment (including filtration) is mandatory; awareness of environmental sources of infection is also important.

Microsporidia

Microsporidia can infect every living animal, some of which probably serve as reservoir hosts for human infection. However, host specificity has not been well defined. The spores are environmentally resistant and can survive for years if kept moist.

Infection with microsporidial spores usually occurs through ingestion; however, inhalation of spores and direct inoculation from the environment almost certainly occur.

Preventive measures include increased attention to personal hygiene and sanitation measures, increased awareness of environmental exposure possibilities, and adequate water treatment.

Infection occurs through contact with contaminated water; organisms enter through the nasal mucosa and may travel via the olfactory nerve to the brain. Disease can be very severe and life-threatening; keratitis is also caused by these organisms, and infection can be linked to blindness or severe corneal damage. Eye infections can be linked to contaminated lens solutions or direct accidental inoculation of the eye from environmental water and/or soil sources.

Prevention includes avoidance of contaminated environmental water and soil sources and adequate care of contact lens systems.

Protozoa, Other Sites Amebae

Free-living amebae are associated with warm, freshwater environments; they are also found in soil. Although humans can harbor these organisms, person-to-person transfer is rare. Environmental sources are the primary link to human infection. Contaminated eye care solutions have been linked to organisms that cause keratitis.

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TABLE 46.4    Epidemiology of the More Common Groups of Human Parasites—cont’d

Parasite Group Habitat (Reservoir)

Mode of Transmission

Prevention

Flagellates

T. vaginalis is found in the genitourinary system and is usually acquired by sexual transmission.

Prevention involves awareness of sexual transmission; treatment of all partners is necessary when infection is diagnosed in an individual patient.

Trichomonas vaginalis infection is found in a large percentage of humans; humans may present as symptomatic or asymptomatic. Person-to-person transfer is very common; reinfection is also common, particularly if sexual partners are not treated.

Protozoa, Blood and Tissue Malaria, babesiosis

Humans harbor the five species of malaria (Plasmodium vivax, P. ovale, P. malariae, P. knowlesi, and P. falciparum). Other animals can carry Babesia spp., and animal reservoir hosts play a large role in human transmission.

These organisms are borne by arthropods, Plasmodium spp. by the female anopheline mosquito and Babesia spp. by one or more genera of ticks. Infections can be transmitted transplacentally, via shared needles, through blood transfusions, and from organ transplants.

Prevention involves vector control and awareness of transmission through blood transfusions, shared drug needles, congenital infections, and organ transplants. Careful monitoring of the blood supply is necessary. Malaria prophylaxis is recommended for persons traveling to endemic areas.

Flagellates (leishmaniae)

Some strains of leishmaniae have reservoir hosts (e.g., dogs for the Mediterranean strain of Leishmania donovani and wild rodents for the African strains of L. donovani). Leishmania tropica has been linked to the same animal reservoirs.

Transmission is through the bite of infected sandflies. Infection can be spread person-to-person contact (cutaneous lesions), blood transfusions, shared needles, and organ transplants.

Prevention includes vector control, avoiding environmental sources (e.g., dogs, wild rodents), and careful handling of all clinical specimens from infected patients.

Flagellates (trypanosomes)

Humans are the only known hosts for Trypanosoma brucei gambiense (West African trypanosomiasis); Trypanosoma brucei rhodesiense (East African trypanosomiasis) infections are found in a number of antelope and other ungulates that act as reservoir hosts. Rodents and some mammals are reservoir hosts for Trypanosoma cruzi.

Transmission is through the bite of the infected tsetse fly and through blood transfusion, shared needles, and organ transplants. Transmission of T. cruzi is through the infected feces of the triatomid bug; the bug takes a blood meal, immediately defecates, and the human host scratches the infected feces into the bite site; bug saliva contains an irritant that stimulates scratching.

Prevention relies on vector control and awareness of potential exposure/infection from blood sources (transfusions, shared needles, organ transplants). Laboratory accidents while handling infected blood have been reported.

Nematodes, intestinal

These roundworms generally do not have animal reservoirs relevant to human infection. One exception is the pig ascarid; human infections have been reported. These worms are found worldwide; Ascaris lumbricoides is probably the most common parasite in humans. Strongyloides stercoralis is particularly important as the causative agent of severe disease in the compromised host.

A. lumbricoides and Trichuris trichiura eggs must undergo development in the soil before they are infective; thus, children who play in the dirt are a particularly high-risk group. Ingestion of food and water contaminated with infective eggs is the primary route of infection. Hookworm and S. stercoralis infections are initiated by larval penetration of the skin from contaminated soil. Pinworm infection (E. vermicularis) is acquired through ingestion of infective eggs from the environment (hand-to-mouth).

Prevention includes avoiding ingestion of contaminated soil and/or avoiding frequenting soil contaminated with hookworm eggs (pets, soil, water, warmth, warm weather); treatment for pinworm is recommended, but reinfection is common.

Continued

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TABLE 46.4    Epidemiology of the More Common Groups of Human Parasites—cont’d

Parasite Group Habitat (Reservoir)

Mode of Transmission

Prevention

Nematodes, tissue

Trichinella spp. have a number of animal reservoir hosts, including bears, walruses, pigs, rodents, and other animals. Dog and cat hookworms cause cutaneous larva migrans (CLM), and the dog and cat ascarid, Toxocara spp., causes visceral and ocular larva migrans (VLM, OLM). These infections can be serious and cause severe disease if not treated.

Trichinella organisms are acquired by ingestion of raw or poorly cooked infected meat. CLM is caused by skin penetration of infective larvae from the soil; children should avoid sandboxes where dogs and cats are known to defecate. Larval migration is limited to the skin. VLM and OLM are caused by accidental ingestion of Toxocara spp. eggs from contaminated soil; larval migration occurs throughout the body, including the eyes.

Preventive measures include adequate cooking of infected meat; awareness of possibility of contaminated soils for dog and cat hookworms and/or ascarids; and covering of all sandboxes where pets have access to defecate and children play.

Nematodes, filarial

Wuchereria bancrofti, Loa loa, and Onchocerca volvulus have no animal reservoirs and are found only in humans, whereas Brugia spp. can be found in cats and monkeys. Dracunculus medinensis can infect dogs, cats, monkeys, and humans.

Filarial nematodes are transmitted through the bite of a bloodsucking arthropod (midges, mosquitoes, and flies). Dracunculus infections are acquired through ingestion of water contaminated with small crustaceans, Cyclops spp., which contain infective larvae.

Prevention involves vector control and protection of well-water sources.

Cestodes, intestinal

The human serves as the definitive host for beef (Taenia saginata) and pork (Taenia solium) tapeworms; cows/ camels and pigs serve as intermediate hosts, respectively. Humans also serve as the intermediate host for T. solium (cysticercosis). Diphyllobothrium latum adult tapeworms can be found in a number of wild animals, the most important being dogs, bears, seals, and walruses, which serve as reservoir hosts; humans are the definitive host. Hymenolepis nana (dwarf tapeworm) can occur in rodents; humans can serve as both intermediate and definitive hosts, with development from the egg to adult worm occurring in the human intestine.

Human infection with the adult worm occurs through ingestion of raw or poorly cooked meat (beef, camel, pork) containing the intermediate forms, the cysticerci. Humans become the accidental intermediate host when eggs from an adult T. solium tapeworm are ingested. The cysticerci develop in the muscle and tissues of the human rather than the pig. Infection with the adult D. latum tapeworm occurs through ingestion of poorly cooked freshwater fish containing the sparganum or plerocercoid larval form. Infection with H. nana is primarily acquired through accidental ingestion of eggs from an adult tapeworm.

Preventive measures involve adequate cooking of infected meat and treatment of patients harboring adult tapeworms (accidental ingestion of eggs can lead to infection).

Cestodes, tissue

Adult worms are found in a variety of animals; the human becomes the accidental intermediate host after ingestion of eggs from the adult worms. Reservoir hosts include dogs, cats, and rodents.

Ingestion of certain tapeworm eggs or accidental contact with certain larval forms can lead to tissue infection with Taenia solium, Echinococcus spp., and several others.

Preventive measures involve increased attention to personal hygiene and sanitation measures.

Trematodes, intestinal

Fish-eating wild and domestic animals serve as reservoir hosts. The definitive host of Fasciolopsis buski is the pig.

Ingestion of water chestnut and caltrop (raw, peeled with the teeth) is the source of infection; metacercariae are encysted on the plant material. Pig feces are used to fertilize various water plant crops.

Preventive measures include avoiding eating raw water plants that may contain encysted larval forms of the flukes and adequate waste disposal of farm animal feces (pigs).

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TABLE 46.4    Epidemiology of the More Common Groups of Human Parasites—cont’d

Parasite Group Habitat (Reservoir)

Mode of Transmission

Prevention

Trematodes, liver, lung

Cats, dogs, and wild fish-eating mammals can serve as reservoir hosts for Opisthorchis spp., Clonorchis sinensis, and Paragonimus spp. Fasciola hepatica is normally a parasite of sheep, and Fasciola gigantica is a parasite of cattle; humans are accidental hosts.

Infection occurs through ingestion of encysted metacercariae in raw or poorly cooked fish, crabs, crayfish, and on plants. Infection with Fasciola spp. is not easily acquired (the parasite is not that well adapted to the human host).

Prevention requires thorough cooking of potentially infected fish, crabs, crayfish and avoiding eating raw water plants that may contain encysted metacercariae.

Trematodes, blood

Schistosoma mansoni and S. haematobium appear to be restricted to the human host; S. japonicum can be found in cattle, deer, dogs, and rodents; S. mekongi is found in dogs and rodents. The worms mature in the blood vessels, and eggs make their way outside the body in stool and/ or urine. The freshwater snail is a mandatory part of the life cycle (contains developmental forms of schistosome).

Infection occurs through skin penetration by infected cercariae released from a freshwater snail containing the intermediate stages of the schistosome life cycle. Cercariae can be released from the snail intermediate host singly or in groups.

Prevention involves protection from potentially contaminated water sources; awareness of mode of transmission; and proper handling of human waste containing eggs (continued infection of snail intermediate hosts).

TABLE 46.5    Parasitic Infections: Clinical Findings in Normal and Compromised Hosts

Organism

Normal Host

Compromised Host

Entamoeba histolytica

Asymptomatic to chronic-acute colitis, extraintestinal disease may also occur (primary site: right upper lobe of liver).

Diminished immune capacity may lead to extraintestinal disease.

Free-living amebae Naegleria fowleri Acanthamoeba spp. Balamuthia mandrillaris Sappinia spp.

Patients tend to have eye infections with Acanthamoeba spp. linked to poor lens care.

Primary amoebic meningoencephalitis (PAM); granulomatous amebic encephalitis (GAE).

Giardia duodenalis

Asymptomatic to malabsorption syndrome.

Certain immunodeficiencies tend to predispose an individual to infection.

Toxoplasma gondii

Approximately 50% of individuals have antibody and organisms in tissue but are asymptomatic. It is important to note that a developing fetus may be severally affected if the mother is infected; this is highly dependent on the time (trimester) when the infection occurs.

Disease in compromised hosts tends to involve the central nervous system (CNS), with various neurologic symptoms; it can mimic neurologic symptoms of infection with the human immunodeficiency virus (HIV).

Cryptosporidium spp. Cryptosporidium hominis (humans) Cryptosporidium parvum (humans and animals)

Self-limiting infection with diarrhea and abdominal pain.

Because of the autoinfective nature of the life cycle, infection is not self-limiting and may produce fluid loss of more than 10 L/day; multisystem involvement may occur. No effective therapy is available.

Cyclospora cayetanensis

Self-limiting infection with diarrhea (3–4 days); relapses common.

Diarrhea may persist for 12 weeks or longer; biliary disease has also been reported in this group, particularly those with acquired immunodeficiency syndrome (AIDS).

Continued

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TABLE 46.5    Parasitic Infections: Clinical Findings in Normal and Compromised Host—cont’d

Organism

Normal Host

Compromised Host

Cystoisospora belli

Self-limiting infection with mild diarrhea or no symptoms.

May lead to severe diarrhea, abdominal pain, and possibly death (rare case reports); diagnosis occasionally may be missed because of failure to recognize the oocyst stage; is not seen when concentrated from polyvinyl alcohol (PVA) fixative.

Sarcocystis spp.

Self-limiting infection with diarrhea or mild symptoms.

Symptoms may be more severe and last for a longer period.

Microsporidia Anncaliia Nosema Vittaforma Encephalitozoon Enterocytozoon Pleistophora Trachipleistophora “Microsporidium”

Little is known about these infections in the normal host. Most infections have been identified as causing intestinal symptoms (Enterocytozoon, Encephalitozoon) or eye infections (Vittaforma, Encephalitozoon).

Organisms infect various parts of the body; diagnosis often depends on histologic examination of tissues; routine examination of clinical specimens (e.g., stool, urine) is becoming more common; infection can probably cause death.

Leishmania spp.

Asymptomatic to mild disease. Depending on species, infection can result in cutaneous, diffuse cutaneous, or mucocutaneous disease.

More serious manifestations of visceral leishmaniasis; some cutaneous species manifest visceral disease; infection is difficult to treat and manage; definite coinfection with AIDS.

Strongyloides stercoralis

Asymptomatic to mild abdominal complaints; can remain latent for many years because of low-level infection maintained by internal autoinfective life cycle.

Can result in disseminated disease (hyperinfection syndrome resulting from autoinfective nature of life cycle); abdominal pain, pneumonitis, sepsis-meningitis with gram-negative bacilli, eosinophilia; distinct link to certain leukemias or lymphomas; can be fatal.

Crusted (Norwegian) scabies

Infections can range from asymptomatic to moderate itching.

Severe infection with reduced itching response; hundreds of thousands of mites on the body; infection is very easily transferred to others; secondary infection is common.

TABLE 46.6    Pathogenesis and Spectrum of Parasitic Diseases

Parasite Group

Pathogenesis

Spectrum of Disease

Protozoa, Intestinal Amebae

Pathogens can cause severe disease; however, exposure does not always lead to disease; infection may be self-limiting; disease more likely in the compromised host.

Nonpathogens cause no disease, patients are asymptomatic; Entamoeba histolytica causes intestinal symptoms (bloody diarrhea) and the potential for amoebic liver abscess; other tissues may be involved, especially in the immunocompromised patient. “Blastocystis hominis” comprises a number of strains, some considered pathogenic; patients’ conditions range from asymptomatic to severe diarrhea.

Flagellates

Not all patients are infected upon exposure; disease spectrum varies; some patients may remain asymptomatic; if nonexposed patients become infected, symptoms are much more likely to occur.

Nonpathogens cause no disease, patients are asymptomatic; Giardia duodenalis (malabsorption syndrome) and Dientamoeba fragilis cause intestinal symptoms ranging from “indigestion” to nonbloody diarrhea, cramping, gas, and so on.

Ciliates

Neobalantidium coli infection is rare in the United States; people who have regular contact with pigs are much more likely to become infected; wide range of symptoms.

N. coli causes intestinal symptoms, including severe watery diarrhea, similar to coccidial and microsporidial infections.

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TABLE 46.6    Pathogenesis and Spectrum of Parasitic Diseases—cont’d

Parasite Group

Pathogenesis

Spectrum of Disease

Coccidia

All coccidia infective to humans can cause severe disease, particularly in the immunocompromised patient; infections in the immunocompetent patient tend to be self-limiting; Cryptosporidium spp. can maintain the infective cycle in the patient as a result of the autoinfective portion of the life cycle (the immunocompromised patient cannot produce antibody to limit this autoinfective cycle); huge water-borne outbreaks have been documented for Cryptosporidium spp.; infecting dose is low for Cryptosporidium spp.

Cryptosporidium spp., Cyclospora cayetanensis, and Cystoisospora belli cause intestinal symptoms, including severe watery diarrhea; infections are more severe in immunocompromised patients. Life-threatening infections can be seen with Cryptosporidium spp.; the organisms can disseminate to other tissues, primarily the lung. Sarcocystis can cause intestinal symptoms and/or muscle pain, depending on the mode of infection (ingestion of oocysts or infected meat).

Microsporidia

A number of genera are pathogenic for humans and for animals; wide range of body sites; disease varies, depending on patient’s immune status; disease outcome is complicated by lack of treatment for some genera; albendazole is effective for Encephalitozoon spp.

Every human tissue may be infected; Enterocytozoon bieneusi and Encephalitozoon intestinalis are the most common and are found in the intestinal tract; the latter can also disseminate to other tissues, including the kidneys. Eye infections have been seen in both healthy and compromised patients; severe corneal infections seen.

Protozoa, Other Sites Amebae

Pathogenic for humans; disease ranging from acute meningoencephalitis to chronic encephalitis, cutaneous infections to keratitis, and the potential for other body sites; disease spectrum depends on patient’s immune capacity and the organism involved; disease can be mild (Acanthamoeba spp.) to fatal (Naegleria fowleri).

Infection occurs through contact with contaminated water; organisms enter through the nasal mucosa and may travel via the olfactory nerve to the brain. Disease caused by N. fowleri can be severe and life-threatening (primary amoebic meningoencephalitis [PAM)]; chronic granulomatous amoebic encephalitis (GAE) can be caused by Acanthamoeba spp. and Balamuthia mandrillaris; keratitis is also caused by these organisms, and infection can be linked to blindness or severe corneal damage. Eye infections can be linked to contaminated lens solutions or to direct, accidental inoculation of the eye from environmental water and/or soil sources.

Flagellates

Trichomonas vaginalis causes genitourinary disease, depending on vaginal pH, presence or absence of other organisms, sexual practices, and other factors. Disease can vary from mild to severe.

T. vaginalis is found in the genitourinary system and is usually acquired by sexual transmission. Disease can be asymptomatic in the male but can cause pain, itching, and discharge in females; some strains of drug-resistant Trichomonas have been documented.

Protozoa, Blood and Tissue Malaria, babesiosis

Plasmodium vivax, Plasmodium falciparum, Plasmodium ovale, Plasmodium knowlesi, and Plasmodium malariae are pathogenic for humans; P. falciparum malaria is the leading cause of death in endemic areas; although the host can develop antibody, protection is strain specific and short-lived.

Malaria can cause a range of symptoms, with lifethreatening illness caused by P. falciparum/P. knowlesi; symptoms include fever, chills, nausea, and central nervous system (CNS) symptoms; Babesia infections often mimic those seen with malaria but without the fever periodicity.

Flagellates (leishmaniae)

Leishmania donovani invades the spleen, liver, and bone marrow and can cause serious disease, particularly in compromised hosts.

Leishmaniasis can infect the skin and mucous membranes and the organs of the reticuloendothelial system; symptoms can be mild to life threatening.

Flagellates (trypanosomes)

Humans are the only known hosts for Trypanosoma brucei gambiense (West African trypanosomiasis); Trypanosoma brucei rhodesiense (East African trypanosomiasis) infections are found in a number of antelope and other hoofed mammals that serve as reservoir hosts. Trypanosoma cruzi (American trypanosomiasis) can be found in rodents and chickens.

T. brucei gambiense and T. brucei rhodesiense cause African sleeping sickness, with eventual invasion of the CNS, leading to coma and death; Chagas disease (T. cruzi) causes acute to chronic problems, primarily linked to cardiac disease and diminished cardiac capacity; the muscles of the gastrointestinal (GI) tract are also infected, leading to loss of function in terms of movement of food through the GI tract. Continued

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TABLE 46.6    Pathogenesis and Spectrum of Parasitic Diseases—cont’d

Parasite Group

Pathogenesis

Spectrum of Disease

Nematodes, intestinal

These worms can cause mild to severe disease, depending on the worm burden (original number of eggs ingested or infective larvae penetrating the skin); young children and debilitated patients are more likely to be symptomatic; severe infections are seen in hyperinfections caused by Strongyloides stercoralis (autoinfective life cycle and immunocompromised patients); outcome varies tremendously from patient to patient and depends on the original infective dose.

Ascaris lumbricoides, Trichuris trichiura, and hookworm symptoms range from none to diarrhea, pain, and so on, depending on the worm burden; anemia may be seen with severe hookworm infection; S. stercoralis infections can involve many body tissues (disseminated disease) in the immunocompromised patient and can cause death; pinworm infection (Enterobius vermicularis) symptoms range from none to anal itching, irritability, loss of sleep, and so on.

Nematodes, tissue

Depending on the infective dose, Trichinella spp. can cause mild to severe disease; both cutaneous larva migrans (CLM) (dog/cat hookworm larvae) and toxocariasis (visceral larva migrans [VLM], ocular larva migrans [OLM]) through ingestion of dog/cat ascarid eggs cause serious disease if not treated; CLM, VLM, and OLM are seen in children more than adults.

Trichinella spp. can cause eosinophilia, muscle aches and pains, and death, depending on the worm burden; CLM can cause severe itching and eosinophilia as a result of larval migration in the skin; VLM and OLM are caused by larval migration throughout the body, including the eyes (mimics retinoblastoma).

Nematodes, filarial

Wuchereria bancrofti, Loa loa, and Onchocerca volvulus cause human disease; however, some filarial infections are not well adapted to humans and require many years of exposure before disease is evident; some infections are not evident, some cause multiple disease manifestations.

Symptoms range from asymptomatic to elephantiasis, blindness, skin changes, lymphadenitis, and lymphangitis; in some cases, Loeffler syndrome may also be seen.

Cestodes, intestinal

The beef (Taenia saginata), pork (Taenia solium), and freshwater fish (Diphyllobothrium latum) tapeworms infect humans and are generally found in the intestine as a single worm. In the case of cysticercosis, ingestion of T. solium eggs can cause mild to severe disease, depending on the infecting dose and body site (muscle, CNS). Hymenolepis nana (dwarf tapeworm) infection can lead to many worms in the intestinal tract (autoinfective cycle; the organism can go from egg to larval form to adult in the human host).

Human infection with the adult tapeworm can cause no symptoms, or mild intestinal symptoms may occur. When the human becomes the accidental intermediate host for T. solium, CNS symptoms may occur, including epileptic seizures. Infection with the adult D. latum tapeworm can cause intestinal symptoms, such as pain, diarrhea, and so on, but the patient may also be asymptomatic; a vitamin B12 deficiency may be seen. Infection with H. nana is primarily acquired from accidental ingestion of eggs from an adult tapeworm; symptoms may be absent, or diarrhea may be present.

Cestodes, tissue

Echinococcus spp. can cause severe disease, depending on the original infecting dose of tapeworm eggs; multiple organs can be involved, including brain, liver, lung, and bone; some cysts grow like a metastatic tumor; surgical removal can be very difficult if not impossible.

Depending on the body site, hydatid cysts can cause pain, anaphylactic shock (fluid leakage), or CNS symptoms. The patient may be unaware of infection until a cyst begins to press on other body organs or a large fluid leak occurs.

Trematodes, intestinal

Many genera and species are pathogenic for humans; disease severity depends on the infective dose of metacercariae; some patients may be unaware of the infection.

Intestinal trematodes can cause pain and diarrhea; intestinal toxicity can sometimes be seen in heavy infections with Fasciolopsis buski.

Trematodes, liver and lung

Many genera and species are pathogenic for humans; disease severity depends on the infective dose of metacercariae; some patients may be unaware of the infection.

Paragonimus spp. infection in the lungs can be severe, resulting in coughing, shortness of breath, and other symptoms; liver fluke infection can involve the bile ducts and gallbladder; symptoms depend on worm burden.

Trematodes, blood

Schistosomes are pathogenic for humans; however, the loading dose of cercariae from infected water sources determines the outcome of disease; very light infections may not produce symptoms; heavy infections can lead to death.

Symptoms may range from asymptomatic in light infections to severe organ failure resulting from deposition of eggs and subsequent granuloma formation in the tissues; “pipe-stem” fibrosis is seen in blood vessels; collateral circulation may develop; severe disease can cause death.

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TABLE 46.7    Recommendations for Stool Testing

Patient and/or Situation

Test Ordereda

Follow-Up Test

Patient with diarrhea and acquired immunodeficiency syndrome (AIDS) or other cause of immune deficiency. Potential water-borne outbreak (municipal/city water supply).

Cryptosporidium or Giardia/Cryptosporidium immunoassay.

If immunoassays are negative and symptoms continue, special tests for microsporidia (modified trichrome stain) and other coccidia (modified acid-fast stain), in addition to ova and parasites (O&Ps) examination, should be performed.

Patient with diarrhea (nursery school, day care center, camper, backpacker). Patient with diarrhea and potential waterborne outbreak (resort setting). Patient with diarrhea from areas where Giardia sp. is the most common parasite found.

Giardia or Giardia/Cryptosporidium immunoassay (perform testing on two stools before reporting as negative). Particularly relevant for areas of the United States where Giardia sp. is the most common organism found.

If immunoassays are negative and symptoms continue, special tests for microsporidia and other coccidia (see previous entry) and O&P examination should be performed.

Patient with diarrhea and relevant travel history. Patient with diarrhea who is a past or present resident of a developing country. Patient in an area of the United States where parasites other than Giardia sp. are found.

O&P examination, Entamoeba histolytica immunoassay; various tests for Strongyloides spp. may be relevant (even in the absence of eosinophilia).

If examinations are negative and symptoms continue, special tests for coccidia and microsporidia should be performed.

Patient with unexplained eosinophilia and possible diarrhea; if chronic, patient may also have history of respiratory problems (larval migration) and/or sepsis or meningitis (hyperinfection).

Although the O&P examination is a possibility, the agar plate culture for Strongyloides stercoralis is recommended (it is more sensitive than the O&P examination).

If tests are negative and symptoms continue, additional O&P examinations and special tests for microsporidia and other coccidia should be performed.

Patient with diarrhea (suspected food-borne outbreak).

Test for Cyclospora cayetanensis (modified acid-fast stain, autofluorescence).

If tests are negative and symptoms continue, special procedures for microsporidia and other coccidia and O&P examination should be performed.

aDepending

on the particular immunoassay kit used, various single or multiple organisms may be included. Selection of a particular kit depends on many variables, such as clinical relevance, cost, ease of performance, training, personnel availability, number of test orders, training of physician clients, sensitivity, specificity, equipment, and time to result. Very few laboratories handle this type of testing in exactly the same way. Many options are clinically relevant and acceptable for good patient care. It is critical that the laboratory report indicate specifically which organisms could be identified using the kit; a negative report should list the organisms relevant to that particular kit.

Note: Two ordering/collection/processing/examination situations are considered STAT orders (i.e., they require immediate attention for potentially life-threatening situations): central nervous system (CNS) specimens to be examined for free-living amebae and blood films in potential malaria or other cases involving blood parasites. 

Specimen Processing Diagnostic parasitology procedures designed to detect organisms in clinical specimens typically depend on morphologic criteria and visual identification (Evolve Procedures 46.1 to 46.10). Many clinical specimens, such as those from the intestinal tract, contain numerous artifacts that complicate the differentiation of parasites from surrounding debris. Specimen preparation may require concentration methods designed to increase the chance of finding the organism(s) by removing some fecal debris. Microscopic examination requires review of the prepared clinical specimen using multiple magnifications; organism identification also depends on the skill of the microbiologist. Final identification is

based on microscopic examination of stained preparations to identify key characteristics of the parasite form. (Table 46.3 includes specific details on specimen processing.) 

Approach to Identification Protozoa are small, ranging from 1.5 μm (microsporidia) to approximately 80 μm (Neobalantidium coli, a ciliate) in size. Some are intracellular and require multiple detection and staining methods for identification. Helminth infections are usually diagnosed by finding eggs, larvae, and/or adult worms in various clinical specimens, primarily from the intestinal tract (Fig. 46.2). Identification to the species level may require microscopic examination of the specimen. Recovery and identification of blood parasites can require concentration, culture, and microscopy. Confirmation of suspected parasitic infections depends on the proper collection, processing, and examination of clinical specimens; often multiple specimens may be necessary to find and confirm the existence of a parasitic infection and the identity of the suspected organism or organisms (Table 46.10).

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TABLE 46.8    Stool Specimen Collection and Testing Options

Option

Pros

Cons

Rejection of stools from inpatients who have been in house for longer than 3 days.

Data suggest that patients who begin to have diarrhea after they have been inpatients for a few days are not symptomatic from parasitic infections but generally from other causes.

The chance always exists that the problem is related to a health care–associated (nosocomial) parasitic infection (rare); Cryptosporidium spp. and microsporidia may be possible considerations.

Examination of a single stool (ova and parasites [O&Ps]). Data suggest that 40%–50% of organisms present are found with only a single stool examination. Two O&Ps examinations (concentration, permanent stained smear) are acceptable but not always as good as three specimens (may be a relatively cost-effective approach); any patient remaining symptomatic requires additional testing.

Some believe that most intestinal parasitic infections can be diagnosed from examination of a single stool. If the patient becomes asymptomatic after collection of the first stool, subsequent specimens may not be necessary.

Diagnosis from a single stool examination depends on the experience of the laboratory scientist, proper collection, and the parasite load in the specimen. In a series of three stool specimens, commonly all three specimens are not positive and/or may be positive for different organisms.

Examine a second stool only after the first is negative and the patient is still symptomatic.

With additional examinations, yield of protozoa increases (Entamoeba histolytica, 22.7%; Giardia lamblia, 11.3%; and Dientamoeba fragilis, 31.1%).

Assumes the second (or third) stool is collected within the recommended 10-day period for a series of stools; protozoa are shed periodically. May be inconvenient for patient.

Examination of a single stool and an immunoassay (enzyme immunoassay [EIA], fluorescent antibody [FA], lateral or vertical flow cartridge). This approach is a mix: one immunoassay may be acceptable; however, immunoassay testing of two separate specimens may be required to confirm the presence of Giardia antigen. One O&P examination is generally insufficient.

If the examinations are negative and the patient’s symptoms subside, probably no further testing is required.

Patients may show symptoms (off and on), so ruling out parasitic infections with only a single stool and one fecal immunoassay may be difficult. If the patient remains symptomatic, then even if two Giardia immunoassays are negative, other protozoa may be missed (Entamoeba histolytica/Entamoeba dispar group, Dientamoeba fragilis, Cryptosporidium spp., microsporidia). It is not recommended to perform both the O&P and fecal immunoassay automatically as a stool examination for parasites.

Pool three specimens for examination; perform one concentration and one permanent stain (the laboratory pools the specimens).

Three specimens are collected by the patient (three separate collection vials) over 7–10 days; pooling by the laboratory may save time and expense.

Organisms present in low numbers may be missed because of the dilution factor once the specimens have been pooled.

Microscopic Examination High-quality clinical-grade binocular compound brightfield microscopes, kept in good working condition, are essential for examining specimens for parasites. Organism identification depends on morphologic differences found after careful examination using a regular brightfield microscope at low (100×), high dry (400×), and oil immersion (1000×) magnifications. The use of a 50× or 60× oil-immersion objective for scanning can be very helpful, particularly when the 50× oil and 100× oil-immersion objectives are side by side. The microscope should be set up using Köhler illumination, which maximizes the brightness and uniformity of the light that contacts the specimen, providing optimal viewing conditions. A stereoscopic microscope is recommended for larger specimens (e.g., arthropods, tapeworm proglottids,

various artifacts). The total magnification usually varies from approximately 10× to 45×, either with a zoom capacity or with fixed objectives (0.66×, 1.3×, 3×) used with 5× or 10× oculars. Depending on the density of the specimen or object to be examined, the light source must be directed from under the stage or onto the top of the stage. Because size is an essential factor for the accurate description and identification of parasites, especially for the characterization of fecal protozoa, a stage micrometer and calibrated ocular must be used for final identification. The stage micrometer typically has a 0.1-mm line with increments every 0.01 mm. This stage micrometer line is then viewed using the ocular at each objective magnification on the microscope to calibrate the ocular’s scaled line. This allows the microbiologist to determine the exact size of the parasitic element using the ocular (Fig. 46.3).

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

619

TABLE 46.9    Fecal Fixatives Used in Diagnostic Parasitology (Intestinal Tract Specimens)

Fixative

Concentration

Permanent Stained Smear Trichrome, Iron-Hematoxylin, Special Stains/Coccidia and Microsporidia

Immunoassays: Giardia lamblia Cryptosporidium spp.

5% or 10% formalin

Yes

No

Yes

Concentrations and IAs (EIA, FA, Rapids)

5% or 10% buffered formalin

Yes

No

Yes

Concentrations and IAs (EIA, FA, Rapids)

MIF

Yes

Polychrome IV stain

ND

No published data

SAF

Yes

Iron-hematoxylin best

Yes

Concentrations, permanent stains, and IAs (EIA, FA, Rapids)

Schaudinn’s (Hg base) no PVAa

Rare

Yes

No

Permanent stains; Hg interferes with IAs; primarily used with fresh stool specimens (no fixative collection vials)

Schaudinn’s (Hg base) with PVAa

Rare

Yes

No

Permanent stains; Hg and PVA interfere with IAs; considered gold standard fixative for permanent stains

Schaudinn’s (Cu base) with PVAb

Rare

Yes

No

Permanent stains; PVA interferes with IAs; stains not as good as with Schaudinn’s fixative using Hg or Zn

Schaudinn’s (Zn base) with PVAc

Rare

Yes

No

Permanent stains; PVA interferes with IAs; the same fixative as TOTAL-FIX without PVA (see Commentary)

Ecofriendly ECOFIX (PVA)d

Rare

Yes

No

Permanent stains; PVA interferes with IAs; works best with ECOSTAIN; Wheatley’s trichrome second best

Universal fixative,e ecofriendly TOTAL-FIX

Yes

Yes

Yes

No formalin, no mercury, no PVA; concentrations, permanent stains, special stains, fecal IAs

Comments

Commentary The most common collection option (original public health approach) is a two-vial system: one vial of 5% or 10% formalin or buffered formalin and one vial of fixative containing the plastic adhesive polyvinyl alcohol (PVA). The formalin vial is used for concentration and fecal immunoassays, and the PVA vial is used for the permanent stained smear. Regulations for formalin originally were developed for industry, not the clinical laboratory, where amounts of formalin tend to be quite low. However, a laboratory using any amount of formalin must be monitored. SEMIUNIVERSAL FIXATIVES Examples of a semiuniversal fixative include sodium acetate–acetic acid–formalin (SAF) (no mercury or PVA; contains formalin) and ECOFIX (no mercury or formalin; contains PVA). UNIVERSAL FIXATIVE Currently, TOTAL-FIX is the only fixative that contains no formalin, no PVA, and no mercury. TOTAL-FIX can be used without adding PVA to the fixative; adequate drying time for smears before staining is the most important step (minimum of 1 hour in 37°C incubator; more time is required for thicker fecal smears). This fixative can be used for concentration, permanent stained smear, special stains for coccidia or microsporidia, and fecal immunoassays for Giardia and Cryptosporidium spp. FORMALIN FIXATIVE Formalin has been used for many years as an all-purpose fixative that is appropriate for helminth eggs and larvae and for protozoan cysts, oocysts, and spores. Two concentrations are commonly used: 5%, which is recommended for preservation of protozoan cysts, and 10%, which is recommended for helminth eggs

Continued

620 PA RT I V    Parasitology

TABLE 46.9    Fecal Fixatives Used in Diagnostic Parasitology (Intestinal Tract Specimens)—cont’d and larvae. Although 5% is often recommended for all-purpose use, most commercial manufacturers provide 10%, which is more likely to kill all helminth eggs. To help maintain organism morphology, the formalin can be buffered with sodium phosphate buffers (i.e., neutral formalin). Selection of specific formalin formulations is at the user’s discretion. Aqueous formalin permits examination of the specimen as a wet mount only, a much less accurate technique than a permanent stained smear for identifying intestinal protozoa. However, the fecal immunoassays for Giardia duodenalis and Cryptosporidium spp. can be performed from the aqueous formalin vial. Fecal immunoassays for the Entamoeba histolytica/E. dispar group are limited to fresh or frozen fecal specimens or Cary-Blair transport medium. After centrifugation, special stains for the coccidia (modified acid-fast stains) and microsporidia (modified trichrome stains) can be performed from the concentrate sediment obtained from formalin-preserved stool material. Use of the sediment provides a more sensitive test. OCCUPATIONAL SAFETY AND HEALTH ADMINISTRATION REGULATIONS ON THE USE OF FORMALDEHYDE Formaldehyde has been in use for more than a century as a disinfectant and preservative, and it is found in a number of industrial products. Disagreement exists about the carcinogenic potential of lower levels of exposure, and epidemiologic studies of the effects of formaldehyde exposure among humans have given inconsistent results. Studies of industry workers with known exposure to formaldehyde report little evidence of increased cancer risk. In addition, people with asthma appear to respond no differently than healthy individuals after exposure to concentrations of formaldehyde up to 3 ppm. The federal Occupational Safety and Health Administration (OSHA) requires all workers to be protected from dangerous levels of vapors and dust. Formaldehyde vapor is the most likely air contaminant to exceed the regulatory threshold in a laboratory, particularly in anatomic pathology. Current OSHA regulations require vapor levels not to exceed 0.75 ppm (measured as a time-weighted average [TWA]) and 2 ppm (measured as a 15-minute short-term exposure). OSHA requires monitoring for formaldehyde vapor wherever formaldehyde is used in the work place. The laboratory must have evidence at the time of inspection that formaldehyde vapor levels have been measured. Both 8-hour and 15-minute exposures must have been determined. If each measurement is below the permissible exposure limit and the 8-hour measurement is below 0.5 ppm, no further monitoring is required as long as laboratory procedures remain constant. If the 0.5-ppm, 8-hour TWA or the 2-ppm, 15-minute level is exceeded, monitoring must be repeated semiannually. If either the 0.75-ppm, 8-hour TWA or the 2-ppm, 15-minute level is exceeded (very unlikely in a routine microbiology laboratory setting), employees must be required to wear respirators. Accidental skin contact with aqueous formaldehyde must be prevented with the use of proper clothing and equipment (gloves, laboratory coats). The amendments of 1992 add medical removal protection provisions to supplement the existing medical surveillance requirements for employees suffering significant eye, nose, or throat irritation and for those experiencing dermal irritation or sensitization from occupational exposure to formaldehyde. In addition, these amendments establish specific hazard-labeling requirements for all forms of formaldehyde, including mixtures and solutions composed of at least 0.1% formaldehyde in excess of 0.1 ppm. Additional hazard labeling, including a warning label that formaldehyde presents a potential cancer hazard, is required where formaldehyde levels, under reasonably foreseeable conditions of use, may potentially exceed 0.5 ppm. The final amendments also provide for annual training of all employees exposed to formaldehyde at levels of 0.1 ppm or higher. Note: The use of monitoring badges may not be a sensitive enough method to correctly measure the 15-minute exposure level. Contact the OSHA office in your institution for monitoring options. Usually, the accepted method involves monitoring airflow in the specific area or areas in the laboratory where formaldehyde vapors are found. POLYVINYL ALCOHOL ADHESIVE (NOT A FIXATIVE) PVA is a water-soluble, synthetic polymer used as a viscosity-increasing agent in pharmaceuticals, as an adhesive in parasitology fecal fixatives, and as a lubricant and protectant in ophthalmic preparations. PVA is defined as a water-soluble polymer made by hydrolysis of a polyvinyl ester (e.g., polyvinyl acetate). It is used in adhesives, as a textile and paper sizer, and for emulsifying, suspending, and thickening solutions. PVA is not a fixative, but rather an adhesive to help glue stool material onto the slide; this is the only purpose of PVA as an additive to parasitology fecal fixative formulations. PVA is a plastic resin that is incorporated into Schaudinn’s fixative. Although some laboratories may perform a fecal concentration from a PVA-preserved specimen, some parasites do not concentrate well, and some do not exhibit the typical morphology that would be seen in concentration sediment from a formalinbased fixative. PVA fixative solution is highly recommended as a means of preserving cysts and trophozoites for later examination. Use of PVA fixative also allows specimens to be shipped (by regular mail service) from any location in the world to a laboratory for examination. PVA fixative is particularly useful for liquid specimens and should be used in the ratio of three parts PVA to one part fecal specimen. Note: Very detailed information on all fixative options can be found in Garcia LS. Diagnostic Medical Parasitology. 6th ed. Washington, DC: ASM Press; 2016. Cu, Copper; EIA, enzyme immunoassay; FA, fluorescent antibody; Hg, mercury; IA, immunoassay; MIF, merthiolate-iodine-formalin fixative; ND, no data; PVA, polyvinyl alcohol; Rapids, cartridge-format, membrane-flow IAs; SAF, sodium acetate–acetic acid–formalin; Zn, zinc. aThese two fixatives use the mercuric chloride base in the Schaudinn’s fixative; this formulation is still considered the gold standard against which all other fixatives are evaluated (organism morphology after permanent staining). bThis modification uses a copper sulfate base rather than mercuric chloride; the morphology of stained organisms is not as good as with Hg or Zn. cThis modification (proprietary formula) uses a zinc base rather than mercuric chloride and works well with both trichrome and iron-hematoxylin. dThis fixative uses a combination of ingredients but is prepared from a proprietary formula (contains PVA). eThis modification uses a combination of ingredients, including zinc, but is prepared from a proprietary formula. The aim is to provide a universal fixative that can be used for the fecal concentration, permanent stained smear, and available immunoassays for Giardia duodenalis, Cryptosporidium spp., and Entamoeba histolytica. However, currently, fecal immunoassays for the E. histolytica require fresh or frozen specimens; testing can also be performed from stool submitted in Cary-Blair transport medium.

Microscopy for parasitology is labor intensive and requires significant knowledge and skills for the identification of the organisms. Techcyte Inc. (Lindon, Utah), has developed a digital artificial intelligence system for identifying O&Ps from stool samples. The system preclassifies the slides by screening out negative samples before the technologist reviews the positive slides for organism identification. The system is designed to reduce technologist time and cost.

Intestinal Tract Stool specimens are the most common specimen submitted to the diagnostic laboratory for parasite identification.

As a result, the most commonly performed procedure in parasitology is the traditional examination for O&Ps, although the use of rapid fecal immunoassays and molecular techniques has greatly increased over the past 10 years. Several other diagnostic techniques are available for the recovery and identification of parasitic organisms from the intestinal tract. Although many laboratories do not routinely offer all these relatively simple and inexpensive techniques, the clinician should be familiar with the relevance of information obtained from them. It is rarely necessary to examine stool specimens for scolices and proglottids of cestodes and adult nematodes and trematodes in order to

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

60 30

0 Opisthorchis viverrini

Clonorchis sinensis

Capillaria philippinensis

Taenia spp.

Hymenolepis nana

90 60 30

0 Trichuris trichiura

Ascaris lumbricoides fertile

Enterobius vermicularis

Hookworm

Diphyllobothrium latum

90 60 30

0 Hymenolepis diminuta

Trichostrongylus spp.

Nanophyetus salmincola

Paragonimus westermani

Ascaris lumbricoides infertile

150 120 90 60 30 0 Dipylidium caninum

Echinostoma spp.

Fasciolopsis buski

Fasciola hepatica

150 120 90 60

30 0 Schistosoma japonicum

•

Schistosoma mekongi

Schistosoma haematobium

Schistosoma intercalatum

Schistosoma mansoni

Fig. 46.2  Relative egg size of helminths that infect humans. Measurements are in micrometers (μm). (From Centers for Disease Control and Prevention. Intestinal Parasites: Comparative Morphology Figures. Figure 6: Relative Sizes of Helminth Eggs. https://www.cdc.gov/dpdx/diagnosticprocedures/stool /morphcomp.html. Accessed October 26, 2020.)

621

Positive Specimen

Comments

Stool; sigmoidoscopy specimens.

O&Ps examination; stained sigmoidoscopy slides; stool immunoassays.

Trophozoites and/or cysts.

Many of the protozoa can look very much alike; see diagnostic tables for details; immunoassays for E. histolytica/E. dispar group and E. histolytica (fresh, frozen stool, Cary Blair required).

CNS; eye.

CSF, corneal scrapings, biopsy, eye care solutions; CSF examination STAT request.

Stains, culture, FA, biopsy; B. mandrillaris cannot be grown on agar culture, whereas N. fowleri and Acanthamoeba species can.

Trophozoites or cysts.

CNS disease life threatening with N. fowleri (PAM); other CNS infections more chronic (GAE); keratitis can lead to blindness.

Ingestion of food or water contaminated with infective cysts or trophozoites (D. fragilis, T. hominis); fecal-oral transmission.

Intestinal tract.

Stool; duodenal specimens or Entero-Test capsule (string test) for Giardia duodenalis.

O&P examination; wet preparations or stains of duodenal material; stool immunoassays (can use fresh or formalin fixed specimens; no PVA); need two specimens for IA for G. duodenalis.

Trophozoites or cysts

G. duodenalis is very difficult to recover; stool immunoassays are more sensitive than routine O&Ps examinations; D. fragilis requires permanent stain for identification.

Sexually transmitted; wet towels less likely but possible.

Urinary tract; genital system; males may be asymptomatic.

Vaginal secretions, prostatic fluid, often recovered in urine sediment

Wet preparations, culture, immunoassays; molecular testing.

Trophozoites

Often diagnosed by motility in urine sediment or wet preparations.

Ingestion of food or water contaminated with infective cysts; fecal-oral transmission.

Intestinal tract.

Stool.

O&Ps examination; wet preparations better than permanent stained smear.

Trophozoites and/or cysts

Not common in the United States; associated with pigs; seen in proficiency testing specimens.

Infection Acquired

Location in Host

Intestinal Amebae

Ingestion of food or water contaminated with infective cysts; fecal-oral transmission.

Intestinal tract; E. histolytica infection may disseminate to the liver (extraintestinal amebiasis); Blastocystis strains are pathogenic or nonpathogenic; cannot differentiate on morphology.

Contaminated water or soil; dust, contaminated eye solutions; organisms may enter through nasal mucosa, travel to brain via olfactory nerve.

Entamoeba histolytica Entamoeba bangladeshi Entamoeba dispar Entamoeba hartmanni Entamoeba coli Entamoeba moshkovskii Entamoeba polecki Endolimax nana Iodamoeba bütschlii Blastocystis hominis

Free-Living Amebae Naegleria fowleri Acanthamoeba spp. Balamuthia mandrillaris Sappinia spp.

Intestinal Flagellates Giardia duodenalis Dientamoeba fragilis Chilomastix mesnili Pentatrichomonas hominis

Urogenital Flagellates Trichomonas vaginalis

Intestinal Ciliate Neobalantidium coli

Diagnostic Specimen

Diagnostic Testa

Organism

622 PA RT I V    Parasitology

TABLE 46.10    Common Human Parasites: Diagnostic Specimens, Tests, and Positive Findings

TABLE 46.10    Common Human Parasites: Diagnostic Specimens, Tests, and Positive Findings—cont’d

Comments

Stool; biopsy; duodenal specimen; sputum.

Modified acid-fast stains; stool immunoassays; concentration, wet prep for C. belli.

Oocysts in stool or scrapings; other developmental stages in tissues.

Cryptosporidium spp. cause severe diarrhea in compromised patient; nosocomial transmission.

Intestinal tract; organisms can disseminate to other body sites (kidney).

Stool; biopsy.

Modified trichrome stains; optical brightening agents; experimental immunoassays; biopsy and histology (tissue Gram stains).

Spores in stool; other developmental stages in tissues.

Can cause serious diarrhea in compromised host.

Ingestion of food or water contaminated with infective spores; fecal-oral transmission; inhalation; direct environmental contact to eyes; probably hands to eyes.

All tissues.

All body fluids and/ or tissues relevant, depending on body site.

Modified trichrome stains; optical brightening agents; experimental immunoassays; biopsy and histology (tissue Gram stains).

Spores in stool, urine, other body fluids; developmental stages in tissues.

Can cause serious diarrhea in compromised host; a number of eye infections documented in immunocompetent patients.

Ingestion of raw meat; oocysts from cat feces.

Eye, CNS in compromised patient.

Biopsies (any tissue), CSF.

Serology, tissue culture; recovery from CSF.

Positive serology; recovery of trophozoites in CSF.

Many people have positive serologies for T. gondii; infections are serious in immunocompromised patients.

Ingestion of food or water contaminated with infective eggs; penetration of skin by infective larvae in soil.

Intestine; S. stercoralis may disseminate (hyperinfection), primarily in immunocompromised patients.

Stool; duodenal contents (S. stercoralis); cellophane tape preps or paddles for E. vermicularis.

O&Ps examination; special concentrates and cultures; examination of tapes for E. vermicularis.

Adult worms, eggs and/or larvae, depending on the roundworm involved.

Review direct and indirect life cycles (migration through heart, lung, trachea to intestine), 4–6 consecutive tapes required to rule out Enterobius infection.

Location in Host

Intestinal Coccidia

Ingestion of food or water contaminated with infective oocysts; fecal-oral transmission.

Intestinal tract; Cryptosporidium spp. can disseminate to other tissues in compromised host (lung, gallbladder).

Ingestion of food or water contaminated with infective spores; fecal-oral transmission.

Intestinal Microsporidiab Enterocytozoon bieneusi Encephalitozoon intestinalis

Microsporidia; Other Body Sites Encephalitozoon spp. Anncaliia vesicularum Microsporidium Pleistophora Trachipleistophora Vittaforma Tubulinosema

Tissue Protozoa Toxoplasma gondii

Intestinal Nematodes Enterobius vermicularis Trichuris trichiura Ascaris lumbricoides Hookworm Strongyloides stercoralis

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Continued

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

Positive Specimen

Infection Acquired

Cryptosporidium spp. Cyclospora cayetanensis Cystoisospora belli

Diagnostic Specimen

Diagnostic Testa

Organism

Diagnostic Specimen

Diagnostic Testa

Positive Specimen

Comments

Migration through tissues.

Serum.

Serology.

Positive serology.

Skin penetration of larvae.

Skin tracks, migration.

Visual inspection.

Presence of tracks/ skin.

Eosinophilia, visual tracks.

Human is accidental host for VLM, OLM, and CLM.

Trichinella spp.

Ingestion of raw pork.

Muscle.

Serum, muscle biopsy.

Serology, squash prep.

Positive serology, larvae.

Outbreaks still occur.

Anisakis, others

Ingestion of raw marine fish.

Intestine.

Submission of larvae.

ID of larvae.

Positive larval ID.

Sometimes identified only after surgical removal.

Intestinal Cestodes

Ingestion of:

Intestine.

Taenia saginata (beef)

Raw beef.

Stool and/or proglottids.

O&Ps, India ink proglottids.

Eggs, proglottid branches.

Taenia solium (pork)

Raw pork.

Stool and/or proglottids.

O&P, India ink proglottids.

Eggs, proglottid branches.

Diphyllobothrium latum

Raw freshwater fish.

Stool and/or proglottids.

O&P.

Eggs, proglottid shape.

Hymenolepis nana

Tapeworm eggs.

Stool.

O&Ps.

Eggs.

Hymenolepis diminuta

Grain beetles.

Stool.

O&Ps.

Eggs.

Dipylidium caninum

Fleas from dogs/cats.

Stool and/or proglottids.

O&Ps.

Eggs, proglottid shape.

Tissue Cestodes

Ingestion of:

Echinococcus granulosus

Eggs from dog tapeworm.

Serology, centrifugation of fluid; histology.

Eggs from fox tapeworm.

Positive serology; hydatid sand, tapeworm. Tissue.

E. granulosus (enclosed cyst).

Echinococcus multilocularis

Serum, hydatid cyst aspirate; biopsy.

Taenia solium (pork)

Eggs from human tapeworm.

Serum, scans, biopsy.

Serology, films, histology.

Positive serology, positive scans, tapeworm tissue.

Small, enclosed cysticerci (cysticercosis).

Organism

Infection Acquired

Location in Host

VLM, OLM (Toxocara spp.)

Ingestion of infective eggs.

CLM (dog/cat hookworm)

Tissue Nematodes

Liver, lung, and so on.

CNS, subcutaneous tissues.

Eggs for the two Taenia spp. look alike; gravid proglottid or scolex is needed for identification.

E. multilocularis (cyst wanders through tissue).

624 PA RT I V    Parasitology

TABLE 46.10    Common Human Parasites: Diagnostic Specimens, Tests, and Positive Findings—cont’d

TABLE 46.10    Common Human Parasites: Diagnostic Specimens, Tests, and Positive Findings—cont’d

Organism

Infection Acquired

Intestinal Trematodes

Ingestion of ­metacercariae:

Fasciolopsis buski

On water chestnuts.

Metagonimus yokogawai

In raw fish.

Location in Host

Diagnostic Specimen

Intestine.

Diagnostic Testa

Positive Specimen

Comments

Stool.

O&Ps examination; M. yokogawai, H. heterophyes eggs very small; use high dry power.

Eggs in stool.

Eggs of F. buski look identical to those of the liver fluke, Fasciola hepatica.

Eggs of F. hepatica look almost identical to those of F. buski; lung fluke eggs in sputum resemble brown metal filings.

Heterophyes heterophyes

Liver and Lung Trematodes

Ingestion of metacercariae.

Fasciola hepatica

On watercress.

Liver.

Stool.

O&Ps.

Eggs in stool.

Clonorchis sinensis

In raw fish.

Liver, bile ducts.

Stool, duodenal drainage.

O&Ps.

Eggs in stool, and so on.

Paragonimus spp.

In raw crabs.

Lung.

Stool, sputum.

O&Ps.

Eggs in stool. and/or sputum.

Skin penetration of cercariae released from the freshwater snail intermediate host.

Veins over the large intestine.

Because the adult worms may become located in the “incorrect” veins, both urine and stool (unpreserved) should be examined.

O&Ps; hatching test for egg viability (all specimens collected with no preservatives); concentrates performed with saline, not water.

Eggs in stool and/or urine.

When schistosomiasis is suspected, stool, random urine, and 24-h urine specimen (collected with no preservatives).

Drawn immediately: STAT request. Blood draw every 6 h until confirmed as positive or negative.

Thick, thin blood films; rapid immunoassay methods (not yet FDA approved in United States); concentration methods.

Parasites present.

P. falciparum and P. knowlesi infections are medical emergencies; complete patient history mandatory (travel, prophylaxis, prior history); Giemsa or other bloodstain recommended.

Blood.

Thick, thin blood films.

Parasites present.

Can mimic ring forms of P. falciparum; patient will have no travel history outside of United States.

Blood Trematodes

Schistosoma haematobium

Bladder.

Schistosoma japonicum; Schistosoma mekongi

Small intestine.

Malaria

Preerythrocytic

Plasmodium vivax Plasmodium ovale Plasmodium malariae Plasmodium knowlesi

Infection through mosquito bite, blood transfusion, shared drug needles; transplacental.

Babesia spp.

Blood. Blood. Blood. Blood. Blood plus capillaries of deep tissues (spleen, liver, bone marrow).

Plasmodium falciparum

Babesiosis

Liver.

Tick borne; transfusion; organ transplants.

Blood.

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Continued

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

Schistosoma mansoni

Infection Acquired

Location in Host

Diagnostic Specimen

Trypanosoma brucei gambiense

Bite of tsetse fly.

Blood, lymph nodes, CNS.

Blood, node aspirate, CSF.

Trypanosoma brucei rhodesiense

Bite of tsetse fly.

Blood, lymph nodes, CNS.

Trypanosoma cruzi

Feces of triatomid bug (kissing bug) (bug feces scratched into bite site).

Blood, striated muscle (e.g., heart, GI tract).

Bite of sand fly.

Diagnostic Testa

Positive Specimen

Comments

Thick, thin films.

Trypomastigotes.

Thick, thin films.

Trypomastigotes.

Blood, cardiac changes, muscle biopsy.

Thick, thin films, histology; culture.

Trypomastigotes in blood, amastigotes in tissue.

African sleeping sickness more common with T. brucei gambiense. Chagas disease (American trypanosomiasis) (xenodiagnosis an option).

Macrophages of skin.

Skin biopsy.

Stained smears, cultures.

Leishmania braziliensis complex (mucocutaneous)

Skin, mucous membranes.

Skin, membrane biopsy.

Stained smears, cultures.

Amastigotes in clinical specimens indicated.

Animal inoculation rarely used; some research labs now using PCR.

Leishmania donovani complex (visceral)

Spleen, liver, bone marrow (RE system).

Blood, bone marrow, liver/spleen biopsy.

Thick, thin blood films; stained smears, cultures.

Lymphatics (adults), blood (microfilariae), nodules (adults), skin, eye (microfilariae).

Blood. Skin snips, blood; biopsy nodule.

Thick and thin films; various concentrations. Biopsy, tease skin apart in water; thick and thin films.

Microfilariae.

Elephantiasis possible; periodicity a factor in finding microfilariae; some microfilariae sheathed (S), some not (NS) “African eye worm.”

Eye (adults), lymphatics (adults), blood (microfilariae) for all three.

Blood.

Thick, thin films; various concentrations.

Microfilariae, adult worm. Microfilariae.

Organism Trypanosomes

Leishmaniae Leishmania tropica complex (cutaneous)

Filarial Nematodes Wuchereria bancrofti (S)

Bite of mosquito.

Onchocerca volvulus (NS)

Black fly.

Microfilariae; adult in tissue nodules.

Less Common Loa loa (S)

Black gnat.

Brugia malayi (S)

Mosquito.

Mansonella spp. (NS)

Mosquito.

CLM, Cutaneous larva migrans; CNS, central nervous system; CSF, cerebrospinal fluid; FA, fluorescent antibody; FDA, Food and Drug Administration; GAE, granulomatous amoebic encephalitis; IA, immunoassay; ID, identification; NS, not sheathed; OLM, ocular larva migrans; O&P, ova and parasite; PAM, primary amoebic meningoencephalitis; PCR, polymerase chain reaction; RE, reticuloendothelium system; S, sheathed; VLM, visceral larva migrans aAlthough serologic tests are not always mentioned, they are available for a number of parasitic infections. Unfortunately, most are not routinely available. Contact your state Public Health Laboratory or the Centers for Disease Control and Prevention (CDC) in Atlanta, Georgia. bThe microsporidia are classified with the fungi.

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TABLE 46.10    Common Human Parasites: Diagnostic Specimens, Tests, and Positive Findings—cont’d

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• BOX 46.1 Direct Smear: Review

Principle To detect the presence of motile forms of parasites (primarily protozoan trophozoites or larvae); to diagnose organisms that cannot be identified from the permanent stain methods; to allow quick diagnosis of heavily infected patients; to estimate the worm burden of a patient. 

Specimen

• Fig. 46.3  Stage and ocular micrometer.

Any fresh liquid or soft stool specimen that has not been refrigerated or frozen. 

Reagents

confirm the diagnosis or to identify the organism to the species level. Other specimens from the intestinal tract—such as duodenal aspirates or drainage, mucus from the Entero-Test Capsule technique, and sigmoidoscopy material—can also be examined as wet preparations and as permanent stained smears after processing with trichrome or iron-hematoxylin staining. Ova and Parasite Examination

The traditional O&Ps examination comprises three separate protocols: the direct wet mount, the concentration technique and indirect wet mount examination, and the permanently stained smear. The direct wet mount, which requires fresh stool, is designed to enable the detection of motile protozoan trophozoites. The specimen is examined microscopically at low and high dry magnifications (100×, entire 22- by 22-mm cover slip; 400×, one-third to one-half of a 22- by 22-mm cover slip) (Box 46.1). However, because of potential problems resulting from the lag time between specimen passage and receipt in the laboratory, the direct wet examination has been eliminated from the routine O&Ps examination in many laboratories in the United States in favor of specimens collected in stool preservatives. The direct wet preparation is not performed for specimens received in the laboratory in preservatives. The various fixatives available are included in Table 46.9. Each preservative or combination of preservatives has advantages and disadvantages for different types of parasites and for different laboratory methods. Each laboratory should consider the types of parasites most commonly seen in their location and population and should select the preservatives that maximize the detection of such parasites. The second part of the O&Ps is the indirect examination after specimen concentration, which is designed to facilitate recovery of protozoan cysts, coccidian oocysts, microsporidial spores, and helminth eggs and larvae (Box 46.2). Both flotation and sedimentation methods are available; the most common procedure is the formalin–ethyl acetate sedimentation method (previously the formalin-ether method) (Fig. 46.4). The concentrated specimen sediment can be examined as a wet preparation with or without iodine using low and high dry magnifications (100×, 400×) as indicated for the direct wet smear examination.

0.85% NaCl; Lugol’s or D’Antoni’s iodine. 

Examination Low-power examination (100×) of entire 22- by 22-mm cover slip preparation (both saline and iodine); high dry power examination (400×) of at least one-third of the cover slip area (both saline and iodine). 

Results Results from the direct smear examination should be considered presumptive. However, some organisms can be identified definitively (Giardia duodenalis cysts and Entamoeba coli cysts, helminth eggs and larvae, Cystoisospora belli oocysts). The report should be considered “preliminary”; the final report includes the results of the concentration and permanent stained smear. 

Notes and Limitations When iodine is added to the preparation, the organisms are killed and motility is lost. Specimens submitted in stool preservatives and fresh, formed specimens should not be examined using this procedure; the concentration and permanent stained smear techniques should be performed instead. Oil immersion examination (1000×) is not recommended (the organism morphology is not that clear).

The third part of the O&Ps examination is the permanently stained smear, which is designed to facilitate the identification of intestinal protozoa (Box 46.3). The permanently stained smear is the most important procedure performed to confirm the diagnosis of intestinal protozoan infections. Several staining methods are available; the two most commonly used are the Wheatley modification of the Gomori tissue trichrome stain and the iron-hematoxylin stain (Fig. 46.5). This part of the O&PS examination is critical for confirmation of suspicious objects seen in the wet examination and for identification of protozoa that may not have been visible in the wet preparation. Permanently stained smears are examined using oil immersion objectives (600× or 800× for screening and 1000× for final review of 300 or more oil-immersion fields). The permanently stained smears also provide a permanent record for reexamination or confirmation. Modified acid-fast stains are necessary for the identification of intestinal coccidia (Box 46.4), and modified trichrome stains are recommended for identification of

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• BOX 46.2 Concentration: Review

Principle To concentrate the parasites present, either through sedimentation or by flotation. The concentration is specifically designed to allow recovery of protozoan cysts, coccidian oocysts, microsporidian spores, and helminth eggs and larvae. 

Specimen Any stool specimen that is fresh or preserved in formalin (most common), polyvinyl alcohol (PVA; mercury based or non–mercury based), sodium acetate–acetic acid–formalin (SAF), merthiolateiodine-formalin (MIF), or the newer fixatives for the single-vial system—universal fixatives containing no mercury, formalin, or PVA. 

Reagents Formalin, ethyl acetate, zinc sulfate 5% or 10% (specific gravity, 1.18 for fresh stool and 1.20 for preserved stool); 0.85% sodium chloride (NaCl); Lugol’s or D’Antoni’s iodine. 

Examination Low-power examination (100×) of entire 22- by 22-mm cover slip preparation (iodine recommended but optional); high dry power examination 400×) of at least one-third of the cover slip area (both saline and iodine). 

Results Results from the concentration examination should be considered presumptive. However, some organisms can be identified definitively (Giardia duodenalis cysts and Entamoeba coli cysts, helminth eggs and larvae, Cystoisospora belli oocysts). The report should be considered “preliminary” and should be coordinated with the findings from the permanent smear. 

Notes and Limitations Formalin–ethyl acetate sedimentation concentration is most commonly used. Zinc sulfate flotation may not detect operculated or heavy eggs (Clonorchis eggs, unfertilized Ascaris eggs). For the flotation technique, both the surface film and sediment must be examined before a negative result is reported. Smears prepared from concentrated stool are normally examined at low power (100×) and high dry power (400×). Oil-immersion examination (1000×) may be used with caution because the morphology of some organisms may not be clear. The addition of too much iodine may obscure helminth eggs (i.e., it produces an effect that mimics debris).

•

Fig. 46.4  Formalin–ethyl acetate concentration test tube after centrifugation showing four layers; from the bottom: sediment, formalin layer, debris plug, and ethyl acetate layer.

through a series of wire screens (graduated from coarse to fine mesh) to look for scolices and proglottids. The appearance of scolices after therapy is an indication of successful treatment. If the scolex has not been passed, it may still be attached to the mucosa. Because the scolex can continue to produce new proglottids, the infection will continue. If this occurs, the patient must be retreated to eradicate the infection completely.  Examination for Pinworm

intestinal microsporidia (Box 46.5). These stains are specifically designed to enable the identification of coccidian oocysts and microsporidian spores, respectively. Fig. 46.6 provides a diagrammatic overview of the processing of fecal specimens for parasite identification. 

Enterobius vermicularis, pinworm or seatworm, a roundworm parasite commonly found in children worldwide. The adult female worm migrates out of the anus, usually at night, and deposits her eggs in the perianal area. The adult female (8 to 13 mm long) may occasionally be found on the surface of a stool specimen or on the perianal skin. Because the eggs are usually deposited around the anus, they are not commonly found in feces and must be detected by other diagnostic means. Diagnosis of pinworm infection is usually based on the recovery of eggs, which are described as thickshelled, football-shaped eggs with one slightly flattened side. Often, each egg contains a fully developed embryo and is infective within a few hours after being deposited (Fig. 46.7). 

Recovery of the Tapeworm Scolex

Sigmoidoscopy Material

The procedure for the recovery of the tapeworm scolex is rarely requested in the United States and is no longer clinically relevant because of the effective use of medication to treat tapeworm infections. However, stool specimens may be examined for scolices and gravid proglottids of cestodes for species identification. This procedure requires mixing a small amount of feces with water and straining the mixture

Material obtained from sigmoidoscopy can be helpful in the diagnosis of amebiasis that has gone undetected by routine fecal examination. However, a series of at least three routine stool examinations for parasites should be completed before a sigmoidoscopy examination is performed. A sigmoidoscopy specimen should be processed immediately. All three methods of examination are recommended (direct,

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• BOX 46.3 Permanent Stained Smear: Review

Principle To allow the examination and recognition of detailed organism morphology under oil-immersion examination (100× objective for a total magnification of 1000×), primarily enabling the recovery and identification of intestinal protozoa. 

Specimen Any stool specimen that is fresh or preserved in polyvinyl alcohol (PVA; mercury based or non–mercury based), sodium acetate– acetic acid–formalin (SAF), merthiolate-iodine-formalin (MIF), or the newer single vial–system fixatives (universal fixatives). 

A

Reagents Trichrome, iron-hematoxylin, modified iron-hematoxylin, polychrome IV, or chlorazol black E stains and their associated solutions; dehydrating solutions (alcohols and xylenes or xylene substitutes); mounting fluid (optional). The use of true absolute alcohol (100% ethanol) is preferred over the use of 95%/5% alcohol. 

Examination The entire smear should be examined on low power (100×) for the presence of large parasite forms such as larvae or helminth eggs. Oil-immersion examination (1000×) should include at least 300 fields; additional fields may be required if suspect organisms have been seen in the wet preparations from the concentrated specimen. 

B

Results Most suspected protozoa and/or human cells can be confirmed by examination of the permanently stained smear. These reports should be categorized as “final” and are reported as such (the direct wet smear and the concentration examination provide “preliminary” results). 

Notes and Limitations The most commonly used stains are trichrome and ironhematoxylin. Unfortunately, helminth eggs and larvae may take up the stain inconsistently and are not easily identified from the permanent stained smear. Coccidian oocysts and microsporidian spores also require other staining methods for identification. Permanent stained smears are examined under oilimmersion examination (1000×). The slide may be screened using the 50× or 60× oil-immersion objectives, but the results should not be reported until the examination has been completed using the 100× oil-immersion lens. Confirmation of intestinal protozoa (both trophozoites and cysts) is the primary purpose of this technique. 

Important Reminder When nonmercury fixatives or one of the single-vial options (usually a zinc-based proprietary formula or one of the universal fixatives) is used, the iodine-alcohol step can be eliminated. After drying, the slides can be placed directly into the stain (trichrome or hematoxylin). However, if fecal specimens have been preserved with mercury-based fixatives, the iodine-alcohol step must be included in the routine staining protocol as well as subsequent rinse steps to remove the mercury and iodine. Some laboratories leave the staining protocol as is, including the iodine-alcohol step, which does not harm smears preserved with non–mercury based fixatives.

C •

Fig. 46.5 Stool material stained with Wheatley’s trichrome stain. (A) Charcot-Leyden crystals. (B) Polymorphonuclear leukocytes. (C) Blastocystis hominis central body forms (larger objects) and yeast cells (smaller, more homogeneous objects).

concentration, permanent smear). However, depending on the availability of trained personnel, proper fixatives, or the amount of specimen obtained, one or two procedures may be used. It is important to note that even the most thorough examination will be meaningless if the specimen has been improperly collected, fixed, and transported.  Duodenal Drainage

In patients infected with G. duodenalis or S. stercoralis, routine stool examinations may not be sufficient to identify the infecting organisms. Duodenal drainage material may increase the likelihood of identifying the parasites. However, the “falling leaf ” motility often described for Giardia

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• BOX 46.4 Modified Acid-Fast Permanent Stained

• BOX 46.5 Modified Trichrome Permanent Stained

Principle

Principle

To provide contrasting colors for background debris and parasites. This technique is designed to allow examination and recognition of acid-fast characteristics of organisms under high dry power (40× objective, for a total magnification of 400×), primarily allowing recovery and identification of intestinal coccidian oocysts. The internal morphology of sporozoites can be seen within some Cryptosporidium oocysts under oil immersion (1000×); Cyclospora oocysts do not have a specific internal morphology. 

This technique is designed to allow examination and recognition of organism morphology under oil immersion examination (100× objective, for a total magnification of 1000×), primarily allowing recovery and identification of intestinal microsporidial spores. The internal morphology (horizontal or diagonal “stripes”) may be seen in some spores under oil immersion. 

Smear: Review

Specimen Any stool specimen that is fresh or preserved in formalin, sodium acetate–acetic acid–formalin (SAF), or the newer fixatives for the single-vial system (universal fixatives). 

Reagents Kinyoun’s acid-fast stain, modified Ziehl-Neelsen stain, and their associated solutions; dehydrating solutions (alcohols and xylenes or xylene substitutes); mounting fluid (optional). The decolorizing agents are less intense than the acid alcohol used in routine acid-fast staining (this is what makes these “modified” acid-fast procedures). The recommended decolorizer is 1%–3% sulfuric acid. Many laboratories use 1% so that the Cyclospora oocysts retain more color. 

Examination High dry examination (400×) of at least 300 fields; additional fields may be required if suspicious organisms have been seen but are not clearly acid-fast stained. 

Results Identification of Cryptosporidium and Cystoisospora oocysts should be possible. Cyclospora oocysts, which are twice the size of Cryptosporidium oocysts, should be visible but tend to be more acid-fast variable. Although microsporidia are acid-fast, their small size makes recognition very difficult. Final laboratory results depend heavily on the appearance of the quality control (QC) slides and comparison with positive patient specimens. 

Notes and Limitations Both the cold and hot modified acid-fast methods are excellent for staining coccidian oocysts. Some clinicians believe that the hot method may result in better stain penetration, but the differences are probably minimal. Procedure limitations are related to specimen handling (proper centrifugation speeds and time, use of no more than two layers of wet gauze for filtration, and complete understanding of the difficulties in recognizing microsporidial spores). In addition, some controversy exists over whether the organisms lose the ability to take up acid-fast stains after long-term storage in 10% formalin. The organisms are more difficult to find in specimens from patients who do not have the typical watery diarrhea (more formed stool contains more artifact material).

trophozoites in fresh stool samples is rarely seen in these preparations. The organisms may be caught in mucus strands, and the movement of the flagella on the Giardia trophozoites may be the only subtle motility visible. Strongyloides larvae are usually very motile. It is important to

Smear: Review

Specimen Any stool specimen that is fresh or preserved in formalin, sodium acetate–acetic acid–formalin (SAF), or the newer fixatives for the single-vial system (universal fixatives). 

Reagents Modified trichrome stain (using high dye–content chromotrope 2R) and associated solutions; dehydrating solutions (alcohols, xylenes, or xylene substitutes); mounting fluid (optional). 

Examination Oil immersion examination (1000×) of at least 300 fields; additional fields may be required if suspect organisms have been seen but are not clearly identified. 

Results Identification of microsporidial spores may be possible; however, their small size makes recognition difficult. Final laboratory results depend heavily on the appearance of the quality control (QC) slides and comparison with positive patient specimens. 

Notes and Limitations Because of the difficulty in getting dye to penetrate the microsporidial spore wall, this staining approach can be helpful. Procedure limitations are related to specimen handling (proper centrifugation speeds and time, use of no more than two layers of wet gauze for filtration, and complete understanding of the difficulties in recognizing microsporidial spores because of their small size [1–3 μm]). 

Commercial Suppliers Suppliers must be asked about specific fixatives and whether the fecal material can be stained with the modified trichrome stains and modified acid-fast stains. They should also be asked whether the fixatives prevent the use of any of the newer fecal immunoassay methods available for several of the intestinal amebae, flagellates, coccidia, and microsporidia.

keep the light intensity low for visualization of the moving parasites. The duodenal fluid typically contains mucus, and the organisms tend to be located in the mucus threads. Therefore, centrifugation of the specimen before examination is important. Fluorescent antibody or immunoassay detection kits (Cryptosporidium or Giardia) can also be used with fresh or formalinized material. It is important to check the package insert of each kit to see which specimen types are acceptable. If a presumptive diagnosis of giardiasis is reached as a result of a wet prep examination, the cover slip can be

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Fecal specimens (O&P)

10% formalin

PVA

Wet mount (Helminths and protozoa)

Trichrome stain (Evolve Procedure 46.2) (Protozoa)

ELISA* (Giardia and Cryptosporidium) Modified trichrome stain (Evolve Procedure 46.5) (Microsporidia)

Formalin-ethyl acetate sedimentation (Evolve Procedure 46.1) Wet mount (Helminths and protozoa) Direct mount epifluorescence* (Cyclospora and Cystoisospora)

Acid–fast stain (Evolve Procedure 46.4) (Cryptosporidium, Cyclospora, and Cystoisospora)

Safranin stain* (Cyclospora)

• Fig. 46.6  Processing of fecal specimens for ova and parasites (modified according to diagnostic guidelines established by the Centers for Disease Control and Prevention). *Indicates a special test procedure.

removed and the specimen fixed with Schaudinn’s fluid, other fixatives containing polyvinyl alcohol (PVA), or a “universal fixative” (i.e., no formalin, mercury, or PVA) for subsequent staining with trichrome or iron-hematoxylin. If the amount of duodenal material submitted is very small, permanent stains can be prepared in lieu of using a portion of the specimen for a wet prep. This approach provides a permanent record; it may also improve the visualization of parasites using an oil-immersion examination of the stained specimen at 1000× compared with the examination of unstained organisms with minimal motility at a lower magnification. 

capsule. The cord protrudes through one end of the capsule and is taped to the side of the patient’s face. The capsule is swallowed. The gelatin dissolves in the stomach, and the weighted cord is carried by peristalsis into the duodenum. The cord is attached to the weight by a slipping mechanism; the weight is released and passes out in the stool when the cord is retrieved after 4 hours. The mucus collected on the cord is examined using the direct wet mount method for parasites, including S. stercoralis, G. duodenalis, Cryptosporidium spp., microsporidia, and the eggs of Clonorchis sinensis. 

Duodenal Capsule Technique (Entero-Test)

T. vaginalis is typically identified by the examination of wet preparations of vaginal or urethral discharges, prostatic secretions, or urine sediment. Multiple specimens may be needed to detect the organisms. The specimens should be diluted with a drop of saline and examined for motile

The duodenal capsule technique is a simple, convenient method for collecting duodenal contents, thus eliminating the need for intestinal intubation. The technique involves the use of a length of nylon cord coiled inside a gelatin

Urogenital Tract Specimens

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A. Use a piece of clear (not frosted) cellophane tape approximately 4 inches (10 cm) long.

B. Hold the tape between thumbs and forefingers with sticky side facing outward.

C. Before the patient has arisen from bed in the morning (preferably while the child is still asleep), press the sticky side of the tape against the skin across the anal opening with even, thorough pressure.

D. Gently place the sticky side of the tape down against the surface of a clear glass slide. Label the slide with the patient’s name.

•

Fig. 46.7 Method of collecting a cellophane (Scotch) tape preparation for pinworm diagnosis. This method dispenses with the tongue depressor, requiring only tape and a glass microscope slide. The tape must be pressed deep into the anal folds.

organisms under low power (100×) with reduced illumination. As the jerky motility begins to diminish, the undulating membrane may often be observed under high dry power (400×). Unfortunately, the overall sensitivity of wet mount examinations is limited compared with culture and/ or molecular testing, such as polymerase chain reaction (PCR) analysis. Stained smears are usually not necessary for the identification of T. vaginalis. The number of false-positive and false-negative results reported from stained smears supports the value of confirmation by observation of motile organisms from the direct mount, culture media, or more sensitive direct antigen detection methods. 

Sputum Although not a common specimen, expectorated sputum may be submitted for parasitic examination. Organisms found in sputum that may cause pneumonia, pneumonitis, or Loeffler syndrome include the migrating larval stages of A. lumbricoides, S. stercoralis, and hookworm; the eggs of Paragonimus spp.; Echinococcus granulosus hooklets (Fig. 46.8); and the protozoa Entamoeba histolytica, Entamoeba gingivalis, Trichomonas tenax, Cryptosporidium spp., and possibly the microsporidia. Some of the smaller organisms must be differentiated from fungi such as

• Fig. 46.8  Echinococcus granulosus, cyst. (Enlarged for detail, 500×).

Candida spp., Pneumocystis jiroveci, and Histoplasma capsulatum. In a Paragonimus infection, the sputum may be

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viscous, streaked with blood, and tinged with brownish flecks, which are clusters of eggs (“iron filings”). Induced sputa are collected after patients have used appropriate cleansing procedures to reduce oral contamination. The induction protocol is critical for the success of the procedure, and well-trained individuals such as respiratory therapists are needed to recover the organisms. 

Aspirates Examination of aspirated material for the diagnosis of parasitic infections may be extremely valuable, particularly when routine testing methods have failed to demonstrate the presence of the organisms. These specimens should be transported to the laboratory immediately after collection and processed as quickly as possible. Aspirates include liquid specimens collected from a variety of body sites. Aspirates most commonly processed in the parasitology laboratory include fine-needle aspirates and duodenal aspirates. Fluid specimens collected by bronchoscopy include bronchoalveolar lavages and bronchial washings. Fine-needle aspirates may be submitted for slide preparation, culture, or both. Aspirates of cysts and abscesses for amebae may require concentration by centrifugation, digestion, microscopic examination for motile organisms in direct preparations, cultures, and microscopic evaluation of stained preparations. Aspiration of cyst material (usually liver or lung) for diagnosis of hydatid disease usually is performed when surgical techniques are used for cyst removal. The aspirated fluid is submitted to the laboratory and examined for hydatid sand (scolices) or hooklets; absence of this material does not rule out the possibility of hydatid disease because some cysts are sterile. Bone marrow aspirates for Leishmania and Trypanosoma cruzi amastigotes or Plasmodium spp. require staining with any of the blood stains (Giemsa, Wright’s, Wright-Giemsa combination, rapid stains, or Field’s stain). Giemsa stain is the preferred stain for blood parasites because it provides better visibility of intracellular details for optimal morphology. Examination of specimens may confirm an infection previously missed by examination of routine blood films. Cases of primary amoebic meningoencephalitis (PAM) are rare, but examination of spinal fluid may reveal the causative agent, Naegleria fowleri, one of the free-living amebae (Fig. 46.9). Although rare, this disease has a very high mortality rate. For this reason, a cerebrospinal fluid (CSF) sample with a request for examination for parasites is always considered a STAT procedure. 

•

Fig. 46.9  Naegleria fowleri in brain tissue (hematoxylin and eosin stain).

in Chapter 6. Tissue for examination using permanent sections or electron microscopy should be fixed as specified by the processing laboratory. In certain cases, a biopsy may be the only means of confirming a suspected parasitic problem. Specimens examined as fresh material rather than as tissue sections should be kept moist in saline and submitted to the laboratory immediately. Detection of parasites in tissue depends in part on specimen collection and adequate material to perform the recommended diagnostic procedures. Biopsy specimens are usually small and may not be representative of the diseased tissue. Multiple tissue samples often improve diagnostic results. To optimize the yield from any tissue specimen, all areas should be examined by as many procedures as possible. Tissues are collected using invasive procedures, many of which are very expensive and lengthy; consequently, these specimens deserve the most comprehensive procedures possible. A muscle biopsy for diagnosis of infection with Trichinella spp. can be processed as a routine histology slide or can be examined as a squash preparation (Fig. 46.10). Tissue submitted in a sterile container on a sterile sponge dampened with saline may be used for cultures of protozoa after mounts for direct examination or impression smears for staining have been prepared. If cultures will be processed for parasites, sterile slides should be used for smear and mount preparation. Examination of tissue impression smears is detailed in Table 46.11. 

Biopsy Specimens

Blood

Biopsy specimens are recommended for diagnosis of tissue parasite infections. Impression smears, teased, and squash preparations of biopsy tissue from skin, muscle, cornea, intestine, liver, lung, and brain can be used for this purpose, in addition to standard histologic preparations. An impression smear is a collection of cells, microorganisms, or fluids produced by pressing the surface of the tissue specimen against a slide for review. Squash preparations are described

Depending on the life cycle, a number of parasites may be recovered from blood specimens. Although organisms may be motile in fresh whole blood, species identification is typically accomplished from examination of permanently stained thick and thin blood films (Fig. 46.11). Blood films can be prepared from fresh, whole blood containing no anticoagulants, anticoagulated blood, or sediment from the various concentration procedures. The two most

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• Fig. 46.10  Trichinella spp. larvae encysted in muscle.

commonly used hematology stains are Wright’s and Giemsa stains. Many clinical laboratories use a commercial stain for hematology that is a combination of Wright’s and Giemsa. The stain of choice is Giemsa stain because it provides optimal detail of intracellular malarial and other blood-borne parasites. However, blood parasites can also be seen on blood films stained using Wright’s or other stains, including rapid staining options. Delafield’s hematoxylin stain is often used to improve visibility of the microfilarial sheath. Multiple smears should be made as soon as the specimen is received in the laboratory, but initially only one should be stained and observed for parasites. The remaining slides can be stained with routine or specialized stains depending on what is observed on the first slide. A request for examination of blood films for parasites is always a STAT request. Examination of any blood smear for parasitology should be performed by a highly experienced laboratory scientist. Thin Blood Films

In any examination of thin blood films for parasitic organisms, the initial screening should be completed using the low-power microscope objective (10×). Depending on the training and experience of the laboratory scientist, examination of a thin film for parasites usually takes 15 to 20 min (300 or more oil immersion fields at a magnification of 1000×). Some use a 50× or 60× oilimmersion objective to screen stained blood films; however, the chance is greater that small parasites (e.g., Plasmodium spp., Babesia spp., Leishmania donovani) will be missed at the lower total magnification (500× or 600×) compared with the 1000× total magnification achieved using the 100× oil-immersion objective. Microfilarial organisms are rarely present in large numbers. Frequently only a few organisms are identified in each thin film preparation. Because microfilariae are carried with the smear during preparation and typically located at the edges or feathered end of the thin film, the entire film should be scanned to ensure that no microfilariae are missed. The feathered end of the film should be examined for intracellular and extracellular parasites. Because the cells

in the feathered end are spread further apart, the morphology and size of the infected red blood cells (RBCs) may be more clearly visible (Box 46.6). Before a smear is reported as negative for the presence of parasites, a minimum of 300 fields should be examined. The request for blood film examination should be considered a STAT procedure, and all results (negative and positive) should be reported by telephone to the physician as soon as possible. If a result is positive, the appropriate government agencies (local, state, and federal) should be notified within a reasonable time, in accordance with guidelines and laws. It is important to note that one negative set of blood films is not sufficient to rule out blood parasites. Both malaria and Babesia infections have been missed with automated differential instruments, delaying treatment. Although these instruments are not designed to detect intracellular blood parasites, the inability of the automated systems to distinguish between uninfected RBCs and those infected with parasites may pose diagnostic problems. Because automated systems may not be able to detect intracellular parasites, a manual differential should always be performed whenever a blood sample is submitted for parasitology.  Thick Blood Films

In the preparation of a thick blood film, the greatest concentration of blood cells is in the center of the film. The examination should be performed at low magnification to detect microfilariae. Examination of a thick film usually requires 5 to 10 min (approximately 100 oil-immersion fields). A search for malarial organisms and trypanosomes should be completed using oil immersion (total magnification of 1000×). Intact RBCs are commonly seen at the very periphery of the thick film; if infected, such cells may prove useful in the diagnosis of malaria (Box 46.7). However, a thin film preparation should always be used to confirm speciation of malarial and other blood parasite organisms.  Buffy Coat Films

L. donovani, trypanosomes, and H. capsulatum (a fungus with intracellular elements resembling those of L. donovani) may be detected in the peripheral blood. The parasite or fungus is detected in the neutrophils and large mononuclear cells found in the buffy coat (a layer of white blood cells seen just above the red cell layer after centrifugation of whole citrated blood). With L. donovani, the nuclear material stains dark red-purple, and the cytoplasm is light blue. In contrast, H. capsulatum is visible as a large dot of nuclear material (dark red-purple) surrounded by a clear halo. Trypanosomes and microfilariae may also concentrate with the buffy coat cells due to their size. 

Direct Detection Methods Significant progress has been made in the development and application of molecular methods for the diagnosis of parasitic infections, including the use of purified or

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

TABLE 46.11    Examination of Impression Smears

Tissue

Possible Parasite

Staina

Lung

Microsporidia

Modified trichrome, acid-fast stain, Giemsa, tissue Gram stain, optical brightening agent (calcofluor), methenamine silver, electron microscopy (EM)

Toxoplasma gondii

Giemsa, immune-specific reagent

Cryptosporidium spp.

Modified acid-fast stain, immune-specific reagent

Entamoeba histolytica

Giemsa, trichrome

Toxoplasma gondii

Giemsa

Leishmania donovani

Giemsa

Cryptosporidium spp.

Modified acid-fast stain, immune-specific reagent

Entamoeba histolytica

Giemsa, trichrome

Naegleria fowleri

Giemsa, trichrome

Acanthamoeba spp.

Giemsa, trichrome

Balamuthia mandrillaris

Giemsa, trichrome

Sappinia spp.

Giemsa, trichrome

Entamoeba histolytica

Giemsa, trichrome

Toxoplasma gondii

Giemsa, immune-specific reagent

Microsporidia

Modified trichrome, acid-fast stain, Giemsa, tissue Gram stain, optical brightening agent (calcofluor), methenamine silver, EM

Encephalitozoon spp.

Modified trichrome, acid-fast stain, Giemsa, optical brightening agent (calcofluor), methenamine silver, EM

Leishmania spp.

Giemsa

Onchocerca volvulus

Giemsa

Mansonella streptocerca

Giemsa

Acanthamoeba spp.

Giemsa, trichrome

Microsporidia

Modified trichrome, acid-fast stain, Giemsa, optical brightening agent (calcofluor), methenamine silver, EM

Acanthamoeba spp.

Giemsa, trichrome

Naegleria fowleri

Giemsa, trichrome

Small intestine

Cryptosporidium parvum (both small and large intestine)

Modified acid fast, immune-specific reagent

Jejunum

Cyclospora cayetanensis

Modified acid-fast

Microsporidia Enterocytozoon bieneusi Encephalitozoon intestinalis

Modified trichrome, acid-fast stain, Giemsa, optical brightening agent (calcofluor), methenamine silver, EM

Duodenum

Giardia duodenalis

Giemsa, trichrome

Colon

Entamoeba histolytica

Giemsa, trichrome

Various genera of microsporidia Acanthamoeba spp.

Acid-fast stain, Giemsa, modified trichrome, methenamine silver, optical brightening agent (calcofluor), EM Giemsa, trichrome, calcofluor (cysts)

Trichinella spiralis

Wet examination, squash preparation

Microsporidia Pleistophora sp., Anncaliia sp., Trachipleistophora sp.

Modified trichrome, acid-fast stain, Giemsa, optical brightening agent (calcofluor), methenamine silver, EM

Liver

Brain

Skin

Nasopharynx, sinus cavities

Intestine

Cornea, conjunctiva Muscle

aWhenever

Giemsa stain is mentioned in the table, any bloodstain is acceptable: Giemsa, Wright’s, Wright-Giemsa combination, rapid blood stains.

635

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• BOX 46.6 Thin Blood Films: Review

Principle

•

Fig. 46.11 Thin and thick blood smear for the identification of blood parasites. Most commonly used for identifying Plasmodium or Trypanosoma spp.

recombinant antigens and nucleic acid probes. The presence of parasite-specific antigen indicates current disease. Nucleic acid–based parasitic diagnostic tests are primarily available in specialized research or reference centers. PCR and other nucleic acid probe tests have been reported for almost all species of parasites. The demand for implementation of direct detection using molecular methods will continue to increase as the cost of these tests decreases and automation increases.

Intestinal Parasites Immunoassays are generally simple and enable the simultaneous performance of many tests, thereby reducing overall costs. Antigen detection in stool specimens is often limited to the detection of one or two pathogens simultaneously. A routine O&P examination should also be performed to detect other parasitic pathogens. The current commercially available antigen tests (direct fluorescent antibody [DFA], enzyme immunoassay [EIA], indirect fluorescent antibody [IFA], and the cartridge formats) have excellent sensitivity and specificity compared with routine microscopy. Available antigen detection tests are listed in Table 46.12. The most common immunoassays are designed to confirm infection with E. histolytica, G. duodenalis, and Cryptosporidium spp. 

Blood Parasites Several new blood parasite antigen detection systems are available and have been effectively tested in field trials. Several quick screening tests are available for malaria using an immunochromogenic or antigen-capture EIA system (Table 46.12). Although these simple rapid methods are appropriate for population screening, a blood film from a positive patient must be examined to confirm the diagnosis and properly identify the species. Commercial PCR testing kits are currently available for Plasmodium spp. and Wuchereria bancrofti. Some of the rapid test kits for Plasmodium can detect all of the species that commonly infect humans but may not be able to differentiate the individual species. Other PCR kits are designed to specifically identify P. falciparum. Because PCR techniques

This technique is designed to allow examination and recognition of detailed organism morphology under oil-immersion examination (100× objective, for a total magnification of 1000×), primarily allowing recovery and identification of Plasmodium spp., Babesia spp., Trypanosoma spp., Leishmania donovani, and filarial blood parasites. The thin blood film is routinely used for specific parasite identification, although the number of organisms per field is significantly reduced compared with the thick blood film. The primary purpose is to allow malarial parasites to be visualized in the RBCs and to assess the size of infected RBCs compared with that of uninfected RBCs. RBC morphology is preserved using this method. 

Specimen Finger-stick blood, whole blood, or anticoagulated blood (ethylenediaminetetraacetic acid [EDTA] recommended). Multiple smears should be made within 1 h of collection, but initially only one smear should be stained. Additional smears can be stained if needed by different methods to enhance identification. 

Reagents Giemsa stain (films must be prefixed with absolute methanol before staining), Wright’s stain (the stain contains the fixative), or Wright-Giemsa stains and their associated solutions; mounting fluid (optional). 

Examination Oil-immersion examination (1000×) of at least 300 fields; additional fields may be required if suspect organisms have been seen in the thick blood film. The slide may be screened using the newer 50× or 60× oil-immersion objectives, but the results should not be reported until the examination has been completed using the 100× oil-immersion lens. A blood film must be examined totally at a lower power to rule out the presence of microfilariae, which tend to be found near the edges of the smear. 

Results The thin blood film is routinely used for parasite identification to the species level (Plasmodium spp.). Both the thick and thin films should be examined before the final result is reported. 

Notes and Limitations The thin blood film is prepared exactly as one used for a differential count. A well-prepared film is thick at one end and thin at the other. The use of clean, grease-free slides is mandatory; long streamers of blood indicate that the slide used as a spreader was dirty or chipped. Streaks in the film usually are caused by dirt; holes in the film indicate grease on the slide. Although Giemsa stain is the stain of choice, blood parasites can be seen using other stains; however, the parasite morphology and color may not be consistent with that described for Giemsa-stained organisms. Giemsa stain does not stain the sheath of Wuchereria bancrofti; hematoxylin-based stains (e.g., Delafield’s hematoxylin) are recommended for these organisms. The WBCs on the stained blood film serve as the quality control; if the WBC morphology and color are acceptable, then any parasites present will also appear normal and will be acceptable. Alternately, previously positive patient smears (if available) can be fixed with methanol and air dried, then stored in a sealed container at −70°C. These QC slides should be brought to room temperature before staining.

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• BOX 46.7 Thick Blood Films: Review

Cultivation

Principle

Parasite culture techniques are not routinely used to detect parasite infections in the United States. Any laboratory providing these types of cultures must maintain stock quality control (QC) cultures of specific organisms, often obtained from the American Type Culture Collection (ATCC). The relevant QC organisms are cultured simultaneously with the patient specimen, thus providing some assurance that the culture system functioned properly. The methods for in vitro culture are often complex, and QC is difficult and not feasible in the routine diagnostic laboratory. Some techniques may be available in certain institutions, particularly those in which research and consulting services are available. The cultivation of parasites is an essential technique for laboratories developing diagnostic testing methods, for epidemiologic analysis of outbreaks, for testing for drug resistance, or for the analysis of parasite life cycles. Although T. vaginalis can be easily detected in vaginal or urethral secretions as well as urine sediment, accurate identification may require an experienced laboratory scientist. Several test kits are commercially available that allow quick culture confirmation of Trichomonas infection. OSOM Trichomonas and InPouch TV are two of the methods commonly used in physicians’ offices or clinics. Trichomonas culture is very sensitive and specific, but it may take up to 7 days for growth. Physicians may request cultures when a patient is suspected to have a Trichomonas infection but is negative by microscopic examination. This is an example of an axenic culture method, in which the parasites are grown in pure culture without augmentation with bacteria. Keratitis caused by Acanthamoeba may be confirmed by microscopic observation but also by culture of the parasite. Corneal scrapings or contact lenses are inoculated onto nutrient agar that has an overlay of Escherichia coli or Enterobacter spp., which serves as a food source for the ameba. The plate is observed for up to 7 days to detect the organism’s growth on the agar surface. Identification of the organism is confirmed by microscopic morphology or PCR. This culture technique is called monoxenic because a single known bacterial species is used to help cultivate the organism. This same culture technique is used to recover and confirm granulomatous amebic encephalitis (GAE) caused by Acanthamoeba or PAM caused by another free-living amebic parasite, N. fowleri. In both cases, because of the critical nature of the infection, the organism identification must be confirmed by PCR or other antigen confirmation methods.

This technique is designed to allow examination and recognition of detailed organism morphology under oil-immersion examination (100× objective, for a total magnification of 1000×), primarily allowing recovery and identification of Plasmodium spp., Babesia spp., Trypanosoma spp., Leishmania donovani, and filarial blood parasites. The thick blood film is routinely used for detection of parasites, because the number of organisms per field is much greater than with the thin blood film. The primary purpose is to allow examination of a larger volume of blood than is seen with the thin blood film. RBC morphology is not preserved using this method. 

Specimen Finger-stick blood, whole blood, or anticoagulated blood (ethylenediaminetetraacetic acid [EDTA] recommended). 

Reagents Giemsa stain, Wright’s stain, or Wright-Giemsa stains and their associated solutions; mounting fluid (optional). 

Examination Oil-immersion examination (1000×) of at least 300 fields; additional fields may be required if suspect organisms have been seen in the thin blood film. The slide may be screened using the newer 50× or 60× oil-immersion objectives, but the results should not be reported until the examination has been completed using the 100× oil-immersion lens. A blood film must be examined at a lower power to rule out the presence of microfilariae. 

Results The thick blood film is used to detect the presence of parasites; final identification may require examination of the thin blood film. Both should be examined before the final result is reported. 

Notes and Limitations The thick blood film is prepared by spreading a few drops of blood (using a circular motion) over an area approximately 2 cm in diameter. If whole blood is used, the examiner should continue stirring about 30 s to prevent the formation of fibrin strands. The use of clean, grease-free slides is mandatory. The film is allowed to air-dry at room temperature (heat is never applied to these films). Although Giemsa stain is the stain of choice, blood parasites can be seen using other stains; however, the parasite morphology and color may not be consistent with that described for Giemsa-stained organisms. Giemsa stain does not stain the sheath of Wuchereria bancrofti; hematoxylin-based stains (e.g., Delafield’s hematoxylin) are recommended for these organisms. The WBCs on the stained blood film serve as the quality control; if the WBC morphology and color are acceptable, then any parasites present will also appear normal and will be acceptable. Alternately, previously positive patient smears (if available) can be held fixed with methanol and air-dried, then stored in a sealed container at −70°C. The QC slides should be brought to room temperature before staining.

can amplify the parasite’s DNA, it can detect infection even when the number of parasites is low. It can also be used to confirm species identification when the morphology is unclear. 

Larval-Stage Nematodes The use of certain fecal culture methods (sometimes referred to as coproculture) is especially helpful for detecting light infections of hookworm, S. stercoralis, and Trichostrongylus spp. The rearing of infective-stage nematode larvae improves the diagnosis of hookworm and trichostrongyle infections, because the eggs of these species are identical and differentiation is based on larval morphology. These techniques are also useful for obtaining infective-stage larvae for research

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TABLE a 46.12    Example Antigen Detection Kits for Stool or Vaginal Discharge Specimens

Manufacturer or Distributorb

Test Format/ Methodology

PARA-TECT Cryptosporidium Antigen 96

Medical Chemical Corporation

EIA

ProSpecT Rapid (Cryptosporidium)

Remel

EIA

Xpect Crypto

Remel

RAPID

Cryptosporidium

TechLab

EIA

Crypto CELISA

Cellabs

EIA

Crypto CEL

Cellabs

IFA

ColorPAC Giardia/Cryptosporidium RAPID

Becton Dickinson

RAPID

PARA-TECT Cryptosporidium/Giardia

Medical Chemical

DFA

Merifluor

Meridian Bioscience

DFA

ImmunoCard STAT Cryptosporidium/Giardia

Meridian Bioscience

RAPID

ProSpecT Giardia/Cryptosporidium

Remel

EIA

Xpect Giardia/Cryptosporidium RAPID

Remel

RAPID

Crypto/Giardia CEL

Cellabs

IFA

Organism/Kit Name Cryptosporidium spp.

Cryptosporidium spp. and Giardia sp.

Cryptosporidium spp., Giardia sp., and Entamoeba histolytica Triage (fresh, frozen)

BioSite/Alere

RAPID

Entamoeba histolytica II

TechnLab

EIA

Entamoeba CELISA

Cellabs

EIA

E. histolytica

Wampole

EIA

Remel

EIA

Giardia CELISA

Cellabs

EIA

PARA-TECT Giardia Antigen 96

Medical Chemical Corporation

EIA

ProSpecT Giardia

Remel

EIA or RAPID

Giardia

Wampole

EIA

Giardia II

TechLab

EIA

Giardia EIA

Antibodies, Inc.

EIA

Giardia CEL

Cellabs

IFA

Simple-Read Giardia

Medical Chemical Corporation

Rapid

Affirm VPIII

Becton Dickinson

Probe

T. vaginalis

Chemicon

DFA

OSOM Trichomonas

Sekisui Diagnostics

Rapid

Quik-Trich

PanBio

LA

Entamoeba histolytica

Entamoeba histolytica/E. dispar ProSpecT

Giardia duodenalis

Trichomonas vaginalis

DFA, Direct fluorescent antibody; EIA, enzyme immunoassay; IFA, indirect fluorescent antibody; RAPID, rapid immunochromatographic assay. aThe kits are available commercially in the United States for immunodetection of parasitic organisms or antigens in stool or vaginal discharge. This is a representative list; not every available kit is listed.

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TABLE 46.12    Example Antigen Detection Kits for Stool or Vaginal Discharge—cont’d bAntibodies,

Inc., P O Box 1560, Davis, CA 95617-1560; Becton Dickinson, 1 Becton Dr., Franklin Lakes, NJ 07417; BioSite, 11030 Roselle St., San Diego, CA 92121; Cellabs, P O Box 421, Brookvale, NSW 2100, Australia; Chemicon, 28835 Single Oak Dr., Temecula, CA 92590; PanBio InDx, 1756 Sulfur Spring Rd., Baltimore, MD 21227; Genzyme Virotech, Gmbh, Lowenplatz 5, 66248, Russelheim, Germany; Medical Chemical Corporation, 19430 Van Ness Avenue, Torrance, CA 90501; Meridian Bioscience, Inc., 3471 River Hills Dr., Cincinnati, OH 45244; Novocastra, 30 Ingold Rd., Burlingame, CA 94010; Panbio Inc, 9075 Guilford Rd, Columbia, MD, 21046 Remel, 12076 Santa Fe Drive, Lenexa, KS 66215; Sekisui Diagnostics, LLC, One Wall Street, Burlington, MA 01803; TechLab, VPI Research Park, 1861 Pratt Dr., Blacksburg, VA 24060; Wampole Laboratories, P O Box 1001, Cranbury, NJ 08512; Adapted from the Centers for Disease Control and Prevention. Laboratory Identifications of Parasitic Diseases of public Health Concerns (website). www.cdc. gov/dpdx/diagnosticprocedures/stool/antigendetection.html. https://www.cdc.gov/dpdx/diagnosticprocedures/other/vaginalswabs.html. Accessed June 29, 2019.

purposes. Diagnostic methods available include the HaradaMori filter paper strip culture, the Petri dish/slant filter paper culture, the agar plate method, and the charcoal culture. 

Blood Protozoa Leishmaniasis is often diagnosed by observation of the nonmotile amastigote stage of the parasite on blood smears, particularly as intracellular forms within monocytes and macrophages. The extracellular motile form, or promastigote stage found in tissue specimens, can also be cultured in specialized media (Novy-MacNeal-Nicolle [NNN] medium). This method is not part of a routine analysis in the clinical laboratory. Recent breakthroughs have allowed scientists to cultivate the malarial parasites that infect humans and a number of other animal species. Although not used for diagnosis of malaria in the clinical laboratory, the ability to cultivate Plasmodium species has led to a number of significant developments, including a better understanding of the life cycle, development of analytical testing methods such as PCR, and research leading to the development of protective vaccines. A unique cultivation method involves the use of intermediate hosts to isolate a parasitic organism from a human host. This technique is called xenodiagnosis. It was primarily used for the detection of chronic Chagas disease caused by T. cruzi. The insect vector Triatomid bug was allowed to take a blood meal from the patient and was examined for the presence of the trypanosomes in its gut. In addition to the detection of Chagas disease, other now archaic xenodiagnostic methods have been used to detect leishmaniasis and onchocerciasis. 

Serodiagnosis Serodiagnosis, or testing patient serum for the presence of antibodies, has been available for many years. However, serologic methods for the detection and identification of parasitic infections are not routinely offered by most clinical laboratories because of their high cost, difficulty of interpretation, low test volume, and limited sensitivity and specificity. Direct parasite detection and/or detection of parasite antigens are the methods of choice in all but a few situations. In parasitic infections where the organism reproduces in the host tissues, continuous antigenic stimulation of the host’s immune system occurs as the infection progresses. In these cases, there is a positive correlation between clinical symptoms and serologic test results. Serodiagnosis

is recommended when the collection of a direct specimen may cause significant risk to the patient, as with echinococcosis or cysticercosis. Serodiagnosis is also recommended when the infection may be widespread, making specimen collection difficult, as with infections caused by Toxoplasma or Toxocara spp. Standard serologic techniques used in the laboratory for parasitic diagnostics include latex agglutination, enzyme-linked immunosorbent assay (ELISA)/EIA, IFA, and immunoblot techniques. The Centers for Disease Control and Prevention (CDC) offers a number of serologic procedures for diagnostic purposes, some of which are not available elsewhere. Regulations for submitting specimens to the CDC may vary from state to state. Each laboratory should check with the appropriate county or state department of public health for specific instructions. Additional information on procedures and the interpretation of test results may be obtained directly from the CDC: http://www.cdc.gov/parasites/. 

Prevention The prevention of human parasitic infections is directly linked to understanding the life cycles of the various organisms and their modes of infection (Table 46.4). Preventive measures include avoiding direct exposure, as by improving personal hygiene, ensuring proper sanitation, and eliminating sexual activities that may involve fecal-oral contact. Adequate water treatment (including filtration) may be required, in addition to overall awareness of environmental sources of infection. In some cases, avoiding contaminated environmental water and soil sources may be important; for example, this is mandatory in dealing with systems of contact lens care, methods of sinus irrigation methods, potential infection with free-living amebae from municipal water supplies. Chemoprophylactic agents to prevent clinical symptoms are given to individuals traveling to areas where malaria is endemic. These medications are effective against the erythrocytic forms but do not actually prevent infection with malaria; that is, the drugs do not prevent sporozoites from entering the host, traveling to the liver, and beginning the preerythrocytic developmental cycle. In general, chloroquine is the drug of choice, although different regimens may be used for chloroquine-resistant strains of malaria and for different species of malaria. Vector control and awareness of transmission through blood transfusions, shared drug needles, congenital infections, and organ transplants are also important

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considerations in preventing human parasitic disease. Careful monitoring of the blood supply is required to prevent the transmission of parasites. This is particularly important in areas of the world where blood-borne parasites play a large role in human disease or in the case of blood donors who have recently traveled to endemic areas. Adequate cooking of meat that may be infected is also important; cultural habits may influence the handling and eating of raw or poorly cooked foods. Prevention depends on a thorough understanding of the life cycle and epidemiology of all parasites that cause human disease. This information is critical to the prevention of human disease caused either by parasites limited to the human host or by parasites that can cause disease in humans and other animal hosts. An example of a global initiative to eliminate parasitic infection can be seen in the management of Guinea worm disease, caused by the parasite Dracunculus medinensis. Individuals become infected with the worms by drinking contaminated water. In 1980, the CDC, World Health Organization (WHO), and other partners began a campaign to provide access to clean drinking water as well as identifying sources of potential infection. In 1986, there were 3.5 million cases of Guinea worm disease, primarily in Africa. Thanks to the efforts of the global Guinea Worm Eradication Program, only

25 human cases from Chad, Ethiopia, and South Sudan were reported worldwide in 2016. The ultimate goal is complete elimination of the parasite. (WHO Collaborating Center for Research Training and Eradication of Dracunculiasis. Guinea Worm Wrap Up #245Cdc-pdf External, 2017, Centers for Disease Control and Prevention [CDC]: Atlanta, Georgia.) 

Ectoparasites Ectoparasites, member of the phylum Arthropoda, can affect human health in several ways. Arthropods can act as biologic vectors of disease, as when mosquitoes transmit malaria or when ticks transmit Lyme disease during a blood meal. They can also act as mechanical vectors—for example, when flies or cockroaches help transmit bacteria to food or water, causing enteric diseases. Mites and lice can act as biologic vectors of disease, but they can also cause disease directly by their presence (lice infestation) and the body’s reaction to them (scabies). Although not true parasites, members of the Arthropoda can also affect humans through venomous bites (spiders) or stings (scorpions). Any arthropod that breaks the skin can also cause a secondary bacterial infection at the injury site (Table 46.13).

TABLE 46.13    Arthropods (Ectoparasites)

Arthropods

Organisms Transmitted

Other Impacts

Ticks (Four pairs of legs; two pairs of mouth parts; no antennae; larval, nymph, and adult stages; separate sexes)

Anaplasma Arboviruses Babesia spp. Borrelia burgdorferi (Lyme disease) Borrelia sp. (relapsing fever) Ehrlichia spp. Francisella sp. Rickettsia spp. Flavivirus (tick-borne encephalitis)

Secondary bite wound infections Tick paralysis (toxin)

Mites, infestation

St. Louis encephalitis western equine encephalitis Rickettsia sp.

Scabies (infestation) pruritus

Arachnida

Spiders

Venom reaction to bite (e.g., brown recluse, black widow)

Scorpions

Venom reaction to sting (e.g., bark scorpion)

Insecta Fleas (three pairs of legs; rear legs much larger)

Dipylidium caninum Hymenolepis nana Hymenolepis diminuta Yersinia pestis Rickettsia typhi (murine typhus) Francisella sp.

Pruritus, site of bite

Flies 1–2 pairs of wings; separate head, thorax, and abdomen)

Trypanosoma rhodesiense Trypanosoma gambiense Leishmania spp. Onchocerca volvulus Loa loa Bartonella (Oroya fever)

Mechanical transfer of disease; poor sanitation (dysentery, cholera)

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TABLE 46.13    Arthropods (Ectoparasites)—cont’d

Arthropods

Organisms Transmitted

Other Impacts

Bugs (three pairs of legs; some have wings)

Trypanosoma cruzi (Triatomids including bed bugs)

Bite wound reactions; pruritus Mechanical transfer of disease; poor sanitation (dysentery, cholera, e.g., cockroaches)

Lice (wingless; three segments)

Borrelia (relapsing fever) Bartonella (trench fever) Rickettsia prowazeki (typhus)

Pruritus reaction to infestation; waste products

Mosquitoes (three pairs of legs, three segments; long abdomen; 3000+ species)

Plasmodium spp. Wuchereria bancrofti Brugia malayi Arboviruses Flaviviruses (dengue, yellow fever) Togaviruses (encephalitis)

Bite reaction—pruritus

•

Fig. 46.12 Dorsal view of a male Rocky Mountain wood tick, Dermacentor andersoni. This tick species is a known North American vector of Rickettsia rickettsii, which is the etiologic agent of Rocky Mountain spotted fever (RMSF). (Courtesy the Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

Ectoparasites can be directly observed without a microscope, but it is better to place them in 70% to 95% ethanol to maintain their morphology and color and examine them using a stereomicroscope or low power on a light microscope. Permanent fixation can be obtained using Permount (Fisher Scientific, Pittsburgh, PA). Arthropods have four characteristics: a chitinized exoskeleton, pairs of jointed legs or appendages, bilateral symmetry, and a hemocele. Members of the Arachnida (ticks, mites, spiders, and scorpions) have four pairs of appendages (legs) (Fig. 46.12), whereas members of the Insecta (flies, lice, fleas, bugs, and mosquitoes) have three pairs of appendages (legs and wings) (Figs. 46.13 and 46.14). Identification is based on recognizing the morphologic characteristics of the arthropod and carefully describing the head, thorax, and abdomen as well as noting the number of legs or other appendages and the presence of antennae.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

• Fig. 46.13  Xenopsylla cheopis, the Oriental rat flea, the primary vector of bubonic plague and murine typhus. (400×)

• Fig. 46.14  Pediculus sp., louse that infests humans and transmits a variety of diseases including endemic typhus, trench fever, and relapsing fever.

Chapter Review 1. Which of the following is essential for a complete and accurate fecal examination for O&Ps? a. Antigen detection using specific antibodies b. Direct wet mount for motility of the organisms c. Permanent stained slide d. Concentration wet mount e. Culture for specific organisms 2. Which specimen is most likely to provide recovery of Trichomonas vaginalis? a. Urine b. Urethral discharge c. Vaginal discharge d. Feces e. Biopsy tissue 3. Examination of sputum may be necessary to diagnose infection with a. Paragonimus spp. b. Trichinella spiralis c. Wuchereria bancrofti d. Giardia duodenalis e. Taenia saginata 4. Which of the following is a key characteristic of the thick blood film? a. It allows the parasite to be seen inside the RBCs. b. It allows identification of parasites to the species level. c. Less slide area must be examined compared with a thin blood film. d. It should not be made using anticoagulated or fresh blood. e. It allows counting of the infected RBCs to determine the relative parasite burden.

5. True or False _____ When using fecal fixatives, it is mandatory to thoroughly mix the stool specimen with the fixative. _____ Collection of a single stool is sufficient to confirm a negative result for the O&PS examination. _____ One of the more common intestinal nematodes is Ascaris lumbricoides. _____ The most relevant diagnostic tests for the recovery and identification of Babesia spp. are the thick and thin blood films. _____ Parasitic serology for antibody detection is critical for the diagnosis of intestinal protozoa. 6. Matching: Match each term with the correct description. _____ free-living amebae _____ fecal concentration _____ Trichinella spp. _____ Cryptosporidium spp. _____ trichrome _____ microsporidia _____ leishmaniasis _____ pinworm _____ amebiasis _____ 10% formalin

a. sedimentation method b. permanent stain for fecal specimens c. impression smears d. CSF examination e. sigmoidoscopy examination f. a fecal fixative used in O&PS examination g. modified acid-fast stain h. modified trichrome stain i. cellophane (Scotch) tape preparation j. examination of biopsy specimen

PROCEDURE 46.1

Formalin-Ether (Formalin–Ethyl Acetate) Sedimentation Techniques Principle Formalin fixes eggs, larvae, oocysts, and spores so that they are no longer infectious and also preserves their morphology. Fecal debris is extracted into the ethyl acetate phase of the solution, freeing the sedimented parasitic elements from at least some of the artifact material in the stool. Numerous ether substitutes are available; the term ethyl acetate used throughout this chapter is used in the general sense (ether substitute). 

Method 1. Transfer ¼ to ½ teaspoon of fresh stool into 10 mL of 5% or 10% formalin in a 15-mL shell vial, unwaxed paper cup, or 16- by 125-mm tube (the container used may vary depending on individual preference) and mix thoroughly. Let stand 30 min for adequate fixation. 2. Filter this material (use a funnel or pointed paper cup with the end cut off) through two layers of gauze into a 15-mL centrifuge tube. 3. Add physiologic saline or 5% or 10% formalin to within ½ inch (1.5 cm) of the top and centrifuge for 10 min at 500 g.

4. Decant; 0.5–1 mL of sediment should have been produced. Resuspend the sediment in saline to within ½ inch (1.5 cm) of the top and centrifuge again for 10 min at 500 g. This second wash may be eliminated if the supernatant fluid after the first wash is light tan or clear. 5. Decant and resuspend the sediment in 5% or 10% formalin (with the tube only half full). If the amount of sediment left at the bottom of the tube is very small, do not add ethyl acetate in step 6; merely add the formalin, then spin, decant, and examine the remaining sediment. 6. Add approximately 3 mL of ethyl acetate, stopper the tube, invert, and then shake vigorously for 30 s. Hold the tube so that the stopper is directed away from your face; remove the stopper carefully to prevent spraying of material caused by pressure in the tube (best performed under a biohazard hood). 7. Centrifuge for 10 min at 500 g. Four layers should result: a small amount of sediment in the bottom of the tube, containing the parasites; a layer of formalin; a plug of fecal debris on top of the formalin layer; and a layer of ether substitute at the top (Fig. 46.9).

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8. Free the plug of debris by ringing with an applicator stick and decant all the fluid. Use a cotton-tipped applicator to remove debris from the sides of the test tube. After proper decanting, a drop or two of fluid remaining on the side of the tube will drain down to the sediment. Mix the fluid with the sediment and prepare a wet mount for examination. The formalin–ethyl acetate sedimentation procedure may be used on material preserved with polyvinyl alcohol (PVA), although steps 1 and 2 differ as follows: 1. Fixation time with PVA should be at least 30 min. Mix the contents of the PVA bottle (stool-PVA mixture: one part stool to two or three parts PVA) with applicator sticks. Immediately after mixing, pour approximately 2–5 mL of the stool-PVA mixture (the amount varies, depending on the viscosity and density of the mixture) into a 15-mL shell vial, 16- by 125-mm tube, or similar container and add approximately 10 mL of physiologic saline or 5% or 10% formalin. 2. Filter this material (use a funnel or paper cup with the pointed end cut off) through two layers of gauze into a 15-mL centrifuge tube. Steps 3 through 8 are the same for both fresh and PVApreserved material.

Note: Tap water may be substituted for physiologic saline throughout this procedure; however, saline is recommended. Some workers prefer to use 5% or 10% formalin for all the rinses (steps 3 and 4). When the sediment in the bottom of the tube is being examined, 1. Prepare a saline mount (one drop of sediment and one drop of saline solution mixed together) and scan the whole 22- by 22-mm cover slip under low power for helminth eggs or larvae. 2. Iodine may be added to aid detection of protozoan cysts; the specimen should be examined under high dry power. If iodine is added before low power scanning, make sure that the iodine is not too strong; otherwise, some of the helminth eggs will stain so darkly that they will be mistaken for debris. 3. Occasionally, a precipitate is formed when iodine is added to the sediment obtained from a concentration procedure with PVA-preserved material. The precipitate is formed from the reaction between the iodine and excess mercuric chloride that has not been thoroughly rinsed from the PVA-preserved material. The sediment can be rinsed again to remove any remaining mercuric chloride, or the sediment can be examined as a saline mount without the addition of iodine.   

PROCEDURE 46.2

Wheatley’s Trichrome Stain for Fecal Specimens (Modification of Gomori’s Trichrome Stain for Tissue) Principle The internal elements that distinguish among cysts and trophozoites can best be visualized with a stain that enhances the morphologic features. In addition, such a stained smear provides a permanent record of the results. 

Reagents

Expected Results

A. Formula Chromotrope 2R Light green SF Phosphotungstic acid Acetic acid (glacial) Distilled water

0.6 g 0.3 g 0.7 g 1 mL 100 mL

B. Stain Preparation

8. Place in two changes of 100% ethanol for 2–5 min each.a 9. Place in two changes of xylene or toluene for 2–5 min each.a 10. Mount in Permount or other mounting medium; use a No. 1 thickness coverglass. 

1. Add 1 mL of glacial acetic acid to the dry components. 2. Allow the mixture to stand for 15–30 min to “ripen”; then add 100 mL of distilled water. This preparation gives a uniform and reproducible stain; the stain should be purple. Store in Coplin jars. 

Method 1. Prepare fresh fecal smears or polyvinyl alcohol (PVA) smears as described. 2. Place in 70% ethanol for 5 min. (This step may be eliminated for PVA smears.) 3. Place in 70% ethanol plus D’Antoni’s iodine (dark reddish brown) for 2–5 min. 4. Place in two changes of 70% ethanol: one for 5 mina and one for 2–5 min. 5. Place in trichrome stain solution for 10 min. 6. Place in 90% ethanol, acidified (1% acetic acid), for up to 3 s (do not leave the slides in this solution any longer). 7. Dip once in 100% ethanol.

Background debris will be green, and protozoa will show bluegreen to purple cytoplasm. The nuclei and inclusions will be red or purple-red and sharply delineated from the background. Note: If you are currently using one of the stool fixatives that contains a mercuric chloride substitute (e.g., zinc sulfate), remember that the proficiency testing specimens you receive for permanent staining may have been preserved in PVA using the mercuric chloride fixative base. If you use the trichrome staining method for your mercuric chloride substitute fixatives, you may have eliminated the 70% alcohol/iodine step and the following 70% alcohol rinse steps from your method. However, when you stain the proficiency testing fecal smears, you must incorporate the iodine step and the next 70% alcohol rinse steps back into your staining protocol before placing your slides into the trichrome stain. These two steps are designed to remove the mercury from the smear and then to remove the iodine; when your slide is placed into the trichrome stain, both the mercury and iodine are no longer present in the fecal smear. If you fail to incorporate these two steps into your staining protocol, the quality of your proficiency testing stained smears will be poor. Although you may be using mercuric chloride substitute fixatives, both the iodine/70% alcohol and subsequent 70% alcohol rinse steps before the trichrome stain can be used with no damage to the slides. However, some proficiency testing specimens have been fixed using mercuric chloride and will require the iodine and subsequent alcohol rinses before the trichrome staining step. All steps after the trichrome stain would remain the same for either type of fixative.    aAt

this stage, slides can be held several hours or overnight.

  

PROCEDURE 46.3

Modified Iron-Hematoxylin Stain (With Carbolfuchsin Step) Principle

H. Carbolfuchsin

Reagents

Basic fuchsin (solution A): Dissolve 0.3 g basic fuchsin in 10 mL of 95% ethanol. Phenol (solution B): Dissolve 5 g of phenol crystals in 100 mL distilled water. (Gentle heat may be needed.) 1. Mix solution A with solution B. 2. Store at room temperature. Solution is stable for 1 year. 

A.  Mayer’s Albumin

Method

The internal elements that distinguish among cysts and trophozoites can be visualized with a stain that enhances the morphologic features. In addition, the stained smear provides a permanent record of the results. 

Add glycerin to an equal quantity of fresh egg white. Mix gently and thoroughly. Store at 4°C and indicate an expiration date of 3 months. Mayer’s albumin from commercial suppliers can normally be stored at 25°C for 1 year (e.g., Product #756, E.M. Diagnostic Systems, 480 Democrat Road, Gibbstown, NJ 08027; [800] 443–3637). 

B.  Stock Solution of Hematoxylin Stain Hematoxylin powder Ethanol (95% or 100%)

10 g 1000 mL

1. Mix well until dissolved. 2. Store in a clear glass bottle, in a light area. Allow to ripen for 14 days before use. 3. Store at room temperature with an expiration date of 1 year. 

C. Mordant Ferrous ammonium sulfate [Fe(NH4)2(SO4)2 • 6 H2O] Ferric ammonium sulfate [FeNH4(SO4)2 • 12 H2O] Hydrochloric acid (HCl) (concentrated) Add distilled water to 1000 mL

10 g

10 g

10 mL

 D.  Working Solution of Hematoxylin Stain

1. Mix equal quantities of stock solution of stain and mordant. 2. Allow mixture to cool thoroughly before use (prepare at least 2 h before use). The working solution should be made fresh every week. 

E.  Picric Acid

Mix equal quantities of distilled water and an aqueous saturated solution of picric acid to make a 50% saturated solution. 

F.  Acid-Alcohol Decolorizer Hydrochloric acid (HCl) Alcohol to 1000 mL

 G. 

70% Alcohol and Ammonia

70% alcohol Ammonia (enough to bring the pH to approximately 8.0)

 

30 mL (concentrated)

50 mL 0.5–1 mL

1. Slide preparation: a. Place 1 drop of Mayer’s albumin on a labeled slide. b. Mix the sediment from the sodium acetate–acetic acid–formalin (SAF) concentration well with an applicator stick. c. Add approximately 1 drop of the fecal concentrate to the albumin and spread the mixture over the slide. 2. Allow the slide to air dry at room temperature (the smear appears opaque when dry). 3. Place the slide in 70% alcohol for 5 min. 4. Wash in a container of tap water (not running water) for 2 min. 5. Place the slide in Kinyoun’s stain for 5 min. 6. Wash the slide in running tap water (i.e., a constant stream of water into a container) for 1 min. 7. Place the slide in acid/alcohol decolorizer for 4 min.a 8. Wash the slide in running tap water (a constant stream of water into a container) for 1 min. 9. Place the slide in the iron-hematoxylin working solution for 8 min. 10. Wash the slide in distilled water (in a container) for 1 min. 11. Place the slide in the picric acid solution for 3–5 min. 12. Wash the slide in running tap water (a constant stream of water into a container) for 10 min. 13. Place the slide in 70% alcohol plus ammonia for 3 min. 14. Place the slide in 95% alcohol for 5 min. 15. Place the slide in 100% alcohol for 5 min. 16. Place the slide in two changes of xylene for 5 min. 

Procedure Notes 1. The first 70% alcohol step acts with the Mayer’s albumin to “glue” the specimen to the glass slide. The specimen may wash off if insufficient albumin is used or if the slides are not completely dry before staining. 2. The working hematoxylin stain should be checked each day by adding a drop of stain to alkaline tap water. If a blue color does not develop, prepare a fresh working stain solution. 3. The picric acid differentiates the hematoxylin stain by removing more stain from fecal debris than from the protozoa and removing more stain from the organism’s cytoplasm than from the nucleus. 4. When properly stained, the background should be various shades of gray-blue, and protozoa should visible. The cytoplasm will stain a medium-blue cytoplasm and the nuclei will stain a dark blue–black.   

aThis

step can also be performed as follows: a. Place slide in acid/alcohol decolorizer for 2 min. b. Wash slide in running tap water (constant stream of water into container) for 1 min. c. Place slide in acid/alcohol decolorizer for 2 min. d. Wash slide in running tap water (constant stream of water into container) for 1 min. e. Continue staining sequence with step 9 above (iron hematoxylin working solution).

  

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PROCEDURE 46.4

Modified Acid-Fast Stain for Coccidia Reagents A. Carbolfuchsin Basic fuchsin Phenol Alcohol (95%) Distilled water

4g 8 mL 20 mL 100 mL

Dissolve the basic fuchsin in the alcohol; then add the water slowly while shaking. Melt the phenol in a 56°C water bath and add 8 mL to the stain, using a pipette with a rubber bulb. 

B. Decolorizer Ethanol (95%) Concentrated HCl

97 mL 3 mL

Working under a chemical fume hood, add the hydrochloric acid to the alcohol slowly. 

C. Counterstain Methylene blue Distilled water

0.3 g 100 mL

  “Cold” Modified Acid-Fast Stain Method (Kinyoun) 1. Spin an aliquot of 10% formalinized stool for 10 min at 500 g. 2. Remove the upper layer of sediment with a pipette and place a thin layer on a microscope slide. Note: If the stool specimen contains a large amount of mucus, 10 drops of 10% potassium hydroxide (KOH) can be added to the sediment (step 2), which is vortexed, rinsed with 10% formalin, and respun before smear preparation. Some laboratories use this approach routinely before smear preparation. 3. Heat-fix the smear at 70°C for 10 min. 4. Stain the fixed smear for 3–5 min (no heat necessary). 5. Wash in distilled, filtered water and shake off excess water. 6. Flood with decolorizer for approximately 1 min. Check to see that no more red color runs when the slide is tipped.

Add a bit more decolorizer for very thick slides or those that continue to bleed red dye. 7. Wash thoroughly with filtered water as previously and shake off excess. 8. Flood with counterstain for approximately 1 min. 9. Wash with distilled water and drain by standing slides upright. Do not blot dry. The staining of acid-fast organisms may be accelerated by the addition of a detergent or wetting agent. Tergitol No. 7 (Sigma Chemical Co., St. Louis, MO) may be used. Add 1 drop of Tergitol No. 7 to every 30–40 mL of the Kinyoun’s carbolfuchsin stain. Acid-fast bacteria stain red with carbolfuchsin stains. The background color depends on the counterstain: methylene blue imparts a blue color to non–acid-fast material, brilliant green results in a green background, and picric acid results in yellow. 

“Hot” Modified Acid-Fast Stain Method 1. Spin an aliquot of 10% formalinized stool for 10 min at 500 g. 2. Remove the upper layer of sediment with a pipette and place a thin layer on a microscope slide. Note: If the stool specimen contains a large amount of mucus, 10 drops of 10% potassium hydroxide (KOH) can be added to the sediment (step 2), which is vortexed, rinsed with 10% formalin, and respun before smear preparation. Some laboratories use this approach routinely before smear preparation. 3. Heat fix the smear at 70°C for 10 min. 4. Place the slide on a staining rack and flood with carbolfuchsin. 5. Heat to steaming and stain for 5 min. If the slide begins to dry, add more stain without additional heating. 6. Rinse the smear with tap or distilled water. 7. Decolorize with 5% aqueous sulfuric acid for 30 s (thicker smears may require a longer time). 8. Rinse the smear with tap or distilled water, drain, and flood the smear with methylene blue counterstain for 1 min. 9. Rinse with tap or distilled water, drain, and air dry.   

From Kinyoun JJ. A note on Uhlenhuths method for sputum examination, for tubercle bacilli. Am J Pub Health. 1915;5:867.

  

PROCEDURE 46.5

Modified Trichrome Stain for Microsporidia (Weber-Green) Principle

Reagents

The oval shape, spore wall, and diagonal or horizontal “stripe” that distinguish microsporidia spores can best be visualized with a stain that enhances the morphologic features. In addition, such a stained smear provides a permanent record of the results. 

A.  Modified Trichrome Stain Chromotrope 2R Fast green Phosphotungstic acid Acetic acid (glacial) Distilled water

6 ga 0.15 g 0.7 g 3 mL 100 mL

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

1. Prepare the stain by adding 3 mL of acetic acid to the dry ingredients. Allow the mixture to stand (ripen) for 30 min at room temperature. 2. Add 100 mL of distilled water. A properly prepared stain is dark purple. 3. Store in a glass or plastic bottle at room temperature. The shelf life is at least 24 months. 

B. Acid-Alcohol 90% ethyl alcohol Acetic acid (glacial)

995.5 mL 4.5 mL

Prepare by combining the two solutions. 

Method 1. Using a 10-μL aliquot of unconcentrated, preserved liquid stool (5% or 10% formalin or sodium acetate– acetic acid–formalin [SAF]), prepare the smear by spreading the material over an area 45 × 25 mm. 2. Allow the smear to air-dry. 3. Place the smear in absolute methanol for 5 min. 4. Allow to air-dry.

5. Place in trichrome stain for 90 min. 6. Rinse in acid-alcohol for no longer than 10 s. 7. Dip the slides several times in 95% alcohol. Use this step as a rinse. 8. Place in 95% alcohol for 5 min. 9. Place in 100% alcohol for 10 min. 10. Place in xylene substitute for 10 min. 11. Mount with a cover slip (No. 1 thickness), using mounting medium. Check the specimen for adherence to the slide. 12. Examine the smears under oil immersion (×1000) and read at least 100 fields; the examination time probably will be at least 10 min per slide. 

Expected Results Known parasites should be detected readily. If the smear is thoroughly fixed and the stain has been performed correctly, the spores are ovoid and refractile and the spore wall is bright pinkish red. Occasionally the polar tube can be seen either as a stripe or as a diagonal line across the spore. Most of the bacteria and other debris tend to stain green. However, some bacteria and debris stain red.    a10

times the normal trichrome stain formula.

  

PROCEDURE 46.6

Modified Trichrome Stain for Microsporidia (Ryan-Blue) Principle The oval shape, spore wall, and diagonal or horizontal “stripe” that distinguish microsporidia spores can be visualized with a stain that enhances the morphologic features. In addition, such a stained smear provides a permanent record of the results. Numerous variations to the modified trichrome (WeberGreen) stain were attempted to improve the contrast between the color of the spores and the background staining. Optimal staining was achieved by modifying the composition of the trichrome solution. This stain is also available commercially from several suppliers. The specimen can be fresh stool or stool that has been preserved in 5% or 10% formalin, sodium acetate–acetic acid–formalin (SAF), or some of the newer single vial–system fixatives. Actually, any specimen other than tissue believed to contain microsporidia can be stained by this method. 

Reagents A.  Trichrome Stain (Modified for Microsporidia) (Ryan-Blue) Chromotrope 2R Aniline blue Phosphotungstic acid Acetic acid (glacial) Distilled water

6 ga 0.5 g 0.25 g 3 mL 100 mL

1. Prepare the stain by adding 3 mL of acetic acid to the dry ingredients. Allow the mixture to stand (ripen) for 30 min at room temperature. 2. Add 100 mL of distilled water and adjust the pH to 2.5 with 1 M hydrochloric acid (HCl). A properly prepared stain is dark purple. The staining solution should be protected from light.

3. Store in a glass or plastic bottle at room temperature. The shelf life is at least 24 months.

B. Acid-Alcohol (Procedure 46.5) 

Method 1. Using a 10-μL aliquot of unconcentrated preserved liquid stool (5% or 10% formalin or sodium acetate– acetic acid–formalin [SAF]), prepare the smear by spreading the material over an area 45 by 25 mm. 2. Allow the smear to air-dry. 3. Place the smear in absolute methanol for 5 or 10 min. 4. Allow the smear to air-dry. 5. Place in trichrome stain for 90 min. 6. Rinse in acid-alcohol for no longer than 10 s. 7. Dip slides several times in 95% alcohol. Use this step as a rinse (no longer than 10 s). 8. Place in 95% alcohol for 5 min. 9. Repeat step 8. 10. Place in 100% alcohol for 10 min. 11. Place in xylene substitute for 10 min. 12. Mount with a cover slip (No. 1 thickness), using mounting medium. 13. Examine smears under oil immersion (1000×) and read at least 100 fields; the examination time probably will be at least 10 min per slide. 

Expected Results Known parasites should be detected readily. If the smear is thoroughly fixed and the stain has been performed correctly, the spores are ovoid and refractile and the spore wall is bright pinkish red. Occasionally the polar tube can be seen either as a stripe or as a diagonal line across the spore. Most of the bacteria and other debris tend to stain blue. However, some bacteria and debris stain red. 

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Procedure Notes for Modified Trichrome Staining Methods (Weber or Ryan) 1. Positive control smears must be stained and examined each time patient specimens are stained and examined. 2. Because of the difficulty in achieving stain penetration through the spore wall, prepare thin smears and do not reduce the staining time in trichrome. Also, make sure the slides are not left too long in the decolorizing agent (acidalcohol). If the control organisms are too light, leave them in the trichrome longer and shorten the time to two dips in the acid-alcohol solution. Also, remember that the 95% alcohol rinse after the acid-alcohol should be performed quickly to prevent additional destaining from the acidalcohol reagent. 3. When purchasing the chromotrope 2R, obtain the highest dye content available. Two sources are Harleco (Gibbstown, NJ) and Sigma Chemical Co. (dye content among the highest [85%]). Fast green and aniline blue can be obtained from Allied Chemical and Dye (New York City, NY). 4. In the final stages of dehydration, the 100% ethanol and the xylenes (or xylene substitutes) should be kept as free of water as possible. Coplin jars must have tightfitting caps to prevent both evaporation of reagents and absorption of moisture. If the xylene becomes cloudy after the addition of slides from 100% alcohol, return the slides to 100% alcohol and replace the xylene with fresh stock. 

Procedure Limitations for Modified Trichrome Staining Methods (Weber or Ryan) 1. Although this staining method stains the microsporidia, the range of stain intensity and the small size of the spores cause some difficulty in identifying these organisms. Because this procedure results in the staining of many other organisms or objects in stool specimens,

differentiation of the microsporidia from surrounding material is still difficult. In addition, some slight size variation occurs among the spores. 2. If the patient has severe watery diarrhea, less artifact material is present in the stool to confuse with the microsporidial spores. If the stool is semiformed or formed, the amount of artifact material is much greater, and the spores are much harder to detect and identify. Also, remember that the number of spores varies according to the stool consistency (the more diarrheic the stool, the more spores that are present). 3. Those who developed some of these techniques believe that concentration procedures result in an actual loss of microsporidial spores; therefore, the use of unconcentrated, formalinized stool is strongly recommended. However, no data indicate the centrifugation speeds and other parameters used in the study. 4. In the University of California Los Angeles Clinical Microbiology Laboratory, unpublished data have indicated that centrifugation at 500 g for 10 min increases dramatically the number of microsporidial spores available for staining (from the concentrate sediment). This is the same protocol used for centrifugation of all stool specimens, regardless of the suspected organism. 5. Do not use wet gauze filtration (an old standardized method of filtering stool before centrifugation) with too many layers of gauze that may trap organisms and prevent them from flowing into the fluid to be concentrated. No more than two layers of gauze should be used. Commercially available concentration systems that use metal or plastic screens for filtration also may be used.   

aTen

times the normal trichrome stain formula.

  

PROCEDURE 46.7

Modified Trichrome Stain for Microsporidia (Kokoskin; Hot Method) Principle The oval shape, spore wall, and diagonal or horizontal “stripe” that distinguish microsporidia spores can be visualized with a stain that enhances the morphologic features. Changes in temperature (from room temperature to 50°C) and in the staining time (from 90 to 10 min) have been recommended as improvements for the modified trichrome staining methods. In addition, such a stained smear provides a permanent record of the results. 

Method 1. Using a 10-μL aliquot of unconcentrated, preserved liquid stool (5% or 10% formalin or sodium acetate– acetic acid–formalin [SAF]), prepare the smear by spreading the material over an area 45 by 25 mm.

2. Allow the smear to air-dry. 3. Place the smear in absolute methanol for 5 min. 4. Allow the smear to air-dry. 5. Place in trichrome stain for 10 min at a temperature of 50°C. 6. Rinse in acid-alcohol for no longer than 10 s. 7. Dip the slides several times in 95% alcohol. Use this step as a rinse (no longer than 10 s). 8. Place in 95% alcohol for 5 min. 9. Place in 100% alcohol for 10 min. 10. Place in xylene substitute for 10 min. 11. Mount with a cover slip (No. 1 thickness), using mounting medium. 12. Examine smears under oil immersion (1000×) and read at least 100 fields; the examination time probably will be at least 10 min per slide.   

CHAPTER 46  Overview of the Methods and Strategies in Parasitology

PROCEDURE 46.8

Staining Thin Films: Giemsa Stain Principle Spreading the blood cells in a thin layer allows easier visualization of the size of red cells, inclusions, and extracellular forms. 

Method 1. Fix blood films in absolute methanol (acetone-free) for 30 s. 2. Allow the slides to air-dry. 3. Immerse the slides in a solution of 1 part Giemsa stock (commercial liquid stain or stock prepared from powder) to 10–50 parts of Triton-buffered water (pH 7.0–7.2). Stain 10–60 min (see Procedure note). Fresh working stain should be prepared from stock solution each day. 4. Dip slides briefly in Triton X-100 buffered water. 5. Drain thoroughly in the vertical position and allow to air-dry.

Note: A good general rule for stain dilution and staining time is as follows: if the dilution is 1:20, stain for 20 min; if 1:30, stain for 30 min; and so forth. However, a series of stain dilutions and staining times should be tested to determine the best dilution and time for each batch of stock stain. 

Expected Results Giemsa stain colors the components of blood as follows: erythrocytes, pink-orange to pale gray-blue depending on the pH of the stain; nuclei of white blood cells, purple and pale purple cytoplasm; eosinophilic granules, bright purple-red; neutrophilic granules, deep pink-purple. Parasitic forms are blue to purple, with reddish nuclei. Their characteristic morphologies are used for differentiation. Inexperienced workers may confuse platelets with parasites.   

PROCEDURE 46.9

Staining Thick Films: Giemsa Stain Principle A large amount of blood can be examined for parasitic forms by lysing the red blood cells and staining for parasites. The lack of methanol fixation allows lysis of red blood cells by the aqueous stain solution. Although parasites can be found in the larger volume of blood, definitive morphologic criteria necessary for specific organism identification may be more difficult to see. 

Method The procedure followed for thick films is the same as for thin films, except that the first two steps are omitted. If the slide has a thick film at one end and a thin film at the other, fix only the thin portion and then stain both parts of the film simultaneously.

Note: Although Giemsa stain has been used for many years, a number of bloodstains can be used for blood parasites (e.g., Giemsa, Wright’s, and Wright-Giemsa combination, rapid bloodstains). Different stains produce somewhat different colors for the cell nuclei and cytoplasm; however, color variability is also seen with Giemsa stain depending on pH. Regardless of the stain used, the quality control is built into the slide; the colors seen in any parasites present mimic those seen in the white blood cells (WBCs). Therefore, the WBCs serve as the quality control organisms. It is not mandatory to use actual blood parasites as the quality control organisms. Alternately, previously positive patient smears (if available) can be held fixed with methanol and air-dried, then stored in a sealed container at −70°C. The QC slides should be brought to room temperature before staining.   

PROCEDURE 46.10

Calibration of the Ocular Micrometer Principle Exact determination of size is a critical standard in the correct identification of parasite elements, especially protozoa found in fecal specimens. For that reason, each microscope used for parasite identification should have a calibrated ocular micrometer. 

Materials 1. Ocular micrometer disk installed in one of the eyepieces of the microscope (ocular). The ocular micrometer is used to measure items seen through the microscope. 2. Stage micrometer used to calibrate the ocular micrometer. For highest accuracy, the micrometer should

be certified by an agency such as National Institute of Standards and Technology (NIST). 

Method 1. Insert the ocular micrometer disk into one of the oculars on the microscope. 2. Place the stage micrometer slide on the microscope stage. 3. Align the ocular scale to the stage micrometer and focus so that you are able to clearly view both the large (0.1 mm) and the small (0.01 mm) increments 4. Move the stage micrometer so that the “0” line on the ocular micrometer aligns with the “0” line on the stage micrometer.

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5. Find a point as far down the scale as possible where the ocular and stage micrometer scales match. 6. Count the number of ocular micrometer spaces that align with the number of millimeters on the stage micrometer. • For example: If 0.5 mm on the stage micrometer aligns with 42 ocular micrometer spaces, then 0.5 mm/42 ＝ 0.0119 mm/ocular micrometer space. • Because parasites are typically reported in micrometers, this should be converted to micrometers by multiplying by 1000.

• Therefore 0.0119 mm/ocular space × 1000 μm/mm ＝ 11.9 μm/ocular micrometer space. • This procedure should be repeated for each objective on the microscope. 7. The calibration is microscope-specific, so if more than one microscope is used, each microscope must be calibrated. 8. Calibrations should be posted by the microscope for each magnification. 9. Calibration should be repeated annually, after each cleaning, or when an objective or ocular lens has been changed.   

47

Intestinal Protozoa OBJECTIVES 1. Describe the basic life cycle, distinguishing morphologic characteristics, clinical disease (if pathogenic), laboratory diagnosis, and prevention for the organisms discussed in this chapter. 2. Define and identify the following parasitic structures: trophozoite, cyst, oocyst, spore, pseudopodia, flagella, cilia, chromatoidal bars, karyosome, central vacuole, cyst form, axoneme, cytostome, spiral groove, undulating membrane, ventral disc, shepherd’s crook, axostyle, macronucleus, micronucleus, apical complex, sporocyst, sporozoite, spore, sarcocyst, and polar tubule. 3. Define life cycle processes, including merogony, gametogony, sporogony, schizogony, and the associated organism(s) and stages. 4. Correlate the parasitic life cycles with the specific diagnostic stages for the organisms listed. 5. Distinguish pathogenic from nonpathogenic protozoa.

PARASITES TO BE CONSIDERED Protozoa Amoebae (intestinal) Entamoeba bangladeshi Entamoeba histolytica Entamoeba dispar* Entamoeba coli Entamoeba gingivalis Entamoeba hartmanni Entamoeba moshkovskii Entamoeba polecki Endolimax nana Iodamoeba bütschlii (buetschlii) Blastocystis spp. Flagellates (intestinal) Giardia duodenalis Chilomastix mesnili Pentatrichomonas hominis Retortamonas hominis Amoeba flagellate Dientamoeba fragilis Ciliates (intestinal) Neoblantidium coli

E

ntamoeba histolytica is considered a true pathogen, and Entamoeba dispar is used to designate nonpathogens. Additional morphologically identical species, Entamoeba moshkovskii and Entamoeba bangladeshi, have 642

been identified which are associated with noninvasive diarrhea. Although it was previously reported that 16% of identified E. dispar trophozoites contain ingested red blood cells, nucleic acid–based methods have been used to resolve this morphological discrepancy. All trophozoites containing red blood cells can be distinctly identified as E. histolytica using nucleic acid–based methods. Immunoassay kits and molecular assays are available for identifying the E. histolytica/E. dispar group and for differentiating E. histolytica, E. dispar, and E. moshkovskii. E. bangladeshi is a species that is indistinguishable from E. histolytica both physically and pathologically; little more is known regarding this species. The protozoa are unicellular eukaryotic organisms, most of which are microscopic. They have a number of specialized organelles that are responsible for life functions and that allow further division of the group into classes. Most protozoa multiply by binary fission and are ubiquitous worldwide. The important characteristics of the intestinal protozoa are presented in Tables 47.1–47.7. The clinically relevant intestinal protozoa are generally considered to be Entamoeba histolytica, Entamoeba moshkovskii, Blastocystis hominis, Giardia duodenalis, Dientamoeba fragilis, Neobalantidium coli, Cystoisospora belli, Cryptosporidium spp., Cyclospora cayetanensis, and the microsporidia. Nonpathogenic intestinal protozoa are listed in various figures and tables but are not discussed in detail.

Amoebae The class Sarcodina, or Amoebae, includes the organisms capable of movement by means of cytoplasmic protrusions called pseudopodia. This group includes free-living organisms, in addition to nonpathogenic and pathogenic organisms found in the intestinal tract and other areas of the body (Tables 47.1 and 47.2). Occasionally, when fresh stool material is examined as a direct wet mount, motile trophozoites may be seen, as well as other, nonparasitic structures (Fig. 47.1).

Entamoeba histolytica General Characteristics Living trophozoites (motile feeding stage) of E. histolytica vary in size from about 12 to 60 μm in diameter. Organisms recovered from diarrheic or dysenteric stools generally are larger than those in formed stool from an asymptomatic individual. The motility has been described as rapid

TABLE 47.1    Intestinal Protozoa: Trophozoites of Common Amoebae

Characteristic

Entamoeba histolytica

Entamoeba dispar/ moshkovskii/ bangladeshi

Entamoeba hartmanni

Entamoeba coli

Endolimax nana

Iodamoeba bütschlii

12–60 μm (usual range, 15–20 μm); invasive forms may be >20 μm

Same size range as E. histolytica

5–12 μm (usual range, 8–10 μm)

15–50 μm (usual range, 20–25 μm)

6–12 μm (usual range, 8–10 μm)

8–20 μm (usual range, 12–15 μm)

Motility

Progressive, with hyaline, fingerlike pseudopodia; motility may be rapid

Same motility as E. histolytica

Usually nonprogressive

Sluggish, nondirectional; blunt, granular pseudopodia

Sluggish, usually nonprogressive

Sluggish, usually nonprogressive

Nucleus (single) and visibility

Difficult to see in unstained preparations

Difficult to see in unstained preparations

Usually not seen in unstained preparations

Often visible in unstained preparation

Occasionally visible in unstained preparations

Usually not visible in unstained preparations

Peripheral chromatin (stained)

Fine granules, uniform in size and usually evenly distributed; may have beaded appearance.

Fine granules, uniform in size and usually evenly distributed; may have beaded appearance.

Nucleus may stain more darkly than in E. histolytica, although morphology is similar; chromatin may appear as solid ring rather than beaded (trichrome).

May be clumped and unevenly arranged on the membrane; may also appear as solid, dark ring with no beads or clumps.

Usually no peripheral chromatin; nuclear chromatin may be quite variable.

Usually no peripheral chromatin.

Karyosome (stained)

Small, usually compact; centrally located but may also be eccentric

Small, usually compact; centrally located but may also be eccentric

Usually small and compact; may be centrally located or eccentric

Large, not compact; may or may not be eccentric; may be diffuse and darkly stained

Large, irregularly shaped; may appear blotlike; many nuclear variations are common; may mimic E. hartmanni or Dientamoeba fragilis

Large, may be surrounded by refractile granules that are difficult to see (“basket nucleus”)

Cytoplasm appearance (stained)

Finely granular, “ground glass” appearance; clear differentiation of ectoplasm and endoplasm; if present, vacuoles are usually small.

Finely granular, “ground glass” appearance; clear differentiation of ectoplasm and endoplasm; if present, vacuoles are usually small.

Finely granular.

Granular, with little differentiation into ectoplasm and endoplasm; usually vacuolated.

Granular, vacuolated.

Granular, may be heavily vacuolated.

Inclusions (stained)

Noninvasive organism may contain bacteria or red blood cells.

Organisms usually contain bacteria; in cytoplasm.

May contain bacteria; no RBCs.

Bacteria, yeast, other debris.

Bacteria.

Bacteria.

sizes refer to wet preparation measurements. Organisms on a permanent stained smear may be 1 to 1.5 μm smaller as a result of artificial shrinkage. RBC, Red blood cell. aThese

CHAPTER 47  Intestinal Protozoa

Sizea (diameter or length)

643

Entamoeba histolytica/ dispar/moshkovskii/ bangladeshi

Entamoeba hartmanni

Entamoeba coli

Endolimax nana

Iodamoeba bütschlii

Sizea

10–20 μm (usual range, 12–15 μm)

5–10 μm (usual range, 6–8 μm)

10–35 μm (usual range, 15–25 μm)

5–10 μm (usual range, 6–8 μm)

5–20 μm (usual range, 10–12 μm)

Shape

Usually spherical

Usually spherical

Usually spherical; may be oval, triangular, or other shapes; may be distorted on permanent stained slide because of inadequate fixative penetration

Usually oval, may be round

May vary from oval to round; cyst may collapse because of large glycogen vacuole space

Nucleus (number and visibility)

Mature cyst: 4 nuclei Immature cyst: 1–2 nuclei; nuclear characteristics difficult to see on wet preparation

Mature cyst: 4 nuclei; Immature cyst: 1–2 nuclei (2-nucleated cysts very common)

Mature cyst: 8 (occasionally 16 or more nuclei may be seen). Immature cysts with 2 or more nuclei are occasionally seen

Mature cyst: 4 Immature cysts: 2 very rarely seen and may resemble cysts of Enteromonas hominis

Mature cyst: 1

Peripheral chromatin (stained)

Peripheral chromatin present; fine, uniform granules, evenly distributed; nuclear characteristics may not be as clearly visible as in trophozoite.

Fine granules evenly distributed on the membrane; nuclear characteristics may be difficult to see.

Coarsely granular; may be clumped and unevenly arranged on membrane; nuclear characteristics not as clearly defined as in trophozoite; may resemble E. histolytica.

No peripheral chromatin.

No peripheral chromatin.

Karyosome (stained)

Small, compact, usually centrally located but occasionally may be eccentric

Small, compact, usually centrally located

Large, may or may not be compact and/or eccentric; occasionally may be centrally located

Smaller than karyosome seen in trophozoites but generally larger than those of genus Entamoeba

Larger, usually eccentric refractile granules may be on one side of karyosome (“basket nucleus”)

Cytoplasm, chromatoidal bodies (stained)

May be present; bodies usually elongate, with blunt, rounded, smooth edges; may be round or oval

Usually present; bodies usually elongate with blunt, rounded, smooth edges; may be round or oval

May be present (less frequently than in E. histolytica); splinter shaped with rough, pointed ends

Rare chromatoidal bodies present; occasionally small granules or inclusions seen; fine linear chromatoidals may be faintly visible on wellstained smears

No chromatoidal bodies present; occasionally small granules may be present

Glycogen (stained with iodine)

May be diffuse or absent in mature cyst; clumped chromatin mass may be present in early cysts (stains reddish brown with iodine)

May or may not be present, as in E. histolytica

May be diffuse or absent in mature cyst; clumped mass occasionally seen in mature cysts (stains reddish brown with iodine)

Usually diffuse if present (stains reddish brown with iodine)

Large, compact, welldefined mass (stains reddish brown with iodine)

Characteristic (diameter or length)

aWet

preparation measurements; in permanent stains, organisms usually are 1–2 μm smaller.

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TABLE 47.2    Intestinal Protozoa—Cysts of Common Amoebae

TABLE 47.3    Intestinal Protozoa—Trophozoites of Flagellates

Number of Nuclei and Visibility

Number of Flagella (Usually Difficult to See)

Shape and Size

Motility

Other Features

Dientamoeba fragilis

Shaped like amebae; 5–15 μm (usual range, 9–12 μm)

Usually nonprogressive; pseudopodia are angular, serrated, or broad lobed and almost transparent.

Percentage may vary, but 40% of organisms have 1 nucleus and 60% have 2 nuclei; not visible in unstained preparations; no peripheral chromatin; karyosome is composed of a cluster of 4–8 granules.

Internal flagella; not visible

Cytoplasm is finely granular and may be vacuolated with ingested bacteria, yeasts, and other debris; may be great variation in size and shape on a single smear.

Giardia duodenalis

Pear-shaped; length 10–20 μm; width, 5–15 μm

“Falling leaf” motility may be difficult to see if organism is in mucus; slight flutter of flagella may be visible using low light (duodenal aspirate or mucus from Entero-Test capsule).

2; not visible in unstained mounts.

4 lateral; 2 ventral, 2 caudal

Sucking disk occupies one half to three fourths of ventral surface; pear-shaped front view, spoon-shaped side view.

Chilomastix mesnili

Pear-shaped; length 6–24 μm (usual range, 10–15 μm); width, 4–8 μm

Stiff, rotary.

1; not visible in unstained mounts.

3 anterior, 1 in cytostome

Prominent cytostome extending one-third to one-half the length of the body; spiral groove across ventral surface.

Pentatrichomonas hominis

Pear-shaped; length 5–15 μm (usual range, 7–9 μm); width 7–10 μm

Jerky, rapid.

1; not visible in unstained mounts.

3–5 anterior, 1 posterior

Undulating membrane extends the length of the body; posterior flagellum extends free beyond end of body.

Trichomonas tenax

Pear-shaped; length 5–12 μm; average of 6.5–7.5 μm; width, 7–9 μm

Jerky, rapid.

1; not visible in unstained mounts.

4 anterior, 1 posterior

Seen only in preparations from mouth; axostyle (slender rod) protrudes beyond the posterior end and may be visible; posterior flagellum extends only halfway down the body; no free end.

Enteromonas hominis

Oval; 4–10 μm (usual range, 8–9 μm); width, 5–6 μm

Jerky.

1; not visible in unstained mounts.

3 anterior, 1 posterior

One side of the body is flattened; posterior flagellum extends free posteriorly or laterally.

Retortamonas intestinalis

Pear-shaped or oval; 4–9 μm (usual range, 6–7 μm); width, 3–4 μm

Jerky.

1; not visible in unstained mount.

1 anterior, 1 posterior

Prominent cytostome extends approximately one half the length of the body.

CHAPTER 47  Intestinal Protozoa

Protozoa

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TABLE 47.4    Intestinal Protozoa—Cysts of Flagellates

Protozoa

Size

Shape

Number of Nuclei

Other Features

Dientamoeba fragilis

5–8 um

Oval to round

1–2

Distinct cyst wall with inner irregular cyst wall. Clear space apparent between outer and inner cyst wall.

Pentatrichomonas hominis, Trichomonas tenax

No cyst stage

Giardia duodenalis

8–19 μm (usual range, 11–14 μm); width, 7–10 μm

Oval, ellipsoidal, or may appear round

4; not distinct in unstained preparations; usually located at one end

Longitudinal fibers in cysts may be visible in unstained preparations; deep-staining median bodies usually lie across the longitudinal fibers. Shrinkage is common, with the cytoplasm pulling away from the cyst wall; “halo” effect may be seen around the outside of the cyst wall because of shrinkage caused by dehydrating reagents.

Chilomastix mesnili

6–10 μm (usual range, 7–9 μm); width, 4–6 μm

Lemon or pearshaped with anterior hyaline knob

1; not distinct in unstained preparations

Cytostome with supporting fibrils, usually visible in stained preparation; curved fibril alongside of cytostome, usually referred to as a “shepherd’s crook.”

Enteromonas hominis

4–10 μm (usual range, 6–8 μm); width, 4–6 μm

Elongate or oval

1–4; usually 2 lying at opposite ends of cyst; not visible in unstained mounts

Resembles Endolimax nana cyst; fibrils or flagella usually not seen.

Retortamonas intestinalis

4–9 μm (usual range, 4–7 μm); width, 5 μm

Pear-shaped or slightly lemonshaped

1; not visible in unstained mounts

Resembles Chilomastix cyst; shadow outline of cytostome with supporting fibrils extends above nucleus; “bird beak” fibril arrangement.

TABLE 47.5    Intestinal Protozoa—Ciliates

Protozoa

Shape and Size

Motility

Number of Nuclei

Other Features

Neobalantidium coli trophozoite

Ovoid with tapering anterior end; 50–100 μm long, 40–70 μm wide (usual range, 40–50 μm)

Ciliates: rotary, boring; may be rapid

1 large kidney-shaped macronucleus; 1 small round micronucleus, which is difficult to see even in stained smear; macronucleus may be visible in unstained preparation.

Body covered with cilia, which tend to be longer near cytostome; cytoplasm may be vacuolated.

Cyst

Spherical or oval; 50–70 μm (usual range, 50–55 μm)

1 large macronucleus visible in unstained preparation; micronucleus difficult to see.

Macronucleus and contractile vacuole are visible in young cysts; in older cysts, internal structure appears granular; cilia difficult to see in cyst wall.

CHAPTER 47  Intestinal Protozoa

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TABLE 47.6    Morphologic Criteria Used to Identify Intestinal Protozoa (Coccidia, Blastocystis spp.)

Protozoa

Shape and Size

Other Features

Cryptosporidium spp. Cryptosporidium parvum (humans and animals) Cryptosporidium hominis (humans)

Oocyst generally round, 4–6 μm; each mature oocyst contains four sporozoites.

Oocyst, diagnostic stage in stool, sporozoites occasionally visible within oocyst wall; acid-fast positive using modified acid-fast stains; various other stages in life cycle can be seen in biopsy specimens taken from gastrointestinal tract (brush border of epithelial cells) and other tissues; disseminated infection well documented in compromised host; oocysts immediately infective (in both formed and/ or watery specimens); nosocomial infections documented; use enteric precautions for inpatients.

Cyclospora cayetanensis

Oocyst generally round, 8–10 μm; oocysts are not mature, no visible internal structure; oocysts may appear wrinkled.

Oocyst, diagnostic stage in stool; acid-fast variable using modified acid-fast stains; color range from clear to deep purple (tremendous variation); best results obtained with decolorizing solution consisting of 1% acid, 3% maximum; oocysts may appear wrinkled (like crumpled cellophane); mimic Cryptosporidium oocysts but are twice as large.

Cystoisospora belli

Ellipsoidal oocyst; range 20–30 μm long, 10–19 μm wide; sporocysts rarely seen broken out of oocysts but measure 9–11μm.

Mature oocyst contains two sporocysts with four sporozoites each; usual diagnostic stage in feces is immature oocyst containing spherical mass of protoplasm (intestinal tract). Oocysts are modified acid-fast positive. Whole oocyst may stain pink, but just the internal sporocysts stain if the oocyst is mature.

Sarcocystis hominis Sarcocystis suihominis

Oocyst thin-walled and contains two mature sporocysts, each containing four sporozoites; commonly thin oocyst wall ruptures; ovoid sporocysts each measure 10–16 μm long and 7.5–12 μm wide.

Thin-walled oocyst or ovoid sporocysts occur in stool (intestinal tract).

Blastocystis spp.

Organisms are generally round, measure approximately 6–40 μm, and are usually characterized by a large, central body (looks like a large vacuole); this stage has been called the central body form.

The more amebic form can be seen in diarrheal fluid but is difficult to identify. The central body forms vary tremendously in size, even on a single fecal smear; this is the most common form seen. Routine fecal examinations may indicate a positive rate much higher than other protozoa; some laboratories report figures of 20% and higher.

and unidirectional. Although this characteristic motility is described, amebiasis rarely is diagnosed based on motility seen in a direct wet mount. The cytoplasm is differentiated into a clear outer ectoplasm and a more granular inner endoplasm. E. histolytica has directional and progressive motility, whereas the other amebae tend to move more slowly and at random. Motility is rarely seen even in a fresh direct wet mount from a patient with diarrhea or dysentery. The cytoplasm is generally more finely granular, and the presence of red blood cells (RBCs) in the cytoplasm is considered diagnostic for E. histolytica (Fig. 47.2); however, most patient samples do not demonstrate trophozoites with ingested red blood cells. Permanent stained smears demonstrate accurate morphology compared with other techniques. When the organism is examined on a permanent stained smear (trichrome or iron-hematoxylin stain), the morphologic characteristics of E. histolytica, E. dispar,

E. moshkovskii, and E. bangladeshi are readily seen. The nucleus is characterized by evenly arranged chromatin on the nuclear membrane and a small, compact, centrally located karyosome (condensed chromatin). As mentioned, the cytoplasm usually is described as finely granular, with few ingested bacteria and scant debris in vacuoles. In organisms isolated from a patient with dysentery, RBCs may be visible in the cytoplasm (Fig. 47.3), and such organisms should be identified as E. histolytica. Most often, infection with E. histolytica/dispar group is diagnosed on the basis of the organism’s morphology, without the presence of RBCs. Newer techniques, including nucleic acid–based testing, are now available that provide a more specific identification and differentiation of Entamoeba spp. and are discussed later in this chapter. As part of the life cycle, the trophozoites may condense into a round mass (precyst), and a thin wall is secreted around the immature cyst (Fig. 47.4). Two types of inclusions may be found in this immature cyst: a glycogen mass

Microsporidia

Immunocompromised Patient

Immunocompetent Patient

Comments

Enterocytozoon bieneusi

Chronic diarrhea; wasting syndrome, cholangitis, acalculous cholecystitis, chronic sinusitis, chronic cough, pneumonitis; cause of diarrhea in organ transplant recipients

Self-limiting diarrhea in adults and children; traveler’s diarrhea; asymptomatic carriers.

Short-term culture only; three strains identified but not named; AIDS patients with chronic diarrhea (present in 5%–30% of patients when CD4 lymphocyte counts are very low); pigs, nonhuman primates

Encephalitozoon hellem

Disseminated infection; keratoconjunctivitis; sinusitis, bronchitis, pneumonia, nephritis, ureteritis, cystitis, prostatitis, urethritis

Possibly diarrhea.

Cultured in vitro; detected in people with traveler’s diarrhea and coinfection with E. bieneusi; pathogenicity unclear; spores not reported yet from stool; psittacine birds

Encephalitozoon intestinalis

Chronic diarrhea, cholangiopathy; sinusitis, bronchitis, pneumonitis; nephritis, bone infection, nodular cutaneous lesions

Self-limiting diarrhea; asymptomatic carriers.

Cultured in vitro; formerly Septata intestinalis; AIDS patients with chronic diarrhea; dogs, donkeys, pigs, cows, goats

Encephalitozoon cuniculi

Disseminated infection; keratoconjunctivitis, sinusitis, bronchitis, pneumonia; nephritis; hepatitis, peritonitis, symptomatic and asymptomatic intestinal infection; encephalitis

Not described; two HIV–serologically negative children with seizure disorder (suspect E. cuniculi infection) presumably were immunocompromised.

Cultured in vitro; wide mammalian host range

Pleistophora sp.

Myositis (skeletal muscle)

Not described.

Tend to infect fish

Pleistophora ronneafiei

Myositis

Not described.

Trachipleistophora hominis

Myositis; myocarditis keratoconjunctivitis; sinusitis

Keratitis.

Cultured in vitro; AIDS patients

Trachipleistophora anthropophthera

Disseminated infection; keratitis

Not described.

AIDS patients

Anncaliia connori

Disseminated infection

Not described.

Formerly Nosema connori; often infects insects; disseminated in infant with SCID

Anncaliia vesicularum

Myositis

Not described.

Formerly Brachiola vesicularum

Anncaliia algerae

Myositis; nodular cutaneous lesions

Keratitis.

Formerly Nosema algerae or Brachiola algerae; cultured in vitro; skin nodules in boy with acute lymphocytic leukemia; found in arthropods

Nosema ocularum

Not described

Keratitis.

HIV–serologically negative individual

Vittaforma corneae

Disseminated infection; urinary tract infection

Keratitis.

Formerly Nosema corneum; cultured in vitro; non-HIV patient

Microsporidium ceylonensisa

Not described

Corneal ulcer, keratitis.

HIV–serologically negative individual, autopsy

Not described

Corneal ulcer, keratitis.

HIV–serologically negative individual, autopsy

Common

Uncommon

Microsporidium

africanuma

Microsporidia (not classified) aMicrosporidium

Keratoconjunctivitis in a contact lens wearer.

is a collective generic name for microsporidia that cannot be classified. AIDS, Acquired immunodeficiency syndrome; HIV, human immunodeficiency virus; SCID, severe combined immunodeficiency.

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TABLE 47.7    Microsporidia That Cause Human Infection

CHAPTER 47  Intestinal Protozoa

1

2

5

6

7

9

4

8

10

12

14

3

649

11

13

15

16

• Fig. 47.1  Various structures that may be seen in stool preparations. (1, 2, and 4) Blastocystis spp. (3 and 5–8) Various yeast cells. (9) Macrophage with nucleus. (10 and 11) Deteriorated macrophage without nucleus. (12 and 13) Polymorphonuclear leukocytes. (14 and 15) Pollen grains. (16) Charcot-Leyden crystals. (Modified from Markell EK, Voge M. Medical Parasitology. 5th ed. Philadelphia: WB Saunders; 1981.)

and highly refractile chromatoidal bars (refractile chromatin structure) with smooth, rounded edges. As the cyst matures (metacyst) (Fig. 47.5 and Fig. 47.3), nuclear division occurs, with the production of four nuclei. Often chromatoidals may be absent in the mature cyst. Cyst morphology does not differentiate E. histolytica, E. dispar, E. moshkovskii, and E. bangladeshi; it may also resemble Entamoeba coli. Cyst formation occurs only in the intestinal tract; once the stool has left the body, cyst formation does not occur. The one-, two-, and four-nucleated cysts are infective and represent the mode of transmission from one host to another. 

Epidemiology • Fig. 47.2  Entamoeba histolytica trophozoite containing ingested red blood cells.

Amebiasis is caused by infection with the true pathogen, E. histolytica. Evidence from molecular studies confirms

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Pathogenesis and Spectrum of Disease

the differentiation of pathogenic E. histolytica and nonpathogenic E. dispar (Fig. 47.6) as two distinct species. E. histolytica is considered the etiologic agent of amebic colitis (Fig. 47.7A) and extraintestinal abscesses (amebic liver abscess; Fig. 47.7B), whereas nonpathogenic E. dispar produces no intestinal symptoms and is not invasive in humans. Trophozoites found in tissue are diagnostic of extraintestinal E. histolytica infection. E. moshkovskii has been shown to cause diarrhea in school-age children and immunocompromised individuals. E. bangladeshi has been recovered from the feces of symptomatic and asymptomatic children. Infection is acquired through the fecal-oral route from infective cysts contained in the feces. The cysts can be ingested in contaminated food or drink or contracted from fomites or various sexual practices that include accidental ingestion of fecal organisms. Flies and cockroaches have been implicated as mechanical vectors of contaminated fecal material. The infection with all Entamoeba spp. occurs worldwide, particularly in areas with poor sanitation. It is estimated that E. histolytica infection kills more than 100,000 people each year. 

The pathogenesis of E. histolytica is related to the organism’s ability to directly lyse host cells and cause tissue destruction. Amebic lesions show evidence of cell lysis, tissue necrosis, and damage to the extracellular matrix. Evidence indicates that E. histolytica trophozoites interact with the host through a series of steps: adhesion to the target cell, phagocytosis, and cytopathic effect. Numerous other parasite factors also play a role. From the perspective of the host, E. histolytica induces both humoral and cellular immune responses; cell-mediated immunity is the major human host defense against this complement-resistant cytolytic protozoan. The presentations of disease are seen with invasion of the intestinal mucosa or dissemination to other organs (most often the liver) or both. However, it is estimated that a small proportion (2% to 8%) of infected individuals have invasive disease beyond the lumen of the bowel. In addition, organisms may be spontaneously eliminated with no disease symptoms. Asymptomatic Infection

Individuals harboring E. histolytica may have either a negative or a weak antibody titer and negative stools for occult

Nucleus

Ingested RBCs

Nucleus

1

2

Chromatoidal bars

Nucleus Nucleus Central karyosome

3

5

4

6

7

• Fig. 47.3  (1) Trophozoite of Entamoeba histolytica (note ingested red blood cells [RBCs]). (2) Trophozoite

of Entamoeba histolytica/Entamoeba dispar group (morphology does not allow differentiation between the Entamoeba species). (3 and 4) Early cysts of E. histolytica/E. dispar group. (5–7) Cysts of E. histolytica/E. dispar group. (8 and 9) Trophozoites of Entamoeba coli. (10 and 11) Early cysts of E. coli. (12–14) Cysts of E. coli. (15 and 16) Trophozoites of Entamoeba hartmanni. (17 and 18) Cysts of E. hartmanni. (From Garcia LS. Diagnostic Medical Parasitology. 4th ed. Washington, DC: ASM Press; 2001. Illustrations 4 and 11 by Nobuko Kitamura.)

CHAPTER 47  Intestinal Protozoa

651

Ingested debris or bacteria

Clumped peripheral chromatin

8

9

10

11 Chromatoidal bars

Nuclei with eccentric karyosome

12

13

15

14

16

17

18

• Fig. 47.3, cont’d 

blood. They also may be passing cysts detectable by a routine ova and parasite (O&P) examination. However, these cysts cannot be morphologically differentiated from those of the nonpathogenic species, most of the asymptomatic infections are actually due to E. dispar. Although trophozoites may be identified, they will not contain any phagocytized RBCs and cannot be differentiated from the nonpathogenic species. Molecular analyses of organisms isolated from asymptomatic individuals have indicated that the isolates typically identified are E. dispar or E. moshkovskii, and up to 80% belong to E. histolytica. Generally, asymptomatic patients never become symptomatic and may excrete cysts for a short period. Asymptomatic patients identified with E. histolytica are at risk for the development of invasive amebiasis.  Intestinal Disease

The incubation period varies from a few days to a much longer time; in an area where E. histolytica is endemic, it is impossible

to determine exactly when exposure to the organism occurred. Normally, the incubation time ranges from 1 to 4 weeks. Tissue invasion by E. histolytica requires a contact-dependent process that involves colonic mucins and amebic lectins in the plasma membrane that mediate adherence to the host mucosa. After adherence, destruction of the host cells involves a process referred to as trogocytosis-like, in which the amebae essentially take bites out of the host cellular membrane. This process is enhanced by E. histolytica adherence that induces membrane blebbing that facilitates the trogocytosis. E. histolytica produces cysteine proteinases utilized in the degradation of colonic mucin glycoproteins, digestion of hemoglobin and villin, inactivation of immune modulators such as interleukin (IL)-18, and the digestion of the host’s extracellular matrix. The digestive action of these proteases contributes to the development of amebic ulcers and tissue damage in the intestinal tract. Amebic ulcers often develop in the cecum, appendix, or adjacent portion of the ascending colon; however, they can also be found

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Ingestion of fecally contaminated water or food containing Entamoeba histolytica cysts

Pleural and pericardial effusions Liver abscess Self-limiting, asymptomatic infection 80% of cases Colon Excystation in lumen of small intestine Multiplication of trophozoites by binary fission

Brain

Invasive disease 20% of cases Extraintestinal disease 10/μL of CSF), increased CSF protein, and neurologic manifestations. Occasionally the larvae may migrate to the eye, causing blindness. Most often, the disease caused by Parastrongylus spp. is self-limiting. 

Laboratory Diagnosis Definitive diagnosis relies on histologic identification of the adult female worm, which has a distinctive morphologic appearance with a spiral, winding uterus resembling a barber pole. Highly specific serologic assays are available for patients with elevated eosinophils. Some cross reactivity has been reported between A. cantonensis and trichinosis, making diagnosis less specific. 

Therapy Anthelmintic therapy, such as mebendazole, may be helpful. It is important to monitor therapy closely because the therapy may actually exacerbate the inflammatory response of the host and cause more systemic damage. If larvae are located within the eye, surgical removal is recommended. 

CHAPTER 51  Tissue Nematodes

Parastrongylus costaricensis (Abdominal Angiostrongyliasis) General Characteristics Parastrongylus costaricensis is found primarily in the cotton rat and black rat. 

Epidemiology The parasite is endemic in areas of Central and South America, including Mexico and Costa Rica. 

Pathogenesis and Spectrum of Disease The life cycle is very similar to that of P. cantonensis. Human infection is typically by ingestion of salad contaminated with excretions from infected slugs or snails. The larvae create inflammatory lesions in the wall of the bowel, resulting in tissue inflammation, necrosis, vomiting, and diarrhea. The patient may experience lower-right-quadrant abdominal pain similar to that manifested in appendicitis. The eggs of P. costaricensis may also remain embedded in the tissue of the human host and are not passed in the patient’s feces. 

Laboratory Diagnosis Histologic identification of the adult worm, larvae, or eggs in tissue sections results in definitive diagnosis. Patients often present with leukocytosis and eosinophilia. Radiologic imaging may be useful. Currently no molecular tests specific for this infection are available. However, for research purposes, A. costaricensis can be identified in tissue by conventional PCR followed by DNA sequencing analysis. 

Therapy Traditional anthelmintic therapy is recommended. 

Gnathostoma spinigerum General Characteristics Gnathostoma spp., a gastric Spirurida, is found in a variety of mammals worldwide. Dogs and cats serve as the definitive host for Gnathostoma spinigerum. Although G. spinigerum is the most common species identified in humans, G. hispidum, G. nipponicum, G. binucleatum, G. procyonis, G. binucleatum and G. doloresi have also been associated with infection. 

Epidemiology The adult worms reside in the stomach of the definitive host, where they mate and produce eggs that are passed in the feces. When the feces are deposited in water, the larvae hatch and infect copepods. The larvae mature in the

741

copepods and are then ingested by a variety of intermediate hosts including fish, snakes, and frogs. Inside the intermediate host, the larvae then migrate to the musculature and encyst until the tissue is ingested by the definitive host. The intermediate hosts may serve as a food staple for a paratenic host, such as a bird. Once ingested by the bird, the larvae can remain viable and be passed to the definitive host or to humans. Once in the definitive host, the larvae excyst and penetrate the gastric wall, migrating and maturing in the stomach. Humans act as accidental hosts when they ingest larvae in contaminated fish. 

Pathogenesis and Spectrum of Disease The worms are incapable of maturation within the human host and migrate aimlessly, causing tissue damage and inflammation, thus causing a disease that resembles visceral larva migrans. The infection is not typically fatal; however, it depends on the migration pattern and organs infected. Any organ system can be involved, but the most common manifestation of infection is localized, intermittent, migratory swelling in the skin and subcutaneous tissues. Such swellings may be painful, pruritic, and/or erythematous. In addition, Gnathostoma spp. commonly cause a parasitic eosinophilic meningitis due to larval migration into the CNS. Systemic infection is typically associated with peripheral eosinophilia, in which the percentage of eosinophils may exceed 50% of the circulating white blood cells (WBCs). 

Laboratory Diagnosis Identification of the larvae in tissue is definitive for diagnosis. The larval head contains four rows of cephalic hooklets, and the body is covered with transverse rows of spines that diminish anteriorly to posteriorly. 

Serologic Testing Both ELISA to detect IgG antibodies and immunoblot testing for neurologic disease has been described; however, these tests are not widely available in the United States and many other countries. 

Therapy Supportive corticosteroid treatment is recommended. Although anthelmintics are not lethal to this organism, they are often recommended. Surgical excision of the larvae is optimal. 

Capillaria hepatica General Characteristics Capillariasis is a parasitic disease in humans caused by two different species of capillarids: Capillaria hepatica and ­Capillaria philippinensis (Chapter 50).

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C. hepatica is transferred through the fecal matter of infected animals and can lead to hepatic capillariasis. Infection is rare but has been reported worldwide. Infective eggs hatch in the intestine of the human host, releasing larvae. The larvae migrate via the portal vein to the liver, where humans are considered a terminal host for the parasite. 

Epidemiology C. hepatica is often found in the liver of such animals as small rodents, monkeys, and prairie dogs and can cause cirrhosis in the animal hosts. When these animals are eaten by larger carnivores, capillarid eggs are ingested and passed in the feces. When this fecal matter is deposited in the soil, the eggs become infective in about 30 days. Accidental ingestion of the eggs from contaminated soil occurs in both animals and humans. The eggs hatch and the larvae migrate to the liver and mature into adult worms. The adult worms lay eggs in the liver. Humans are usually infected after ingesting embryonated eggs in fecal-contaminated food, water, or soil. 

Pathogenesis and Spectrum of Disease C. hepatica can be transmitted human to human when eggs are deposited through human fecal matter into the soil. The eggs then become infective in the soil, and humans ingest infective soil directly by eating soil (pica) or consuming it accidentally or indirectly through contaminated food or water. When a human is infected with only one C. hepatica worm, there are often no signs or symptoms. The adult worms lay eggs in the liver, resulting in clinical manifestations including hepatitis, anemia, fever, hypereosinophilia, and even death. Larvae may also migrate to the lungs, kidneys, or other organs. Table 51.2 summarizes the associated diseases.

Laboratory Diagnosis No eggs are excreted in the feces in true human infections. C. hepatica can be diagnosed by performing a liver biopsy, needle biopsy, or after death at autopsy to identify the adult worm or eggs. 

Therapy Anthelmintic agents including albendazole and mebendazole are recommended. Steroids have been used with C. hepatica infections to help control the inflammation of the liver. 

Prevention Adequate preparation and cooking of seafood—­including fish, snails, crabs, and shrimp—in endemic areas is encouraged. 

Dirofilaria immitis and Other Species General Characteristics The filarial dog heartworm Dirofilaria immitis causes human pulmonary dirofilariasis worldwide. Although D. immitis is the most common species isolated from humans in the United States, additional species infect other mammals. D. repens infects wolves, coyotes, and foxes. Other species include D. ursi in bears; D. striata, wildcats; D. tenuis, raccoons; D. striata, bobcats and pumas; and D. subdermata, porcupines. These organisms are occasionally associated with human infections. D. repens is found in cats and dogs in Europe, Africa, and Asia. Dirofilaria hongkongensis is a newly identified species from Hong Kong and Asia. The parasite is transmitted by dogs and cats, causes ocular dirofilariasis, and is closely related to D. repens. 

Epidemiology The adult worms reside in the right ventricle of the heart of the infected mammal. The adult worm releases microfilariae into the bloodstream, which are then ingested by a mosquito. The microfilariae mature into infective larvae in the insect and are then transferred to another host when the mosquito feeds. The larvae migrate through the host, eventually reaching the heart, where they mature into adult worms. Humans are accidental hosts, and the worms are unable to reach maturity. The organisms die and are swept into the pulmonary circulation, where they become lodged in arteries or arterioles. This obstruction of the pulmonary circulation results in thrombosis, infarction, and inflammation. Eventually a wall of fibrous tissue is deposited around the worm, creating a granulomatous reaction. Species of Dirofilaria that manifest subcutaneously produce nodules that are often tender and may be fixed or migratory. D. repens–associated lesions can occur in a variety of locations, the most typical being exposed sites (e.g., scalp, arms, legs, eyelids, chest), but occasionally lesions have been found in deeper tissue such as the breast, epididymis, spermatic cord, and subconjunctiva. Many reports of D. tenuis involve the facial region (e.g., ocular and periocular sites, oral mucosa, cheek) and breast. There are very rare reports of subcutaneous infections with D. striata, D. ursi, and possibly D. subdermata in humans. D. hongkongensis larvae migrate to the ocular region and cause recurrent eyelid swelling in both eyes and conjunctival inflammation with watery discharge. 

Pathogenesis and Spectrum of Disease Approximately 50% of patients with dirofilariasis are asymptomatic and present with subcutaneous nodules or lung disease. Symptomatic patients present with generalized symptoms such as cough, chest pain, fever, malaise, chills, and hemoptysis. Some patients may present with a peripheral eosinophilia. The respiratory granulomas may

CHAPTER 51  Tissue Nematodes

be identified radiographically and are typically removed to rule out malignancies. Excision of the nodule or granuloma is typically sufficient treatment, resulting in no long-term pathology. 

Laboratory Diagnosis Definitive diagnosis of the parasite from a nodule may be extremely difficult because of the worm’s degeneration in the granuloma. However, the presence of a worm within a pulmonary artery is usually supportive of the diagnosis. Serologic tests are available; however, cross-reactivity may occur with other nematodes; therefore, a negative reaction

Bibliography Baheti NN, Sreedharan M, Krishnamoorthy T, et  al.: Eosinophilic meningitis and an ocular worm in a patient from Kerala, south India, J Neurol Neurosurg Psychiatry 79(3):271, 2008. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Bussaratid V, Dekumyoy P, Desakorn V, et al.: Predictive factors for Gnathostoma seropositivity in patients visiting the Gnathostomiasis Clinic at the Hospital for Tropical Diseases, Thailand during 20002005, Southeast Asian J Trop Med Public Health 41(6):1316–1321, 2010. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Chen J, Liu Q, Liu GH, et  al.: Toxocariasis: a silent threat with a progressive public health impact, Infect Dis Poverty 7(1):59, 2018. Fuller AJ, Munckhof W, Kiers L, et al.: Eosinophilic meningitis due to Angiostrongylus cantonensis, West J Med 159(1):78–80, 1993. Gamble HR, Pozio E, Bruschi F, et al.: International Commission on Trichinellosis: recommendations on the use of serological tests for the detection of Trichinella infection in animals and man, Parasite 11(1):3–13, 2004. Garcia LS: Diagnostic medical parasitology, ed 6, Washington, DC, 2016, ASM Press. Gottstein B, Pozio E, Nöckler K: Epidemiology, diagnosis, treatment, and control of trichinellosis, Clin Microbiol Rev 22(1):127–145, 2009. Hombu A, Yoshida A, Kikuchi T, et al.: Treatment of larva migrans syndrome with long-term administration of albendazole, J Microbiol Immunol Infect 52(1):100–105, 2019. Intapan PM, Khotsri P, Kanpittaya J, et al.: Immunoblot diagnostic test for neurognathostomiasis, Am J Trop Med Hyg 83(4):927–929, 2010. Leung AK, Barankin B, Hon KLE: Cutaneous larva migrans, Recent Pat Inflamm Allergy Drug Discov 11(1):2–11, 2017.

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does not exclude infection. Currently no molecular methods are available in the United States for diagnosis of human dirofilariasis. 

Therapy As previously noted, excision of the granuloma or adult worm is typically sufficient, and additional treatment is not needed.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Li CD, Yang HL, Wang Y: Capillaria hepatica in China, World J Gastroenterol 16(6):698–702, 2010. Miller CL, Kinsella JM, Garner MM, et  al.: Endemic infections of Parastrongylus (Angiostrongylus) costaricensis in two species of nonhuman primates, raccoons, and an opossum from Miami, Florida, J Parasitol 92(2):406–408, 2006. Ogdee JL, Henke SE, Wester DB, et al.: Assessing potential environmental contamination by Baylisascaris procyonis eggs from infected raccoons in Southern Texas, Vector Borne Zoonotic Dis 17(3):185– 189, 2017. Pradeep RK, Nimisha M, Pakideery V, et  al.: Whether Dirofilaria repens parasites from South India belong to zoonotic Candidatus Dirofilaria hongkongensis (Dirofilaria sp. hongkongensis)? Infect Genet Evol 67:121–125, 2019. Schuster A, Lesshafft H, Talhari S, et  al.: Life quality impairment caused by hookworm-related cutaneous larva migrans in resource-poor communities in Manaus, Brazil, PLoS Negl Trop Dis 5(11):e1355, 2011. Simón F, Siles-Lucas M, Morchón R, et al.: Human and animal dirofilariasis: the emergence of a zoonotic mosaic, Clin Microbiol Rev 25(3):507–544, 2012. Veraldi S, Angileri L, Parducci BA, et al.: Treatment of hookwormrelated cutaneous larva migrans with topical ivermectin, J Dermatolog Treat 3:263, 2017. Winkler S, Pollreisz A, Georgopoulos M, et al.: Candidatus Dirofilaria hongkongensis as causative agent of human ocular filariosis after travel to India, Emerg Infect Dis 23(8):1428–1431, 2017. Wu Z, Nagano I, Pozio E, et al.: The detection of Trichinella with polymerase chain reaction (PCR) primers constructed using sequences of random amplified polymorphic DNA (RAPD) or sequences of complementary DNA coding excretory-secretory (E-S) glycoproteins, Parasitology 117(Pt 2):173–183, 1998. Wu Z, Nagano I, Pozio E, et al.: Polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLLP) for the identification of Trichinella isolates, Parasitology 118(Pt 2):211–218, 1999.

Chapter Review 1.  Removal and gradual retraction of the adult gravid female worm is recommended in infections with: a. Ancylostoma braziliense b. Dracunculus medinensis c. Trichinella spiralis d. Toxocara cati 2. The following infection may resemble acute appendicitis: a. Parastrongylus costaricensis b. Gnathostoma sp. c. T. cati d. Parastrongylus caninum 3. After ingesting a roast duck, a 52-year-old male began to experience inflammation of the skin and a small spidery pruritic rash. Which of the following is most likely the cause of the patient’s discomfort? a. D. medinensis b. T. cati c. Gnathostoma spinigerum d. T. spiralis 4. This organism is capable of causing severe neurological damage that often results in death. a. Trichinella sp. b. Toxocara spp. c. Baylisascaris sp. d. Gnathostoma sp.

5. True or False _____ Anthelmintic therapy results in serious complications in all tissue nematode infections. _____ Larvae located in the eye should always be surgically removed, if possible, to prevent blindness. _____ Dirofilariasis often results in severe chest pain, pulmonary embolism, and sudden death. 6. Matching: Match each term with the correct description. _____ Gnathostoma sp. _____ P. cantonensis _____ A. braziliense _____ VLM _____ Toxocara cati _____ trichinosis _____ NLM

a. ocular larva migrans b. roundworm encephalitis c. creeping eruption d. barber pole e. cephalic hooklets f. 1:32 titer g. calcification

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52

Blood and Tissue Filarial Nematodes OBJECTIVES 1. Describe the distinguishing morphologic characteristics and basic life cycle (vectors, hosts, and stages of infectivity) for each of the parasites listed. 2. Define microfilariae, hydrocele, chyluria, and sheath. 3. Describe the diseases and mechanism of pathogenicity, including route of transmission, for each of the species listed. 4. Explain periodicity, including nocturnal and diurnal, as it relates to infection with microfilariae and correlate it with the life cycle of the associated arthropod vector. 5. Differentiate the microfilariae based on the presence or absence of a sheath and the arrangement and number of tail nuclei. 6. Describe the two methods of concentrating blood specimens for the identification of organisms within the peripheral blood. 7. Determine the cause of infection based on patient history, signs and symptoms, and laboratory results.

PARASITES TO BE CONSIDERED NEMATODES Blood and Tissues (Filarial Worms) Brugia beaveri Brugia leporis Brugia malayi Brugia pahangi Brugia timori Dirofilaria spp. Loa loa Mansonella ozzardi Mansonella perstans Mansonella streptocerca Onchocerca volvulus Wuchereria bancrofti

B

lood and tissue filarial nematodes are roundworms that infect humans. These organisms are transmitted via a blood-sucking arthropod vector such as a mosquito, midge, or fly. The filarial nematodes infect the subcutaneous tissues, deep connective tissues, body cavities, and lymphatic system. The life cycles of the filarial nematodes are complex (Fig. 52.1). The infective larval stage resides in the 744

insect vector, and the adult worm stage, which is the pathogenic form, resides in humans. When the arthropod vector feeds on a human blood meal, the infective larvae are injected into the bloodstream. The larvae are motile and migrate to the lymphatic vessels. The infective larvae grow and develop into the adult gravid worm in the human host over a period of months. The male and female adult worms mate in the definitive human host. The female worm produces large numbers of larvae called microfilariae. Depending on the species, the microfilariae may maintain the egg membrane as a sheath or may rupture the egg membrane, resulting in an unsheathed form. These parasites can reside in the host for many years and cause chronic, debilitating conditions and severe inflammatory responses. Identification of the various species is based on the morphology of the microfilaria, the defined circadian rhythm, and the location within the human host. The morphologic characteristics of the microfilariae are important in the identification process and include the presence or absence of the sheath and the presence and arrangement of the nuclei in the worm’s tail (Fig. 52.2). A comparison of the morphologic characteristics of the pathogenic filarial worms is depicted in Fig. 52.3. Diagnosis of infection is based on the identification of the microfilariae in the host’s blood or tissues.

Wuchereria bancrofti General Characteristics Wuchereria bancrofti is transmitted via a mosquito, the Culex fatigans, Anopheles, or Aedes spp. The adult worm, or microfilaria, has a sheath that stains faintly or not at all. It may grow to approximately 298 μm long and 7.5 to 10 μm wide. The tail is pointed and no nuclei are present (Fig. 52.4). 

Epidemiology W. bancrofti is the most commonly identified species of filarial worms that infect humans. It is widely distributed in the tropics and subtropics, including Africa, South America, Asia, the Pacific Islands, and the Caribbean. The mosquito vectors have complex life cycles that include laying eggs and developing larvae on the surface of a water source. When

CHAPTER 52  Blood and Tissue Filarial Nematodes

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Lymphatic Filariasis Wuchereria bancrofti 1 Mosquito stages

8

Mosquito takes a blood meal (L3 larvae enter skin)

Human stages

Migrate to head and mosquito’s proboscis

7

i

L3 larvae

2

6

L1 larvae

3 4

5

Adults in lymphatics

Microfilariae shed sheaths pentrate mosquito’s midgut, and migrate to thoracic muscles

Mosquito takes a blood meal (inests micorfilariae)

Adults produce sheathed microfilariae that migrate into lymph and blood channels

d i

= Infective stage

d

= Diagnostic stage

• Fig. 52.1  Life cycle of Wucheria bancrofti. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

Microfilariae isolated from host

Sheath present

Sheath absent

Tail nuclei No tail nuclei Do not extend to tip Wuchereria bancrofti

Extend to tip

Continuous row Loa loa

Tail nuclei

Tail bent or flexed, O. volvulus

Not continuous, two nuclei at tip Brugia malayi

Do not extend to tip M. ozzardi

Extend to tip M. perstans M. streptocerca (Shepherd’s hook)

• Fig. 52.2  Identification of microfilariae.

the larvae mature into adult mosquitos, the males and females will swarm in the evening and mate. The female requires a blood meal to reproduce. The mosquito becomes the intermediate host for the microfilarial parasite. Humans are the definitive host and the reservoir for W. bancrofti. The parasite has two forms that demonstrate different periodicities. The nocturnal periodic form is found in the peripheral blood during the night between 10 p.m. and 4 a.m. The

second form is found only in the Pacific Islands and is present in the blood at all times, but more frequently during the day in the afternoon hours. 

Pathogenesis and Spectrum of Disease Microfilarial clinical disease varies geographically based on the species of nematode causing the infection. The disease

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A

B

C

D

E

F

G

•

Fig. 52.3 Anterior and posterior ends of microfilariae found in humans. (A) Wuchereria bancrofti. (B) Brugia malayi. (C) Loa loa. (D) Onchocerca volvulus. (E) Mansonella perstans. (F) Mansonella streptocerca. (G) Mansonella ozzardi.

infection. Obstruction of the lymphatic vessels causes fibrosis and a proliferation of dermal and connective tissue, resulting in the wrinkled, dry appearance of an “elephant” extremity. Lymphedema may also occur in the arms, female breasts, and scrotum of infected individuals. Acute lymphatic filariasis results from worms residing within the lymph nodes. The lymph nodes swell, and lymphangitis may appear peripherally from the infected node. A hydrocele—a fluid-filled sac within the scrotum—may form when adult worms block the retroperitoneal or subdiaphragmatic lymphatic vessels. Obstruction of the lymphatic vessels may result in a condition referred to as chyluria, which is a result of lymphatic rupture and fluid entering the urine. The urine will appear milky white. Resulting infection and changes in the skin may lead to increased bacterial infections. Patients residing in tropical regions where filarial parasites are endemic may present with a syndrome referred to as tropical pulmonary eosinophilia (TPE). The microfilariae migrate through the pulmonary blood vessels to the lungs, causing an allergic hypersensitivity in the host. The patients develop a strong immune response to the presence of the parasites with an elevated serum immunoglobulin E (IgE) level. Symptoms of TPE include weight loss, lowgrade fever, cough and wheezing at night, and lymphadenopathy. Without treatment, patients may develop chronic and progressive respiratory complications leading to death.

Endosymbiont W. bancrofti, Brugia spp., and Onchocerca volvulus harbor Wolbachia sp., an endosymbiotic alpha-proteobacterium. Wolbachia is an obligate intracellular organism. The parasites require the endosymbiont for larval development, viability, and fertility. The bacteria have been implicated in the pathogenesis of the infection with filarial parasites. The bacterial antigens enhance the host’s inflammatory response, leading to increased scarring and damage within the host’s lymphatic system. The bacterium is sensitive to tetracycline, doxycycline, azithromycin, and rifampin. Combination antibiotic treatment in conjunction with treatment for the parasite infection improves clearance of the filarial parasite. 

Laboratory Diagnosis Direct Detection • Fig. 52.4  Microfilaria of Wuchereria bancrofti in thick blood film.

may present as acute or asymptomatic for many years. W. bancrofti causes bancroftian filariasis and elephantiasis. The adult worm resides in the lymphatic vessels distal to the lymph nodes. The presence of organisms within the host results in an immunologic response including inflammation, lymphedema, and hyperplasia. Lymphedema most often occurs in the lower extremities. Elephantiasis is a crippling condition that results from extended periods of filarial

Definitive laboratory diagnosis is based on identification of the parasites in blood, fluids, or tissue. Blood samples should be drawn in accordance with the periodicity of the infection to optimize the likelihood of isolating the infecting organism. Direct examination of blood, urine, hydrocele fluid, or chyle may serve to identify the parasite. The fluid is placed on a slide and air-dried to prevent distortion of the parasite. The specimen should be stained with Giemsa, Wright, or hematoxylin stain and examined microscopically. Ultrasound may be used to visualize the organisms within the tissues.

CHAPTER 52  Blood and Tissue Filarial Nematodes

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Filariasis (Brugia malayi) Mosquito Stages

8

Mosquito takes a blood meal (L3 larvae enter skin)

Human Stages

Migrate to head and mosquito’s proboscis

7

6

1

L3 larvae

i 2

Adults in lymphatics

3

Adults produce sheathed microfilariae that reach the blood stream

L1 larvae

4 5

Microfilariae shed sheaths pentrate mosquito’s midgut, and migrate to thoracic muscles

Mosquito takes a blood meal (ingests micorfilariae)

d i

= Infective stage

d

= Diagnostic stage

• Fig. 52.5  Life cycle of Brugia spp. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

Nucleopore filtration or the Knott concentration may be used to increase the likelihood of isolating a filarial parasite from blood. The blood is passed through a polycarbonate filter that contains a 2-μm pore. Distilled water is passed through the filter, lysing the red blood cells and improving visualization of the parasites. The filter is air-dried, stained with Wright or Giemsa, and examined for the presence of microfilaria. The Knott concentration uses centrifugation to concentrate the organisms onto a slide. One milliliter of anticoagulated blood is placed in 9 mL of 2% formalin and centrifuged at 1500 rpm for 1 minute; the sediment is applied to a microscope slide. The slide is stained and examined microscopically. With high-frequency ultrasound, adult worms can sometimes be visualized moving within the lymphatics. 

infections. The commercial testing formats available are not approved by the US Food and Drug Administration (FDA). 

Serologic Testing

Nucleic Acid Detection

Serologic assays that measure antibody response have limited utility in the diagnosis of infections with microfilariae. The antibodies tend to demonstrate a high cross reactivity with other antibodies made in response to a wide variety of parasitic worm infections. The absence of an antibody reaction indicates the lack of infection by a microfilarial species. Laboratory detection of W. bancrofti–circulating antigens has demonstrated high specificity (>97%) and sensitivity (from 70% to 80%) in detecting parasitic

Nucleic acid–based methods have been shown to be the most sensitive diagnostic tests for definitive diagnosis of infection with microfilariae. A variety of formats have been developed, including DNA hybridization, polymerase chain reaction (PCR), multiplex PCR, restriction length polymorphism (RFLP), quantitative PCR, and a PCR enzyme-linked immunosorbent assay. These methods can discriminate past and current infection and can be used to monitor therapy. PCR amplification is available in reference

Antigen Detection A variety of rapid antigen tests are capable of detecting circulating antigens of W. bancrofti from whole blood, serum, or plasma. These tests can be used on blood drawn at any time, thus avoiding the need for specimen collection that depends on the periodicity of the microfilariae. Two rapidformat immunochromatographic tests have been shown to be useful and sensitive for the detection of W. bancrofti and are being used widely by lymphatic filariasis elimination programs.

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laboratories for the rapid diagnosis of an infection with blood microfilariae including W. bancrofti and Brugia spp. Multiplex PCR-RFLP can differentiate W. bancrofti, Dirofilaria imminitis, Dirofilaria repens, Brugia pahangi, and Brugia malayi in blood. B. pahangi is a common parasite of dogs and cats in Malaysia and has been implicated in human infections. A loop-mediated isothermal application assay has also been developed that may be useful in the field or as a point-of-care diagnostic test. These tests are not currently commercially available. 

Brugia malayi and Brugia timori General Characteristics The Brugia spp. are lymphatic filarial parasites resembling W. bancrofti. The adult parasites, microfilariae, vary in size (Brugia timori, 310 μm long and 5 to 6 μm wide; Brugia malayi, 177 to 230 μm long and 5 to 6 μm wide), have a different geographic distribution, and do not typically cause lymphadenitis in the genital regions. 

Epidemiology The Brugia spp. are distributed throughout the Far East including China, Indonesia, Korea, Malaysia, Japan, India, and the Philippines. The distribution of B. timori is limited to the two islands of Timor, an island of Indonesia. The organism is transmitted via mosquitos included in the genus Anopheles and Mansonia. 

Pathogenesis and Spectrum of Disease As in infections with W. bancrofti, two periodic forms exist. The nocturnal form is the most common and is located near areas of coastal rice fields, whereas the nonperiodic form is associated with infections in areas near swampy forests. The pathogenesis and spectrum of disease is essentially the same as for W. bancrofti, with the exception that involvement of the genital lymphatic vessels is predominantly associated with W. bancrofti. Urogenital involvement with chyluria does not occur in infections with Brugia spp. Clinical disease progresses faster after infection with B. malayi than with W. bancrofti. Microfilariae may appear in the blood in as little as 3 to 4 months. Brugia spp. have been implicated in zoonotic infections of dogs, cats, rabbits, and raccoons worldwide. Mosquitos transmit the infection by feeding on an infected animal and then a human host in approximately 2 weeks. Cases of human infection have occurred in the United States in the northeastern region with B. beaveri. B. beaveri infect raccoons, bobcats, or mink and B. leporis infects rabbit. Clinical disease is typically asymptomatic but may present with a tender region in the cervical, axillary, or inguinal region. The lymphatic mass may contain either a live or a dead worm. If the worm is no longer viable, the mass may be surrounded by a granulomatous reaction. 

Laboratory Diagnosis Definitive diagnosis is generally by identification of microfilariae in the blood of infected individuals. The microfilariae can be distinguished from W. bancrofti morphologically. The B. malayi microfilariae are sheathed and contain four to five subterminal and two terminal nuclei in the tail. B. timori also contains five to eight subterminal and terminal nuclei in the tail, but they are much larger than B. malayi. The B. malayi sheath will stain bright pink with Giemsa, whereas the B. timori sheath does not stain. The microfilariae of B. timori tend to be somewhat longer. High-frequency ultrasound has been useful in identifying adult worms in various locations within the patient, such as lymphatic vessels of the legs, inguinal area (groin or lower abdomen), lymph nodes, and female breasts. Nucleic acid–based methods have been developed as previously indicated in the diagnosis of W. bancrofti and can differentiate microfilariae. 

Therapy Diethylcarbamazine (DEC) is the treatment of choice for lymphatic filarial parasites including W. bancrofti and Brugia spp. Additionally, ivermectin and albendazole may be used. Death of the microfilarial worms may result in an increased hypersensitivity reaction requiring the need for treatment with antihistamines to limit the inflammatory symptoms. 

Prevention The use of insect repellent is recommended for travelers in areas where the parasites are endemic. DEC has also been used for prophylactic treatment before travel. Vector control studies in combination with mass drug administration of DEC and ivermectin have successfully decreased the population of the arthropod (insect) vectors and decreased filarial infection in the human hosts. 

Loa loa General Characteristics Loa loa, commonly referred to as the eye worm, is a microfilaria that circulates in the bloodstream with a diurnal periodicity that peaks in the afternoon between 12:00 p.m. and 2:00 p.m. and resides in the subcutaneous tissue in the human host. The microfilariae may grow up to 300 μm. 

Epidemiology The parasite is found within the rain forests of West and Central Africa. The organism is transmitted through a bite of the tabanid fly or deer fly of the genus Chrysops. The female lays her eggs on the leaves of small plants near the water. The larvae feed on small insects and develop in wet

CHAPTER 52  Blood and Tissue Filarial Nematodes

soil. The male fly feeds on pollen, and the female feeds on blood. 

Pathogenesis and Spectrum of Disease

L. loa. These have been adapted to loop-mediated isothermal amplification (LAMP), allowing the potential for point-ofcare and in-field use. A single PCR assay test for loiasis has been approved for diagnostic use in the United States. 

The organism is often associated with asymptomatic infection. The larvae develop into adult worms in approximately 6 to 12 months but can persist in the human host for up to 17 years. The infection is typically identified when the adult worm is seen migrating within the subconjunctiva of the host. Symptoms associated with infection include episodic calabar swellings, which are localized areas of transient angioedema in response to the production of parasitic metabolic products. Predominant swelling on the extremities with inflammation of nearby joints and peripheral nerves may occur. Immune-mediated encephalopathy, nephropathy, and cardiomyopathy may occur. 

Therapy

Laboratory Diagnosis

Onchocerca volvulus

Infections with Loa loa may be asymptomatic for many years before the appearance of microfilariae in the peripheral blood. Therefore patient diagnosis is often made based on the patient’s clinical symptoms, including calabar swelling, eosinophilia, and travel or residency in an endemic area.

General Characteristics

Direct Detection Definitive diagnosis is made by identification of adult worms in the eye or in tissue or by identification of microfilariae in the peripheral blood. Microfilariae have a sheath that does not stain with Giemsa. The adult females are larger than the adult males, and the nuclei extend to the tail in an irregularly arranged fashion. 

Serologic Testing As with other filarial infections, serologic assays have limited use for diagnosis. A Loa-specific recombinant protein (LLSXP-1) has been used in the development of an enzymelinked immunosorbent assay (ELISA) and has demonstrated improved specificity but limited sensitivity. A rapid detection test for LLSXP-1 IgG has been developed that demonstrates a sensitivity greater than 90% and a specificity of approximately 95%. Detection of IgG may be useful in confirming the diagnosis in travelers to areas of endemicity in the presence of unexplained eosinophilia and appropriate clinical symptoms. 

Nucleic Acid Detection PCR-based techniques have been developed for the detection of Loa loa, with specificities and sensitivities as high as 100% and 95%, respectively. However, these techniques yield negative results during prepatency, the period between initial infection and detectability in the host. These tests also demonstrate false-positive results in individuals with previous L. loa infections, as they also detect DNA from dead microfilariae in blood. Real-time qPCR assays have also been developed with high species specificity and sensitivity (96%) for

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DEC is the treatment of choice. In heavy infections, inflammation and allergic reactions may occur, requiring the administration of antiinflammatory medications. Allergic responses can result in central nervous system damage, encephalitis, coma, and death. 

Prevention Prophylactic treatment with DEC has been used to prevent infection. 

Onchocerca volvulus predominantly resides in tissue nodules within the host. Microfilariae measure approximately 300 μm long by 5 to 9 μm wide. 

Epidemiology O. volvulus is found throughout Africa, Central America, and South America. The parasite is transmitted by the black fly, Simulium spp. The black fly lays its eggs in running water where the larvae attach to rocks. The larvae feed on algae and bacteria. The adults emerge as a flying insect. The females require a blood meal, whereas the males are nectar feeders. The flies feed predominantly during the day. 

Pathogenesis and Spectrum of Disease Onchocerciasis, commonly referred to as river blindness, is a result of subcutaneous infection with the parasite. The infections are typically localized to the skin, lymph nodes, and eyes. Skin infections result in pruritus, edema, and erythema. Hypopigmentation or hyperpigmentation can occur after a lengthy infection. Nodules containing the adult worms vary in size and are firm and tender. Lymphadenopathy may be found in the inguinal or femoral regions. Enlargement of the lymph node may result in a condition referred to as “hanging groin,” which may develop into a hernia. Onchocercal eye disease may be seen in moderate to heavy infections. Infections of the eye may lead to serious damage and blindness. Mortality increases among adults who experience blindness and systemic infection. 

Laboratory Diagnosis Direct Detection Definitive diagnosis is made from the identification of the microfilariae from tissue such as in a nodule or skin snip.

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Prevention Black fly larval control using insecticides in endemic areas has been used to assist in the control of transmission of O. volvulus. Community-based administration of ivermectin every 6 to 12 months is also being used to interrupt the transmission of the parasite in endemic areas and has been successful for the elimination of the parasite in Latin America. 

•

Fig. 52.6 Microfilaria of Onchocerca volvulus. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

Skin samples are placed in physiologic buffered saline for up to 24 hours. After incubation, the microfilariae will emerge from the tissue and can be visualized microscopically. Occasionally the microfilariae may be found in blood or urine after treatment. Microfilariae may also be visible in the cornea and the anterior chamber of the eye. The microfilariae lack a sheath. The tail is tapered, appears bent or flexed, and does not include extension of nuclei to the tip (Fig. 52.6). 

Serologic Testing Although serologic tests generally lack specificity, recombinant ELISAs using multiple antigens have demonstrated increased sensitivity and specificity for the diagnosis of onchocerciasis. A rapid test, the SD Bioline Onchocerciasis, for the detection of an O. volvulus antigen known as OV-16, is commercially available. The test is used predominantly for surveillance and elimination of infection but may also be used for the diagnostic confirmation of infection. 

Nucleic Acid Detection PCR amplification assays using material from skin snips or skin scratches provides high sensitivity and specificity; these have been developed, but the limited availability of technical expertise as well as the high cost of the test restrict its use in resource-limited settings and to research laboratories. 

Therapy Ivermectin is the recommended treatment and is effective against both adult worms and microfilariae. However, in Africa, where O. volvulus and L. loa are coendemic, ivermectin treatment is often associated with encephalopathy in patients with heavy microfilaria infections. Surgical excision of nodules containing adult worms is recommended when they are located on the head. There is evidence that a 6-week course of doxycycline is useful in targeting the endosymbiotic proteobacteria Wolbachia spp. This causes sterility in the female adult worms for extended periods, reducing the infection in the host. 

Mansonella spp. (M. ozzardi, M. streptocerca, M. perstans) General Characteristics Mansonella spp. are generally not associated with serious infections. The microfilariae of all species are very similar in size, ranging from approximately 200 to 225 μm long and 4 to 6 μm wide. 

Epidemiology Mansonella spp. are distributed throughout varied geographic regions in Africa and South America. Mansonella ozzardi is limited to Central and South America and the Caribbean islands. The parasites are transmitted by biting midges of the genus Culicoides. The female requires a blood meal for the maturation of eggs and typically bites in the early evening or morning hours. Transmission of M. ozzardi has also been associated with bites from the blackfly (Simulium amazonicum). 

Pathogenesis and Spectrum of Disease Mansonella streptocerca is found in the skin; however, most infected individuals appear to be asymptomatic. Patients may present with a pruritic or papular rash and pigmentation changes. In addition, lymphadenitis may occur. Mansonella perstans resides in the pericardial, pleural, and peritoneal cavities. The location of the M. ozzardi adult worms is unknown. Symptomatic patients present with swelling of the arms, face, or other body parts similar to the calabar swelling identified in infections with L. loa, M. ozzardi, or M. perstans found in the blood. M. perstans and M. ozzardi do not demonstrate periodicity when circulating within the bloodstream. M. ozzardi infections are often asymptomatic and therefore are not well characterized. Some infections with M. ozzardi demonstrate headache, joint pain, fever, pulmonary symptoms, adenopathy, hepatomegaly, and pruritus. 

Laboratory Diagnosis Laboratory diagnosis for M. perstans and M. ozzardi is generally made by identifying the microfilariae in blood or other body fluids. M. perstans and M. streptocerca can be diagnosed by identifying the microfilariae in skin snips.

CHAPTER 52  Blood and Tissue Filarial Nematodes

Mansonella spp. microfilariae do not possess sheaths. M. streptocerca and M. perstans tails contain nuclei that extend to the end of the tip. The tail of M. streptocerca is often referred to as a shepherd’s crook. M. ozzardi organisms have tails with nuclei that do not extend to the tip. 

Nucleic Acid Detection A few studies have evaluated the use of PCR for the detection of Mansonella spp. from venous and capillary blood samples. The tests appear to be more sensitive and specific than microscopy. Currently there are no commercially available nucleic acid tests for Mansonella spp. 

Therapy Ivermectin is effective in the treatment of M. ozzardi infections. DEC is effective in treating both the adult and microfilarial forms of M. streptocerca. Treatment of M. perstans infections has not been effective in most cases. However, because of infection with the endosymbiotic bacteria, Wolbachia spp., treatment with doxycycline has demonstrated some limited success. 

Prevention Prevention relies on the use of insect repellents and adequate clothing. 

Dirofilaria spp. (D. immitis, D. repens, D. tenuis) General Characteristics Filarial nematodes of the genus Dirofilaria cause dirofilariasis. There are many species of Dirofilaria, but human infection is most commonly associated with three species, D. immitis, D. repens, and D. tenuis. Female adult worms grow up to 10 to 12 in in length, whereas males are 4 to 6 in long and have spirally coiled tails. Microfilariae are white, long, and threadlike. 

Epidemiology The definitive natural hosts for these three species are dogs and wild canids, such as foxes and wolves (D. immitis and D. repens) and raccoons (D. tenuis). D. immitis is also known as heartworm and is the cause of human dirofilariasis in the eastern and southeastern United States. D. repens is the leading cause of human dirofilariasis in Europe and is found in Africa and Asia. D. tenuis is found in raccoons in the United States. The adult worms produce microfilariae within the definitive animal host; these circulate in the blood and are ingested by mosquitoes during a blood meal. In mosquitoes, the microfilariae develop into larvae that migrate to the

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proboscis (the long tubular part of the mosquito’s mouth). During a blood meal, the larvae are released into the skin of the human host. The worms are unable to reach maturity in the human host, therefore no microfilariae are detectable. Occasionally larvae will migrate to the pulmonary vessels. Several types of mosquitoes are capable of transmitting Dirofilaria infection, including Aedes, Anopheles, and Mansonia. 

Pathogenesis and Spectrum of Disease Humans and a wide range of other mammals are accidental hosts that play no role in the transmission of Dirofilaria. In these hosts, Dirofilaria larvae can develop into adult worms but the worms remain sexually immature and no microfilariae are produced. The number of human dirofilariasis cases reported has increased dramatically in recent years. The worms produce an inconspicuous granulomatous reaction in the subcutaneous tissue, resulting in the formation of a nodule, or they may migrate to the lung, causing human dirofilariasis. Pulmonary infections are usually asymptomatic but may cause chest pain, cough, fever, and pleural effusion. If the worm lodges in the pulmonary artery, an infarct may occur. Eye infections with Dirofilaria have been identified in North America, Europe, Australia, Africa, Asia, and the Middle East. These infections present with moderate to severe inflammation, blurred vision, and swelling of the eyes. All Dirofilaria examined have contained the Wolbachia endosymbiont. Rarely, D. immitis worms have been identified in the brains and testicles of humans. 

Laboratory Diagnosis Dirofilariasis is diagnosed most frequently by the examination of inflammatory lung tissue or skin nodules. The worm’s cuticle contains chitin, which can be visualized in tissue sections by staining with calcofluor white. 

Therapy The only method of treatment of human dirofilariasis is surgical removal of the lesion or extraction of the worm. Extraction is not essential, however, as the worms die and are often cleared or sequestered in granulomata without treatment. 

Prevention Routine methods for avoiding mosquito vectors will help to prevent infection.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

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Bibliography Abbas KF, El-Monem SG, Malik Z, et  al.: Surgery still opens an unexpected bag of worms! An intraperitoneal live female Dirofilaria worm: case report and review of the literature, Surg Inf ect(3)323–325, 2006. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Boatin BA, Toé L, Alley ES, et  al.: Diagnostics in onchocerciasis: future challenges, Ann Trop Med Parasitol 92(suppl 1):S41–S45, 1998. Boussinesq M: Loiasis, Ann Trop Med Parasitol 100(8):715–731, 2006. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Drame PM, Fink DL, Kamgno J, Herrick JA, Nutman TB: Loopmediated isothermal amplification for rapid and semiquantitative detection of Loa Loa infection, J Clin Microbiol 52(6):2071–2077, 2014. Ettinger S, Feldman E: Textbook of veterinary internal medicine, ed 7, St Louis, 2010, Saunders. Fischer P, Wibowo H, Pischke S, et  al.: PCR-based detection and identification of the filarial parasite Brugia timori from Alor Island, Indonesia, Ann Trop Med Parasitol 96:809–821, 2002. Garcia LS: Diagnostic medical parasitology, ed 6, Washington, DC, 2016, ASM Press. Hamlin KL, Moss DM, Priest JW, et  al.: Longitudinal monitoring of the development of antifilarial antibodies and acquisition of Wuchereria bancrofti in a highly endemic area of Haiti, PLoS Negl

Trop Dis 6(12):e1941, 2012, https://doi.org/10.1371/journal. pntd.0001941. Johnstone C: Heartworm. In Parasites and parasitic diseases of domestic animals, Philadelphia, 1998, University of Pennsylvania. Medeiros JF, Pires Almeida TA, Tavares Silva LB, et al.: A field trial of a PCR-based Mansonella ozzardi diagnosis assay detects high-levels of submicroscopic M. ozzardi infections in both venous blood samples and FTA card dried blood spots, Parasit Vectors 8:280, 2015. Mishra K, Raj DK, Hazra RK, et al.: The development and evaluation of a single step multiplex PCR method for simultaneous detection of Brugia malayi and Wuchereria bancrofti, Mol Cell Probes 21:355–362, 2007. Nolan TJ: Dirofilaria immitis, Philadelphia, 2004, University of Pennsylvania. Pani SP, Hoti SL, Elango A, et al.: Evaluation of the ICT whole blood antigen card test to detect infection due to nocturnally periodic Wuchereria bancrofti in South India, Trop Med Int Health 5:359– 363, 2000. Simón F, López-Belmonte J, Marcos-Atxutegi C, Morchón R, MartínPacho JR: What is happening outside North America regarding human dirofilariasis? Vet Parasitol 133(2–3):181–189, 2005. Sunish IP, Rajendran R, Mani TR, et  al.: Vector control complements mass drug administration against bancroftian filariasis in Tirukoilur, India, Bull WHO 85:138–145, 2007. Theis JH: Public health aspects of dirofilariasis in the United States, Vet Parasitol 133(2–3):157–180, 2004. Udall DN: Recent updates on onchocerciasis: diagnosis and treatment, Clin Infect Dis 44(1):53–60, 2007.

CASE STUDY 52.1 A 45-year-old male returned to the United States after a 3-week safari in Central Africa. He presented to his physician complaining of a tender area near his groin and discomfort during urination. In addition, he was having difficulty sleeping at night because of intermittent periods of fever. A complete blood count was drawn, and the patient exhibited a mild eosinophilia. All other results appeared normal. Urinalysis revealed no abnormal laboratory results. A computed tomography scan of the patient’s lower abdomen and groin showed an unusual mass in his inguinal region. The physician admitted the patient for further observation and tests. Subsequent testing included additional peripheral blood collection during the periodic fevers. Thick and thin smears disclosed no unusual organisms. After concentration of a blood sample and staining with Giemsa, the organism depicted in Fig. 52.6 was identified.

Questions 1. Identify the parasite depicted in Fig. 52.7.

• Fig. 52.7  Identified organism from a patient. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

2. What is the recommended treatment for this patient? 3. What additional parasite could be associated with this patient’s symptoms?   

Chapter Review 1. Periodicities associated with filarial infection are a. Periods of increased microfilariae in the peripheral circulation b. Always regular during the nighttime hours between 10 p.m. and 4 a.m. c. Present in all filarial infections d. Correlated directly with the patient’s symptoms 2. Lymphatic vessel involvement within the retroperitoneal region is associated with infection with a. M. perstans b. B. malayi c. W. bancrofti d. L. loa

3. The endosymbiont Wolbachia sp. is required for parasite reproduction in all of the following except: a. O. volvulus b. B. timori c. L. loa d. W. bancrofti 4. A 25-year-old female patient residing in the Amazon Basin presented to the mobile clinic complaining of intermittent fever, swollen lymph nodes, and a nonproductive cough. Her symptoms persisted for several months. Laboratory results indicated a normal white blood cell count. Serum protein electrophoresis demonstrated an elevated gamma fraction. What additional laboratory tests would assist the physician’s diagnosis? a. Serologic assay b. Serologic assay coupled with PCR for microfilariae identification c. Fractionation of the gamma fraction to identify her antibody abnormality d. Nasal aspirate and sputum specimen for viral and bacterial culture 5. Treatment for microfilariae can be complicated because of: a. Diethylcarbamazine resistance b. Ivermectin resistance c. Allergic reactions to dying parasites d. Drug toxicity 6. True or False _____ L. loa requires surgical removal from the infected host. _____ Heavy microfilaria infections are associated with the development of onchocerciasis. _____ All species of Mansonella contain a sheath and terminal nuclei. _____ The definitive host for dirofilariae causes severe infection in the lungs and eyes. 7. Matching: Match each term with the correct description. _____ Knott concentration _____ chyluria _____ B. timori _____ L. loa _____ B. malayi _____ O. volvulus _____ M. streptocerca _____ D. immitis

a. unsheathed, five to eight subterminal nuclei b. shepherd’s crook c. sheathed, four to five subterminal nuclei d. centrifugation e. dog heart worm f. lymph fluid g. calabar swelling h. river blindness

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Intestinal Cestodes OBJECTIVES 1. Describe the distinguishing morphologic characteristics, clinical disease, basic life cycle (vectors, hosts, and stages of infectivity), and laboratory diagnosis for the intestinal cestodes included in this chapter. 2. Define and identify (where appropriate) the following parasitic structures: scolex, proglottids, rostellum, hermaphroditic, oncosphere, hexacanth embryo, strobila, bothria, and coracidium. 3. Compare and contrast autoinfection and hyperinfection. 4. List several methods of control and prevention of tapeworm infection. 5. Correlate the life cycles with the specific diagnostic stage(s) for each organism.

PARASITES TO BE CONSIDERED Intestinal Cestodes (Tapeworms) Diphyllobothrium latum Diphyllobothrium nihonkaiense Diphyllobothrium spp. Dipylidium caninum Hymenolepis nana Hymenolepis diminuta Taenia asiatica Taenia crassiceps Taenia saginata Taenia solium Taenia spp.

T

he intestinal cestodes are commonly referred to as tapeworms. Tapeworms have a long, segmented, ribbonlike body with a specialized structure for attachment, or scolex, at the anterior end. The adult tapeworm consists of a chain of segments: proglottids, which develop posteriorly from the neck region of the scolex forming the body or strobila. Proglottids may be classified as immature, mature, or gravid containing the uterus and eggs. The crown of the scolex, the rostellum, may be smooth or armed with hooks. The body of the worm (proglottids) varies in the geometric characteristics or number of segments according to the genus and species of the cestode. The mature cestode is hermaphroditic. In other words, the organism contains both male and female reproductive organs. Food is absorbed from the host through the worm’s integument, the outer covering or skin of the organism. Adult worms typically inhabit the small intestine; however, humans may be host to

either the adult or the larval forms, depending on the infecting species. Humans infected with a cestode pass the eggs in the feces. The embryo may be visible within the tapeworm egg as an oncosphere (larva tapeworm within an embryonic envelope, infective stage) or hexacanth embryo. The intermediate host ingests feces containing the adult tapeworm eggs, which further develop into the larva of the cestode. Cestodes generally require one or more intermediate hosts for the completion of their life cycle. Intestinal tapeworm infections are generally asymptomatic. However, if the larval stage develops in human organs outside the intestine, they may cause additional life-threatening complications. Fresh or preserved stools are the specimens of choice for ova and parasites (O&P) examination and cestode identification. Preserved stool containing adult worms or a string of segments (strobila) or the scolex may also be used for diagnosis. Development of serologic testing and molecular assays are more sensitive than stool examination. Although this type of testing may be useful for primary screening, it is most commonly used for confirmation testing for parasitic disease. Chapter 46 describes the methods and specimen requirements in more detail as they relate to parasitology.

Diphyllobothrium latum General Characteristics Diphyllobothrium latum, the freshwater broad fish tapeworm, is the largest human tapeworm and the most common species identified within this genus. Adult worms have been known to reach up to 15 m in length, with more than 3000 to 4000 proglottids, and reside within a host for 30 years or more. The proglottids are characteristically wider than long, with a central rosette-shaped uterine structure (Fig. 53.1). The scolex is spatulate and contains two shallow sucking grooves referred to as bothria (Fig. 53.2A and B). Diphyllobothrium spp. have unembryonated eggs. The eggs are operculated (appears as a lid) with a terminal knob, similar to trematode eggs (Fig. 53.2C ). The intermediate hosts include crustaceans and freshwater fish. 

Epidemiology Diphyllobothrium is not found in the tropics; it is commonly found worldwide where cool lakes are contaminated 753

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by sewage. Diphyllobothrium species can be found wherever freshwater or marine fish are eaten raw, pickled, or marinated. This includes such freshwater fish as burbot, pike, perch, ruff, and salmon. The parasite is found in North America, including the upper Midwest around the Great Lakes, Alaska, and Canada. D. nihonkaiense, originally endemic in Japan, is an emerging parasite in other areas of Europe. There are 14 species of Diphyllobothrium, all capable of infecting humans. Other species of Diphyllobothrium that have been identified in human infection include D. pacificum (Southern Pacific coast of South America), D. cordatum, D. ursi, D. dendriticum (New Guinea and Australia), D. lanceolatum, D. dallieae, and D. yonagoensis.  •

Fig. 53.1 Proglottid demonstrating rosette-shaped uterus in Diphyllobothrium latum. (Courtesy Division of Parasitic Diseases/ Centers for Disease Control and Prevention.)

A

B

Pathogenesis and Spectrum of Disease Diphyllobothrium spp. is the only cestode to have an aquatic life cycle (Fig. 53.3). Fish serve as the reservoir host, with humans serving as the definitive host for D. latum, D. nihonkaiense, D. dendriticum, and D. pacificum. D. latum eggs are found in the feces of infected humans and other fish-eating mammals. Once passed into a water source, such as a lake, the life cycle requires two intermediate hosts. After incubation in freshwater for approximately 2 weeks, the mature eggs release the first larval stage (coracidium). The coracidium larva is ciliated and bears six terminal hooks. The coracidium larvae are ingested by copepods. The coracidium larva sheds its epithelium and further develops into a procercoid larva (infective form). The fish feed upon the small crustaceans ingesting the procercoid larvae. Within the freshwater fish, the larvae develop into the plerocercoid ribbonlike organism with an undivided scolex. The procercoid may pass through multiple paratenic hosts until consumed by a mammal or human. Diphyllobothrium infection occurs through the ingestion of infected fish containing the plerocercoid larval form. Diphyllobothrium latum and other species mature to an adult tapeworm within the human small intestine. Infection is usually asymptomatic, but mild gastrointestinal symptoms may occur, such as diarrhea, abdominal pain, fatigue, vomiting, or dizziness. Symptoms vary depending on the worm burden and the host’s immune response to the organism. The tapeworm nutritional requirements may decrease the host’s vitamin B12 level, resulting in pernicious anemia. 

Laboratory Diagnosis

C • Fig. 53.2 (A) Diphyllobothrium latum scolex. (B) D. latum scolex, bothria visible. (C) D. latum ovum. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

Both eggs and proglottids may be found in the patient’s feces. Visualization of the eggs is enhanced using a wet preparation of the patient’s stool sample. Diagnosis is made by identification of the ovoid, operculated, yellow-brown eggs (58 to 75 μm × 40 to 50 μm) passed in abundance in the stool. They are sometimes confused with the eggs of Paragonimus. The mature gravid proglottids are wider than long (3 mm × 11 mm), often in chains, and contain a rosette-shaped central uterus (Fig. 53.1). Diphyllobothrium identification is through assessment of the morphologic characteristics of

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•

Fig. 53.3 Life cycle of Diphyllobothrium latum. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

the proglottids as previously described. Speciation within the genus Diphyllobothrium is difficult to impossible using morphological characteristics. Nucleic acid amplification and sequencing is the only reliable tool for speciation. Because all of the species cause similar infections and the therapy is similar, speciation is not necessary for treatment but remains important for epidemiological reasons. Molecular approaches to differential identification of D. latum and D. nihonkaiense have been successful using restriction fragment length polymorphisms (RFLP) utilizing the ribosomal DNA sequence of the mitochondrial cytochrome c oxidase I (cox1) gene. No serologic tests are available. 

Therapy Humans infected with Diphyllobothrium develop little to no protective immunity. Reinfection is common. Treatment with praziquantel or niclosamide is effective and nontoxic. Subsequent stool specimens should be reexamined 6 weeks after treatment. The patient may require a vitamin B12 supplement if anemia develops. 

Prevention Prevention simply includes avoiding the consumption of raw fish. The larval stage is destroyed when food is thoroughly cooked at 55°C for 5 minutes or frozen at −20°C for 7 days or flash freezing to −35°C for 15 hours, if the flesh is less than 15 cm thick. Treatment of patients infected with adult tapeworms is indicated to prevent accidental autoinfection. Good hygiene and proper sanitation measures will also prevent reinfection. Treatment of sewage before it enters lakes may help reduce the prevalence of infection. 

Dipylidium caninum General Characteristics D. caninum, the cat or dog tapeworm (Fig. 53.4), is a double-pored (genital pores) tapeworm consisting of many small proglottids. As the tapeworm matures, the proglottids separate and pass in the stool. They may be recognized based on their characteristic “cucumber seed” appearance when

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they are wet, as well as their resemblance to a dried grain of rice. Adult tapeworms measure 10 to 70 cm in length. The scolex contains four suckers and an armed rostellum. Egg packets may also be found in the feces of the host. 

Epidemiology Infection is common worldwide. In the case of D. caninum, human infection is acquired through the accidental

ingestion of fleas. Infection is most often seen in young children because of close contact with infected pets. The tapeworms are found in both wild and domestic dogs and cats. 

Pathogenesis and Spectrum of Disease Ingestion of an infected flea may result in D. caninum infection (Fig. 53.5). The flea is the intermediate host in which infective cysticercoids (larval form) develop; humans, dogs and cats are the reservoir hosts. The cysticercoid larval stage is ingested by a dog or cat and develops into cysticercoid metacestode larvae. The adult worm develops and matures within the reservoir host. An infected human host will usually pass proglottids in a bowel movement, or they may stick to the skin around the anal area. This may result in misdiagnosis of the infection as Enterobius vermicularis. Humans usually have very mild symptoms, such as indigestion, appetite loss, weight loss, perianal itching, persistent diarrhea, and vague abdominal pain. The severity of the disease is dependent on the worm burden. Human infection is usually self-limited. 

Laboratory Diagnosis • Fig. 53.4  Dipylidium caninum tapeworm. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

Symptoms of Dipylidium infection are similar to those of pinworm infection; however, the treatments are very

• Fig. 53.5  Life cycle of Dipylidium caninum. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

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different. The laboratory should confirm suspected infections. Proglottids (8 to 23 μm) may be seen in the stool. D. caninum is also referred to as the “cucumber seed” tapeworm, as previously described (Fig. 53.6). The first sign of infection may be the appearance of seedlike particles in the stool or undergarments of the patient. These particles are the egg-bearing segments of the tapeworm. Groups of egg packets may be found in the stool (Figs. 53.7E). The adult worms have a scolex with four suckers and a conical/ retractile rostellum armed with four to seven rows of small hooklets (Fig. 53.8). Patients may also develop a moderately elevated eosinophilia. Serologic tests are typically performed because of the asymptomatic and self-limiting nature of the infection. Nucleic acid–based testing, testing using RFLP analysis, and hydrolysis probe-based genotyping assays have been developed and validated for genotyping D. caninum; currently, however, they are not utilized for clinical diagnostics. 

Therapy Praziquantel and niclosamide are typically effective for treating D. caninum infection. The medication causes the tapeworm to dissolve within the intestine. The drugs are generally well tolerated by the patient. Household pets should be treated simultaneously to prevent reinfection. 

Prevention To reduce the risk of infection, flea control of pets in the household will reduce exposure to humans via the intermediate host. Keeping cats indoors will help prevent infection by limiting their exposure to fleas by household cats. 

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Hymenolepis nana General Characteristics Hymenolepis nana, also known as the dwarf tapeworm, is very small compared with other tapeworms. The organism has a worldwide distribution and may reach up to 4 cm in length. The proglottid contains a scolex with a short-armed rostellum. It is the most common tapeworm. An intermediate host is not required, thus making person-to-person spread possible. An adult dwarf tapeworm can live within the host for approximately 4 to 6 weeks. 

Epidemiology H. nana is generally found in children. Although it is most prevalent in the southern United States, it has a wide distribution—particularly in crowded areas. It is more common in populations living in conditions of poverty or poor hygiene, in daycare centers, and in persons living in institutional settings or prisons. 

Pathogenesis and Spectrum of Disease H. nana has an unusual life cycle; ingestion of the egg can lead to the development of the adult worm in humans, thus bypassing the need for an intermediate host. Humans can serve as both intermediate and definitive hosts. Infection occurs by accidentally ingesting dwarf tapeworm eggs. This happens most commonly through direct fecal-oral transmission or accidental ingestion of an infected arthropod. The worm resides within the upper ileum of the intestinal tract. Once infection occurs, the dwarf tapeworm may reproduce inside the body, thus causing autoinfection. Autoinfection is essentially a reinfection or constant reproduction of the parasite within the host. Massive infection with several thousand worms may follow autoinfection, resulting in hyperinfection. Hyperinfection refers to a large parasitic burden within the host. Autoinfection appears to initiate a cellular and humoral immune response. The immune response will provide the host with some protective immunity. Most patients are asymptomatic. Symptomatic patients may experience weight loss, nausea, and weakness, loss of appetite, diarrhea, and abdominal discomfort. Young children, especially those with a heavy infection, may develop headaches, an itchy perianal area, or have difficulty sleeping. 

Laboratory Diagnosis • Fig. 53.6  This photomicrograph reveals ultrastructural details exhib-

ited by a single Dipylidium caninum reproductive proglottid. These D. caninum proglottids, which when mature average 12 mm × 3 mm, are pumpkin seed-shaped, are passed with the animal’s feces, and often resemble rice grains when dried. Each proglottid contains egg packets that contain 8 to 15 ova, which are held together by an outer embryonic membrane. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

Adult worms and proglottids are rarely seen in stool specimens. Diagnosis is typically through the identification of eggs in stool specimens. Eggs are characterized by the presence of a thin shell enclosing an embryo (oncosphere) with six hooklets contained within two layers of membrane that is separated from the outer shell. The eggs are spheroidal, pale, and thin-shelled (30 to 47 μm in diameter). The eggs of H. nana and Hymenolepis diminuta are very similar. However,

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A

C

B

D

E • Fig. 53.7 (A) Taenia spp. egg iodine preparation (400× total magnification). (B) Diphyllobothrium latum egg iodine preparation (400× total magnification). (C) Hymenolepis diminuta egg. (D) Hymenolepis nana egg. (E) Dipylidium caninum egg packet iodine preparation (500× total magnification).

H. nana eggs are smaller and have 4 to 8 polar filaments present in the space between the oncosphere and the eggshell (Fig. 53.7). The egg morphology is easily distinguishable in fresh or formalin-fixed fecal samples. It is important to note that eggs are infectious, and therefore unpreserved specimens should be handled carefully. Concentration techniques and repeated examinations will increase the likelihood of detecting light infections. Some patients may demonstrate a low-grade eosinophilia. Nucleic acid–based methods and serological techniques are being developed; however, further evaluation is needed to determine the efficacy and use of these assays in clinical applications. 

Therapy Praziquantel remains the therapy of choice. Niclosamide is also effective and can be repeated with reinfection. Human adults living in endemic areas are provided some immunity as a result of their cellular and humoral immunologic responses. 

Prevention Good hygiene is the best method for control and prevention. Preventing fecal contamination of food and water is the first line of defense. General sanitation measures, along

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oncosphere and the eggshell (Fig. 53.7). The eggs are clearly differentiated from H. nana because of the absence of polar filaments. Nucleic acid–based methods and serological techniques are being developed; however, further evaluation is needed to determine the efficacy and use of these assays in clinical applications. 

Therapy H. diminuta is readily treated with praziquantel, although the disease is self-limiting and treatment is often not necessary. 

• Fig. 53.8  Dipylidium caninum scolex demonstrating the armed rostellum. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

with rodent control, are helpful in controlling the flea population. 

Prevention Prevention is dependent on controlling the populations of infected mice and rats, along with good hygiene and sanitation. 

Taenia solium

Hymenolepis diminuta

General Characteristics

General Characteristics

H. diminuta is an uncommon tapeworm in humans and is typically found in rodents, including rats and mice. Rats are typically the natural reservoir. H. diminuta can infect humans after contamination of grains and flours with rodent feces. 

T. solium, the pork tapeworm, is the intestinal cestode capable of causing serious pathologic damage to the human host. Humans serve as the definitive host, whereas pigs serve as the intermediate host. Humans can also serve as the intermediate host. T. solium may result in an intestinal infection in which the larvae mature and reside in the small intestine for up to 25 years. The organisms can grow to be 2 to 7 m long and produce more than 1000 proglottids, each containing about 50,000 eggs. Cysticercosis (larval forms throughout the body) is the extraintestinal form of the disease and can be much more severe. The disease is life threatening if the organism invades the central nervous system causing neurocysticercosis. 

Pathogenesis and Spectrum of Disease

Epidemiology

The life cycle of H. diminuta involves insects, similar to the life cycle of H. nana. H. diminuta rarely infects humans but may do so if a human accidentally ingests an arthropod infected with cysticercoids. Multiple adult worms may mature in the human intestine. Infections are usually well tolerated by the host because of the small size of the organism. Symptoms may include diarrhea, anorexia, nausea, headache, and dizziness. The infection is more common in children, causing mild diarrhea, remittent fever, and abdominal pain. 

T. solium has a worldwide distribution. Higher rates of illness have been seen in Latin America, Asia, sub-Saharan Africa, and parts of Oceania. The parasite is found in the United States, typically among immigrants from zones of endemicity. The tapeworm is more prevalent in underdeveloped communities with poor sanitation, and when pork is ingested as undercooked or raw. 

H. diminuta, the rat tapeworm, is larger than H. nana and can measure 20 to 60 cm in length. Outbreaks of human infection are rare. 

Epidemiology

Laboratory Diagnosis Proglottids are rarely seen in the stool; diagnosis is made by the identification of eggs. The eggs (70 to 85 μm × 60 to 80 μm) are large, ovoid, yellowish, and moderately thickshelled. The eggs contain a six-hooked oncosphere with the absence of polar filaments in the space between the

Pathogenesis and Spectrum of Disease T. solium infection can result in the presence of both adult and larval stages in the human host (Fig. 53.9). Infection begins when the intermediate host ingests embryonated eggs in feces. Once the egg is ingested, the hexacanth embryo, armed with three pairs of hooks, is released into the intestine where the embryo penetrates the mucosa. The embryo then matures into a cysticercus larvae (containing a fluidfilled bladder) in the tissue. Humans may become infected

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• Fig. 53.9  Life cycle of Taenia spp. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

when they eat raw or undercooked pork containing embedded larvae with cysts. Pork tapeworm infection is usually caused by ingestion of multiple worms. During ingestion and subsequent digestion of the infected meat, the cysticercus is released and attaches to the mucosa within the small intestine of the human host. The cysticercus matures into an adult worm within approximately 5 to 12 weeks. The eggs are then released in the host’s feces. Accidental ingestion of the eggs by the human host may also result in migration of the embryo through the intestine to other areas of the body, including the eyes, brain, muscle, or bone. In addition, the proglottids are motile and may migrate out of the anus. Infection of the adult tapeworm causes few clinical symptoms, although abdominal pain, diarrhea, indigestion, and loss of appetite may be present because of irritation to the mucosa of the intestinal wall. The major complication with T. solium is cysticercosis, in which the human host becomes the intermediate host and harbors the larvae in tissues as

previously described. This infection is further discussed in Chapter 74. 

Laboratory Diagnosis Diagnosis of Taenia tapeworm infection is through the examination of stool samples. Individuals suspected of infection with T. solium should be asked if they have passed any notable tapeworm segments in their stool. Stool specimens should be collected on 3 different days and microscopically examined for the presence of Taenia eggs (Fig. 53.7A). Tapeworm eggs can be detected in the stool 2 to 3 months after the tapeworm infection is established. Eggs are round or slightly oval (31 to 43 μm in diameter) and yellowbrown with a thick, striated shell containing a six-hooked oncosphere. Diagnosis is based on the recovery of eggs or proglottids in stool or from the perianal area. T. solium, T. saginata, and T. asiatica cannot be differentiated based on

CHAPTER 53  Intestinal Cestodes

• Fig. 53.10  This photomicrograph reveals some of the ultrastructural

morphology exhibited by a gravid proglottid from a Taenia solium tapeworm. The number of primary lateral uterine branches, represented by the dark, India ink-stained irregularities, allows differentiation between the two species. T. solium shows 7 to 13 branches on each side. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

• Fig. 53.11  This photomicrograph reveals some of the ultrastructural

morphology exhibited by a gravid proglottid from a Taenia saginata tapeworm. The number of primary lateral uterine branches, represented by the dark, India ink-stained irregularities, allows differentiation between the two species. T. saginata shows 15 to 20 branches on each side. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

egg morphology. Speciation requires the examination of gravid proglottids or the scolices. T. solium gravid proglottids are longer than wide (19 mm × 17 mm) and may be distinguished from T. saginata and T. asiatica according to the number of uterine branches. T. solium contains 7 to 13 lateral uterine branches along the proglottid (Fig. 53.10), whereas T. saginata and T. asiatica contain more than 13 branches. Uterine branches may be visualized by staining the proglottids with India ink (Fig. 53.11). The scolex contains a neck region that is typically short and half the width of the scolex, and differs from that of T. saginata by the presence of four suckers with hooks in a double row (Fig. 53.12). The adult worm is usually 3 to 5 m long. Extreme care should be taken when handling infectious stool, because T. solium proglottids and eggs are extremely infectious. Additional

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• Fig. 53.12  This photomicrograph reveals some of the ultrastructural

morphology exhibited by the cephalic end of a Taenia solium, highlighting the worm’s scolex, and its four suckers and two rows of hooks. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

laboratory findings may include a low-grade eosinophilia, increased serum IgE level, and the presence of atypical lymphocytes in the cerebrospinal fluid. Serologic assays specific to the adult T. solium tapeworm have been developed for the diagnosis of neurocysticercosis. Complement fixation, hemagglutination, enzyme-linked immunosorbent assay (ELISA), and immunoblot can be used for the detection of anticysticercal antibodies in serum, cerebrospinal fluid, and saliva. The enzyme-linked immunoelectrotransfer blot assay is the most effective method for the detection of antibodies, with sensitivity and specificity as high as 100% and 98%, respectively. However, the sensitivity decreases to about 70% in patients with a single cyst or in those with only calcified lesions. ELISA is more reliable when performed in cerebrospinal fluid than in serum, but the accuracy depends on the viability and location of the cysticerci. These advanced serologic tests are not yet commercially available and are generally only available in specialized laboratories. Stool antigen testing detects at least 2 to 3 times more cases of Taenia infection than stool microscopy. 

Therapy Adult worms can be eradicated with praziquantel or niclosamide followed by use of a laxative. Expulsion of the scolex must be verified to assume satisfactory treatment. 

Prevention Good hygiene and immediate treatment are essential for the prevention of autoinfection. Pork should be cooked or frozen thoroughly. Cysticerci do not survive temperatures below −10°C or above 50°C. Educational programs concerning the hazards associated with living near human sewage and contaminated drinking water are important in populations at risk. When traveling in countries where food is likely to be contaminated, all raw vegetables and fruits should be washed, peeled, or cooked with clean water before eaten. 

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Taenia saginata General Characteristics T. saginata, or beef tapeworm, has a worldwide distribution and is more common than T. solium. The worm can grow 4 to 12 m and contain 1000 to 2000 segments. T. saginata may produce 100,000 eggs and live up to 25 years in the human intestine. 

Epidemiology T. saginata has a similar life cycle to that of T. solium. Cattle are the intermediate hosts, and humans are infected through the ingestion of cysticerci (larval form with an unarmed scolex) in raw or undercooked beef (Fig. 53.9). 

Pathogenesis and Spectrum of Disease The life cycle of T. saginata begins with human ingestion of undercooked or raw meat infected with larvae. The larvae are ingested in the meat and, after digestion, released into the small intestine, where the worm attaches to the mucosa and matures. In about 3 months, the worm may grow up to 4 to 5 m in length, and gravid segments will begin to break off and pass in stool. After deposition of gravid segments in the soil, an intermediate bovine host may ingest the segments. The segments are digested and the eggs hatch, releasing an oncosphere that penetrates the muscle tissue. After penetration of the mucosa, the organisms are carried via the lymphatic vessels and bloodstream throughout the intermediate host. Humans then ingest the infected meat of the intermediate host, as previously indicated. Humans typically are asymptomatic or experience very mild indigestion, loss of appetite, vomiting, and abdominal discomfort. A rare case of severe infection may result in intestinal obstruction and appendicitis. Patients are often unaware of their infection until gravid motile segments are passed in the feces and cause psychological distress. 

Laboratory Diagnosis The stool should be examined for proglottids and eggs; eggs may also be present on anal swabs. The eggs of T. saginata are indistinguishable from other Taenia spp. The adult worms can reach up to 25 m in length. The uterus of T. saginata is longer than wide and typically contains 15 to 18 lateral branches on each side (Fig. 53.11). The scolex has four suckers and is unarmed or does not contain any hooklets (Fig. 53.13). Stool specimens should be handled with care, because the eggs cannot be distinguished from other Taenia spp. Slight eosinophilia may develop. 

Therapy Recommended treatment includes praziquantel or niclosamide. Treatment of T. saginata can be considered

•

Fig. 53.13 Under a low magnification, this photomicrograph highlights the scolex, or head region of the cestode, Taenia saginata. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

successful when no proglottids are passed for 4 consecutive months. 

Prevention Beef should be inspected for cysticerci and thoroughly cooked before it is ingested. 

Taenia asiatica General Characteristics T. asiatica, or Asian tapeworm, is found primarily in remote areas of the East and Southeast Asian countries to include Taiwan, Indonesia, Korea, Vietnam, the Philippines, Thailand, China, and Japan. The morphology of T. asiatica’s gravid proglottids and adult worm is difficult to distinguish from that of T. saginata. 

Epidemiology T. asiatica has a similar life cycle to that of T. solium. It has been suggested that T. asiatica is possibly misdiagnosed as T. solium since both species share some of the same hosts, and co-infections are thought to occur more frequently than reported. Pigs, cattle, and goats are the intermediate hosts, and humans are infected through the ingestion of cysticerci (larval form with an unarmed scolex) in raw or undercooked liver from cattle or pork. 

Pathogenesis and Spectrum of Disease The pathogenesis of T. asiatica is identical to that of T. saginata; which begins with human ingestion of undercooked or raw liver infected with larvae (Fig. 53.9). After penetration of the mucosa, the organisms are carried via the lymphatic vessels and bloodstream throughout the intermediate host. Humans then ingest the infected liver of the

CHAPTER 53  Intestinal Cestodes

intermediate host, as previously indicated. Humans typically are asymptomatic or may experience abdominal pain, nausea, weakness, weight loss, headache, and changes in appetite. Patients are often unaware of their infection until gravid motile segments are passed in the feces. Eosinophilia is seen in some patients. 

Laboratory Diagnosis The stool should be examined for proglottids and eggs; eggs may also be present on anal swabs. The eggs of T. asiatica are indistinguishable from those of T. solium or T. saginata. The uterus of T. asiatica is longer than wide and typically contains 12 to 26 lateral branches on each side. The scolex has two rows of rudimentary hooklets that are lost in the mature worm. The adult worm is smaller than T. saginata, measuring 4 to 8 m long, and contains fewer proglottids. Stool specimens should be handled with care, because the eggs cannot be distinguished from those of T. solium or T. saginata. 

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Pathogenesis and Spectrum of Disease T. crassiceps rarely infects humans, as a healthy adult human’s immune system typically removes the parasite before permanent damage occurs. However, individuals who were infected tended to be immunosuppressed patients. Patients are generally asymptomatic but may present with symptoms, such as headache, nausea, and vomiting. Parasites may accumulate in skeletal muscle and subcutaneous tissue, and some patients present with intraocular infections referred to as ocular larva migrans that can cause serious damage to the eyes, even blindness. 

Laboratory Diagnosis Diagnosis is made by the observation of cysticerci in biopsy or autopsy specimens. Positive ELISA for anticysticercal antibodies helps confirm the diagnosis; however, negative test results do not exclude cysticercosis. The patient may present with eosinophilia. 

Therapy

Therapy

Recommended treatment includes praziquantel or niclosamide, followed by a laxative. Treatment of T. asiatica is not usually required; however, if prescribed, it can be considered successful when no proglottids are passed for 4 consecutive months. 

Surgical removal is mandatory for individuals with intraocular cysts. Ocular cysticercosis can be effectively treated with anthelmintics, such as albendazole or praziquantel, and oral corticosteroids for inflammation. 

Prevention Pork and beef liver should be inspected for cysticerci and thoroughly cooked before ingested. 

Taenia crassiceps General Characteristics Taenia crassiceps is a parasitic organism whose adult form infects the intestines of carnivores and has been isolated in human cases of cysticercosis. 

Epidemiology It is commonly found in the Northern Hemisphere, especially throughout Canada and the northern United States. The larval stages of T. crassiceps develop subcutaneously or in their body cavities as cysticerci. T. crassiceps inhabits the intestines of carnivores. Inside the carnivore, the tapeworm reproduces. The eggs are passed in the feces, with the natural intermediate hosts of this organism usually being small rodents and moles. When the intermediate host is eaten by another carnivore, the parasite’s life cycle repeats. Humans serve as intermediate hosts when food or water contaminated with feces from infected hosts is consumed. Close contact with infected domestic dogs has been associated with several human infections. 

Prevention Avoid food and water sources that could be contaminated with feces. Animal products consumed should be inspected for cysticerci and thoroughly cooked before ingested. 

Nucleic Acid Detection (All Species) All three species of Taenia, and two genotypes of T. solium, can be differentiated using nucleic acid base excision sequence scanning thymine-base reader analysis for mitochondrial genes. Polymerase chain (PCR) amplification of the cytochrome oxidase subunit 1 (cox1) has been successfully used to identify several Taenia solium infections from fixed tissue associated with cases of human cysticercosis. Additional nucleotide sequences used to differentiate the organisms from clinical material include the mitochondrial 12S rRNA, ND1 (NADH dehydrogenase 1), and ITS2 (ribosomal internal transcribed spacer 2). Two additional species were also identified using PCR: T. serialis and T. crassiceps. T. serialis (Chapter 54) was identified in a cystic parasitic larva sample from a subcutaneous sample of a patient’s jaw. T. crassiceps was identified in sternocleidomastoid muscle of the patient’s neck. There have been 12 reported cases of human infection with this species. Four cases of T. martis cysticercosis have been identified in human infections.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

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Bibliography Beugnet F, Labuschagne M, de Vos C, Crafford D, Fourie J: Analysis of Dipylidium caninum tapeworms from dogs and cats, or their respective fleas - part 2. Distinct canine and feline host association with two different Dipylidium caninum genotypes, Parasite 25:31, 2018. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Conlan JV, Vongxay K, Fenwick S, Blacksell SD, Andrew Thompson RC: Does interspecific competition have a moderating effect on Taenia solium transmission dynamics in Southeast Asia? Trends Parasitol 25:398–403, 2009. Deplazes P, Eichenberger RM, Grimm F: Wildlife-transmitted Taenia and Versteria cysticercosis and coenurosis in humans and other primates, Int J Parasitol Parasites Wildl 9:342–358, 2019. Flisser A: State of the art of Taenia solium as compared to Taenia asiatica, Korean J Parasitol 51:43–49, 2013, https://doi.org/10.3347/ kjp.2013.51.1.43. François A, Favennec L, Cambon-Michot C, et al.: Taenia crassiceps invasive cysticercosis: a new human pathogen in acquired immunodeficiency syndrome? Am J Surg Pathol 22(4):488–492, 1998. Galán-Puchades MT, Fuentes MV: Taenia asiatica: the most neglected human taenia and the possibility of cysticercosis, Korean J Parasitol 51(1):51–54, 2013. Galindo M, Gonzalez MJ, Galanti N: Echinococcus granulosus protoscolex formation in natural infections, Biol Res 35(3–4):365–371, 2002. Garcia HH, Del Brutto OH, Nash TE, et al.: New concepts in the diagnosis and management of neurocysticercosis (Taenia solium), Am J Trop Med Hyg 72(1):3–9, 2005. Garcia LS: Diagnostic medical parasitology, ed 6, Washington, DC, 2016, ASM Press. Heldwein K, Biedermann HG, Hamperl WD, et al.: Subcutaneous Taenia crassiceps infection in a patient with non-Hodgkin’s lymphoma, Am J Trop Med Hyg 75(1):108–111, 2006. Hoberg EP: Taenia tapeworms: their biology, evolution and socioeconomic significance, Microbes Infect 4(8):859–866, 2002. Hyneman D: Cestodes. In Baron S, eds: Medical microbiology, ed 4, Galveston, TX, 1996, Addison-Wesley. Jeon HK, Eom KS: Molecular approaches to Taenia asiatica, Korean J Parasitol 51(1):1–8, 2013. Jeon HK, Kim KH, Huh S, et al.: Morphologic and genetic identification of Diphyllobothrium nihonkaiense in Korea, Korean J Parasitol 47(4):369–375, 2009.

Mayta H, Talley A, Gilman RH: Differentiating Taenia solium and Taenia saginata infections by simple hematoxylin-eosin staining and PCRrestriction enzyme analysis, J Clin Microbiol 38:133–137, 2000. Metwally DM, Al-Enezy HA, Al-Turaiki IM, El-Khadragy MF, Yehia HM, Al-Otaibi TT: Gene-based molecular characterization of cox1 and pnad5 in Hymenolepis nana isolated from naturally infected mice and rats in Saudi Arabia, Biosci Rep 39(2), 2019, BSR20181224. Patamia I, Cappello E, Castellano-Chiodo D, Greco F, Nigro L, Cacopardo B: A human case of Hymenolepis diminuta in a child from eastern Sicily, Korean J Parasitol 48:167–169, 2010. Roberts L, Schmidt G: Foundations of parasitology, ed 8, New York, 2009, McGraw-Hill, pp. 351–352. Samkari A, Kiska DL, Riddell SW, Wilson K, Weiner LB, Domachowske JB: Dipylidium caninum mimicking recurrent Enterobius vermicularis (pinworm) infection, Clin Pediatr (Phila). 47:397–399, 2008. Schenone H: Praziquantel in the treatment of Hymenolepis nana infections in children, Am J Trop Med Hyg 29:320, 1980. Schmid S, Grmm F, Huber M, et  al.: JPLL investigator catalog, Cytopathology 25(5):340–341, 2013. Scholz T, Garcia H, Kuchta R, Wicht B: Update on the human broad tapeworm (genus Diphyllobothrium), including clinical relevance, Clin Microbiol Rev 22:146–160, 2009. Sorvillo F, Wilkins P, Shafir S, Eberhard M: Public health implications of cysticercosis acquired in the United States, Emerg Infect Dis 17:1–6, 2011. Sundaram PM, Jayakumar N, Noronha V: Extraocular muscle cysticercosis - a clinical challenge to the ophthalmologists, Orbit 23(4):255–262, 2004. Tappe D, Berkholz J, Mahlke U, et  al.: Molecular identification of zoonotic tissue-invasive tapeworm larvae other than Taenia solium in suspected human cysticercosis cases, J Clin Microbiol 54:172– 174, 2016. Tena D, Simón MP, Gimeno C, et  al.: Human infection with Hymenolepis diminuta: case report from Spain, J Clin Microbiol 36(8):2375–2376, 1998. Turner JA: Human dipylidiasis (dog tapeworm infection) in the United States, J Pediatr 61:763–768, 1962. Wicht B, Yanagida T, Ito A, et  al.: Multiplex PCR for differentiated identification of broadworm tapeworms (cestode: Diphyllobothrium) infecting humans, J Clin Microbiol 48:311, 2010. Wiwanitkit V: Overview of Hymenolepis diminuta infection among Thai patients, MedGenMed 22(6):7, 2004.

CASE STUDY 53.1 A 50-year-old male presented to the physician complaining of headaches and difficulty maintaining his balance. On physical examination, the physician noticed a lump in the man’s left calf. Initial laboratory results, including a complete blood count and differential, were normal. The erythrocyte sedimentation rate (ESR) was slightly elevated, indicating generalized inflammation. The physician asked the man if he had traveled to any areas outside of the United States in the past 12 months. The man had recently returned from volunteering in Haiti. The physician prescribed niclosamide and had the patient collect three stool

samples over the next 12 days. After treatment, the parasite seen in Figs. 53.10 and 53.12 was recovered from the man’s stool.

Questions 1. Identify the parasite to the species level. 2. Explain the morphologic characteristics that would definitively identify the parasite. 3. What, if any, additional treatments are available for this patient?   

Chapter Review 1. Which tapeworm infects cattle as an intermediate host? a. T. saginata b. D. caninum c. H. diminuta d. H. nana 2. Which of the following can bypass the need for an intermediate host? a. D. latum b. D. caninum c. H. diminuta d. H. nana 3. Which eggs are passed unembryonated and may have a small knob at the end of the operculum? a. D. latum b. D. caninum c. H. diminuta d. H. nana 4. Which tapeworm cannot be identified to the species level based on its egg morphology; instead, proglottids must be examined? a. Diphyllobothrium b. Dipylidium c. Hymenolepis d. Taenia 5. Which of the following are treatment(s) of choice for tapeworm infection? a. Praziquantel b. Niclosamide c. Both A and B d. None of the above

6. Which of the following is not an appropriate prevention measure for cestodes? a. Controlling the flea and rat population b. Avoiding the consumption of raw meat or fish c. Immunization d. Practicing good hygiene 7. True or False _____  Diphyllobothrium latum is the only cestode to have an aquatic life cycle. _____  Hymenolepis diminuta, also known as the dwarf tapeworm, may reach up to 4 cm in length. _____ Food is absorbed from the host through the worm’s scolex. _____ Humans are infected with T. saginata by eating cysticerci in raw or undercooked beef. _____ All Taenia species are indistinguishable morphologically requiring detailed genotypic analysis 8. Matching: Match each term with the appropriate description. _____ cestode _____ scolex _____ proglottids _____ rostellum _____ integument _____ hexacanth embryo _____ bothria

a. oncosphere b. crown of scolex c. head d. shallow sucking grooves e. tapeworm f. outer covering g. segments

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Tissue Cestodes OBJECTIVES 1. Describe and compare the life cycles of the tissue cestodes, including reservoir and intermediate hosts. 2. Describe the clinical manifestations and complications of cysticerci in the human host. 3. List the various methods used to diagnose cysticercosis infection. 4. Define and describe the morphologic characteristics of the following: oncosphere, brood capsule, hydatid cyst, and hydatid sand. 5. Describe hydatid disease, including laboratory diagnosis and the best course of treatment. 6. Compare and contrast the pathogenesis and spectrum of disease associated with direct tissue damage versus the immune response to Echinococcus spp. 7. Describe the tapeworm that causes coenurosis, including hosts and symptoms of disease. 8. Describe the preventive measures recommended to prevent infection with tissue cestodes.

PARASITES TO BE CONSIDERED Cestodes (Tapeworms) Tissue (Larval Forms)

Taenia solium Echinococcus granulosus complex E. granulosus sensu stricto E. equinus E. ortleppi E. canadensis Echinococcus multilocularis Echinococcus oligarthrus Echinococcus vogeli Taenia multiceps Taenia serialis Spirometra mansonoides

T

issue cestodes do not reach the adult stage in the human host. The organisms infect the human in their intermediate or cyst stage. These infections are much more serious than those caused by adult tapeworms. The parasites can cause serious disease or even death. Larval cestodes cause infection after accidental ingestion of eggs excreted from the intermediate host (Table 54.1) and release embryos that migrate and lodge in various organs and tissues in the human body. Diagnosis of larval infections can be problematic.

Taenia solium General Characteristics Taenia solium, also known as the pork tapeworm, causes an intestinal infection contracted when a person eats contaminated pork, as discussed in Chapter 53. The adult worm usually causes no clinical disease. Humans may accidentally become the intermediate host by ingesting eggs from human feces. This typically occurs when an individual is already infected with adult T. solium. Autoinfection occurs when the individual swallows eggs after improper hand washing. Humans may develop the larval infection, which could result in cysticercosis. Cysticercosis is usually asymptomatic unless larvae invade the central nervous system (CNS), the globe of the eye, or other muscle and tissues. 

Epidemiology T. solium is found worldwide, with a higher incidence in Latin America. The larval form of the infection rarely occurs in the United States but may be found among immigrants from Mexico. After ingestion of T. solium eggs, the oncospheres hatch in the intestine and invade the intestinal wall. Once the larvae invade the tissue, the organism is capable of spreading systemically by migration to the brain, liver, and other tissues, causing human cysticercosis. Cysticercosis is defined as larval forms distributed throughout the body. Human cysticercosis may also occur when reverse peristalsis returns gravid segments into the intestine, where the eggs hatch and release oncospheres. Cysticerci develop and may live many years. Cysticerci will eventually die and may calcify, which will aid in diagnosis. 

Pathogenesis and Spectrum of Disease Clinical signs and symptoms depend on the location, viability, and number of the cysticerci present. Cysticerci can develop in any organ or tissue of the body. The severity of the symptoms depends on the body site involved; symptoms may not appear for years after the initial infection. The most severe cases are found in the CNS and the eye. Once cysticerci localize in the brain, the organism causes a condition referred to as neurocysticercosis. Infection can cause epileptic-type seizures, headaches, mental disturbances, meningitis, or sudden death. Cysticerci can also be 765

Organism

Acquired Infection Location in Host

Tissue cestodes

Source

Echinococcus granulosus complex

Eggs from dog tapeworm

Liver, lung, etc.

E. oligarthrus

Eggs from a wild dog tapeworm

E. vogeli

Diagnostic Specimen

Diagnostic Test

Positive Specimen

Comments

Serum, hydatid cyst aspirate; biopsy

Serology, centrifugation of fluid; histology

Positive serology; hydatid sand, tapeworm tissue

E. granulosus (enclosed cyst)

Definitive: Intestine Intermediate: cysts within various tissues and organs

Serum, scans, biopsy,

Serology, microscopic and confirmation with ELISA and PCR

Tapeworm tissue, positive serology

The cysts resemble those of E. vogeli. Exogenous proliferation has not been reported. Unicystic single or multiple metacestodes

Eggs from a bush dog tapeworm

Definitive: Intestine Intermediate: cysts found primarily in the liver and lungs

Serum, scans, biopsy

Serology, microscopic and confirmation with ELISA and PCR

Tapeworm tissue, positive serology

These cysts are often interconnected and can have multiple chambers, resulting in multichambered cysts as well as endogenous daughter cysts. These proliferating cysts are similar to those of E. multilocularis

Echinococcus multilocularis

Eggs from fox tapeworm

CNS, subcutaneous tissues

Serum, scans, biopsy

Serology, films, histology

E. multilocularis (cyst develops throughout tissue)

Taenia solium (pork)

Eggs from human tapeworm

Positive serology, positive scans, tapeworm tissue

Taenia multiceps

Eggs from a dog tapeworm

Definitive: Intestine Intermediate: CNS, subcutaneous tissues

Scans, Coenurosis

Biopsy of coenurosis in subcutaneous tissue Serology, confirmation with ELISA and PCR

Positive scans, presence of oncosphere induced coenurosis

Coenurosis most commonly affects the brain, eyes, and subcutaneous tissues

Taenia serialis

Eggs from a dog tapeworm

Definitive: Intestine Intermediate: Various subcutaneous tissues

Scans, coenurosis

Biopsy of coenurosis in subcutaneous tissue

Positive scans, ­presence of multiple protoscoleces within the coenurosis

Usually identified postmortem

Spirometra mansonoides

Eggs from dog and cat tapeworm

Any organ/tissue in the body

Scans, serum, tissue biopsy

Serology, films, histology, ELISA

Positive serology, positive scans, tapeworm tissue

Adult worm has no scolex, which can help differentiate Spirometra from T. solium

CNS, Central nervous system; ELISA, enzyme-linked immunosorbent assay; PCR, polymerase chain reaction.

Small, enclosed cysticerci (cysticercosis)

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TABLE 54.1    Common Human Parasites, Diagnostic Specimens, Tests, and Positive Findings

CHAPTER 54  Tissue Cestodes

found in the eye and must be removed to prevent permanent eye damage, including blindness. Much of the damage from cysticercosis is caused by the severe inflammatory host response that occurs after the cysticerci have died. Antibodies are produced and offer the patient secondary immunity. 

Laboratory Diagnosis Cysticercosis can be difficult to diagnose. T. solium eggs are found in stools in fewer than half of the patients with cysticercosis. Demonstration of eggs or proglottids in the feces is an indication of Taenia infection but does not provide a diagnosis for cysticercosis. Definitive diagnosis usually requires the identification of cysticercus in the tissue. The organism is surgically removed and microscopically examined for the presence of suckers and hooks on the scolex. The cysticercus is round to oval, translucent, and about 5 mm or more in diameter. The organism has a scolex with four suckers and a rostellum with a circle of hooks. Fine-needle aspiration cytology may be helpful in the diagnosis and eliminates the need for surgical biopsy. Diagnosis may also be made using computed tomography (CT) scans and magnetic resonance imaging (MRI). Radiographs may also be useful in detecting calcifying cysticerci within tissue. Ocular cysticercosis may be diagnosed by visual identification of the larval worm. A highly specific and sensitive enzyme-linked immunoelectrotransfer blot assay is available from the Centers for Disease Control and Prevention (CDC). The assay has demonstrated 100% specificity and 98% sensitivity for the identification of antibodies using serum or CSF, primarily for diagnosis of neurocysticercosis. The assay uses purified antigen from T. solium containing seven different major glycoproteins. In all patients, regardless of their clinical presentation, the immunoblot assay is slightly more sensitive in serum than in CSF specimens; consequently, there is no need to obtain CSF solely for use in the immunoblot assay. Currently available antibody detection tests for cysticercosis do not distinguish between active and inactive infections and thus have not been useful in evaluating the outcomes and prognoses of treated patients. Nucleic acid–based methods and species-specific polymerase chain reaction (PCR) have been described to differentiate Taenia species in CSF, but these are not widely used for clinical laboratory diagnosis of neurocysticercosis. 

Therapy Cysticercosis should be treated with corticosteroids, anticonvulsants, and surgery if deemed appropriate. Treating nonviable cysticerci in the brain of asymptomatic patients has not been proven necessary. Not all patients respond to treatment and not all patients must be treated, because the inflammatory response as a result of the treatment may be more serious than the disease. Symptomatic neurocysticercosis should be managed using treatment that decreases the patient’s symptoms. When treatment is suggested, albendazole is the drug of choice. If praziquantel is used, it should be combined with corticosteroids to reduce the inflammatory response and should not be used for ocular or spinal

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cord infections. Surgery may be required for ocular, spinal, or brain involvement. 

Prevention Education, meat inspection, and improvement of sanitation measures are the key preventive measures. Other prevention methods are discussed in Chapter 53. 

Echinococcus granulosus Complex General Characteristics Echinococcus is the smallest of all tapeworms (3 to 9 mm long) with three to five proglottids. It contains a scolex with four suckers and a rostellum with hooks to attach to the intestinal wall. Echinococcus granulosus sensu lato can be used as a general term for all of these species and strains. The tapeworm is found in the small intestine of the definitive host, the canine. Eggs are ingested by the intermediate hosts, which include a variety of mammals including sheep (E. granulosus sensu stricto), cattle (E. ortleppi), moose (E. canadensis), horse (E. equinus), and humans. Of the s­ everal strains of E. granulosus that have been identified, the dogsheep strain is the most common. Regardless of the strain, humans are typically accidental hosts and are considered a dead end, because the life cycle of the organism is unable to continue in a human host. Oncospheres hatch in the intestine of the intermediate host and invade the circulatory system, where they develop into hydatid cysts. Disease symptoms vary with the site and size of the cyst. Echinococcosis (hydatid disease) results from the presence of one or more cysts, which can develop in any tissue. 

Epidemiology E. granulosus complex is most common in cool, damp areas where the mammalian hosts are prevalent, such as southern South America, Russia, East Africa, and the western United States. The eggs in the definitive host are passed through the feces and contaminate soil, water, or food. The eggs are able to survive freezing conditions and can remain viable within the environment for several years. Adult worms are found only in the definitive canine hosts (Fig. 54.1). 

Pathogenesis and Spectrum of Disease Hydatid disease in humans is potentially dangerous, depending on the size and location of the cyst. Some cysts may remain undetected for many years until they grow large enough to affect other organs. Many people never know they are infected. The cyst is very slow growing in humans. It is usually fluid-filled and has a germinal layer from which many thousands of scolices are budded. These are known as daughter cysts (brood capsules), which attach to the germinal layer or free-float in the cyst. The scolices in the hydatid fluid resemble grains of sand and are called hydatid sand (Figs. 54.2A and 54.3). The result is a unilocular cyst containing

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Adult worms have matured in intestine Ingested by definitive host (dog)

Eggs/proglottids in feces

Develop into infective hydatid cyst (larvae)

Echinococcus granulosus (hydatid disease) Hydatid cysts in liver, lung, brain

Ingested by intermediate host (sheep)

Carried by bloodstream to tissues (liver, lung, brain)

Eggs hatch, penetrate intestine

Eggs ingested by humans (accidental intermediate host)

• Fig. 54.1  Life cycle of Echinococcus granulosus (hydatid disease).

future adult worms. The cyst may resemble a slow-growing tumor. Infection in the liver or lungs may be asymptomatic for many years, but the pressure eventually causes noticeable symptoms. Most hydatid cysts occur within the liver. Cysts within the liver cause chronic abdominal pain and allergic reactions and may result in cholangitis (infection of the common bile duct) and cholestasis (interference with the flow of bile from the liver). Cysts that develop in the lungs may cause infections and abscesses and result in chronic cough, shortness of breath, and chest pain. During the life cycle of the cyst, there may be occasional seepage of fluid into the host tissue and circulation, causing sensitization or activation of the immune response from the presence of the parasite. The rupture and release of the fluid of a hydatid cyst may cause anaphylactic shock as a result of the primary sensitization in a previously asymptomatic individual. If a cyst bursts within the human body, many new cysts may be released that are typically eliminated via the host’s cellular immune response. Leaking fluid from a cyst may cause notable eosinophilia. 

A

Laboratory Diagnosis Clinical symptoms of a slow-growing abdominal tumor with or without eosinophilia are suggestive of infection. Human infection ranges from asymptomatic to severe, including death. Diagnosis of echinococcosis is made through the identification of cysts in the infected organ, accompanied by positive serologic tests. Immunodiagnostic tests can be very helpful in the diagnosis of echinococcal disease and should be used before invasive methods. A variety of serologic tests, including enzyme-linked immunosorbent assay (ELISA) and Western blot serology, are available at the CDC in the United States. However, false-positive reactions may occur in persons with other helminthic infections, cancer, and chronic immune disorders. Negative test results do not rule out echinococcosis because some carriers do not have detectable antibodies. The presence of detectable antibodies in a patient depends on the physical location, integrity, and vitality of the larval cyst. Cysts in the liver are more likely to elicit an antibody response than cysts in the lungs, and, regardless of localization, antibody detection tests are least sensitive in patients with intact hyaline cysts. Cysts in the lungs, brain, and spleen are associated with lowered serodiagnostic reactivity, whereas those in bone appear to regularly stimulate

B

C • Fig. 54.2 (A) Echinococcus granules. (B) Ovum. (C) Scolex. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

a detectable antibody response. Fissuration or rupture of a cyst is followed by an abrupt antibody response. A patient with senescent, calcified, or dead cysts is generally found to be seronegative. Indirect hemagglutination (IHA), indirect fluorescent antibody (IFA) tests, and enzyme immunoassays (EIAs) are sensitive tests for detecting antibodies in serum of patients with cystic disease; sensitivity rates vary from 60% to

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769

cetrimide, or 70% to 95% ethanol) if surgical removal is not feasible. In other cases, the cysts may be surgically removed after thirty minutes of instillation with one of the previously noted chemical treatments. Albendazole or albendazole plus praziquantel have been used effectively to kill the scolices within the cyst, reduce the size of the cyst, and prevent recurrence. Cystic lesions have been known to resolve in some patients without the need for therapy. 

Prevention

• Fig. 54.3  Echinococcus granulosus, hydatid sand (300×). (Inset) Two individual hooklets (1000×).

90%, depending on the characteristics of the cases. At present, the best available serologic diagnosis is obtained by using a combination of tests. EIA or IHA is used to screen all specimens; a positive reaction is confirmed by immunoblot assay or any gel diffusion assay. Although these confirmatory assays give false-positive reactions with sera of 5% to 25% of persons with neurocysticercosis, the clinical and epidemiological presentation of neurocysticercosis is rarely confused with that of cystic echinococcosis. Antibody responses are also useful in monitoring treatment in some cases. Following successful radical surgery, antibody titers decline and sometimes disappear; titers rise again if secondary cysts develop. Tests for Arc 5 or IgE antibodies appear to reflect antibody decline during the first 24 months postsurgery, whereas the IHA and other tests remain positive for at least 4 years. Consistent declines in antibody titers do not follow chemotherapy. Consequently, the usefulness of serology to monitor the course of disease is limited; imaging techniques provide a more accurate assessment of the patient’s condition. Ultrasound, MRI, and CT have improved the diagnosis and may provide visualization of the fluid-filled cysts. Calcified cysts can be visualized using conventional x-ray. Cysts from different species can also be distinguished by the morphology of the protoscolices, if they are present. Microscopic examination of the cyst fluid for the identification of the scolices can be useful in diagnosis. A 1% eosin stain may be added to the fluid to assist in the visualization and determination of whether the cyst is viable. Nonviable scolices will stain with the eosin; viable scolices will not. 

Therapy PAIR (puncture, aspiration, infection, and reinjection) is used for the inactivation of hydatid sand by injecting the cyst with a cysticidal agent (hypertonic 30% saline, 0.5%

Preventive measures include avoiding contact with infected dogs, and deworming animals regularly. Effective control includes educating the population concerning the danger and means of transmission of hydatid disease as well as maintaining good hygiene and practicing safe disposal of dog feces. Slaughtered animals must be disposed of properly to prevent dogs from being exposed to contaminated materials, thus interrupting the Echinococcus life cycle. 

Echinococcus multilocularis General Characteristics Although rarely found in the brains of humans, Echinococcus multilocularis causes alveolar hydatid disease, which is a fatal form of echinococcosis. It is the most lethal of all helminthic diseases. The cyst is extremely dangerous because it lacks a laminated membrane and develops a series of connected chambers. The chambers contain little or no fluid and rarely contain a scolex. The morphology of the cyst is very similar to that of E. granulosus, but the adult organisms are much smaller (1.2 to 3.7 mm). The cysts are very resistant to cold temperatures. 

Epidemiology E. multilocularis is found in Asia, Europe, and northern North America, including areas such as Alaska, Montana, and Minnesota. Foxes, coyotes, and dogs are the definitive host for E. multilocularis, whereas rodents are the intermediate host. The parasite is occasionally transmitted to humans through the ingestion of contaminated food or water and by handling infected animals. Fur trappers and veterinarians are at an increased risk of infection because of exposure to infected animals. The life cycle of E. multilocularis is essentially identical to that of E. granulosus. 

Pathogenesis and Spectrum of Disease Alveolar hydatid disease is a highly lethal, destructive disease. The cyst of E. multilocularis grows slowly and may take years to produce clinical symptoms. Many cysts are asymptomatic during the life of the infected individual and are sometimes found during autopsy, surgery, or imaging scans related to other clinical conditions. The severity of symptoms depends on the location of the cyst and the size, as seen with E. granulosus. Cysts form primarily in the liver and metastasize to

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the lung or brain. Cysts in the liver are not restricted with a laminated cyst wall and are capable of expansion into a multicystic structure. This multilocular (many chambers) hydatid cyst is often mistaken for a hepatic sarcoma, making diagnosis difficult. This disease is often fatal. 

Epidemiology

Laboratory Diagnosis

Pathogenesis and Spectrum of Disease

Ultrasound, CT, and MRI are used to visualize the cyst and can be supported with serologic testing. Serologic tests, such as ELISA, are sensitive and highly specific. Most patients with alveolar disease have detectable antibodies in serologic tests using heterologous E. granulosus or homologous E. multilocularis antigens. With crude Echinococcus antigens, nonspecific reactions create the same difficulties as described previously with E. granulosus complex. However, immunoaffinity-purified E. multilocularis antigens (Em2) used in EIA demonstrate positive antibody reactions in more than 95% of alveolar cases. Using a comparison of serologic reactivity to Em2 antigen with that of antigens containing components from both E. multilocularis and E. granulosus differentiates alveolar from cystic disease. Combining two purified E. multilocularis antigens (Em2 and recombinant antigen II/3-10) in a single immunoassay optimizes the sensitivity and specificity. These antigens are commercially available as an EIA kit in Europe, but not in the United States. As in cystic echinococcosis, Em2 tests are more useful for postoperative follow-up than for monitoring the effectiveness of chemotherapy. 

The definitive hosts for E. vogeli are bush dogs (Speothos venaticus) and the intermediate hosts are South American rodents, especially pacas (Cuniculus paca). Dogs, which may be given the entrails from pacas after a hunt, can also act as definitive hosts. The metacestode is found primarily in the liver of the intermediate host, but it can also occur in the lungs and other organs. In pacas, E. vogeli cysts are fluid-filled, usually 0.5 cm to 6 cm in diameter, and can occur singly or as aggregates. These cysts are often interconnected and can have multiple chambers. E. vogeli undergoes exogenous proliferation in accidental hosts such as primates, resulting in multichambered cysts as well as endogenous daughter cysts. These proliferating cysts, like those of E. multilocularis, are invasive. Exogenous proliferation does not seem to occur in the natural host. The definitive hosts for E. oligarthrus are wild felids, and the intermediate hosts are rodents. This species can mature in experimentally infected housecats. In the intermediate host, cysts develop in the muscles, subcutaneous tissues, and internal organs such as the heart and lungs. The cysts resemble those of E. vogeli and can reach up to 5 cm in diameter. Exogenous proliferation has not been reported. In human infections, unicystic single or multiple metacestodes were found behind the eye or in the heart. E. oligarthrus has not been documented in domesticated intermediate mammalian hosts. Two outbreaks caused by E. vogeli, one affecting nutrias and the other in nonhuman primates, have been reported in zoos. Orangutans and gorillas developed severe clinical signs including very pendulous abdomens. A number of animals died or had to be euthanized. In pacas, E. vogeli does not seem to be symptomatic unless the cysts become very large. 

Therapy The most common treatment is to remove the parasite surgically; however, the disease is usually diagnosed late, when it is inoperable and results in a high rate of fatality. Presurgical treatment with albendazole is recommended to reduce the size of the cyst before surgical removal. For inoperable cases, life-long treatment with mebendazole and albendazole has been used successfully and may be the preferred treatment in many cases. 

Prevention Controlling rodents is an important means of prevention, along with educating the public at risk to avoid exposure to infective feces. Practicing good hygiene and periodically deworming household pets are also helpful. 

Echinococcus oligarthrus and Echinococcus vogeli General Characteristics Infections with E. vogeli and E. oligarthrus are usually known as polycystic echinococcosis (or neotropical polycystic echinococcosis), from the form of the disease that is identified in the intermediate hosts. Because E. oligarthrus manifests as a single or discrete cyst in humans, this disease has also been called unicystic echinococcosis. 

Many strains of Echinococcus are distributed worldwide; however, E. vogeli and E. oligarthrus have been found only in Central and South America. 

Laboratory Diagnosis Echinococcus eggs are morphologically indistinguishable from Taenia spp., and the tiny proglottids are rarely noticed in feces. ELISAs that detect Echinococcus antigens in fecal samples (coproantigen ELISA) can be used to screen definitive hosts. This assay can detect both prepatent and patent infections. A PCR assay designed for fecal samples (coproDNA assay) is mainly used to confirm the infection or to identify eggs from the feces. Echinococcus adults or their proglottids can also be found in the definitive host after purgation with arecoline compounds. E. oligarthrus and E. vogeli adults usually have three segments. Direct examination of the intestines at necropsy may be used in some circumstances; however, subtle differences are present in the mature proglottids. E. oligarthrus is approximately 2 to 3 mm long and E. vogeli is 3.9 to 5.6 mm long. In addition

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to these characteristics, Echinococcus species can also be distinguished by PCR followed by sequencing or restriction fragment length polymorphism analysis. 

Therapy In the definitive host, Echinococcus spp. can be treated with antihelminthic drugs. Praziquantel, which is effective against both juvenile and adult parasites, is often used. In intermediate hosts, surgery is often the treatment of choice. Long-term antihelminthic treatment may also suppress some cysts. Long-term daily albendazole treatment, after surgical resection of the cyst masses, has suppressed parasite growth in some patients. 

Prevention Preventive measures include avoiding contact with infected dogs and deworming animals regularly. Effective control includes educating the population concerning the danger and routes of transmission associated with the disease as well as maintaining good hygiene and safe disposal of dog feces. Slaughtered animals must be disposed of properly to prevent dogs from being exposed to contaminated materials. 

Taenia multiceps and Other Species General Characteristics Taenia multiceps is the most common canid tapeworm that causes coenurosis in humans. Additional dog tapeworm species, Taenia crassiceps and Taenia serialis, have also been associated with human coenurosis (Chapter 53). The coenurus (larval form) may cause destructive damage or death but is an extremely rare disease in humans. The coenurus cyst is a unilocular cyst that contains a transparent fluid, similar to cysticercus, although the worm has multiple scolices. Daughter cysts may also be seen. The adult tapeworm of T. multiceps is 5 to 6 cm long and consists of 200 to 250 segments. The scolex has four suckers and a proboscis (tubular appendage) or rostellum with 22 to 32 hooks arranged in two rows. 

Epidemiology T. multiceps is most often found in Africa, although it may be seen in South America, the United States, and Canada. The adult worm is typically found in dogs and other canids. Many animals serve as the intermediate host, such as sheep, cattle, and deer. The animals become infected through the ingestion of eggs while grazing. Humans can also serve as an intermediate host. Human infection occurs from accidental ingestion of dog feces containing the eggs. 

Pathogenesis and Spectrum of Disease The oncosphere hatches and penetrates the intestinal wall of the intermediate host. The embryo is carried via the bloodstream to various parts of the body including the brain, eyes, and

771

CNS, where the organism lodges and the coenurus develops. The coenurus develops into multiple daughter cysts. Symptoms include headache, vomiting, paralysis, and blindness. The coenurus causes a serious disease in sheep and in dogs that have eaten the brains of infected sheep. Infected sheep lose their balance and rotate in circles until they fall (screw disease). The human clinical condition is known as gid, sturdy, or staggers. 

Laboratory Diagnosis Diagnosis is similar to that for Echinococcus infection. CT and MRI may be useful for detecting the cysts. Microscopic identification can be used if the cyst has been removed surgically. A serologic test available from the CDC provides additional clinically significant information in combination with imaging studies to establish a diagnosis. Nucleic acid–based methods have also been used for the diagnosis of T. multiceps infections. Although different serological methods, including ELISA and indirect hemagglutination assay, have been developed to diagnose coenurosis, the antigens used in these assays are natural worm extracts and cannot be produced commercially. Indirect ELISA assays using stable recombinant antigens Tm7 and heat shock protein 70 have been successfully developed for the diagnosis of coenurosis. Although PCR may be useful to detect nucleic acid in CSF, the current complexity of the concentration, extraction, and amplification process prevent routine use in clinical laboratories. 

Therapy Treatment is similar to that for Echinococcus. The most common treatment is surgery if possible, although the drugs used for cysticercosis may also be effective against coenurus infection. 

Prevention Dogs associated with sheep and other livestock should not be fed the brain or spinal cord from infected animals and should be dewormed regularly. Good hygiene should be practiced and care taken not to eat or drink anything contaminated with dog feces. 

Taenia serialis General Characteristics T. serialis has been found in a number of locations, including North America, South America, Europe, and Africa. 

Epidemiology T. serialis, also known as a canid tapeworm, are parasites of carnivores, particularly dogs, with herbivorous animals such as rabbits serving as intermediate hosts. Humans become accidental hosts when eggs are ingested from food or water contaminated with infected dog feces. 

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Pathogenesis and Spectrum of Disease

Spirometra mansonoides

Hatching of the T. serialis usually occurs only if the eggs have been exposed to gastric secretions. The oncospheres hatch in the intestine, invade the intestinal wall, and are carried in the blood throughout the ­ tissues. Within the tissues, the larvae (called metacestodes) develop into cysticerci or coenuri, which are larvae that group within cysts. The infection with the metacestode larval form (coenurus) of T. serialis is called coenurosis. When humans ingest these eggs from the infected tissue of a definitive host, the eggs develop into coenuri. These coenuri can occur in humans within muscle, brain, eye, or subcutaneous connective tissue. The symptoms are variable, and depend on the location and number of larvae. Coenuri in the skin or subcutaneous tissue usually present as painless nodules. The lesions are often fluctuant and tender. Most subcutaneous nodules manifest on the trunk, sclera, ­subconjunctiva, neck, shoulders, head, and limbs. Coenuri in the neck may affect movement and swallowing. Clinically, coenuri can mimic lymphomas, lipomas, pseudotumors, or neurofibromas. Coenuri in the CNS cause headache, fever, and vomiting. Localizing neurologic symptoms may also develop, including nerve palsies, Jacksonian epilepsy, pachymeningitis, obstructive or communicating hydrocephalus, and intracranial arteritis with transient hemiparesis. Coenuri in the eye can cause both intraocular and orbital infections, and patients may present with varying degrees of visual impairment. If not removed, coenuri in the eye can result in painful inflammation, glaucoma, and eventual blindness. 

General Characteristics Sparganosis is an infection caused by the plerocercoid larvae of Spirometra. The larvae (spargana) are white, wrinkled, and ribbon-shaped. They may be 3 mm wide and up to 30 cm long. The sparganum has bothria (longitudinal grooves) instead of suckers. No scolex is present, which can help differentiate Spirometra from T. solium. 

Epidemiology Spirometra is found worldwide; most human cases of sparganosis are found in Asia. Sparganosis is endemic in animals throughout North America but rare in humans. Adult Spirometra live in the intestine of dogs and cats. Eggs are shed in feces, hatch in water, and release free-swimming ciliated coracidia. The coracidia are then ingested by copepods, which become infected. Reptiles, fish, and amphibians ingest infected copepods containing the procercoid (elongated and globular) larvae. The procercoid larvae develop into plerocercoid (pseudosegmented with a scolex) larvae in the second intermediate host. Humans are accidental hosts; they acquire sparganosis after ingestion of contaminated water or by consuming undercooked fish. The life cycle is identical to that of the broad fish tapeworm, Diphyllobothrium spp. Humans are unable to serve as the definitive host for Spirometra. However, spargana can live up to 20 years in the human host. 

Pathogenesis and Spectrum of Disease

Diagnosis is made by the observation of coenuri in tissue biopsy or autopsy specimens. Coenuri are usually readily distinguished from cysticerci by the presence of multiple protoscoleces (infective form of the immature developing scolex). 

Spargana migrate and lodge anywhere in the human body. Clinical symptoms depend on which organs or tissues are involved. Spargana can live for several years before symptoms develop. Sparganosis is usually asymptomatic until the larvae grow and cause an inflammatory reaction. Painful nodules can develop in the tissues. A variety of symptoms may occur, including seizure, weakness, headache, and eye pain that can lead to blindness if left untreated. 

Therapy and Prevention

Laboratory Diagnosis

Surgical removal or antiparasitic agents can be used to treat coenurus. Oral epsiprantel, praziquantel, or fenbendazole can be used to treat coenuri infection. Discretion should be used when treating this evolving cestode as the dead parasite may cause a significant inflammatory response in the host. The inflammation can be managed with the use of corticosteroids. In many cases surgical removal of the coenurus is a safer option, as leakage of fluid from the cyst during surgery is unlikely to cause a new cyst. Surgical excision is often curative. Prevention is similar to that for T. multiceps. Good hygiene is essential to prevent contamination of food and the environment with feces from infected animals. Proper food preparation is also essential to avoid consumption of infected raw or undercooked meat. 

Definitive diagnosis is usually made by removal and identification of the sparganum from infected tissue. Serodiagnosis utilizing ELISA methodology can be used to target antisparganum IgG antibodies within the blood. ELISA may be positive around 10 to 12 days postinfection and is almost 100% effective at detecting the anti-sparganum antibodies at 14 to 22 days post infection. Clinical history, ELISA, MRI, and CT can all be used together to presumptively diagnose sparganosis. Eosinophilia may also be present. 

Laboratory Diagnosis

Therapy Praziquantel has been used with limited success. Injection of ethanol into the nodule along with surgical removal of the complete sparganum is the treatment of choice. 

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Prevention Prevention strategies should include safe drinking water practices and awareness of the dangers of consuming raw fish and amphibians. Water in contaminated areas should be boiled before it is consumed.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography An XX, Yang GY, Wang YW, et al.: Prokaryotic expression of Tm7 gene of Taenia multiceps and establishment of indirect ELISA using the expressed protein, Chin J Vet Sci 9:99–105, 2011. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Blutke A, Hamel D, Hüttner M, et  al.: Cystic echinococcosis due to Echinococcus equinus in a horse from southern Germany, J Vet Diagn Invest 22(3):458–462, 2010. Budke CM, White Jr AC, Garcia HH: Zoonotic larval cestode infections: neglected, neglected, tropical diseases, Negl Trop Dis 3:e319, 2009. Carroll CL, Connor DH: Sparganosis. In Connor DH, et al.: Pathology of infectious disease, Stamford, CT, 1997, Appleton Lange. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Dahniya MH, Hanna RM, Askelou S, et al.: The imaging appearances of hydatid disease at some unusual sites, Br J Radiol 74:283–289, 2001. Eckert J, Deplazes P: Biological, epidemiological, and clinical aspects of echinococcosis, a zoonosis of increasing concern, Clin Microbiol Rev 17:107–135, 2004. El-On J, Shelef I, Cagnano E, et al.: Taenia multiceps: a rare human cestode infection in Israel, Vet Ital 44:621–631, 2008. Garcia HH, Evans CA, Nast TE, et al.: Current consensus guidelines for treatment of neurocysticercosis, Clin Microbiol Rev 15:747– 756, 2002. Garcia LS: Diagnostic medical parasitology, ed 6, Washington, DC, 2016, ASM Press. Gonzalez LM, Montero E, Harrison LJ, et  al.: Differential diagnosis of Taenia saginata and Taenia solium infection by PCR, J Clin Microbiol 38:737–744, 2000.

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Horton J: Albendazole for the treatment of echinococcosis, Fundam Clin Pharmacol 17:205–212, 2003. Hu DD, Cui J, Wang L, et  al.: Immunoproteomic analysis of the excretory-secretory proteins from Spirometra mansoni sparganum, Iran J Parasitol 8(3):408–416, 2013. Huang X, Xu J, Wang Y, et al.: GP50 as a promising early diagnostic antigen for Taenia multiceps infection in goats by indirect ELISA, Parasit Vectors 9(1):618, 2016. Li M-W, Lin H-Y, Xie W-T, et al.: Enzootic sparganosis in Guangdong People’s Republic of China, Emerg Infect Dis 15(8):1317–1318, 2009. Liance M, Janir V, Bresson-Hadni S, et  al.: Immunodiagnosis of Echinococcus infections: confirmatory testing and species differentiation by a new commercial western blot, J Clin Microbiol 38:3718–3721, 2000. Lightowlers MW, Gottstein B: Echinococcosis/hydatidosis: antigens, immunological and molecular diagnosis. In Thompson RCA, Lymbery AJ, editors: Echinococcus and hydatid disease, Wallingford, UK, 1995, CAB International, pp 355–410. McManus DP, Zhang W, Li J, et  al.: Echinococcosis, Lancet 362:1295–1304, 2003. Oryan A, Amrabadi O, Sharifiyazdi H, et al.: Application of polymerase chain reaction on cerebrospinal fluid for diagnosis of cerebral coenurosis in small ruminants, Parasitol Res 114(10):3741–3746, 2015. Polat P, Kantarci M, Alper F, et al.: Hydatid disease from head to toe, Radiographics 23:475–494, 2003. Pedrosa I, Saiz A, Arrazola J, et al.: Hydatid disease: radiologic and pathologic features and complications, Radiographics 20:795–817, 2000. Rodriguez S, Wilkins P, Dorny P: Immunological and molecular diagnosis of cysticercosis, Pathog Glob Health 106:286–298, 2012. Spickler, Anna R. Echinococcosis. 2011. Available at: http://www.cfsph .iastate.edu/DiseaseInfo/factsheets.php. Suckow MA, Stevens KA, Wilson RP: The laboratory rabbit, guinea pig, hamster, and other rodents, Academic Press, 2012, p 441. Wang Y, Nie H, Gu X, et al.: An ELISA using recombinant TmHSP70 for the diagnosis of Taenia multiceps infections in goats, Vet Parasitol 212(3–4):469–472, 2015. Zhang W, Li J, McManus D: Concepts in immunology and diagnosis of hydatid disease, Clin Microbiol Rev 16:18–36, 2003.

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CASE STUDY 54.1 A 65-year-old man from Montana was admitted to the emergency department in a local hospital. The patient is a sheepherder and owns dogs. He claims to have no travel history outside the local area. He enjoys hunting and trapping in the area and frequently enjoys eating his venison. He was admitted with upper right quadrant pain and vomiting. An abdominal mass was felt and an MRI ordered. MRI showed a 5-cm mass in the liver. Fluid-filled cysts and a scolex were surgically removed (Fig. 54.4).

Suckers

rostellum

Questions 1. What parasite should be considered? 2. What additional testing should be performed to aid in diagnosis? 3. Which preventive measures should be used to control the spread of this parasite?

•

Fig. 54.4 Scolex collected from patient’s liver biopsy. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

  

Chapter Review 1. Which of the following is a characteristic of Echinococcus? a. Longest tapeworm found in humans b. Vitamin B12 deficiency c. Hydatid cysts d. Cysticercus 2. Which of the following cysts is not encased in a capsule? a.  T. solium b. T. multiceps c.  Mansonoides d. E. multilocularis 3. What is the drug of choice for treating hydatid cysts? a. Albendazole and praziquantel b. Praziquantel c. Mebendazole d. Niclosamide 4. Surgery may be the best course of treatment for which tapeworm? a.  E. granulosus b. E. multilocularis c.  T. multiceps d. All of the above 5. Human cysticercosis occurs when: a.  T. solium eggs are ingested b. Reverse peristalsis returns gravid segments into the intestine c. Humans ingest contaminated pork d. All of the above 6. Cysticercosis develops: a. In any organ or tissue in the body b. After ingesting contaminated beef c. And will die and calcify in 2 to 3 weeks d. All of the above

7. Neurocysticercosis is defined as: a. Cysticerci localized in the liver b. Cysticerci localized in the brain c. Cysticerci localized in the eye d. All of the above 8. Definitive diagnosis of cysticercus is made by: a. Identification of cysticerci in tissue b. CT or MRI c. Immunoblot assay d. All of the above 9. The cyst of E. multilocularis: a.  Grows quickly and produces many clinical symptoms b. Is most commonly found in the southern United States c. Is capable of forming a multicystic structure attributable to lack of a laminated membrane d. Causes coenurus in humans 10. Humans become infected with E. multilocularis when they: a. Ingest an oncosphere from an infected deer b. Ingest coracidia from an infected copepod c. Ingest eggs from an infected fox d. Ingest eggs from human feces 11. Humans acquire sparganosis by: a. Accidental ingestion of dog feces b. Handling infected animals c.  Consuming procercoid larvae in undercooked fish d. Ingesting human feces

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12.  True or False _____ Hydatid disease results when humans become the accidental host of T. solium cysts. _____ The presence of a scolex differentiates Spirometra from T. solium. _____ The rupture of a hydatid cyst may cause anaphylactic shock. _____ The cyst of T. solium lacks a laminated membrane.

13.  Matching: Match each term with the correct description. _____ neurocysticercosis _____ hydatid disease _____ hydatids _____ brood capsules _____ hydatid sand _____ multilocular _____ coenurus _____ bothria _____ CT _____ MRI

a. daughter cysts b. many-chambered cysts c. longitudinal grooves d. echinococcosis e. magnetic resonance imaging f. T. multiceps larval form g. cysticerci localized in brain h. computed tomography i. scolices in hydatid fluid j. cysts

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Intestinal Trematodes OBJECTIVES 1. List the clinically significant intestinal trematodes. 2. Describe the general life cycle of the intestinal trematodes, and identify the life cycle stage that is infective for humans. 3. Describe the diagnostic methods used to identify intestinal trematodes. 4. Explain the pathogenesis of intestinal trematode infections. 5. List the drugs of choice for intestinal trematode infections. 6. Describe the natural environment or habitat of intestinal trematodes, the route of transmission, and preventive measures.

PARASITES TO BE CONSIDERED Helminths Trematodes (flukes) Intestinal Echinostoma ilocanum Fasciolopsis buski Gastrodiscoides hominis Heterophyes heterophyes Metagonimus yokogawai Centrocestus spp. Haplorchis spp. Stellantchamus spp. Pygidiopsis spp.

T

he intestinal trematodes (flukes) are members of the phylum Platyhelminthes (flatworms), are dorsoventrally flattened, and require at least one intermediate host (a freshwater snail). Human infection occurs by ingestion of metacercariae (tailless encrusted larvae) encysted on freshwater vegetation or fish. Most trematodes are hermaphroditic (both ovaries and testes are contained within each adult worm). The parasites are typically identified from eggs shed in the feces. The adult worms are located in the small intestine, where they lay eggs that may be embryonated or remain unembryonated until shed from the body via feces. The egg continues developing after reaching the water, and a ciliated, free-swimming miracidium larva is released. The miracidium enters a snail host and develops into a redia (cylindrical larvae), followed by development into tailed cercariae. The cercariae emerge from the snail and encyst as a metacercariae on water plants or fish. A human host ingests raw or undercooked plants (Fasciolopsis buski), fish (Heterophyes, Metagonimus yokogawai), or freshwater mollusks or fish 774

(Echinostoma spp.) containing the metacercariae, which excyst in the intestinal tract, attach, and mature into adults. A representative life cycle for M. yokogawai is shown in Fig. 55.1.

Echinostoma spp. General Characteristics Varieties of species of echinostomes have been reported that infect humans including E. hortense, E. ilocanum, E. macrorchis, E. perforatum, and E. revolutum. The majority of human infections are caused by E. ilocanum. E. ilocanum adult flukes are elongated, leaflike, and approximately 1 cm in length and 0.2 cm in width. Both ends are attenuated, and the posterior end may be slightly pointed. Approximately 50% of adult worms demonstrate 49 to 53 collar spines arranged in alternating rows around the oral sucker. The eggs are passed in the feces of infected patients and are large and oval, with a relatively narrow operculum and small abopercular wrinkles at the posterior end. The eggs measure 89 to 112 μm long and 58 to 69 μm wide. 

Epidemiology Echinostomes infect freshwater mollusks, primarily snails. Infections are common in Russia, Southeast Asia, and the Far East. Rats and dogs serve as reservoir hosts in endemic areas. 

Pathogenesis and Spectrum of Disease Echinostome infections can cause severe clinical manifestations, particularly in cases with heavy worm burdens. Infections can induce mucosal ulceration and bleeding in the duodenum and jejunum through mechanical irritation of the worms (using collar spines, and oral and ventral suckers) leading to severe gastrointestinal discomforts, including epigastric or abdominal pain accompanied by diarrhea, easy fatigue, and malnutrition, for several months’ duration. Patients may appear asymptomatic in light infections and experience mild abdominal pain and diarrhea. 

Therapy and Prevention Praziquantel is the drug of choice for echinostomiasis although it is not included in the US product labeling for

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• Fig. 55.1  This is an illustration of the life cycle of the parasitic trematode, Metagonimus yokogawai, the

causal agent of metagonimiasis. (Photo Courtesy the Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

Fasciolopsis buski General Characteristics The adults of F. buski have an elongated shape and range from 20 to 75 mm long and approximately 8 to 20 mm wide (Fig. 55.2). They have an oral sucker at the anterior end and a ventral sucker located about midway to the posterior end. The eggs, which are indistinguishable from those of Fasciola hepatica (Fig. 55.3), are oval and elongated, transparent, and yellow-brown with an operculum (lid) at one end, and they range in size from 130 to 140 mm long and 80 to 85 mm wide and may be unembryonated. •

Epidemiology

these infections. Although a single dose of 25 mg/kg of praziquantel is the recommended dose for treatment of intestinal fluke infection, echinostome infections can be treated with a slightly lower single oral dose of 10 to 20 mg/kg praziquantel. Proper food preparation and avoiding the consumption of raw, undercooked, or freshly pickled fish prevents infection. Praziquantel is the drug of choice for treatment. In addition, albendazole may also be used. 

F. buski is found in Bangladesh, Cambodia, China, India, Indonesia, Laos, Malaysia, Pakistan, Taiwan, Thailand, and Vietnam and is prevalent in school-aged children. Contaminated feces drain into the water from farmland, where feces is used for fertilization, or defecation occurs in or near water sources. Reservoir hosts include pigs, dogs, and rabbits. F. buski is the largest of the intestinal trematodes, and infection is acquired by ingestion of raw water chestnuts or caltrop (plants with spiny heads or fruit). The definitive host is the pig, and fish-eating wild and domestic animals may

Fig. 55.2 Whole mount of Fasciolopsis buski. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

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Epidemiology Human gastrodiscoidiasis is endemic in Southeast Asia, as well as the Philippines, Guyana, India, and Assam. High incidence can be attributed to low standards of sanitation in regions such as rural farms and villages, where night soils are used for fertilizer. Infection in both humans and animals results from the ingestion of contaminated vegetation. Transmission to humans occurs when undercooked or raw infected fish are ingested. Reservoir hosts include rats and deer mice. Humans are an accidental host with pigs serving as the definitive host.  • Fig. 55.3  Fasciola egg. The eggs of Fasciolopsis buski and Fasciola

hepatica are indistinguishable morphologically. (Courtesy Dr. Henry Travers, Sioux Falls, SD.)

serve as reservoir hosts. The water vegetation may become contaminated when feces is used for fertilization or where disposal of farm animal feces is inadequate. 

Pathogenesis and Spectrum of Disease The intestinal attachment site of the adult worms often becomes locally inflamed and ulcerated and may hemorrhage. Moderate to heavy infections may cause abdominal pain, diarrhea, intestinal obstruction, and edema of the abdomen and lower extremities and may result in inadequate absorption of vitamin B12. Eosinophilia is common. 

Therapy and Prevention The current treatment of choice for F. buski infection is praziquantel; single dose up to three times a day of niclosamide has been reported to have some in vitro efficacy. Infection can be prevented by immersing water plants in boiling water for a few seconds before peeling, and properly cooking water plants and fish before they are eaten. In addition, changes are needed in agricultural practices and health education in the endemic areas. 

Gastrodiscoides hominis General Characteristics The natural habitat for G. hominis, also known as the colonic fluke, is the colon of pigs. The adult worm is vase-shaped and bright pink, averaging 5 to 8 mm long and 3 to 5 mm wide. The anterior region has a prominent oral sucker. The posterior portion is discoidal, and the ventral sucker is close to the posterior end. The tegument is smooth and contains a series of concentric folds bearing numerous tightly packed tubercles. Ciliated and non-ciliated papillae are arranged around the oral sucker on the ventral surface of the worm. Eggs measure approximately 146 by 66 μm, are rhomboidal (parallelogram shape), transparent, and green in color. Each egg contains about 24 vitelline (yolk sac–like) cells and a central unembryonated ovum. 

Pathogenesis and Spectrum of Disease Gastrodiscoidiasis as an infection in the definitive host is usually mild to asymptomatic. Following ingestion by a human host the metacercaria travel through the digestive tract into the duodenum to the cecum, where the larvae selffertilize and lay eggs. Disease presentation is dependent on the worm burden. Patients may be asymptomatic, but heavy infections cause diarrhea, fever, abdominal pain, colic, malnutrition, and anemia. In severe cases, where there are large numbers of eggs present, papular lesions and desquamation lead to necrosis. Inflammatory reactions can occur in the heart or mesenteric lymphatic system. 

Therapy and Prevention Praziquantel is the treatment of choice for infections. Changes in agricultural practices, health education, and proper hygiene can prevent transmission in the endemic areas. The preferred drug, praziquantel, eliminates the parasite with three doses at 25 mg/kg in 1 day. Prevention of this disease is not difficult when simple sanitary measures are taken. Night soil should never be used as a fertilizer because it could contain any number of parasites. Vegetables should be washed thoroughly, and meat properly cooked. 

Heterophyes: Metagonimus yokogawai, Centrocestus spp., Haplorchis spp., Stellantchamus spp., and Pygidiopsis spp. General Characteristics There are 22 genera in the family Heterophyidae, with six medically relevant genera. Heterophyes and M. yokogawai are the most prominent. Additional genera included in the family include Centrocestus spp., Haplorchis spp., Stellantchamus spp., and Pygidiopsis spp.

Heterophyes heterophyes Adult H. heterophyes worms range in size from 1.0 to 1.7 mm in length and 0.3 to 0.4 mm in width and have a broadly rounded posterior. The adult H. heterophyes also has an additional sucker, the genital sucker, which surrounds

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the genital pore. The eggs are small, yellow-brown, embryonated, and operculated and may have minimal opercular shoulders. Eggs range in size from 26 to 30 mm long and 15 to 17 mm wide and may be indistinguishable from M. yokogawai.

Epidemiology H. heterophyes is distributed throughout China, Egypt, India, Iran, Israel, Japan, Korea, Sudan, the Philippines, Tunisia, and Turkey. Reservoir hosts include cats, dogs, and birds. Snails serve as the intermediate host and a variety of freshwater fish serve as the second intermediate host. This very small trematode is acquired through ingestion of pickled, raw, or inadequately cooked fish. 

Metagonimus yokogawai Adult M. yokogawai adult worms range in size from 1.0 to 2.5 mm long and approximately 0.4 to 0.8 mm wide. The eggs are small, yellow-brown, embryonated, and operculated, and may have minimal opercular shoulders. Eggs range in size from 26 to 30 mm long and 15 to 17 mm wide and may be indistinguishable from H. heterophyes. 

Epidemiology M. yokogawai is found in the Balkans (a cultural region of southeastern Europe), China, Indonesia, Israel, Japan, Korea, Russia, Spain, and Taiwan and is considered the most common intestinal fluke infection in the Far East. Reservoir hosts include cats, dogs, and birds. Freshwater snails serve as the intermediate host, and a variety of freshwater fish serve as the second intermediate host. This very small trematode is acquired through ingestion of pickled, raw, or inadequately cooked fish. 

Centrocestus spp. Adult worms range in size from 280 to 330 μm long and approximately 150 to 180 μm wide. The body is covered in scalelike tegumental spines. The worm has a terminal oral sucker with 32 circumoral spines arranged in two rows and a small ventral sucker. The eggs are oval, yellowish brown, with a distinct operculum. The shell surface appears to have a lattice design. The eggs are approximately 33 μm long and 17 μm wide.

Epidemiology Centrocestus spp. is an intestinal foodborne trematode that parasitizes birds and mammals, including humans. The species is native to Asia, with worldwide distribution. The primary molluscan intermediate host, Melanoides spp., is reported to be present in more than 10 countries throughout Asia and the Americas. Many species of fish are the second intermediate hosts. Metacercariae encyst in the gills of the fish causing pathological developmental delay and death, giving rise to economic losses in the fish farming industry. 

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Haplorchis spp. The adult worms of Haplorchis are characterized by the presence of a small, armed ventral sucker and a single testis, which differentiates it from Metagonimus and Heterophyes. A distinct morphological feature of Haplorchis is the ventral sucker that has a semi-lunar (half-moon shaped) group of 12 to 16 long, crescentic, and hollow spines and a sinistral (left side) patch of very minute solid spines. The eggs are oval with a convex operculum. The eggs are very similar in shape, size, and morphology to Opisthorchis viverrini and cannot be morphologically differentiated.

Epidemiology Human infections with Haplorchis spp. are prevalent in Southeast Asia, including countries located in the Indochina Peninsula, Taiwan, the Philippines, and Egypt. The intermediate host is freshwater snails, Melania spp. Fish are the second intermediate hosts, and dozens of species of birds and fish-eating mammals, including dogs, cats, and humans, serve as definitive hosts. 

Stellantchamus spp. The adult flukes are pyriform with a small submedian ventral sucker and an expulsor-type elongated saclike seminal vesicle. Eggs are elongated, ovoid, and slender, from 25.3 to 29.2 μm long and 11.1 to13.4 μm wide.

Epidemiology Human infections with Stellantchamus spp. are prevalent in Southeast Asia, including Korea, Japan, and Kuwait. The intermediate host is the brackish water snail, Melanoides spp. The second intermediate hosts are brackish water fish, primarily mullets. Additional freshwater fish, including the half-beaked fish and climbing perch, can also serve as the second intermediate host. Natural definitive hosts include cats, dogs, pigs, rats, humans, and birds. 

Pygidiopsis spp. The adult flukes have a small concave body with a medial ventral sucker. The worm has a unique ventrogenital apparatus (having two groups of spines around the genital pore; 5 to 6 right side and 7 to 9 left side). The two testes are side by side. The worm produces small (19.8 to 22.9 μm long and 11.1 to 13.4 μm wide), ovoid, pyriform eggs with no distinct pattern on the shell.

Epidemiology Human infections with Pygidiopsis spp. are prevalent in Southeast Asia and other regions, including Korea, Japan, Vietnam, and Egypt. The primary intermediate host is the brackish water snail, Melanoides spp. The second intermediate host is brackish water fish. The definitive hosts include wolves, cats, dogs, foxes, shrews, rats, pelicans, kites, ducks, and cormorants. 

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Pathogenicity and Spectrum of Disease Infections with a small number of worms from any of the heterophyid genera may be asymptomatic. Symptoms in heavy infections may include abdominal pain, diarrhea with a large amount of mucus, and ulceration of the intestinal wall. Eggs may gain entry into the intestinal capillaries and the lymphatic system, where they can be carried to the heart, brain, spinal cord, or other tissues, causing emboli or granuloma formation. 

Prevention Infection can be prevented by avoiding ingestion of raw, inadequately cooked, and pickled or salted fish. The risk of infection could be reduced with improved sanitary conditions and health education programs. 

Laboratory Diagnosis Identification of the intestinal trematodes is made by recovery of eggs, or in rare cases adults, from stool specimens using a sedimentation method such as formalin-ethyl acetate. The sediment may be examined in a wet mount with or without iodine. The eggs of F. buski are identical to those of F. hepatica, E. ilocanum, and G. hominis; and those of H. heterophyes and M. yokogawai are very similar. Diagnosis of heterophyid infections may also require assessment of symptoms, obtaining a travel history, and/or recovery of adult worms.

Nucleic Acid Detection Various polymerase chain reaction (PCR) methods have shown potential in detecting intestinal flukes. These

methods take advantage of the different types of DNA nucleotide sequence variations demonstrated by the different species within a particular genus. Multiplex qPCR has been developed to detect the presence of the major intestinal parasites known to cause gastroenteritis, including the trematodes C. sinensis and M yokogawai. The assays reportedly exhibit 100% sensitivity and 100% specificity. Polymerase chain reaction–restriction fragment length polymorphism (PCR-RFLP) and simple sequence repeat anchored PCR have been reported to be useful in distinguishing species of the Metagonimus genus (including M. yokogawai). Information derived from RFLP involving specific sites in ribosomal RNA and mitochondrial cytochrome oxidase I (mtCOI) genes may help to differentiate M. yokogawai from other Metagonimus species. Six members of the Heterophyidae family can be distinguished with PCR assays based on variations in rDNA polymorphisms among the species. 

Treatment The drug of choice for treatment of intestinal trematode infection is praziquantel, an isoquinoline derivative administered orally in three doses for 1 day. There may be some mild side effects, but these usually disappear within 48 hours and may be more severe in those with heavy infections. An alternative drug is niclosamide and is administered for 1 to 2 days.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Bogitsch BJ, Carter CE, Oeltmann TN, editors: Human parasitology, ed 3, San Diego, 2005, Academic Press. Caroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Chai JY, Jung BK: Fishborne zoonotic heterophyid infections: an update, Food Waterborne Parasitol 8–9:33–63, 2017. Chai JY: Echinostomes in humans. In Bernard F, Rafael T, editors: The biology of echinostomes: from the molecule to the community, New York, 2009, Springer, pp 147–183. Chai JY, Shin EH, Lee SH, Rim HJ: Foodborne intestinal flukes in Southeast Asia, Korean J Parasitol 47(Suppl):S69–S102, 2009. https://doi.org/10.3347/kjp.2009.47.S.S69. Chai JY, Sohn WM, Cho J, et  al.: Echinostoma ilocanum infection in two residents of Savannakhet Province, Lao PDR, Korean J Parasitol 56(1):75–79, 2018. https://doi.org/10.3347/ kjp.2018.56.1.75. Dzikowski R, Levy MG, Poore MF, Flowers JR, Paperna I: Use of rDNA polymorphism for identification of heterophyidae infecting freshwater fishes, Dis Aquat Organ 59(1):35–41, 2004.

Dada-Adegbola HO, Falade CO, Oluwatoba OA, et al.: Gastrodiscoides hominis infection in a Nigerian-case report, West Afr J Med 23(2): 185–186, 2004. Fried B, Graczyk TK, Tamang L: Food-borne intestinal trematodiases in humans, Parasitol Res 93:159–170, 2004. Garcia LS: Diagnostic medical parasitology, ed 6, Washington, DC, 2016, ASM Press. Han ET, Shin EH, Phommakorn S, et  al.: Centrocestus formosanus (Digenea: heterophyidae) encysted in the freshwater fish, Puntius brevis, from Lao PDR, Korean J Parasitol 46(1):49–53, 2008. Jeon HK, Lee D, Park H, et  al.: Human infections with liver and minute intestinal flukes in Guangxi, China: analysis by DNA sequencing, ultrasonography, and immunoaffinity chromatography, Korean J Parasitol 50(4):391–394, 2012. John DT, Petri WA: Markell’s and Voge’s medical parasitology, ed 9, St Louis, 2006, Elsevier. Kajugu PE, Hanna RE, Edgar HW, et al.: Fasciola hepatica: specificity of a coproantigen ELISA test for diagnosis of fasciolosis in faecal samples from cattle and sheep concurrently infected with gastrointestinal nematodes, coccidians and/or rumen flukes (paramphistomes), under field conditions, Vet Parasitol 212(3–4):181–187, 2015.

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Keiser J, Utzinger J: Food-borne trematodiases, Clin Microbiol Rev 22:466–483, 2009. Kumchoo K, Wongsawad C, Vanittanakom P, Chai JY, Rojanapaibul A: Effect of niclosamide on the tegumental surface of Haplorchis taichui using scanning electron microscopy, J Helminthol 81(4):329–337, 2007. Lee SH, Hwang SW, Chai JY, et al.: Comparative morphology of eggs of heterophyids and Clonorchis sinensis causing human infections in Korea, Korean J Parasitol 22(2):171–180, 1984. Lovis L, Mak TK, Phongluxa K, et al.: PCR diagnosis of opisthorchis viverrini and haplorchis taichui infections in a Lao community in an area of endemicity and comparison of diagnostic methods for parasitological field surveys, J Clin Microbiol 47(5):1517–1523, 2009. Quang TD, Duong TH, Richard-Lenoble D, et  al.: [Emergence in humans of fascioliasis (from Fasciola gigantica) and intestinal distomatosis (from Fasciolopsis buski) in Laos], Sante 18(3):119– 124, 2008.

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Won EJ, Kim SH, Kee SJ, et al.: Multiplex real-time PCR assay targeting eight parasites customized to the Korean population: potential use for detection in diarrheal stool samples from gastroenteritis patients, PloS One 11(11):e0166957, 2016. Yang HJ, Guk SM, Han ET, et  al.: Molecular differentiation of three species of Metagonimus by simple sequence repeat anchored polymerase chain reaction (SSR-PCR) amplification, J Parasitol 86(5):1170–1172, 2000. Yu JR, Chai JY: Metagonimus. In Liu D, editor: Molecular detection of foodborne pathogens, Boca Raton, FL, 2010, CRC Press, pp 805–812.

CASE STUDY 55.1 The husband of a 32-year-old woman is employed in the foreign service, and the couple has recently been on assignment to Africa. The wife complained of a 2-month history of abdominal pain, vomiting, diarrhea, and weight loss. While in Africa, she had eaten the locally grown watercress. A stool specimen was collected for culture, ova, and parasite examination. The bacterial culture was negative. A wet mount made during the parasite examination showed large, oval, operculated, and unembryonated helminth eggs.

Questions 1. What parasite is the probable cause of the patient’s symptoms? 2. Another parasitic worm has indistinguishable eggs. How would the infections caused by these two worms differ? 3. How did the patient most likely acquire this infection? 4. What would be the preferred treatment for this infection?   

Chapter Review 1. Fluke eggs are equipped with a lid at the top of the shell called a/an: a. Egress b. Operculum c. Nodule d. Button 2. The infective life cycle stage of a fluke is the: a. Miracidium b. Cercariae c. Metacercariae d. Pleurocercariae 3. What intermediate host is required in the life cycle of all trematodes? a. Freshwater snail b. Crayfish c. Aquatic vegetation d. Freshwater crab

4. Which two of the following are small flukes whose eggs are generally indistinguishable? a. Fasciolopsis b. Heterophyes c. Metagonimus d. Paragonimus 5. What laboratory finding is common in F. buski infections? a. Increased levels of vitamin B12 b. Increased serum bilirubin levels c. Decreased number of red blood cells d. Increased number of eosinophilia 6. The drug of choice for an intestinal trematode infection is: a. Niclosamide b. Praziquantel c. Albendazole d. Tiabendazole

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Liver and Lung Trematodes OBJECTIVES 1. List the clinically significant trematodes capable of infecting the liver and lungs. 2. Describe the general life cycle of the liver and lung flukes and identify the infective stage for humans. 3. Describe the diagnostic methods used to identify the liver and lung flukes, including the microscopic differentiation of eggs and serologic methods. 4. Describe the pathogenesis of the liver and lung flukes, including location and associated disease manifestations. 5. List the drug of choice for infections with liver and lung flukes. 6. Describe the transmission of the liver and lung flukes and discuss how infection may be prevented.

PARASITES TO BE CONSIDERED Trematodes (Flukes) Liver/Lung

Clonorchis sinensis Opisthorchis felineus Opisthorchis viverrini Fasciola gigantica Fasciola hepatica Paragonimus africanus Paragonimus caliensis Paragonimus heterotremus Paragonimus kellicotti Paragonimus mexicanus Paragonimus miyazakii Paragonimus skrjabini Paragonimus uterobilateralis Paragonimus westermani

T

he parasites within this chapter are typically foodborne and may have serious economic implications. Clonorchis spp., Opisthorchis spp., and Fasciola spp. live in the biliary ducts of humans. Paragonimus spp. are found in the lungs and in other body sites.

The Liver Flukes General Characteristics The adults of these trematodes live in the biliary ducts and may be found in the gallbladder in heavy infections. Three of these, Clonorchis sinensis (the Chinese liver fluke), 780

Opisthorchis felineus, and Opisthorchis viverrini (the Southeast Asian liver fluke), are elongated and narrow and much smaller than Fasciola (the sheep liver fluke). These flukes also all require a freshwater snail as an intermediate host. 

Epidemiology and Life Cycle C. sinensis is distributed throughout China, Japan, Korea, Taiwan, and Vietnam. O. viverrini is found in Cambodia, Laos, Thailand, and Vietnam, and O. felineus is found in Northern Europe and Asia. Reservoir hosts include dogs and cats. Fasciola spp. cause very similar disease, differing primarily in geographic distribution, with F. hepatica occurring in Europe and North and South America, and F. gigantica in Asia and Africa. Fasciola spp. affect the economics of the sheep and cattle industries. Reservoir hosts include dogs, hogs, cats, martens, badgers, minks, weasels, pigs, equines, and rats. Infected feces enter the water system because of improper drainage and unsanitary practices. The life cycle of the liver flukes is very similar to that of the intestinal flukes. The adult worms produce eggs in the biliary ducts that are then excreted from the body in the feces. The free-swimming miracidium is released from the egg in fresh water and enters the snail host, where it develops into a redia and then cercariae, which leaves the snail and enters the water (Fig. 56.1). The cercariae of Clonorchis and Opisthorchis are ingested by a second intermediate host, a freshwater fish. The cercariae then encyst and develop into the metacercariae within the intermediate host. The metacercaria is the infective stage for humans. When infected freshwater fish are eaten raw or undercooked, the metacercariae will excyst in the duodenum and then travel to the bile duct, where they mature. The cercariae of Fasciola encyst on freshwater vegetation, such as watercress and water chestnuts, and develop into metacercariae. When the infected vegetation is eaten raw, the metacercariae will excyst in the duodenum and then travel to the bile duct and mature. Fig. 56.2 depicts the general life cycles of the liver and lung flukes. 

Pathogenesis and Spectrum of Disease Light infections with C. sinensis or Opisthorchis spp. are most common and may be asymptomatic. Heavier infections with these flukes may present with fever, abdominal pain, and jaundice. Eosinophilia and increased serum levels

CHAPTER 56  Liver and Lung Trematodes

of immunoglobulin E (IgE) may occur. Severe infections may cause obstruction of the biliary ducts, resulting in enlargement and tenderness of the liver, cirrhosis, cholecystitis (inflammation of the gallbladder), and cholangiocarcinoma (cancerous growth in bile duct epithelium). Even light infections with Fasciola may cause fever, ­abdominal pain, nausea, diarrhea, enlargement and tenderness

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of the liver, jaundice, nonproductive cough, eosinophilia, and elevated serum IgE levels. Leukocytosis, eosinophilia and mild to moderate anemia may also be present. More severe infections may result in obstruction of the biliary ducts, cirrhosis, cholecystitis, and cholangiocarcinoma. During migration in the human body, the larvae may penetrate the peritoneal cavity, and adult flukes may be found in the intestinal walls, lungs, heart, or brain. Many symptoms of infection disappear when the parasite has lodged in the biliary passages resulting in chronic infection. The chronic phase presents with liver abnormalities and eosinophilia. In chronic infections, worms have been identified in the intestinal wall, brain, heart, lungs, and skin. 

Laboratory Diagnosis

• Fig. 56.1  Cercaria of a liver fluke. (Photo courtesy Dr. Henry Travers, Sioux Falls, SD.)

Identification of the liver flukes is primarily made by recovery of the eggs in feces using a sedimentation method and a wet mount with or without iodine staining. The adult worms of Clonorchis are elongated and ­narrow and a transparent reddish-yellow color. Adult Clonorchis may vary in size from 10 to 25 mm × 3 to 5 mm. The eggs

• Fig. 56.2  Life cycle of the trematode, Clonorchis sinensis, the causal agent of clonorchiasis. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

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of Clonorchis are 28 to 30 μm × 14 to 18 μm. The eggs have shouldered opercula and a small knob at the end opposite the operculum, are yellow-brown in color, and are embryonated when they leave the body (Fig. 56.3A). Diagnosis is based on microscopic identification of eggs in stool specimens. However, the eggs of Clonorchis are practically indistinguishable from those of Opisthorchis. The adult fluke can also be surgically recovered. Like Clonorchis, the adult worms of Opisthorchis are elongated and narrow and a transparent reddish-yellow color. Adult worms of Opisthorchis, however, are much smaller: 5 to 10 mm × 0.8 to 1.9 mm. The adult worm of O. felineus is lancet-shaped and is larger than O. viverrini. The size of Opisthorchis eggs is slightly smaller than Clonorchis; Opisthorchis eggs are 19 to 29 μm × 12 to 17 μm. Also like Clonorchis, the eggs have shouldered opercula and a small knob at the end opposite the operculum, are yellow-brown in color, and are embryonated when they leave the body (Fig. 56.3). Diagnosis is made by detecting eggs in a duodenal aspirate (bile fluid), the recovery of adult worms, or by clinical history. Eggs are not excreted in patients with biliary obstruction, requiring needle aspiration, surgery, or an autopsy specimen for confirmation. The adult worm of Fasciola is much larger (2 to 5 cm × 0.8 to 1.3 cm) (Fig. 56.3C), with a cephalic cone at the anterior end, that contains the oral sucker. The adult fluke of F. gigantica is similar to F. hepatica; however, it is somewhat more lancet-shaped and has a less distinct cephalic cone. The eggs are 130 to 150 μm × 70 to 90 μm, operculated, brownish-yellow, and unembryonated when they leave the body. The eggs of F. gigantica tend to be somewhat larger (160 to 190 μm by 70 to 90 μm). Because the eggs of Fasciola spp. and Fasciolopsis are virtually indistinguishable, it may also be necessary to recover eggs from bile specimens or to recover adult worms. The diagnosis of F. gigantica is the same as F. hepatica; however, eggs may even be less likely to be identified in the stool of infected patients. Definitive identification of Fasciola is important, because the treatment is different from that for Fasciolopsis. Fig. 55.3 shows

A

B

the eggs of Fasciola hepatica, and Fig. 55.2 is a Fasciolopsis buski adult fluke.

Serologic Testing C. sinensis and O. viverrini infections elicit a strong immune response. IgE levels are elevated in infections, and IgM is detectable in acute infections, followed by an increase in IgA and IgG. In chronic infections, the IgA level returns to normal, but IgG and IgM remain elevated. Serological diagnosis has been successful for C. sinensis; using ELISA is sensitive and demonstrates low cross-reactivity to related organisms. However, this means of diagnosis is not available for O. viverrini due to a lack of standardization and falsepositive results; this is caused by cross-reactivity with other parasitic diseases. ELISA has been successful using excretory and secretory antigens from adult worms of O. felineus cultivated in vivo. Serologic testing is available for the diagnosis of Fasciola spp. Serologic testing can be useful in the acute phase of infection because specific antibodies to Fasciola may become detectable 2 to 4 weeks following infection, whereas egg production typically is not apparent until 3 to 4 months after exposure. Serologic testing can also be of value for cases of chronic Fasciola infection in persons with low-level or sporadic egg production, as well as in persons with ectopic infection. It may also help rule out pseudofascioliasis associated with ingestion of parasite eggs in sheep or beef liver. The immune diagnosis of human F. hepatica infection includes an enzyme immunoassay (EIA) with excretorysecretory (ES) or recombinant antigens and confirmatory testing of EIA-positive specimens with an immunoblot assay. Enzyme-linked immunosorbent assay (ELISA) serum IgG antibody testing may demonstrate cross-reactivity with other trematodes, such as the schistosomes. 

Nucleic Acid Detection A variety of PCR-based methods including loop-mediated isothermal amplification (LAMP) have been developed for the detection of the liver flukes, including F. hepatica. These

C

• Fig. 56.3  Trematode eggs. (A) Clonorchis sinensis. (B) Paragonimus westermani. (C) Fasciola hepatica

(note the partially open operculum). (From Centers for Disease Control and Prevention. DPDx-Laboratory Identification of Parasitic Diseases of Public Health Concern.)

CHAPTER 56  Liver and Lung Trematodes

methods demonstrate a high sensitivity and specificity for the diagnosis of F. hepatica infection in comparison to routine fecal and indirect serological tests. The LAMP detects DNA of C. sinensis and O. viverrini in freshwater snails, the second intermediate fish hosts, and patient feces. In human fecal samples, LAMP-based technology detects C. sinensis in human infection with as low as 1 egg per 100 mg of feces. Further evaluation of the LAMP-based diagnosis test showed a sensitivity of 97.1% and specificity of 100%. Similar LAMP assays are also available for O. viverrini, with the variation of sensitivity and specificity relating to the repetition of different target genes when detecting DNA. A multiplex PCR assay has been used to detect C. sinensis and Opisthorchis in fish and infected patients in endemic areas. A single-step duplex real-time fluorescence resonance energy transfer (FRET) real-time PCR has been developed to diagnose and differentiate C. sinensis and Opisthorchis infections in human fecal samples. The assay differentiates the two species with 100% specificity and sensitivity. Despite the clinical utility of nucleic acid methods for the detection of the liver flukes, they are not generally available in routine laboratories, even in endemic regions. 

Therapy and Prevention The drug of choice for treatment of infections with Clonorchis and Opisthorchis is praziquantel (25 mg/kg) given orally three times per day for 2 days. The drug of choice for the treatment of Fasciola spp. is triclabendazole (praziquantel is not as effective). Infections with Opisthorchis often require additional treatment with antibiotics due to secondary bacterial infections. Human infection can be prevented by ensuring that fish and aquatic vegetation are properly cooked before consumption, as well as by the improvement of sanitary conditions, along with good personal hygiene. 

The Lung Flukes General Characteristics The genus Paragonimus contains approximately 15 species known to infect humans. Paragonimus westermani is the most common and widely distributed lung fluke. P. mexicanus is located in Central and South America, and P. kellicotti is found in North and South America. Both are important human pathogens. The adult worms live in the lungs and produce eggs that may be present in sputum, or if expectorated and swallowed may be present in feces. Like other trematodes, a freshwater snail is required as an intermediate host. 

Epidemiology and Life Cycle Paragonimiasis is found primarily in the Far East, in India, the Philippines, China, Japan, Korea, Manchuria, Papua New Guinea, and Southeast Asia, following ingestion of uncooked crabs, crayfish, freshwater shrimp, mussels, and

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paratenic hosts. Reservoir hosts for P. westermani include dogs and cats, and those for P. mexicanus include domestic and wild pigs, dogs, and rodents. Species of Paragonimus may also be found in other freshwater crab- or crayfisheating mammals. P. kellicotti is becoming an increasing concern in the United States. It is widely distributed throughout North America and is found in cats, dogs, bobcats, raccoons, foxes, skunks, minks, and coyotes. Transmission in the United States is associated with behavioral activities that include alcohol consumption, dares, and the demonstration of outside survival skills. The adult worms, encapsulated in the lungs, produce eggs that leave the lung via the bronchioles, stimulating a cough response. The eggs are swallowed and eventually excreted in the feces. Egg size varies with species from approximately 80 to 120 μm long and 45 to 70 μm wide (Fig. 56.3B). The free-swimming miracidium is released from the egg in fresh water and enters the snail host, where it develops into a redia and then cercariae, which leaves the snail and enters the water. The cercariae then enter a second intermediate host, a crab or crayfish, where they encyst and develop into metacercariae. The metacercaria is the infective stage for humans. When infected freshwater crabs and crayfish are eaten raw or undercooked, the metacercariae will excyst in the duodenum and then migrate through the intestinal wall, and eventually through the diaphragm and into the lungs where they encapsulate (usually in pairs) and mature (Fig. 56.2). 

Pathogenesis and Spectrum of Disease Light infections may be asymptomatic. The migration of the metacercariae through muscle and tissue may cause local pain and an immune response to the tissue damage. In the lungs, the immune response causes infiltration of eosinophils and neutrophils. Serum IgE levels are usually elevated. Eventually the adult worms are encapsulated in a granuloma. Presence of the worms in the lungs usually results in a chronic cough, with possible production of blood-tinged sputum. The cough provides a mechanism to transport eggs up into the throat, where they are swallowed and then may be excreted in the feces. The larvae of P. mexicanus, P. skrjabini, P. heterotremus, and P. hueitungensis may migrate to other areas of the body, commonly causing subcutaneous or lower abdominal nodules to form. The larvae of Paragonimus may enter other sites such as the liver, intestinal wall, muscles, peritoneum, and the brain, where they can cause severe damage. These ectopic infections are generally associated with P. heterotremus, P. mexicanus, and P. westermani. Computed tomography (CT) and magnetic resonance imaging (MRI) may be used to reveal cysts with edema, migration tracks, and nodules in various areas of the body including the brain, liver, and lungs. 

Laboratory Diagnosis The adult worms of Paragonimus vary in size, 10 to 25 mm × 3 to 5 mm, and are a reddish-brown color. The

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eggs of P. westermani measure 80 to 120 μm × 45 to 60 μm, and those of P. mexicanus are approximately 80 μm × 40 μm. The eggs are unembryonated when they leave the body, operculated with opercular shoulders, thick-shelled, and brownish-yellow. The eggs of Paragonimus are similar to those of Diphyllobothrium (freshwater fish tapeworm) and can be differentiated by the operculum, opercular shoulders, and thickened shell at the end opposite the operculum. Paragonimus eggs may be recovered from sputum, pleural effusions, and occasionally in feces using a sedimentation concentration method. The eggs may be observed in a wet mount (with or without iodine stain) (Fig. 56.4). CharcotLeyden crystals may also be observed in sputum or lung tissue specimens. Charcot-Leyden crystals are slender and pointed at both ends. The crystals normally appear colorless and stain purplish to red with trichrome. Elevated levels of eosinophils may be present in whole blood, and elevated IgE levels may be present in serum. Lesions in the lungs may be observed by x-ray.

•

Serologic Testing Serologic testing is available in the United States for the diagnosis of P. westermani and P. kellicotti. EIA tests and immunoblot (IB) assays and hemagglutination tests can be used to diagnose active infections. Antibody levels detected by EIA and IB do decline after chemotherapy and death of the worms. Pleural effusion samples are more suitable for the detection of antibodies than serum. The Division of Parasitic Disease at the Centers for Disease Control and Prevention (CDC) performs serum IgG EIA and immunoblot testing. Cross-reactivity with other species and trematodes may occur. 

Nucleic Acid Detection The conventional immunological diagnosis is sensitive in human paragonimiasis but unsustainable in e­ pidemiological surveys. A LAMP assay has successfully amplified the gene sequence of P. westermani eggs in sputum and pleural fluid from patients, as well as metacercariae in freshwater crabs and crayfish. LAMP demonstrates a detection limit of 1 ×

Fig. 56.4  Life cycle of the trematode, Paragonimus westermani, the causal agent of paragonimiasis. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

CHAPTER 56  Liver and Lung Trematodes

10-8 ng/μL, approximately 100 times more sensitive than PCR. The LAMP method also yields positive and negative results coinciding with those from parasitology tests, making it an excellent candidate for field surveys and clinical diagnoses of paragonimiasis. 

Treatment and Prevention The drug of choice for treatment of Paragonimus infections is praziquantel given three times a day for 2 to 3 consecutive days. Triclabendazole in a single- or two-dose regimen is also effective but is not available in the United States. Human infection can be prevented by not eating pickled, raw, or undercooked crabs or crayfish. Crustaceans should be cooked to an internal temperature of 145°F. Care should also be taken to properly clean utensils used in the preparation of these foods. Improvement of sanitary conditions and practices may also help to reduce the prevalence of these infections.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

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Bibliography Bogitsch BJ, Carter CE, Oeltmann TN, editors: Human parasitology, ed 3, San Diego, 2005, Academic Press. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Deng MH, Zhong LY, Kamolnetr O, et al.: Detection of helminths by loop-mediated isothermal amplification assay: a review of updated technology and future outlook, Infect Dis Poverty 8(1):20, 2019. Fried B, Abruzzi A: Food-borne trematode infections of humans in the United States, Parasitol Res 106:1263–1280, 2010. Garcia LS: Diagnostic medical parasitology, ed 6, Washington, DC, 2016, ASM Press. John DT, Petri WA: Markell’s and Voge’s medical parasitology, ed 9, St Louis, MO, 2006, Saunders. Keiser J, Utzinger J: Food-borne trematodiases, Clin Microbiol Rev 22:466–483, 2009. Shin SH, Hsu A, Chastain HM, et al.: Development of two FhSAP2 recombinant–based assays for immunodiagnosis of human chronic fascioliasis, Am J Trop Med Hyg 95(4):852–855, 2016. Slemenda SB, Maddison SE, Jong EC, et al.: Diagnosis of paragonimiasis by immunoblot, Am J Trop Med Hyg 39:469–471, 1988. Zarrin-Khameh N, Citron DR, Stager CE, et al.: Pulmonary paragonimiasis diagnosed by fine-needle aspiration biopsy, J Clin Microbiol 46:2137–2140, 2008.

CASE STUDY 56.1 A 50-year-old male visiting from Japan complained of fever, abdominal pain, and jaundice. A stool specimen was collected for ova and parasite examination. A blood specimen was also collected and sent to a clinical laboratory for testing. The blood work showed a slightly increased serum bilirubin level. The parasite examination showed small, oval, operculated eggs (approximately 80 μm × 45 μm), with opercular shoulders and a knob on the opposite end.

Questions 1. What parasite is the probable cause of this infection? 2. Another parasitic worm has eggs that are almost identical. What fact about the patient can aid in deciding which worm is responsible for his infection? 3. What other complications might be expected with this infection? 4. What would be the preferred treatment for this infection?

Chapter Review 1. Which of the following flukes has an operculated flaskshaped egg with prominent shoulders and a knob at the opposite end? a. Opisthorchis b. Paragonimus c. Fasciola d. Fasciolopsis 2. For which one of the following flukes would sputum be the diagnostic specimen? a. Opisthorchis b. Paragonimus c. Fasciola d. Clonorchis 3. The eggs of Fasciola hepatica are indistinguishable from those of: a. Clonorchis sinensis b. Heterophyes heterophyes c. Opisthorchis viverrini d. Fasciolopsis buski 4. Where might Clonorchis be found that Opisthorchis is not found? a. Vietnam b. Thailand c. Korea d. Cambodia

5. Infection with Clonorchis or Opisthorchis may result from eating raw or undercooked: a. Aquatic vegetation b. Crabs c. Crayfish d. Freshwater fish 6. What laboratory finding may be observed in Paragonimus infections? a. Increased number of eosinophils b. Decreased serum bilirubin level c. Decreased vitamin B12 level d. Increased number of red blood cells 7. The drug of choice for treatment of Fasciola hepatica infections is: a. Albendazole b. Triclabendazole c. Niclosamide d. Praziquantel

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Blood Trematodes OBJECTIVES 1. List the clinically significant blood trematodes. 2. Describe the general life cycle of the blood trematodes and how human infection occurs. 3. Explain the diagnostic methods used to identify blood trematodes. 4. Differentiate the eggs of the five species of schistosomes. 5. Describe the pathogenesis of the blood trematodes. 6. List the drugs of choice for treatment of blood trematode infections. 7. Describe the natural habitat for blood trematodes and how infection may be prevented.

PARASITES TO BE CONSIDERED Trematodes Blood

Schistosoma guineensis Schistosoma haematobium Schistosoma intercalatum Schistosoma japonicum Schistosoma malayensis Schistosoma mansoni Schistosoma mattheei Schistosoma mekongi Schistosoma spp.

T

here are five main species of blood flukes that are primarily associated with disease in humans (known as schistosomiasis, bilharziasis, or snail fever), all belonging to the genus Schistosoma. These species are Schistosoma haematobium, Schistosoma mekongi, Schistosoma intercalatum, Schistosoma japonicum (Oriental blood fluke), and Schistosoma mansoni. Seven additional species are rarely associated with human infection and have limited geographic distribution. The blood flukes differ in morphology and life cycle characteristics from the other trematodes. They are similar, however, by requiring a freshwater snail as the intermediate host.

General Characteristics Unlike the other trematodes, adult schistosomes are not flattened, but are rather long, thin, and rounded in shape. There is an oral sucker surrounding the mouth and a ventral sucker located just slightly below the oral sucker. The adult 786

male averages 1.5 cm in length and is wider than the female, having a ventral fold that wraps around the female when they mate (Fig. 57.1). The adult female averages 2 cm in length and is very thin. The eggs of each species are distinct and can be distinguished by size, spine morphology, and sometimes specimen type (Fig. 57.2). The size range for eggs of S. haematobium is 110 to 170 μm long by 40 to 70 μm wide, and they have a sharply pointed terminal spine. They are fully embryonated without an operculum. The size range for the eggs of S. japonicum is 70 to 100 μm long by 50 to 65 μm wide, and they have a small lateral spine that is sometimes difficult to detect (Fig. 57.3). S. mekongi eggs are smaller than those of S. japonicum, ranging in size from 50 to 65 μm long by 30 to 55 μm wide. They are fully embryonated without an operculum and have a small lateral spine. The size range for eggs of S. mansoni is 115 to 180 μm long by 40 to 75 μm wide, and they have a large lateral spine. S. mansoni eggs are inoperculate, immature when released, and take up to 8 to 10 days to develop a miracidium. S. intercalatum eggs are fully embryonated without an operculum, have a terminal spine, and range in size from 140 to 240 μm long by 50 to 85 μm wide. S. intercalatum eggs resemble those of S. haematobium and can be differentiated by Ziehl-Neelsen acid-fast positivity. In addition, S. intercalatum eggs are only found in feces, not in urine specimens. Table 57.1 provides a comparison of the schistosome eggs. One of the main differences between the schistosomes and other trematodes is that instead of being hermaphroditic, there are separate male and female adult worms. In human infection, the adult worms live in either the veins that supply the intestine (S. japonicum, S. intercalatum, S. mekongi, and S. mansoni) or the veins that supply the urinary bladder (S. haematobium). The eggs are passed from the body in either feces or urine. To reach the inside of the intestine or bladder, the eggs must penetrate the tissue from the veins. This is accomplished via a spine that is distinctive among the major species. The embryonated egg releases the miracidium. once it reaches fresh water, where it enters the snail host and develops into the infectious cercaria. The free-swimming cercariae are capable of penetrating through human skin and do not encyst on aquatic vegetation or within other aquatic wildlife. The cercariae penetrate host tissue until they reach a vein; then they travel to capillaries near the lungs and then to the portal vein of the liver, where

CHAPTER 57  Blood Trematodes

they mature. When they are mature, the adult males pair with females and then travel to veins of either the intestine or the bladder, where eggs are produced. 

Epidemiology Schistosomes have a worldwide distribution from Egypt and China to Africa and the Americas. S. haematobium is found in Africa and the Arabian Peninsula and has no reservoir hosts. S. mansoni and S. guineensis are found in Africa, the Arabian Peninsula, and Brazil. Reservoir hosts include wild rodents and marsupials. S. japonicum is found in China,

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Indonesia, and the Philippines. Many domestic animals (cats, dogs, cattle, horses, pigs) serve as reservoir hosts, as do some wild animals. S. mekongi primarily exists in the lower Mekong River basin in southern Laos, Cambodia, and Thailand. Reservoir hosts include dogs and domestic pigs. S. intercalatum and S. guineensis are primarily found in central and western Africa. Reservoirs include rodents, marsupials, and nonhuman primates. Human schistosome infection is caused by fecal (and urine) contamination of small bodies of water that favor the growth of the snail hosts. Infection with S. japonicum is especially prevalent in areas where humans work in rice paddies. Reports have indicated that S. haematobium is capable of cross-breeding with other species. This phenomenon has been identified between S. haematobium and S. bovis, as well as between S. guineensis and S. intercalatum. This is important when considering epidemiology and identification of these parasites 

Pathology and Spectrum of Disease

•

Fig. 57.1 This low-power photomicrograph reveals some of the ultrastructural morphology exhibited by coupled male and female Schistosoma mansoni parasites. Unlike the flukes, adult schistosomes are either male or female, with the female residing in a gynecophoral canal within the male. Male worms are robust, tuberculate, and measure 6 to 12 mm in length. Females are longer (7 to 17 mm in length) and slender. Adult S. mansoni reside in the venous plexuses of the colon and lower ileum and in the portal system of the liver of their host. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

A

Infection with only a small number of worms may be asymptomatic. A variety of species cause acute toxemic schistosomiasis resembling serum sickness. S. japonicum causes significant hepatointestinal disease resulting in portal hypertension and splenic and hepatic enlargement. S. intercalatum is primarily associated with rectal schistosomiasis in regions of Africa. S. haematobium is the only species that causes urinary schistosomiasis. Quite often, penetration of the skin by cercariae causes localized swelling and itching (cercarial dermatitis). The migration of the larvae through the body may cause transient symptoms of fever, malaise, cough (when they migrate in the lungs), or hepatitis (when in the liver). The adults are able to acquire some host antigens on their outer surface, and so may not elicit an immune response, although the eosinophil count

B • Fig. 57.2 (A) Schistosoma mansoni adult female; note the long pointed end (400× total magnification). (B)

Schistosoma mansoni adult male. Males are shorter and wider to accommodate the insertion of the female during copulation. (400× total magnification.)

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may be high. Severe tissue damage, with associated pain, fever, and chills, may occur when the eggs travel through tissue to reach the intestines or bladder. There may also be bloody diarrhea or blood in the urine (hematuria). Necrosis, lesions, and granulomas may develop, as well as obstruction

of the bowel or ureters. Urinary schistosomiasis may give rise to calcifications in the bladder and renal failure. S. intercalatum and S. guineensis tend to produce milder symptoms. Eggs are primarily deposited in the colon, resulting in blood and mucus in the stool.

• Fig. 57.3  This is an illustration depicting the life cycle of flatworms of the genus Schistosoma, the causal

agents of the parasitic disease schistosomiasis. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

TABLE 57.1    Diagnostic Characteristics of the Blood Trematodes

Blood Trematode

Adult Location

Size of Egg

Description of Egg

Schistosoma haematobium

Veins surrounding bladder

110–170 μm × 40–70 μm

Pointed terminal spine, no operculum, embryonated

Schistosoma intercalatum/guineensis

Venules of colon

140–240 μm × 50–85 μm

Resembles egg of S. haematobium, but acidfast positive

Schistosoma japonicum

Venules of small intestine

70–100 μm × 50–65 μm

Small lateral spine, no operculum, embryonated

Schistosoma mansoni

Venules of large intestine

115–180 μm × 40–75 μm

Large lateral spine, no operculum, embryonated

Schistosoma mekongi

Venules of small intestine

50–65 μm × 30–55 μm

Resembles egg of S. mansoni, but much smaller

CHAPTER 57  Blood Trematodes

A

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B

• Fig. 57.4 (A) Schistosoma mansoni egg. (B) Schistosoma japoniC

Penetration of human skin by the cercariae of blood flukes that commonly infect other mammals or aquatic birds may cause a schistosomal dermatitis known as “swimmer’s itch.” Erythema, edema, and intense itching may develop that usually disappear within 1 week. The cercariae of these species are not able to complete their life cycle by entering the human bloodstream and are destroyed by the host immune system. 

Laboratory Diagnosis The standard method of diagnosis is by the detection of characteristic eggs in feces or rectal biopsy for S. japonicum, S. mekongi, S. mansoni, S. intercalatum, and S. guineensis (and perhaps S. haematobium if these worms have migrated to a bladder vein that is close to the intestine) and in urine (usually concentrated before examination) or bladder tissue biopsy for S. haematobium. A wet mount with or without iodine from a sedimentation or concentration method can be examined for eggs. Fig. 57.4A–C provides images of three different schistosome eggs. To optimize recovery of S. haematobium in urine, the specimen should be collected between noon and 2 p.m.

Antigen Detection Two circulating schistosome antigens, the anodic and cathodic antigens, can be detected in the urine of patients

cum egg. The arrowhead indicates the presence of the small lateral spine. (C) Schistosoma haematobium egg. (Courtesy Division of Parasitic Diseases/Centers for Disease Control and Prevention.)

with schistosomiasis. A lateral flow rapid detection kit is available for the circulating cathodic antigen and is commercially available. This rapid diagnostic test (RDT) has been used widely in Africa and Brazil but is not FDA approved. Another flow assay is under development for the detection of S. japonicum and S. haematobium. 

Serologic Testing There are serologic assays that are available for diagnosis of schistosomal IgG antibody (enzyme immunoassay [EIA], enzyme-linked immunosorbent assay [ELISA], and immunoblot), but these methods cannot distinguish between current and previous infections. This type of assay may, however, be useful for travelers who have returned from endemic areas. These assays lack sensitivity and specificity during the early stages of disease, or for detecting persistence of antibodies after treatment for schistosomiasis. These assays are performed at the Division of Parasitic Disease at the Centers for Disease Control and Prevention (CDC) and may be available at some private reference laboratories. Several nucleic acid–based methods have been developed that demonstrate high sensitivity and specificity using genomic or mitochondrial sequences. In addition, schistosome DNA has been identified in plasma using real-time polymerase chain

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reaction (PCR). Generally, an etiological method or a simple PCR method cannot distinguish between the species of Schistosoma. Repetitive sequences utilizing nested PCR to identify genes located within the parasitic chromosomes aid in detecting schistosomiasis. Various highly repetitive sequences have been successfully used to identify schistosomiasis: SjR2 for S. japonicum, SM1-7 for S. mansoni, and the Dra I for S. haematobium. In addition, the ribosomal RNA sequences are highly repetitive and conserved in the DNA of schistosomes. Both the 18S and 28S rDNA sequences have been used to detect parasite-derived DNA in fecal, serum, and urine samples in either monoplex or multiplex real-time PCR assays. Eukaryotic organisms, such as schistosomes, contain mitochondria in their cells. Mitochondria contain their own circular DNA molecule and replicate independently of the cell nucleus in a eukaryotic cell. A novel real-time PCR technique was developed that can identify the nicotinamide adenine dinucleotide hydrogen (NADH) 1 and 2, nad1 and nad2, specific for S. japonicum. This method successfully identified S. japonicum DNA in human stool samples with low-intensity infections. Another PCR assay has been used to detect the nad3 in urine samples of patients infected with schistosomes. This approach can accurately differentiate S. haematobium from other schistosomes. Additional genes commonly used for identification of S. japonicum and S. mansoni include cox2 and nad6. A multiplex PCR assay for the cytochrome C oxidase subunit I, cox1, of several human schistosomes was developed that can differentiate S. mansoni, S. haematobium, S. japonicum, and S. mekongi infections. Nucleic acid–based tests are highly sensitive and specific; however, costs associated with instrumentation and

Bibliography Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Bogitsch BJ, Carter CE, Oeltmann TN, editors: Human parasitology, ed 3, San Diego, 2005, Academic Press. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Garcia LS: Diagnostic medical parasitology, ed 6, Washington, DC, 2016, ASM Press. Gobert GN, Chai M, Duke M, et al.: Copro-PCR-based detection of Schistoma eggs using mitochondrial DNA markers, Mol Cell Probes 19:250–254, 2005. Gryseels B, Polman K, Clerinx J, et  al.: Human schistosomiasis, Lancet 368:1106–1118, 2006. He P, Song LG, Xie H, et al.: Nucleic acid detection in the diagnosis and prevention of schistosomiasis, Infect Dis Poverty 5:25, 2016. https://doi.org/10.1186/s40249-016-0116-y. John DT, Petri WA: Markell’s and Voge’s medical parasitology, ed 9, St Louis, MO, 2006, Saunders.

reagents in endemic areas limit use. Improved methods and tools are needed for broad implementation. Recent reports have shown that loop-mediated isothermal amplification (LAMP), a highly sensitive amplification technique that does not require thermal cycling (and its associated costs and equipment), can detect parasite DNA in a variety of platyhelminth- and nematode-infected hosts, including those with schistosome infections. 

Therapy The drug of choice for treatment of schistosome infections is praziquantel, given in two or three doses daily. 

Prevention Because human infection is through direct penetration by the cercariae, prevention of schistosome infection is more difficult to achieve. Educational programs are required to help people in endemic areas understand how to help prevent infection. Sanitary conditions need to be improved, with proper disposal not only of human waste but also that of domestic animals (in areas with S. japonicum and S. mekongi). A safe water supply for bathing and washing clothes is also necessary. Various snail control methods have been implemented in endemic regions, but these methods are very costly and would need to be repeated on a regular basis to have the desired effect.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Pontes LA, Oliveira MC, Dias-Neto E, et al.: Comparison of a polymerase chain reaction and the Kato-Katz technique for diagnosing infection with Schistosoma mansoni, Am J Trop Med Hyg 68:652– 656, 2003. Song J, Liu C, Bais S, Mauk MG, Bau HH, Greenberg RM: Molecular detection of schistosome infections with a disposable microfluidic cassette, PLoS Negl Trop Dis 9(12):e0004318, 2015. https://doi. org/10.1371/journal.pntd.0004318. Urbani C, Sinoun M, Socheat D, et al.: Epidemiology and control of mekongi schistosomiasis, Acta Trop 82:157–168, 2002. Van Dijk K, Starink MV, Bait A, et al.: The potential of molecular diagnosis of cutaneous ectopic schistosomiases, Am J Trop Med Hyg 83:958–959, 2010. Wichmann D, Panning M, Quack T, et al.: Diagnosing schistosomiasis by detection of cell-free parasite DNA in human plasma, PLoS Negl Trop Dis 3:e422, 2009. Wilson M, Schantz PM, Nutman T, editors: Molecular and immunological approaches for diagnosis of parasitic infection, ed 7, Washington, DC, 2006, ASM Press.

CASE STUDY 57.1 An 18-year-old male had recently been on a 1-month long trip to Brazil with a group of volunteer workers, where he had enjoyed swimming in a nearby river with the local teenage volunteers. He complained of crampy abdominal pain and twice noticed a small amount of blood in his feces. A stool specimen was collected for ova and parasite examination. A blood specimen was also collected and sent to a clinical laboratory for testing. The results of the blood tests showed increased eosinophil and IgE levels. The parasite examination showed large (115 μm × 75 μm), inoperculate, oval eggs with large lateral spines.

Questions 1. What parasite is the probable cause of this infection? 2. Where in the human host would the adults of this parasite be found? 3. What recent activity of this patient is probably responsible for his infection? 4. What is the drug of choice for treatment of infection with this parasite?

  

Chapter Review 1. The mode of transmission of schistosomal infection is by: a. Ingestion of contaminated aquatic vegetation b. Direct penetration of the skin by cercariae c. Ingestion of raw fish d. Mosquito bite 2. A diagnostic characteristic of the egg of S. mansoni is: a. A large lateral spine b. No spine c. A pointed terminal spine d. A small lateral spine 3. Infection with S. haematobium may present with which of the following? a. Nausea b. Basophilia c. Hematuria d. Jaundice 4. The drug of choice for treatment of schistosome infections is: a. Metrifonate b. Praziquantel c. Bilarcil d. Niclosamide

5. Which of the following is a major contributing factor to infection with S. japonicum? a. Absence of animal reservoir hosts b. Easily controlled snail population c. Easily passed from person-to-person d. Large number of humans working in rice paddies 6. Matching: Match the location in the body with the adult schistosome worm: _____ S. mansoni _____ S. haematobium _____ S. japonicum _____ S. mekongi _____ S. intercalatum

a. veins surrounding the bladder b. venules of the small intestine c. venules of the large intestine d. venules of the colon

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PART V  Mycology

58

Overview of Fungal Identification Methods and Strategies OBJECTIVES 1. Define the terms mycology; saprophytic; dermatophyte; and polymorphic, dimorphic, and thermally dimorphic fungi. 2. Define and differentiate superficial, cutaneous, subcutaneous, and systemic mycoses, including the tissues involved. 3. Differentiate the colonial morphology of yeasts and filamentous fungi (molds). 4. Define and differentiate anamorph, teleomorph, and synanamorph. 5. Describe three ways in which fungi reproduce. 6. List the media used for optimal recovery of fungi, including their incubation requirements. 7. List the common antibacterial agents used in fungal media. 8. Explain and differentiate the characteristic colonial morphology of fungi, including topography (rugose, umbonate, verrucose), texture (cottony, velvety, glabrous, granular, wooly) and surface described (front, reverse). 9. Describe and differentiate the sexual and asexual reproduction of the Ascomycota. 10. Define and differentiate rapid, intermediate, and slow growth rates with regard to fungal reproduction and cultivation. 11. Describe the proper method of specimen collection for fungal cultures, including collection site, acceptability, processing, transport, and storage. 12. Give the advantages and disadvantages of using screwcapped culture tubes, compared with agar plates, in the laboratory. 13. Describe the chemical principle and methodologies used to identify fungi, including calcofluor white–potassium hydroxide preparations, hair perforation, cellophane (Scotch) tape preparations, saline/wet mounts, lactophenol cotton or aniline blue, potassium hydroxide, Gram stain, India ink, modified acid-fast stain, periodic acid–Schiff stain (PAS), Wright’s stain, Papanicolaou stain, Grocott-Gomori methenamine silver (GMS), hematoxylin and eosin (H&E) stain, Masson-Fontana stain, tease mount, and microslide culture.

M

ycology is a specialized discipline in the field of biology concerned with the study of fungi, including their taxonomy, environmental impact, and genetic and biochemical properties. These microorganisms are recognized as important causes of disease, from superficial infections to those that are life threatening and rapidly fatal. Because of the change in patient profiles, particularly the increase in immunocompromised individuals and the increased use of antifungals, a number of fungal species normally found in the environment have been recognized as important causes of human disease. In addition, the implementation of genome-sequencing technology and the ability to identify polymerase chain reaction (PCR)generated amplicons from nonculturable clinical isolates continually identifies new organisms and increases the understanding of the diversity of fungal infections associated with human disease. It is estimated that more than one billion people worldwide suffer from fungal infections. The modern clinical laboratory, therefore, must provide methods for isolating and identifying the causes of fungal disease. Susceptibility testing of these isolates may also be necessary. This chapter is designed to assist clinical laboratory professionals and microbiologists with the basics of diagnostic clinical mycology and is not considered inclusive. The field of clinical mycology is rapidly evolving with the implementation of new molecular methods for classification and identification. Therefore, it is often necessary to consult other references for more detailed information within this vast field of study.

Epidemiology Fungal infections are an increasing threat to individuals. The number of nosocomial, health care–associated, and community-associated infections has increased dramatically. The major factors responsible for the increase in the number of fungal infections are alterations in the host, particularly

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the growing number of immunocompromised individuals and the increasing use of antifungal medications. Whether associated with the use of antifungal medications, immunosuppressive agents, or serious underlying diseases, these factors may lead to infection by organisms normally nonpathogenic or part of the patient’s normal microbiota. These infections may occur in patients with debilitating diseases, such as progressive infection with the human immunodeficiency virus (HIV) or diabetes mellitus or in patients with impaired immunologic function resulting from corticosteroid or antimetabolite chemotherapy. Other common predisposing factors include complex surgical procedures and antibacterial therapy. More than 135,000 valid species of fungi exist, but it is estimated that the number of undiscovered species ranges from 1 to 10 million with about 1000 to 1500 new species identified annually. With the implementation of molecular technology, the number of clinically relevant fungi and pathogenic species will continue to grow. Many of these organisms normally live a saprophytic existence (living on dead or decayed organic matter) in nature and persist in the environment. Fungal infections generally are not communicable in the usual sense, through person-to-person transmission. Humans become accidental hosts for fungi by inhaling spores or through the introduction of fungal elements into tissue by trauma. Except for disease caused by the dimorphic fungi, healthy individuals are relatively resistant to most infections caused by fungi. Classic infections are now appearing in new forms in patients, and the old “harmless” saprophytic molds are implicated in serious diseases. This ability of normally saprophytic fungi to cause disease in patients means that laboratories must be able to identify and report a wide array of fungi. The primary fungal pathogens appear to have welldefined geographic locations. An example of this is the dimorphic fungi Coccidioides spp. Coccidioides is usually found only in the Southwest United States in the desert, northern Mexico, and Central America. Opportunistic pathogens such as Candida and Aspergillus spp. are found all over the world. 

General Features of the Fungi Fungi seen in the clinical laboratory generally can be categorized into two groups based on the appearance of the colonies formed. The yeasts are unicellular organisms (Chapter 62) and produce moist, creamy, opaque, or pasty colonies on media, whereas the filamentous fungi or molds produce multicellular structures (Chapters 59 and 60) and demonstrate fluffy, cottony, woolly, or powdery colonies. Several systemic fungal pathogens that exhibit either a yeast or yeastlike phase and filamentous forms are referred to as dimorphic. When dimorphism is temperature-dependent, the fungi are designated as thermally dimorphic. In general, these fungi produce a mold form in the environment or when cultured on routine artificial mycology agar at 25°C

to 30°C and are yeastlike in tissue or when cultured on enriched artificial medium at 35°C to 37°C. The medically important dimorphic fungi are Histoplasma sp., Blastomyces spp., Coccidioides spp., Cokeromyces recurvatus, Emergomyces spp., and Paracoccidioides spp. Additionally, some of the medically important yeasts, particularly the Candida species, may produce yeast forms, pseudohyphae, and/or true hyphae (Chapter 62). Fungi that have more than one independent form or spore stage in their life cycle are called polymorphic fungi. The polymorphic features of this group of organisms are not temperature dependent. 

Taxonomy of the Fungi Fungi are composed of a vast array of organisms that are unique compared with plants and animals. They are heterotrophic (saprophytic) and require preformed organic carbon for nutrition. Included among these are the mushrooms, rusts and smuts, molds and mildews, and yeasts. Despite their great variation in morphologic features, most fungi share the following characteristics: • Chitin in the cell wall • Ergosterol in the cell membrane • Reproduction by means of spores, produced asexually or sexually • Lack of chlorophyll • Lack of susceptibility to antibacterial antibiotics Significant changes have resulted in the organization of the groups, taxonomy, and nomenclature within the fungi due to the introduction of molecular methods. Traditionally, the fungi have been categorized based on phenotypic traits, which vary based on temperature, atmospheric conditions, nutrient availability, and humidity, to name a few. Because of this variation, phenotypic classification is unclear, complex, and variable. Molecular analysis of these organisms, which includes extensive DNA sequencing, is now considered the gold standard to determine taxonomic designation and classification. In addition, fungal taxonomy has been complicated by fungi names that describe the sexual (teleomorph) or asexual (anamorph) stage. This system has become obsolete. The International Botanical Congress adopted a one-fungus, one-name policy, published in the International Code of Nomenclature, Article 59. The implementation of DNA sequencing for the classification and taxonomy of fungi is not without challenges. As with other microorganisms, fungi have the ability to exchange genetic material and demonstrate evidence of recombination among populations, making speciation difficult. Currently, no standard exists that provides for a definition of the amount of genetic diversity that is allowable, or that limits the species level in taxonomy of fungi. Some clinically relevant fungi have been identified that include molecular siblings. A molecular sibling is an organism that cannot be distinguished based on phenotypic, metabolic, and clinical presentation but is recognized as a different

CHAPTER 58  Overview of Fungal Identification Methods and Strategies

• Fig. 58.1  A cleistothecium of Pseudallescheria boydii that has opened and is releasing numerous ascospores (×750).

• Fig. 58.2  Scedosporium apiospermum showing asexually produced conidia borne singly on conidiophores (anellophores [arrows]) (×430).

species based on molecular analysis. Molecular siblings that do not differ in traditional characteristics, including disease presentation, are often considered a species complex. The use of the term “complex” does not hold any taxonomic relevance; it provides a system to limit organism identification within the clinical laboratory. Clinical laboratories may choose to report species complexes as preliminary

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• Fig. 58.3  Anamorph form of Pseudallescheria boydii (×500).

identifications until further studies can be performed on the isolate, including antifungal susceptibility profiles when appropriate. Historically, the fungi were categorized into three wellestablished phyla: Zygomycota, Ascomycota, and Basidiomycota. The previous phylum, Zygomycota, contained a very diverse group of organisms. Zygomycota has now been replaced with the subphyla Mucoromycotina and Entomophthoromycotina. The subphyla Mucoromycotina includes the order Mucorales and the genera Lichtheimia, Mucor, Rhizomucor, and Rhizopus; the Entomophthoromycotina, order Entomophthorales, includes the genera Basidiobolus and Conidiobolus. These two subphyla include organisms that produce aseptate or sparsely septate hyphae and exhibit asexual reproduction by sporangiospores. Some species may reproduce sexually and form zygospores in culture. The Ascomycota include many fungi that reproduce asexually by the formation of conidia (asexual spores) and sexually by the production of ascospores. The filamentous ascomycetes are ubiquitous in nature and produce true septate hyphae. They may exhibit a sexual form (teleomorph) but also exist in an asexual form (anamorph) (Figs. 58.1 to 58.3). Fungi that have different asexual forms of the same fungus are called synanamorphs. An example of a clinically important fungus that belongs to the phylum Ascomycota is Histoplasma capsulatum. Numerous opportunistic fungi such as Aspergillus, the atypical fungi, Pneumocystis, and yeast such as the Saccharomyces, Saprochaete, and Candida, also belong to the Ascomycota. The phylum Basidiomycota includes fungi that reproduce sexually through the formation of basidiospores on a specialized structure called the basidia. Medically relevant

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basidiomycetous yeasts include the genera Cryptococcus, Malassezia, Pseudozyma, Rhodotorula, Sporobolomyces, and Trichosporon. The filamentous Basidiomycetes are uncommon causes of human respiratory and systemic disease in immunocompetent and immunosuppressed patients and are poorly characterized. In addition, several organisms that are pathogenic to humans and have been classified as fungal or parafungal organisms that phenotypically resemble protozoans or yeast and remain unculturable are included in Chapter 61. 

Examples of subcutaneous infections include chromoblastomycosis, mycetoma, and phaeohyphomycotic cysts (Chapter 60). Some of the agents of systemic fungal infections include the genera Blastomyces, Coccidioides, Histoplasma, Cokeromyces, Talaromyces, Sporothrix, Emergomyces, and Paracoccidioides. Infections caused by most of these organisms involve the lungs but also may become widely disseminated and involve any organ system. Any of the fungi could be considered an opportunistic pathogen in the appropriate clinical setting. The list of uncommon fungi found to cause disease in humans expands every year. Fungi previously thought to be nonpathogenic may be the cause of infections. The infections these organisms cause occur primarily in patients with some type of compromise of the immune system. This may occur secondary to an underlying disease process, such as diabetes mellitus, or prolonged use of an immunosuppressive agent. Although any fungus potentially can cause disease in these patients, the most commonly encountered genera in this group are Aspergillus, Candida, and Cryptococcus, among others. All of these organisms may cause disseminated (systemic) disease. Some of the dematiaceous fungi may cause deeply invasive phaeohyphomycoses (i.e., produce brownpigmented structures) in this patient population. Classification by type of infection allows the clinician to attempt to categorize organisms in a logical fashion into groups by clinical relevance. Table 58.1 presents an example of a classification of infections and their etiologic agents that is useful to clinicians. The identification of organisms in clinical samples requires examination of the macroscopic and microscopic characteristics of the fungal culture. Although

Clinical Classification of the Fungi The complexity associated with the taxonomic classifications of fungi makes the identification of clinically relevant organisms to the species level inherently difficult. For clinicians, dividing the fungi into four categories of mycoses according to the type of infection in combination with macroscopic and microscopic characteristics is used to provide a systematic approach to identification. The clinical categories of fungi are separated as follows: • S  uperficial (cutaneous) mycoses • S  ubcutaneous mycoses • S  ystemic mycoses • O  pportunistic mycoses The superficial, or cutaneous, mycoses are fungal infections that involve hair, skin, or nails without direct invasion of deeper tissue. The fungi in this category include the dermatophytes (agents of ringworm, athlete’s foot) and agents of infections such as tinea, tinea nigra, and piedra. All of these infect keratinized tissues. Some fungi cause infections that are confined to the subcutaneous tissue without dissemination to distant sites. TABLE 58.1    General Clinical Classification of Pathogenic Fungi

Cutaneous

Subcutaneous

Opportunistic

Systemic

Superficial mycoses Tinea Piedra Candidosis Dermatophytosis

Chromoblastomycosis Sporotrichosis Mycetoma (eumycotic) Phaeohyphomycosis

Aspergillosis Candidosis Cryptococcosis Geotrichosis Mucormycosis Fusariosis Trichosporonosis Entomophthoromycosis Othersa

Aspergillosis Blastomycosis Candidosis Coccidioidomycosis Adiaspiromycosis Emmonsiosis Histoplasmosis Cryptococcosis Geotrichosis Paracoccidioidomycosis Penicilliosis Pneumocystosis Sporotrichosis Pseudoallescheriosis/scedosporiosis Mucormycosis Entomophthoromycosis Fusariosis Trichosporonosis

aVirtually

any fungus may cause disease including systemic infection in a profoundly immunocompromised host.

CHAPTER 58  Overview of Fungal Identification Methods and Strategies

the macroscopic characteristics may be suggestive of particular genera of fungi, the microscopic characteristics tend to provide a more reliable method for identification. This requires a microscopic examination of the isolate for the appearance of hyphal elements, including the presence, absence, and number of septa. If the hyphae appear to be broad and predominantly nonseptate (i.e., cells are not separated by a septum or wall), Mucoromycetes, Basidiobolomycoses, or Entomophthoromycoses should be considered. If the hyphae are septate, they must be examined further for the presence or absence of pigmentation. If a dark pigment is present in the hyphae, the organism is considered to be melanized (dematiaceous), and the conidia are then examined for their morphologic features and their arrangement on the hyphae. If the hyphae are nonpigmented, they are considered to be hyaline. The fungi are then examined for the type and the arrangement of the conidia produced. The molds are identified by recognition of their characteristic microscopic features. It is important that the clinical laboratory evaluate the isolation of a fungus from a clinical sample, because it may be regarded as a contaminant acquired during collection, transportation, processing, or incubation. It is important to consider that all contaminating fungi may be considered pathogenic in the appropriate clinical setting. Each isolate should be carefully evaluated on a caseby-case basis. 

Pathogenesis and Spectrum of Disease Fungal infection is caused by either primary pathogens or opportunistic pathogens. Infections caused by primary pathogens can occur in immunocompetent hosts, are not always as virulent, and may lead to subclinical disease. Opportunistic pathogens primarily infect immunocompromised hosts. Opportunistic pathogens include almost any fungus present in the environment. An increase in the identification of opportunistic fungal infections in humans is in large part a result of the immunocompromised nature of the host but also the increasing use of antifungals and improved diagnostic methods. In addition, organism-specific factors, called virulence factors, make invading tissues and causing disease easier. Some virulence factors include: • The organism’s size (with inhalation, the organism must be small enough to reach the alveoli) • The organism’s ability to grow at 37°C at a neutral pH • Conversion of the dimorphic fungi from the mycelial (mold) form into the corresponding yeast or spherule form in the host • Toxin production Most of the fungi exist in environmental niches as saprophytic organisms (Table 58.2). Perhaps the fungi that cause disease in humans have developed various mechanisms that allow them to establish disease in the human host. Table 58.3 describes some of the known or speculative virulence factors of the fungi known to be pathogenic for humans. 

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Laboratory Diagnosis Collection, Transport, and Culturing of Clinical Specimens The diagnosis of fungal infections depends entirely on the selection and collection of an appropriate clinical specimen for microscopic analysis and culture. Many fungal infections are similar clinically to mycobacterial infections, and often the same specimen is cultured for both fungi and mycobacteria. If a fungal clinical infection is suspected, the sample should always be cultured on fungal media. If the specimen quantity is insufficient for microscopy and culture, the culture should be performed. Many infections have a primary focus in the lungs; respiratory tract secretions are usually included among the specimens selected for culture. It should be emphasized that dissemination to distant body sites may occur, and fungi may be recovered from nonrespiratory sites. Proper collection of specimens and rapid transport to the clinical laboratory are crucial to the recovery of fungi. Specimens often contain not only the etiologic agent but also contaminating bacteria or fungi that rapidly overgrow some of the slower-growing pathogenic fungi. The viability of fungi decreases over time. Specimens should be processed within 2 hours of receipt. Most fungal specimens can be maintained at room temperature. If processing of intravascular catheter tips, other medical devices (stents, surgical implants, replacement joints, etc.), lower respiratory tract or urine specimens will be delayed, the samples can be refrigerated for a short time. Dermatologic specimens (skin, hair, nails) are very sensitive to cold temperatures. Yeasts (e.g., Candida spp.) commonly are recovered on routine bacteriology media and fungal culture media. A few specific comments concerning specimen collection and culturing are discussed later in this chapter. Additional information on specimen collection is included in Table 5.1.

Lower Respiratory Tract Secretions Respiratory tract secretions (sputum, induced sputum, bronchial washings, bronchoalveolar lavage, and tracheal aspirations) are perhaps the most common specimens collected for fungal culture. Viscous lower respiratory tract specimens should be pretreated with a mucolytic agent and concentrated by centrifugation. The sediment should then be plated to media without antibiotic and a media containing antibacterial antibiotics to prevent overgrowth by contaminants and ensure optimal recover of fungi. The antifungal agent cycloheximide prevents overgrowth by rapidly growing molds and should be included in at least one of the culture media. As much specimen as possible (0.5 mL) should be used to inoculate each medium. Two fungal organisms commonly isolated from cystic fibrosis (CF) patients include Candida spp. and Scedosporium spp. Therefore, it is recommended that selective and differential chromogenic medium for the isolation of Candida spp. and Scedosporium-selective medium containing antifungal

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TABLE 58.2    Summary of Common Pathogens

Organism

Natural Habitat

Infectious Form

Mode of Transmission

Common Sites of Infection

Aspergillus spp.

Ubiquitous, plants

Conidia

Inhalation

Lungs, eyes, skin, nails

Hyphae

Blastomyces spp.

Unknown, possibly soil/wood in moist areas

Probably conidia

Usually inhalation

Lungs, skin, long bones

Yeast

Candida spp.

Human microbiota

Yeast, pseudohyphae, and true hyphae

Direct invasion/dissemination

GI and GU tracts, nails, viscera, blood

Yeast, pseudohyphae, and true hyphae

Coccidioides spp.

Soil of many arid regions

Arthroconidia

Inhalation

Lungs, skin, meninges

Spherules, endospores

Cryptococcus spp.

Bird feces, soil

Yeasta

Inhalation

Lungs, skin, meninges

Yeast

Histoplasma capsulatum

Bat and bird feces

Conidia

Inhalation

Lungs, bone marrow, blood

Yeast

Paracoccidioides spp.

Possibly soil, plants

Conidia

Inhalation/trauma

Lungs, skin, mucous membranes

Yeast

Sporothrix spp.

Soil, plants

Conidia/hyphae

Trauma/rarely inhalation

Skin and lymphatics, lungs, meninges

Yeast

Dermatophytes

Human disease, animals, soil

Conidia/hyphae

Contact

Skin, hair, or nails

Hyphae

Clinical Form

aPossibly

the conidia of the teleomorphic stage. GI, Gastrointestinal; GU, genitourinary.

agents dicloran and benomyl should be added to respiratory cultures of CF patients to improve the isolation of these organisms. Specimens may be stored at room temperature if processing is completed within 2 hours; if processing is delayed, specimens should be refrigerated at 4°C. 

Sterile Body Fluids Including Cerebrospinal Fluid Most sterile body fluids are generally collected in heparin blood tubes to prevent clotting. Lysis centrifugation tubes may also be used for the collection of sterile body fluids. Cerebrospinal fluid lumbar puncture tubes collected for culture should be concentrated by centrifugation and the concentrated sediment used to inoculate the culture medium. Cultures should be examined daily. It is recommended that ≥2 mL of specimen should be centrifuged and up to 0.5 mL of sample be inoculated onto each type of fungal culture medium. If less than 1 mL of specimen is submitted for culture, following centrifugation, 1-drop aliquots of the sediment should be placed on several areas on the agar surface. Many laboratories utilize screw-cap tubes containing slants or culture vials to avoid contamination of fungal cultures. Media used for the recovery of fungi from

sterile fluids should contain no antibacterial or antifungal agents. Cryptococcus spp., a pathogenic encapsulated yeast associated with meningitis, is inhibited by the antifungal agent cycloheximide. Once submitted to the laboratory, sterile body fluid specimens should be processed promptly. If prompt processing is not possible, samples should be kept at room temperature. Sterile body fluid specimens should never be refrigerated.

Blood and Bone Marrow Disseminated fungal infections are a major cause of morbidity and mortality in hospitalized patients, and blood cultures provide an accurate method for determining the cause in many instances. Currently several automated blood culture systems that utilize fungal medium modifications for the isolation of fungi, including the BACTEC (Becton Dickinson, Sparks, MD), BacT/ALERT 3D (bioMérieux, Durham, NC), and VersaTREK (Thermo Scientific, Oakwood Village, OH), are adequate systems for the recovery of yeasts, except Malassezia spp. Laboratories that frequently recover dimorphic fungi and molds from blood or bone marrow are encouraged to use the lysis-centrifugation system, the Isolator. A heparinized syringe or pediatric Isolator tube

CHAPTER 58  Overview of Fungal Identification Methods and Strategies

TABLE   Virulence Factors of Medically Important 58.3  Fungi

Fungal Pathogen

Putative Virulence Factor

Aspergillus spp.

Elastase-serine protease Proteases Toxins (gliotoxin, fumagillin, helvolic acid) Elastase-metalloprotease Aspartic acid proteinase Aflatoxin Catalase Lysine biosynthesis p-aminobenzoic acid synthesis

Blastomyces spp.

Cell wall alpha-1,3-glucan BAD-1 an adhesion and immune modulator

Coccidioides spp.

Extracellular proteinases

Cryptococcus spp.

Capsule Phenoloxidase melanin synthesis Varietal differences

Dematiaceous fungi

Phenoloxidase melanin synthesis

Histoplasma capsulatum

Cell wall alpha-1,3-glucan Intracellular growth Thermotolerance CBP, binds calcium

Paracoccidioides spp.

Estrogen-binding proteins Cell wall components Beta-glucan Alpha-1,3-glucan

Sporothrix spp.

Thermotolerance Extracellular enzymes

is recommended for the collection of bone marrow samples. The Isolator has been proven optimal for the recovery of H. capsulatum and other filamentous fungi. Special automated fungal media such as the BACTEC MYCO/F lytic medium or BacT/ALERT media may also be used for the isolation of filamentous molds from blood samples. With this system, red blood cells and white blood cells, which may contain the microorganisms, are lysed, and centrifugation concentrates the organisms before culturing. The concentrate is inoculated onto the surface of appropriate culture media, and most fungi are detected within the first 4 days of incubation. However, occasional isolates of H. capsulatum may require approximately 10 to 14 days for recovery. In addition to the lysis-centrifugation system, special automated fungal media such as the BACTEC MYCO/F lytic medium or BacT/ALERT media may also be used for the isolation of filamentous molds to improve the recovery from blood. Bone marrow samples should not be placed in automated blood culture media. The optimal temperature for fungal blood cultures is 30°C, and the suggested incubation time is 21 days. 

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Eye (Corneal Scrapings or Vitreous Humor) Corneal scrapings collected by a physician should be placed directly onto microscopic slides and inoculated onto noninhibitory media such as Sabouraud dextrose agar, in either an X- or C-shaped pattern. Vitreous humor aspiration should be concentrated by centrifugation, similar to processing a cerebrospinal fluid (CSF) sample, and the sediment should be used for smears and culture. Specimens should be inoculated onto a noninhibitory media, inhibitory mold agar, and brain-heart infusion (BHI) agar with 10% sheep blood. Samples should be processed as soon as possible and stored at room temperature. Media containing cycloheximide should be avoided to prevent inhibition of potential isolates. 

Hair, Skin, and Nail Scrapings Specimens of hair, skin scrapings or biopsies, and nail clippings are usually submitted for dermatophyte culture and are contaminated with bacteria or rapidly growing fungi or both. Samples collected from lesions may be obtained by scraping the skin or nails with a scalpel blade or microscope slide; infected hairs are removed by plucking them with forceps. Only the leading edge of skin lesions should be sampled, because the centers often contain nonviable organisms. These specimens should be placed in a sterile container; they should not be refrigerated. Hair samples may be cut into 1 mm pieces and applied to the medium using a sterile forceps. Skin and nail samples should be cut into smaller pieces, placed on the medium using a sterile forceps and pressed slightly into the agar. Mycosel agar, which contains chloramphenicol and cycloheximide, is satisfactory for the recovery of dermatophytes. If infection with Malassezia furfur is suspected, olive oil or an olive oil–saturated paper disk should be placed in the first quadrant of the agar plate. Cultures should be incubated for a minimum of 21 days at 30°C before being reported as negative. 

Vaginal Candida spp. are considered normal vaginal flora, and therefore, identification without symptoms is not significant. Vaginal lesions may also be present with histoplasmosis or paracoccidioidomycosis. Vaginal samples should be transported to the laboratory within 24 hours of collection using culture transport swabs. Swabs should be kept moist in sterile tubes. This method of collection provides a specimen suitable for a wet preparation. Both selective and inhibitory agars should be plated. Vaginal cultures should be screened for yeasts using chromogenic agars for Candida spp. 

Urine First morning, clean-catch, suprapubic or direct catheterized urine samples are recommended for fungal culture. Urine samples should be processed as soon after collection as possible. The 24-hour urine and Foley catheter samples are unacceptable for culture. Quantitative cultures are not useful. All urine samples should be centrifuged and the sediment cultured using a loop to provide adequate isolation of colonies. Because urine often is contaminated with gram-negative

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bacteria, media containing antibacterial agents must be used to ensure the recovery of fungi. If processing is completed within 2 hours, samples may remain at room temperature. However, if processing is delayed, specimens should be refrigerated at 4°C. If a urine transport system is used, samples may be stored at room temperature for up to 72 hours. 

Tissue All tissues should be processed before culturing by mincing; it is critical not to grind them. Grinding will disrupt, and may damage, a fungal isolate resulting in no growth; except when H. capsulatum is suspected. This pathogen is intracellular, requiring homogenization to release the fungal cells to ensure growth. Tissue pieces should be pressed into the appropriate culture media or partially embedded to provide an oxygen tension gradient, and the cultures should be incubated at 30°C for 21 days (incubation may be extended if clinical suspicion of a mycotic disease is high). 

Culture Media and Incubation Requirements A number of fungal culture media are satisfactory for use in the clinical microbiology laboratory (Table 58.4). Most are adequate for the recovery of fungi. For optimal recovery, a battery of media should be used; the following are recommended: • Media with and without cycloheximide to prevent the overgrowth of slow-growing fungi by more rapidly growing species. It is important to note that cycloheximide may also be inhibitory to some fungi. • Media with and without an antibacterial agent (media with an antibacterial agent are used for specimens likely to contain contaminating bacteria; they are not necessary for specimens from sterile sites). • Inhibitory agar controls bacterial contamination more effectively than Sabouraud dextrose agar. • Chloramphenicol is an inhibitory agent for the growth of contaminating bacteria; however, it is important to note that it is also inhibitory to Nocardia and other aerobic actinomycetes. • Growth of dimorphic fungi is enhanced on enriched media such as BHI containing antibiotics and 5% to 10% sheep blood. This enhances growth but inhibits sporulation. Once isolated, the fungi should be immediately subcultured to blood-free enriched media for identification. • Birdseed agar may be used for the cultivation of Cryptococcus spp. from CSF, pleural fluid, bone marrow, tissue, and lower respiratory specimens. • Specific chromogenic agar may be used to identify some species of yeast. • Additional specialized media may be required for additional fungal isolates that have unique nutritional or incubation requirements. Agar plates (petri dishes), rectangle agar vials, or screwcapped agar tubes are satisfactory for the recovery of fungi; however, plates or vials are preferred, because they provide

better aeration of cultures and a large surface area for better isolation of colonies. Agar plates provide greater ease of handling by laboratory professionals when making microscopic preparations for examination. Agar tends to dehydrate during the extended incubation period required for fungal recovery, but this problem can be minimized by using agar plates or vials containing at least 40 mL of agar and placing them in a humidified incubator. Agar plates should be opened and examined in a certified biologic safety cabinet (BSC). Many laboratories discourage the use of agar plates because of safety considerations; however, the advantages outweigh the disadvantages. Compared with agar plates, screw-capped culture tubes are more easily stored, require less space for incubation, and are easily handled. In addition, they have a lower dehydration rate, and most laboratory workers believe cultures are less hazardous to handle when in tubes. However, disadvantages, such as relatively poor isolation of colonies, a reduced surface area for culturing, and a tendency to promote anaerobiosis, discourage routine use in most clinical microbiology laboratories. If culture tubes are used, the tube should be as large as possible to provide an adequate surface area for isolation. After inoculation, tubes should be placed in a horizontal position for at least 1 to 2 hours to allow the specimen to absorb to the agar surface and prevent settling at the bottom of the tube. Cultures should be incubated at room temperature, or preferably at 30°C, for 21 to 30 days before they are reported as negative. A relative humidity in the range of 40% to 50% can be achieved by placing an open pan of water in the incubator. Cultures should be examined at least three times weekly during incubation. As previously mentioned some clinical specimens are contaminated with bacteria or rapidly growing fungi or both, requiring the use of antifungal and antibacterial agents. The addition of 0.5 μg/mL of cycloheximide and 16 μg/mL of chloramphenicol to media traditionally has been advocated to inhibit the growth of contaminating molds and bacteria, respectively. However, better results have been achieved using a combination of 5 μg/mL of gentamicin and 16 μg/mL of chloramphenicol as antibacterial agents. Ciprofloxacin at a concentration of 5 μg/mL may be used. Cycloheximide may be added to any of the media that contain or lack antibacterial antibiotics. However, if cycloheximide is included in the battery of culture media, a medium lacking this ingredient should also be included. Pathogenic fungi, such as Cryptococcus spp., Candida krusei and other Candida spp., Trichosporon spp., P. boydii, and Aspergillus spp., are partially or completely inhibited by cycloheximide. 

Direct Microscopic Examination Direct microscopic examination of clinical specimens has been used for many years; however, its usefulness should be reemphasized. Because the mission of a clinical microbiology laboratory is to provide a rapid and accurate diagnosis, the mycology laboratory can provide this service in many cases by direct examination (particularly with the

CHAPTER 58  Overview of Fungal Identification Methods and Strategies

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TABLE 58.4    Fungal Culture Media: Indications for Use

Media

Indications for Use

Media Composition

Mode of Action

Primary Recovery Media Brain-heart infusion agar

Primary recovery of saprobic and pathogenic fungi

Brain-heart infusion, enzymatic digest of animal tissue, enzymatic digest of casein, dextrose, sodium chloride

The agar provides a rich medium for bacteria, yeast, and pathogenic fungi.

Brain-heart infusion agar (fungal formulation) with antibiotics

Primary recovery of pathogenic fungi exclusive of dermatophytes

Brain-heart infusion, enzymatic digest of animal tissue, enzymatic digest of casein, dextrose, sodium chloride, 10% sheep blood, antibiotics (chloramphenicol, cycloheximide, and gentamicin)

The agar provides a rich medium for yeast and pathogenic fungi, including systemic dimorphic fungi.

Chromogenic agar

Isolation and presumptive identification of yeast and filamentous fungi

Chromopeptone Glucose Chromogen mix Chloramphenicol

Chromogen mix contains substrates that react with enzymes produced by different organisms that result in the production of characteristic color changes.

Dermatophyte test medium

Primary recovery of dermatophytes; recommended as screening medium

Soy, peptone, dextrose, cycloheximide, gentamicin, chloramphenicol, phenol red

Dermatophytes produce alkaline metabolites, which raise the pH and change the medium from red to yellow.

Inhibitory mold agar

Primary recovery of pathogenic, cycloheximide sensitive fungi exclusive of dermatophytes

Chloramphenicol, casein, dextrose, starch, sodium phosphate, magnesium sulfate, sodium chloride, manganese sulfate

Examine plates for growth. Chloramphenicol inhibits bacterial growth.

Potato flake agar

Primary recovery of saprobic and pathogenic fungi and the stimulation of conidia formation.

Potato flakes, glucose, cycloheximide, chloramphenicol, bromthymol blue

Growth is enhanced by a pH alkaline reaction of fungus. Chloramphenicol and antibiotics inhibit the growth of bacteria and nonpathogenic fungi.

Mycobiotic or mycosel agar

Primary recovery of dermatophytes but may also be used for the recovery of other pathogenic fungi.

Pancreatic digest of soybean meal and dextrose with cycloheximide, chloramphenicol.

Inhibits bacteria and saprophytic fungi

Sabouraud dextrose with brain-heart infusion (SABHI) agar

Primary recovery of saprobic and pathogenic fungi

Sabouraud dextrose, brainheart infusion agar. With chloramphenicol, cycloheximide, penicillin, and/or streptomycin. 10% sheep blood may be added.

Isolates and enhances growth of all fungi including the yeast phase of dimorphic fungi.

Yeast-extract phosphate agar with ammonia

Primary recovery of pathogenic fungi exclusive of dermatophytes

Yeast extract, dipotassium phosphate, chloramphenicol. 1 drop of ammonium hydroxide is applied to the agar surface prior to inoculation and allowed to diffuse into the medium.

Enhances the recovery and sporulation of Blastomyces and Histoplasma capsulatum from contaminated specimens

Differential Test Media Acetate Ascospore agar

Detection of ascospores in ascosporogenous yeasts (e.g., Saccharomyces spp.)

Potassium acetate, yeast extract, dextrose

Potassium acetate is necessary, and yeast extract increases the sporulation of yeasts. Continued

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TABLE 58.4    Fungal Culture Media: Indications for Use—cont’d

Media

Indications for Use

Media Composition

Mode of Action

Christensen’s urea agar

Identification of Cryptococcus, Trichosporon, and Rhodotorula spp. Separation of Trichophyton mentagrophytes from Trichophyton rubrum

2% urea, phenol red

Produces urease and a change in the pH.

Cornmeal agar with Tween 80 and trypan blue

Differentiation of Candida spp. by chlamydospore production

Cornmeal, Tween 80, cornmeal infusion, and trypan blue

Addition of Tween 80 enhances the production of chlamydospores, pseudohyphal and arthrospore formation. The addition of trypan blue provides a contrasting background for observing the morphologic features of yeasts.

Czapek’s agar

Differential identification of Aspergillus spp.

Sodium nitrate, sucrose, yeast extract

Produces characteristic features of yeast and fungus of any organism that can use sodium nitrate.

Niger seed agar (birdseed agar)

Identification of Cryptococcus spp., particularly Cryptococcus neoformans and Cryptococcus gattii

Guizotia abyssinica seeds or niger seeds, dextrose, creatinine, chloramphenicol

C. neoformans and C. gattii produce the enzyme phenol oxidase, resulting in a brown pigment through metabolism of caffeic acid. Creatinine enhances the melanization of some strains of C. neoformans.

Potato dextrose agar

Demonstration of conidia formation and the pigment production by Trichophyton rubrum; preparation of microslide cultures and sporulation of dermatophytes

Potato infusion, dextrose, tartaric acid Note: Some laboratories use potato flake agar, because it may be more stable.

Carbohydrate and potato infusion promotes the growth of yeasts and molds, and the low pH (tartaric acid) partially inhibits bacterial growth.

Trichophyton agars 1–7

Identification of Trichophyton spp.

Dextrose, monopotassium phosphate, magnesium sulfate, amino acids 1. Casamino acids; vitamin free 2. Casamino acids plus inositol 3. Casamino acids plus inositol and thiamine 4. Casamino acids plus thiamine 5. Casamino acids plus niacin 6. Ammonium nitrate 7. Ammonium nitrate plus histidine

Trichophyton spp. may be differentiated by growth in the presence of different amino acids.

Yeast fermentation broth

Identification of yeasts by determining fermentation

Yeast extract, peptone, bromcresol purple, and a specific carbohydrate (e.g., dextrose, maltose, sucrose)

Most yeasts produce acid, which is indicated by a change in the solution from purple to yellow as a positive fermenter.

Yeast nitrogen-base agar

Identification of yeasts by determining carbohydrate assimilation

Ammonium sulfate, carbon source (e.g., glucose, sucrose, raffinose)

Assimilation of carbon by yeast cells produces a positive result.

CHAPTER 58  Overview of Fungal Identification Methods and Strategies

Gram stain) of the clinical specimen submitted for culture. Microbiologists are encouraged to become familiar with the diagnostic features of fungi commonly encountered in clinical specimens and to recognize them when stained by various dyes. This important procedure often can provide the first microbiologic proof of the cause of disease in patients with fungal infection and guide the selection of appropriate media to support growth. Tables 58.5 and 58.6 present the methods available for direct microscopic detection of fungi in clinical specimens and a summary of the characteristic microscopic features of each. Fig. 58.4 presents photomicrographs of some of the fungi commonly seen in clinical specimens. Traditionally the potassium hydroxide preparation has been the recommended method for direct microscopic examination of specimens in dermatological samples. Calcofluor white (CW) (Evolve Procedure 58.1) and lactophenol blue (LPCB) bind specifically to polysaccharides, chitin, and cellulose present in fungal cell walls. In addition, LPCB contains lactic acid that aids in the preservation of fungal structures and phenol that acts as a killing agent. Slides prepared using these methods may be observed using fluorescent (CW) or bright-field microscopy (LPCB). 

Serologic Testing Molecular diagnostics may eventually replace the use of serologic testing for the diagnosis of fungal infections. These methods currently lack standardization for performance when taken directly from patient specimens. No commercially available procedures exist for serologic testing of most fungi. However, serology testing is a useful tool for the diagnosis of invasive fungal infections with select organisms, such as Cryptococcus, Coccidioides, Blastomyces, Histoplasma, and Aspergillus spp. Antibody testing has proven useful but not for immunocompromised patients, who are incapable of producing a measurable humoral response. Acute and convalescent titers need to be monitored during treatment of the fungal infection. Immunodiffusion testing is a simple, cost-effective procedure. Although it is 100% specific, it is relatively insensitive and is not used as a screening tool. This test also requires 2 to 3 weeks to exhibit a positive result. Enzyme immunoassays for both antibody and antigen have been used. These tests are also commonly negative in immunocompromised patients, especially early in the infection. Point-of-care (POC) testing using lateral flow assay-based devices have the potential to improve the diagnosis of fungal infections, especially in developing countries with limited laboratory resources. POC devices generally are inexpensive and portable while providing rapid and reproducible results that are highly sensitive and specific. Undoubtedly, the development of these devices will be useful in monitoring and detecting fungal antibodies.

801

(1,3)-β-d-Glucan Detection (1,3)-β-d-Glucan, a polysaccharide present in the cell wall of some fungi, is found in the blood of patients that have invasive fungal infections. Although detection of the polysaccharide has demonstrated success in the diagnosis and monitoring of fungal meningitis, variation in assay sensitivity and specificity, along with the need for what are considered significant levels in the blood of a patient with fungal infections, indicates further studies are necessary. 

Molecular Methods Phenotypic and biochemical identification methods are extremely time-consuming for the identification of fungal pathogens. One of the most critical risk factors associated with mortality from systemic mycoses is the time to diagnosis, making molecular detection methods ideal in the clinical laboratory. In addition to diagnosis, drug resistance among invasive Candida and Cryptococcus spp. has increased, requiring the development of new assays to detect drug resistance. Ideally, a molecular direct hybridization assay or amplification assay panel of primers specific for the detection of fungi in clinical specimens would include the most common organisms known to cause disease in immunocompromised patients (including the dimorphic fungi and Pneumocystis spp.). The current literature contains references to all of the major human fungal pathogens, describing species and strain-specific primers and probes, yet no commercial methods are available to the clinical laboratory. Sequence-based molecular identification of fungal isolates is a useful diagnostic tool and has resulted in the development of commercial diagnostic assays such as the MicroSEQ D2 rDNA Fungal Sequencing Kit (Thermo Fisher, Grand Island, NY). Currently, DNA sequencing technology, including whole genome sequencing, remains confined to research and reference laboratories due to the large capital investment and expertise required for implementation. Currently, a few FDA-cleared molecular diagnostic assays are available, which include amplification and hybridization techniques. These assays primarily focus on the identification of Candida, Cryptococcus, Aspergillus spp., and the systemic dimorphic fungi. 

Matrix-Assisted Laser Desorption Ionization Mass Spectrometry Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) is a biophysical method that significantly reduces the time required to specifically identify fungal organisms. The major disadvantage of this technique is the need to have pure cultures of the clinical isolates for sample preparation, adding days and weeks to the process. The protein profiles for fungi also vary significantly based on environmental and culture conditions, making standardization of the analytical process extremely difficult in fungal identification. The level of fungal organisms, such as yeast in the bloodstream and other clinical fluid samples is generally too low for direct detection by

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TABLE 58.5    Summary of Methods Available for Direct Microscopic Detection of Fungi in Clinical Specimens

Method

Use

Time Required

Advantages

Disadvantages spp.a

Acid-fast stain and partial acid-fast stain

Detection of mycobacteria and Nocardia spp., respectively

12 min

Detects Nocardia and some isolates of Blastomyces spp.

Tissue homogenates are difficult to observe because of background staining.

Alcian blue or mucicarmine stain.

Mucopolysaccharide stains used to visualize the capsule of Cryptococcus spp. in histological tissue sections.

30 min

Detects encapsulated yeast in tissue sections

Blastomyces dermatitidis and Rhinosporidium seeberi may also react positively with this stain.

Auramine-rhodamine stain

Detection of mycobacteria and Nocardia spp., respectively

10 min

Excellent screening tool; sensitive and affordable.

Not as specific for acid-fast organisms as Ziehl-Neelsen stain.

Calcofluor white stain

Detection of fungi

1 min

Can be mixed with KOH; detects fungi rapidly because of bright fluorescence.

Requires use of a fluorescence microscope; background fluorescence prominent, but fungi exhibit more intense fluorescence; vaginal secretions are difficult to interpret. Nonspecific reactions may be observed, such as cotton fibers from swabs and brain tumor biopsies, both falsely resembling branching hyphae.

Gram stain

Detection of bacteria

3 min

Commonly performed on most clinical specimens submitted for bacteriology; detects most fungi.

Some fungi stain well, but others (e.g., Cryptococcus spp.) show only stippling and stain weakly in some instances; some isolates of Nocardia spp. fail to stain or stain weakly.

India ink (nigrosin) stain

Detection of Cryptococcus spp. in CSF

1 min

Diagnostic of meningitis when positive in CSF.

Positive in fewer than 50% of cases of meningitis; not sensitive in non–HIVinfected patients. Artifacts such as erythrocytes, leukocytes, and talc particles from gloves or bubbles may mimic yeast, resulting in false positives.

Lactophenol cotton (aniline) blue wet mount

Most widely used method of staining and observing fungi

1 min

Lactic acid and glycerol preserves structures; slides can be made permanent.

Mechanical treatment dislodges fungal structures.

Potassium hydroxide

Clearing of specimen using 10%–20% KOH to make fungi more readily visible

5 min; if clearing is not complete, an additional 5–10 min is necessary

Rapid detection of fungal elements. 0.1% thimerosal (Sigma Chemical Co.) may be added to preserve the specimen.

Requires experience, because background artifacts are often confusing; clearing of some specimens may require an extended time.

Masson-Fontana stain

Examination of melanin pigment in fungal cell walls

1 h, 10 min

Aids differentiation of melanin and hemosiderin pigments.

Difficult to interpret when only rare granular staining is present.

CHAPTER 58  Overview of Fungal Identification Methods and Strategies

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TABLE 58.5    Summary of Methods Available for Direct Microscopic Detection of Fungi in Clinical Specimens—cont’d

Method

Use

Time Required

Advantages

Disadvantages

Methenamine silver stain

Detection of fungi in histologic section

1h

Best stain for detecting fungal elements (black) against a pale green or yellow background.

Requires a specialized staining method that is not usually readily available to microbiology laboratories.

Periodic acid–Schiff (PAS) stain

Detection of fungi

20 min; 5 min additional if counterstain is used

Stains fungal elements well; hyphae of molds and yeasts can be readily distinguished.

Nocardia spp. do not stain well. Time-consuming and has been replaced in many laboratories by calcofluor white staining procedures.

Toluidine blue O

Rapid detection of P. jiroveci from lung biopsy and BAL specimens.

1 min

Quickly performed, easy, rapid results, and cost-effective.

Trophozoites are not discernable.

Wright’s stain

Examination of bone marrow or peripheral blood smears

7 min

Detects Histoplasma capsulatum and Cryptococcus spp.

Most often used to detect H. capsulatum and Cryptococcus spp. in disseminated disease.

aPartially

acid-fast bacterium. BAL, Bronchoalveolar lavage; CSF, cerebrospinal fluid; HIV, human immunodeficiency virus; KOH, potassium hydroxide.

MALDI-TOF MS. In addition, proteins and hemoglobin tend to interfere with the spectra analytics making identification problematic. Further clinical studies are needed prior to implementation of direct identification using MALDITOF MS for systemic fungal infections. An overview of the technique is discussed in more detail in Chapter 7. 

General Considerations for the Identification of Yeasts Most often yeasts are identified through the use of a combination of differential test media (Fig. 58.5). Identification factors and techniques include the following: • Colonial morphologic features • Microscopic morphologic features • Physiologic studies • Chromogenic agars (presumptive species identification) • Rapid commercial yeast identification tests or panels • Nucleic acid–based methods (direct hybridization or amplification methods) • Matrix-assisted laser desorption/ionization time-of-flight mass spectroscopy (MALDI-TOF MS) Several key characteristics can be seen macroscopically. Colonies have a wide variety of colors, shapes, and textures. Chromogenic agar or a combination of rapid tests can be used to differentiate some pathogenic yeast presumptively. Presumptive identification is indicated when the test or combination of tests do not identify characteristics that are unique to that species. The results of these tests must be correlated

with the macroscopic and microscopic morphological characteristics and the site of infection (i.e., clinical specimen). Wet preps and lactophenol cotton blue stain can aid microscopic identification by improving the visualization of the fungal reproductive structures. Sexual and asexual characteristics are very important. Often a genus can be determined by the microscopic and macroscopic characteristics. India ink stain is useful when Cryptococcus spp. are suspected. Because carbon and nitrogen source differences are the key to differentiating yeasts, many automated and semiautomated commercial systems have been designed with assimilation and fermentation tests. Supplemental testing takes advantage of a limited set of characteristics to further aid identification. The clinical microbiology laboratory has historically operated under the idea that if the identification of the isolate to the species level is incorrect, the patient will not be adversely affected once the susceptibility is completed. This is problematic, when the specific species identifications are used to determine minimum inhibitory concentrations and breakpoints for treatment for bacterial and fungal infections. Treatment failures, along with the increasing occurrence of resistance to echinocandins, fluconazole, and voriconazole, make accurate identification and susceptibility testing required. Molecular diagnostic techniques, proteomic and genomic, clearly demonstrate more accurate and reliable identification for yeasts than conventional biochemical methods. The use of these methods in the routine clinical laboratory is currently limited by the need for standardization and the expansion of the current databases. 

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TABLE 58.6    Summary of Characteristic Features of Select Fungi Seen in Direct Examination of Clinical Specimens

Morphologic Form Found in Specimens

Organism

Size Range (diameter, mm)

Yeastlike

Histoplasma capsulatum

2–5

Small; oval to round budding cells; often found clustered in histiocytes; difficult to detect when present in small numbers.

Sporothrix spp.

2–6

Small; oval to round to cigarshaped; single or multiple buds present; uncommonly seen in clinical specimens.

Cryptococcus spp.

2–15

Cells exhibit great variation in size; usually spherical but may be football-shaped; buds single or multiple and “pinched off”; capsule may or may not be evident; occasionally, pseudohyphal forms with or without a capsule may be seen in exudates of cerebrospinal fluid.

Malassezia furfur (in fungemia)

1.5–4.5

Small; bottle-shaped cells, buds separated from parent cell by a septum; emerge from a small collar.

Blastomyces spp.

8–15

Cells are usually large, double refractile when present; buds usually single; however, several may remain attached to parent cells; buds connected by a broad base.

Paracoccidioides spp.

5–60

Cells are usually large and are surrounded by smaller buds around the periphery (“mariner’s wheel appearance”); smaller cells may be present (2–5 μm) and resemble H. capsulatum; buds have “pinched-off” appearance.

Coccidioides spp.

10–200

Spherules vary in size; some may contain endospores, others may be empty; adjacent spherules may resemble Blastomyces spp.; endospores may resemble H. capsulatum but show no evidence of budding; spherules may produce multiple germ tubes if a direct preparation is kept in a moist chamber ≥24 h.

Rhinosporidium seeberi (protozoan [parafungal] pathogen that is studied in mycology)

6–300

Large, thick-walled sporangia containing sporangiospores are present; mature sporangia are larger than spherules of Coccidioides; hyphae may be found in cavitary lesions.

Spherules

Characteristic Features

CHAPTER 58  Overview of Fungal Identification Methods and Strategies

TABLE 58.6    Summary of Characteristic Features of Select Fungi Seen in Direct Examination of Clinical Specimens—cont’d

Morphologic Form Found in Specimens

Organism

Size Range (diameter, mm)

Yeast and pseudohyphae or hyphae

Candida and Candida dubliniensis

5–10 (pseudohyphae)

Cells usually exhibit single budding; pseudohyphae, when present, are constricted at the ends and remain attached like links of sausage; hyphae, when present, are septate.

M. furfur (in tinea versicolor)

3–8 (yeast) 2.5–4 (hyphae)

Short, curved hyphal elements are usually present, along with round yeast cells that retain their spherical shape in compacted clusters; “spaghetti and meatballs.”

Pauciseptate hyphae

Mucorales: Mucor, Rhizopus, and other genera

10–30

Hyphae are large, ribbonlike, often fractured or twisted; occasional septa may be present; smaller hyphae are confused with those of Aspergillus spp., particularly Aspergillus flavus.

Hyaline septate hyphae

Dermatophytes, skin and nails

3–15

Hyaline, septate hyphae are commonly seen; chains of arthroconidia may be present.

Dermatophytes, hair

3–15

Arthroconidia on periphery of hair shaft producing a sheath indicate ectothrix infection; arthroconidia formed by fragmentation of hyphae in the hair shaft indicate endothrix infection. Long hyphal filaments or channels in the hair shaft indicate favus hair infection.

Aspergillus spp.

3–12

Hyphae are septate and exhibit dichotomous, 45-degree branching; larger hyphae, often disturbed, may resemble those of Mucorales.

Geotrichum spp.

4–12

Hyphae and rectangular arthroconidia are present and sometimes rounded; irregular forms may be present.

Trichosporon spp.

2–4 by 8

Hyphae and rectangular arthroconidia are present and sometimes rounded; occasionally, blastoconidia may be present.

Dematiaceous septate hyphae

Bipolaris spp., Cladophialophora spp., Cladosporium spp., Curvularia spp., Exophiala spp., Exserohilum spp., Hortaea werneckii, Phialophora spp., and other genera.

2–6

Dematiaceous polymorphous hyphae are seen; budding cells with single septa and chains of swollen rounded cells are often present; occasionally, aggregates may be present in infection caused by Phialophora and Exophiala spp.

Sclerotic bodies

Cladophialophora (formerly Cladosporium) carrionii Fonsecaea spp., Phialophora verrucosa, and Rhinocladiella aquaspersa

5–20

Brown, round to pleomorphic, thick-walled cells with transverse septations; commonly, cells contain two fission planes that form a tetrad of cells (sclerotic bodies).

Characteristic Features

Continued

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TABLE 58.6    Summary of Characteristic Features of Select Fungi Seen in Direct Examination of Clinical Specimens—cont’d

Morphologic Form Found in Specimens

Organism

Size Range (diameter, mm)

Granules

Acremonium spp.

200–300

White, soft granules without a cementlike matrix.

Aspergillus Aspergillus nidulans

500–1000

Black, hard grains with a cementlike matrix at the periphery.

Curvularia Curvularia geniculata Curvularia lunata

65–160

White, soft granule without a cementlike matrix.

Exophiala Exophiala jeanselmei

200–300

Black, soft granules, vacuolated, without a cementlike matrix, made of dark hyphae and swollen cells.

Fusarium Fusarium verticillioides (formerly F. moniliformis)

200–500

White, soft granules without a cementlike matrix.

Fusarium solani

300–600

Trematosphaeria grisea (formerly Madurella grisea)

350–500

Black, soft granules without a cementlike matrix; the periphery is composed of polygonal swollen cells, and the center has a hyphal network.

Madurella mycetomatis

200–900

Black to brown, hard granules; two types: (1) rust-brown, compact, filled with cementlike matrix; (2) deep brown, filled with numerous vesicles, 6–14 μm in diameter, cementlike matrix in periphery, central area of light-colored hyphae.

Neotestudina Neotestudina rosatii

300–600

White, soft granules with cementlike matrix at the periphery.

Pseudallescheria Pseudallescheria boydii

200–300

White, soft granules composed of hyphae and swollen cells at the periphery in a cementlike matrix.

General Considerations for the Identification of Molds Filamentous fungi are also identified by a combination of tests (Fig. 58.6). Molds are identified using a combination of the following: • Growth rate • Colonial morphologic features • Microscopic morphologic features In most cases, the microscopic morphologic features provide the most definitive means of identification. Determination of the growth rate can be most helpful when a mold culture is examined. However, this may have limited value, because the growth rate of certain fungi varies, depending on the amount of inoculum present in a clinical specimen. Slow growers form mature colonies in 11 to 21 days, and intermediate growers form mature colonies

Characteristic Features

in 6 to 10 days. Rapid growers form mature colonies in 5 days or less. The growth of Coccidioides is often rapid and is hazardous to microbiologists. In general, the growth rate for the dimorphic fungi, Blastomyces dermatitidis, H. capsulatum, and P. brasiliensis, is slow; 1 to 4 weeks usually are required before colonies become visible. In some instances, cultures of B. dermatitidis, Talaromyces marneffei, and H. capsulatum may be detected within 3 to 5 days. This is a somewhat uncommon circumstance, encountered only when large numbers of the organism are present in the specimen. Colonies of Mucorales may appear within 24 hours, whereas the other hyaline and melanized (dematiaceous) fungi often exhibit growth in 1 to 5 days. The growth rate of an organism therefore is important, but it must be used in combination with other features before a definitive identification can be made.

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807

A

C

B

• Fig. 58.4  Fungi commonly seen in clinical specimens. (A) This potassium hydroxide preparation of a skin

scraping from a patient with a dermatophyte infection shows septate hyphae intertwined among epithelial cells. (Phase-contrast microscopy; ×500.) (B) This calcofluor white stain of urine demonstrates Candida albicans. (C) The deeply staining, small, uniform yeast cells in this histologic section of lung tissue are typical of Histoplasma capsulatum. (Methenamine silver stain; ×430.)

The colonial morphologic features may have limited value for identifying molds because of natural variation among isolates and colonies grown on different culture media. Although common organisms recovered repeatedly in the laboratory may be more easily recognized, colonial morphology is an unreliable criterion that should be used to supplement the microscopic morphologic features of the organism. The color of the colony and uniformity of the color can be important, along with the presence of diffusible pigments in the media. The examiner must be sure to note the color of both the front and reverse sides of the culture. The colony topography describes the various elevations of the colony on the agar plate. Topography can be described as verrucose (furrowed or convoluted), umbonate (slightly raised in the center), or rugose (furrows radiate out from the center). The colony’s texture should also be noted. Various textures can be seen, such as cottony (loose, high aerial mycelium), velvety (low aerial mycelium resembling a velvet cloth), glabrous (smooth surface with no aerial mycelium), granular

(dense, powdery, resembling sugar granules), and wooly (high aerial mycelium that appears slightly matted down). Incubation conditions and culture media must also be considered. For example, H. capsulatum appears as a whiteto-tan fluffy mold on BHI agar and may have a yeastlike appearance when grown on the same medium containing blood enrichment. In general, the microscopic morphologic features of the molds are stable and show minimal variation. Definitive identification is based on the characteristic shape, method of reproduction, and arrangement of spores; however, the size of the hyphae also provides helpful information. The large, ribbonlike, pauciseptate hyphae of the Mucorales are easily recognized; small hyphae, approximately 2 μm in diameter, may suggest the presence of one of the dimorphic fungi or a dermatophyte. The fungi may be prepared for microscopic observation using several techniques. The procedure traditionally used by most laboratories is the cellophane tape preparation (Evolve Procedure 58.2; Fig. 58.7). It can be prepared

808 PA RT V    Mycology

Identification of yeasts in clinical specimens

Microscopic morphology

Ascospore production

Germ tube production

Pigment production (niger or caffeic agar)

Sugar assimilation reactions

Nitrate reduction

Urease production

Carbohydrate fermentation reactions

Capsule production

Commercial yeast identification kits

• Fig. 58.5  Traditional identification of yeasts in clinical specimens. Manual methods, kits, and panels have been predominantly replaced with proteomic and genomic methods.

easily and quickly and often is sufficient to make the identification for most fungi. However, some laboratories prefer the wet mount (Evolve Procedure 58.3; Fig. 58.8) or tease mount (Evolve Procedure 58.4). A microslide culture method (Evolve Procedure 58.5; Fig. 58.9) may be used when greater detail of the morphologic features is required. 

General Morphologic Features of the Molds Specialized types of vegetative hyphae may be helpful for categorizing an organism into a certain group. For example, dermatophytes often produce several types of hyphae, including antler hyphae, so named because they are curved, freely branching, and have the appearance of antlers (Fig. 58.10). Racquet hyphae are enlarged, club-shaped structures (Fig. 58.11). In addition, certain dermatophytes produce spiral hyphae that are coiled or exhibit corkscrewlike turns in the hyphal strand (Fig. 58.12). These structures are not characteristic for any certain group; however, they are found most commonly in dermatophytes. Some species of fungi (Ascomycota) produce sexual spores in a large, saclike structure called an ascocarp (Fig. 58.13). The ascocarp contains smaller sacs, called asci, each of which contains four to eight ascospores. This type of sexual reproduction is not commonly seen in the fungi recovered in the clinical microbiology laboratory; most exhibit asexual reproduction. It is possible that all fungi have a sexual form, but

for some species, it has not yet been observed on artificial culture media. Conidia, which are produced by most fungi, represent the asexual reproductive cycle. The type of conidia and their morphology and arrangement are important criteria for definitively identifying an organism (Fig. 58.14). The simplest type of sporulation is the development of a spore directly from the vegetative hyphae. Arthroconidia are formed directly from the hyphae by fragmentation through the points of septation (Fig. 58.15). When mature, they appear as square, rectangular, or barrel-shaped thickwalled cells. These result from the simple fragmentation of the hyphae into spores, which are easily dislodged and disseminated into the environment. Chlamydoconidia (chlamydospores) are round, thick-walled spores formed directly from the differentiation of hyphae in which there is a concentration of protoplasm and nutrient material (Fig. 58.16). These appear to be resistant resting spores produced by the rounding up and enlargement of the cells of the hyphae. Chlamydoconidia may be intercalary (within the hyphae) or terminal (on the end of the hyphae). A variety of other types of spores occur with many species of fungi. Conidia are asexual spores produced singly or in groups by specialized hyphal strands, conidiophores. Sometimes the conidia are freed from their point of attachment by pinching off, or abstriction. Some conidiophores terminate in a swollen vesicle. From the surface of the vesicle are formed secondary small, flask-shaped phialides, which in turn give rise to long chains of conidia. This type of fruiting structure is characteristic of the aspergilli. A single,

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809

Identification of filamentous fungi from clinical specimens

Molecular detection

Microslide culture method (Evolve Procedure 58.5) Tease mount (Evolve Procedure 58.4)

Serology

Microscopic examination

Macroscopic examination

Growth on cyclohexamidecontaining media

Surface pigment

Reverse pigment

Wet mount (Evolve Procedure 58.3)

Colonial morphology Adhesive tape preparation (Evolve Procedure 58.2)

•

Fig. 58.6 Traditional identification of filamentous fungi from clinical specimens. Some molds may be identified using proteomic and genomic methods.

• Fig. 58.7  Cellophane tape preparation showing placement of tape on

•

simple, slender, tubular conidiophore (phialide) that produces a cluster of conidia, held together as a gelatinous mass, is characteristic of certain fungi, including the genus Acremonium (Fig. 58.17). In other cases, conidiophores form a branching structure called a penicillus, in which each branch terminates in secondary branches (metulae)

and phialides, from which chains of conidia are borne (Fig. 58.18). Species of Penicillium and Paecilomyces are representative of this type of sporulation. Some fungi may produce conidia of two sizes: microconidia, which are small, unicellular, round, elliptical, or pyriform (pear-shaped), or macroconidia, which are large, usually multiseptate, and

slide containing lactophenol cotton or aniline blue.

Fig. 58.8 Performance of a wet mount showing agar positioned under cover slip before pressure is applied to disperse growth.

810 PA RT V    Mycology

•

Fig. 58.9 Microslide culture showing inoculation of an agar plug (arrow).

• Fig. 58.12  Spiral hyphae (arrow) showing corkscrewlike turns (×430).

• Fig. 58.13  Ascocarp showing dark-appearing ascospores (×430).

•

Fig. 58.10 Antler hyphae showing swollen hyphal tips, resembling antlers, with lateral and terminal branching (favic chandeliers) (×500).

A

B

•

Fig. 58.14 Conidia (asexual spores [A]) produced on specialized structures (conidiophores [B]) of Aspergillus (×430).

•

Fig. 58.11  Racquet hyphae showing swollen areas (arrows) resembling a tennis racquet. B

club- or spindle-shaped (Fig. 58.19). Microconidia may be borne directly on the side of a hyphal strand or at the end of a conidiophore. Macroconidia are usually borne on a short to long conidiophore and may be smooth or roughwalled. Microconidia and macroconidia are seen in some fungal species and are not specific, except as they are used to differentiate a limited number of genera. The hyphae of the Mucorales are sparsely septate. Sporulation takes place by progressive cleavage during maturation

A

•

Fig. 58.15 Arthroconidia formation (A) produced by the breaking down of a hyphal strand (B) into individual rectangular units (×430).

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A

B

•

Fig. 58.16 Chlamydoconidia composed of thick-walled spherical cells (arrows) (×430).

• Fig. 58.19  In this preparation of a Trichophyton species, the numerous, small, spherical microconidia (A) are contrasted with a large, elongated macroconidium (B) (×430).

• Fig. 58.17  Simple tubular phialide with a cluster of conidia at its tip (arrow) characteristic of Acremonium (×430).

•

Fig. 58.20 Large, saclike sporangia that contain sporangiospores (arrow) characteristic of the Mucorales (×250).

Clinical Relevance for Fungal Identification

• Fig. 58.18  Complex method of sporulation in which conidia are borne

on phialides produced on secondary branches (metulae [arrow]) characteristic of Penicillium (×430).

in the sporangium, a saclike structure produced at the tip of a long stalk (sporangiophore). Sporangiospores (spores produced in the sporangium) are produced and released by the rupture of the sporangial wall (Fig. 58.20). In rare cases, some isolates may produce zygospores, rough-walled spores produced by the union of two mating types of a Mucorales; this is an example of sexual reproduction. 

The question of when and how far to go with the identification of fungi recovered from clinical specimens presents an interesting challenge. The current emphasis on cost containment and the ever-increasing number of opportunistic fungi causing infection in compromised patients prompts consideration of whether all fungi recovered from clinical specimens should be thoroughly identified and reported. The extent of identification of yeasts from various specimen sources is discussed in Chapter 62. When and how far to proceed in the identification of a mold is a difficult question to answer. Except for obvious plate contaminants, all commonly encountered molds should be identified and reported, if recovered from patients at risk for invasive fungal disease. Immunocompromised patients may have serious or even fatal disease caused by fungi that were once thought to be clinically insignificant. Organisms that fail to sporulate after a reasonable time should be reported as present, but identification is not required if the dimorphic fungi have been ruled out or if the clinician believes the organism is not clinically significant. Ideally, all laboratories should identify all fungi recovered

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from clinical specimens; however, the limits of practicality and economic considerations play a definite role in the decision-making process. The laboratory director, in consultation with the clinicians, must make this decision after considering the patient population, laboratory practice, and economic implications. An increasing number of fungi may be isolated in the clinical microbiology laboratory. They are considered environmental microbiota but in reality must be regarded as potential pathogens, because infections with a number of these organisms have been reported. The laboratory must identify and report all organisms recovered from clinical specimens so that their clinical significance can be determined. In many instances, the presence of environmental fungi is unimportant; however, that is not always the case. 

Laboratory Safety Although the handling of fungi recovered from clinical specimens poses risks, a common sense approach to the handling of these specimens protects the laboratory from contamination and workers from becoming infected. Without exception, mold cultures and clinical specimens must be handled in a class II BSC. Some laboratory professionals believe that mold cultures must be handled in an enclosed BSC equipped with gloves; however, this is not necessary if a laminar flow BSC is used. Yeast cultures may be handled on the bench top. An electric incinerator is suitable for decontaminating a loop used to transfer yeast cultures. Cultures of organisms suspected of being pathogens should be sealed with tape to prevent laboratory contamination and should be autoclaved as soon as the definitive identification is made. If common safety precautions are followed, few problems should occur with laboratory contamination or infection acquired by laboratory personnel. 

Prevention Preventing and controlling fungal infections continue to be a challenge to individuals, researchers, laboratorians, and hospitals. Very few formal recommendations are available to prevent exposure to community-associated fungal infections. Good personal hygiene may be the best course for prevention. However, many strategies can be followed to prevent health care–associated infections. Hospital staff members should be aware of the pathogenesis of fungal infections. Fungi are easily spread in ventilation systems, water, and skin-to-skin contact. Hospitals should follow an infection control plan that includes periodic monitoring of air-handling systems and regular testing for environmental spores. Staff members, patients, and visitors should practice good personal hygiene to minimize exposure to potential fungal infections. The laboratory also plays an important role in fungal prevention and control. Lack of rapid and specific testing continues to be a factor in a timely diagnosis. Early definitive diagnosis ensures that the appropriate therapy is given promptly and prevents mortality.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Barenfanger J, Arakere P, Dela Cruz R, et  al.: Improved outcomes associated with limiting identification of Candida spp. in respiratory secretions, J Clin Microbiol 41:5645–5649, 2003. Beck MR, Dekoster GT, Cistola DP, et al.: NMR structure of a fungal virulence factor reveals structural homology with mammalian saposin B, Mol Microbiol 72:344–353, 2009. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, 2020, Elsevier-Saunders. Bernstein EF, Schuster MG, Stieritz, et al.: Disseminated cutaneous Pseudallescheria boydii, Br J Dermatol 132:456–460, 1995. Bille J, Stockman L, Roberts GD, et al.: Evaluation of a lysis-centrifugation system for recovery of yeasts and filamentous fungi from blood, J Clin Microbiol 18:469–471, 1983. Bille J, Edson RS, Roberts GD: Clinical evaluation of the lysis-centrifugation blood culture system for the detection of fungemia and comparison with a conventional biphasic broth blood culture system, J Clin Microbiol 19:126–128, 1984. Brandhorst TT, Gauthier GM, Stein RA, et al.: Calcium binding by essential virulence factor BAD-1 of Blastomyces dermatitidis, J Biol Chem 280:42156–42163, 2005. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Cherniak R, Sundstrom JB: Polysaccharide antigens of the capsule of Cryptococcus neoformans, Infect Immun 62:1507–1512, 1994. DeHoog GS, Chaturvedi V, Denning DW, et al.: Name changes in medically important fungi and their implications for clinical practice, J Clin Microbiol 53:1056–1062, 2015. DeHoog GS, Haase G, Chaturvedi V, et al.: Taxonomy of medically important fungi in the molecular era, Lancet Infect Dis 13:385– 386, 2013. Hibbett DS, Binder M, Bischoff JF, et al.: A higher-level phylogenetic classification of the fungi, Mycol Res 111:509–547, 2007. Irmenia C, Pagano L, Martino B, et  al.: Invasive infections caused by Trichosporon species and Geotrichum capitatum in patients with hematological malignancies: a retrospective multicenter study from Italy and review of the literature, J Clin Microbiol 43:1818– 1828, 2005. Jacobson ES: Pathogenic roles for fungal melanins, Clin Microbiol Rev 13:708–717, 2000. Koneman EW, Church DL, Procop GW, et al.: Koneman’s color atlas and textbook of diagnostic microbiology, ed 7, Philadelphia, PA, 2017, Wolters Kluwer. Kwon-Chung KJ, Faser JA, Doering TL, et  al.: Cryptococcus neoformans and Cryptococcus gattii, the etiologic agents of cryptococcosis, Cold Spring Harb Perspect Med 4(7):a019760, 2014. Leber AL: Mycology and antifungal susceptibility testing, clinical microbiology procedures handbook, ed 4, Washington, DC, 2016, ASM Press. Lucas S, da luz Martins M, Flores O, et  al.: Differentiation of Cryptococcus neoformans varieties and Cryptococcus gattii using CAP59-based loop-mediated isothermal DNA amplification, Clin Microbiol Infect 16:711–714, 2010. Madariaga MG, Tenorio A, Proia L: Trichosporon inkin peritonitis treated with caspofungin, J Clin Microbiol 41:5827–5829, 2003. Norvell LL: Melbourne approves a new CODE, Mycotaxon 116:481– 490, 2011.

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Procop GW, Roberts GD: Emerging fungal diseases: the importance of the host, Clin Lab Med 24:691–719, 2004. Richardson M, Page I: Role of serological tests in the diagnosis of mold infections, Curr Fungal Infect Rep 12:127–136, 2018. Singh S, Beena PM: Comparative study of different microscopic techniques and culture media for the isolation of dermatophytes, Indian J Med Microbiol 21:21–24, 2003. Steinbach WJ, Schell WA, Miller JL, et al.: Scedosporium prolificans osteomyelitis in an immunocompetent child treated with voriconazole and caspofungin, as well as locally applied polyhexamethylene biguanide, J Clin Microbiol 41:3981–3985, 2003. Tomee J, Kauffman HF: Putative virulence factors of Aspergillus fumigatus, Clin Exp Allergy 30:476–484, 2000.

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Vilela R, Mendoza L: Human pathogenic entomophthorales, Clin Microbiol Rev 31:e00014–e00018, 2018. Walsh TJ, Groll A, Hiemenz J, et al.: Infections due to emerging and uncommon medically important fungal pathogens, Clin Microbiol Infect 10(Suppl 1):48–66, 2004. Walsh TJ, Hayden RT, Larone DH: Larone’s medically important fungi: a guide to identification, ed 6, Washington, DC, 2018, ASM Press. Wickes BL, Wiederhold NP: Molecular diagnostics in medical mycology, Nat Commun 9:5135, 2018. Wiederhold NP, Gibas CFC: From the clinical mycology laboratory: new species and changes in fungal taxonomy and nomenclature, J Fungi 4:138, 2018.

PROCEDURE 58.1 CALCOFLUOR WHITE–POTASSIUM HYDROXIDE PREPARATION

Method 1. Place a drop of calcofluor white (CW) reagent and a drop of 10% potassium hydroxide (KOH) glycerin in the center of a microscope slide. 2. Add a portion of the clinical specimen to the CW-KOH solution and apply a cover slip. 3. If necessary, dissociate particles by applying gentle pressure to the cover slip with a pencil eraser. Allow to stand for 5 minutes. If particles do not dissociate, repeat this step. 4. Examine the slide for fungal elements using a fluorescent microscope with a 400–500 nm exciter filter and a 500–520 nm barrier filter. Slides are scanned at ×10 magnification for fluorescent fungal elements. The presence and nature of fungal elements are discerned using ×40 magnification. 

Reagents

CW reagent: (0.05 g CW, 0.02 g Evans blue, 50 mL distilled water) Calcofluor white is an industrial textile brightener that nonspecifically binds to chitin and other elements in the fungal cell wall. Calcofluor white fluorescence occurs maximally at an excitation wavelength of 440 nm. Under these conditions, fungal elements fluoresce blue-white. Because CW is a nonspecific stain, an appreciation for fungal element morphology on direct examination is crucial for adequate specimen interpretation. The presence of KOH in the solution dissolves human cellular elements and debris, which allows for easier visualization of fungal elements. 

Limitations Careful interpretation is crucial, because nonspecific fluorescence may be observed with the presence of cotton fibers or certain tissues from patients with tumors.

KOH reagent: (10 g KOH, 10 mL glycerin, 80 mL distilled water)   

PROCEDURE 58.2 CELLOPHANE TAPE PREPARATION

Method 1. Touch the adhesive side of a small length of transparent tape to the surface of the colony. 2. Add a drop of lactophenol cotton or aniline blue to the surface of a microscope slide; then, adhere the length of tape to the surface of the slide (Fig. 58.7). 3. Observe microscopically for the characteristic shape and arrangement of the spores. The transparent adhesive tape preparation allows the laboratorian to observe the organism microscopically approximately the way the fungi sporulates in culture. The relationship of the spores, spore-producing structures (e.g., conidiophores), and the body of the fungus are usually intact, and microscopic identification of an organism can be made easily. If the tape is not pressed firmly enough to the surface of the colony, the sample may consist only of conidia and

may not be adequate for identification. When spores are not observed, a wet mount should be made. In some cases, the macroconidia of Histoplasma capsulatum may be seen in wet mount preparations when the adhesive tape preparation reveals only hyphal fragments. In other instances, cultures may have sporulated and reveal only the presence of conidia when the adhesive tape preparation is observed. In this type of situation, a second adhesive tape preparation should be made from the periphery of the colony where sporulation is not as prominent. Some laboratories prefer to use the microslide culture (Evolve Procedure 58.5) for microscopic identification of an organism. This method might appear to be the most suitable, because it allows the examiner to observe microscopically the fungus growing directly underneath the cover slip. Microscopic features should be easily discerned, structures should be intact, and many representative areas of growth are available for observation.   

PROCEDURE 58.3 WET MOUNT

Method 1. With a wire bent at a 90-degree angle, cut out a small portion of an isolated colony. The portion should be removed from an intermediate point between the center and the periphery of the colony and should contain a small amount of the supporting agar. 2. Add a drop of lactophenol cotton or aniline blue to a slide and then place the portion on the slide (Fig. 58.8). 3. Place a cover slip in position and apply gentle pressure with a pencil eraser or other suitable object

to disperse the growth and the agar. Examine the slide microscopically. 

Limitations The major disadvantage of the wet mount is that the characteristic arrangement of spores is disrupted when pressure is applied to the cover slip. However, this method is suitable in many situations, because characteristic spores are visible, but their arrangement cannot be determined. The wet mount is not always adequate for making a definitive identification.   

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PROCEDURE 58.4 TEASE MOUNT

Method 1. Place 1 drop of lactophenol cotton or aniline blue stain on a clean microscope slide. 2. With an inoculating needle, gently remove a small portion of a fungal colony. Pick a colony from an area that is neither in the very center nor on the very edge. 3. Place the fungus in the lactophenol cotton or aniline blue stain.

4. With a second needle, tease the fungus apart to form a thin layer. 5. Place a cover slip over the fungus and examine the slide under a microscope. 6. Seal the edges of the cover slip with clear fingernail polish to preserve the mount. Note: Adding 10% polyvinyl alcohol (PVA) preserves the specimen as a permanent stain for mounting.   

PROCEDURE 58.5 MICROSLIDE CULTURE

Method 1. Cut a small block of a suitable agar medium that has been previously poured into a culture dish to a depth of approximately 2 mm. The block may be cut using a sterile scalpel blade or with a sterile test tube that has no lip (which produces a round block). 2. Place a sterile microscope slide on the surface of a culture dish containing sterile 2% agar. Alternatively, place a round piece of filter paper or paper towel in a sterile culture dish, add two applicator sticks, and position the microscope slide on top. 3. Add the agar block to the surface of the sterile microscope slide. 4. With a right-angle wire, inoculate the four quadrants of the agar plug with the organism (Fig. 58.9). 5. Apply a sterile cover slip to the surface of the agar plug. 6. If the filter paper–applicator stick method is used, add a small amount of sterile water to the bottom of the culture dish. Replace the lid of the culture and allow it to incubate at 30°C. 7. After a suitable incubation period (and working inside a biologic safety cabinet), remove the cover slip and place it on a microscope slide containing a drop of lactophenol cotton or aniline blue. Some suggest placing the cover slip near the opening of an incinerator-burner to allow the organism to dry rapidly on the cover slip before adding it to the stain.

8. Observe microscopically for the characteristic shape and arrangement of spores. 9. If the microslide culture is unsatisfactory for microscopic identification, the remaining agar block may be used later if it is allowed to incubate further. The agar plug is then removed and discarded, and a drop of lactophenol cotton or aniline blue is placed on the area of growth and a cover slip is positioned in place. Many laboratorians like to make two cultures on the same slide so that if characteristic microscopic features are not observed on examination of the first culture, the second is available after an additional incubation period. 

Limitations Although this method is ideal for definitive identification of an organism, it is the least practical of all the methods described. It should be reserved for cases in which an identification cannot be made based on an adhesive tape preparation or a wet mount. Caution: Do not make slide cultures of slow-growing organisms suspected to be dimorphic pathogens (e.g., Histoplasma capsulatum, Blastomyces spp., Coccidioides spp., Paracoccidioides brasiliensis, Emmonsia spp., or Sporothrix spp.). Microslide cultures must be observed only after a cover slip has been removed from the agar plug and not while it is in position on top of the agar plug. This method of observation is very dangerous, because it could cause a laboratory-acquired infection.   

CASE STUDY 58.1 A 5-year-old male recently adopted from Haiti is seen by a primary care physician. The boy appears to have an infection of the hair and scalp. The physician suspects a dermatophyte and removes infected hairs by plucking them with forceps. The sterile container with the specimen is sent to the laboratory for culture.

Questions 1. What agar and incubation temperature might the laboratorian choose for primary culture? 2. The laboratorian notices that the fungus is a slow grower,

taking 12 days to grow. What other details should be noted? 3. The colony appears downy, flat, and spreading. It has a light tan front and red-brown reverse. What is the next step for identifying this fungus? 4. Rare club-shaped microconidia and sterile hyphae are noted with terminal chlamydospores. The laboratorian decides the dermatophyte is a Microsporum species. How might Microsporum canis be differentiated from Microsporum audouinii?   

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Chapter Review 1. All of the following are medically dimorphic fungi except: a.  Histoplasma capsulatum b. Blastomyces dermatitidis c.  Coccidioides spp. d. Aspergillus niger 2. Cutaneous mycoses are fungal infections that involve: a. Hair b. Skin c. Nails d. All of the above 3. Which fungus is most often acquired by traumatic implantation into the skin? a.  Histoplasma capsulatum b. Sporothrix spp. c.  Coccidioides spp. d. Talaromyces marneffei 4. What are the optimal temperature and incubation time before a fungal blood culture is reported as negative? a. 37°C; 21 days b. 37°C; 7 days c. 30°C; 21 days d. 30°C; 7 days 5. What is a disadvantage of using the tease mount? a. The preparation cannot be preserved. b. Lactophenol cotton or aniline blue stains uniformly. c. Conidia may be disrupted in the preparation. d. Contamination is possible. 6.  Which of the following organisms is/are partially or completely inhibited by media containing cycloheximide? a.  Cryptococcus spp. b. Candida krusei c.  Aspergillus niger d. All of the above 7. Which stain uses a bleaching agent for fungal detection? a. Gram stain b. Lactophenol cotton blue c. Calcofluor white d. India ink

8.  Which criterion applies to the identification of Histoplasma capsulatum? a. Red pigment on cornmeal agar b. Produces yeast form at 25°C and mold form at 36°C c. Slow-growing fungus (1–4 weeks) d. All of the above 9.  True or False _____ All yeasts should be screened for the presence of Cryptococcus spp. _____ Rice grain agar is used to differentiate Microsporum canis from Trichosporon rubrum. _____ India ink is used to identify Aspergillus niger. _____ Cryptococcus spp. can be confirmed by antibody testing. 10.  Matching: Match each term with the correct description. _____ saprophytic _____ dermatophytes _____ dimorphic _____ aerial mycelium _____ melanized _____ hyaline _____ conidia _____ ascocarp _____ phialide _____ microconidia

a. yeast and filamentous form b. pigmented hyphae c. asexual spores d. saclike structure containing sexual spores e. living on dead or decayed organic matter f. small, round conidia g. tubular conidiophore h. nonpigmented hyphae i. fungus that infects the skin j. hyphae above the surface

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Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses OBJECTIVES 1. Describe where Mucorales, Basidiobolales, and Entomophthorales are found, how they are transmitted to humans, and the diseases they cause. 2. Describe the characteristic colony morphology of the Mucorales, Basidiobolales, and Entomophthorales. 3. Outline the tests needed to diagnose a Trichophyton species. 4. List the key features that distinguish Trichophyton rubrum and Trichophyton mentagrophytes. 5. Compare and contrast the ways Microsporum audouinii and Microsporum canis are spread and the populations at risk. 6. Define ectothrix and endothrix. 7. Explain why diagnosing an opportunistic fungal infection in an immunocompromised patient is difficult. 8. Differentiate mucormycosis and entomophthoromycosis, including the disease presentation, causative agents, predominant patient populations, and treatment. 9. Compare and contrast Aspergillus and Penicillium spp., both macroscopically and microscopically. 10. Discuss the dimorphic molds in relation to their endemic areas, disease states, and associated diagnostic methods for identification.

HYALINE MOLDS TO BE CONSIDERED Current Name Mucorales

Rhizopus spp. Mucor spp. Actinomucor sp. Apophysomyces spp. Cokeromyces sp. Lichtheimia spp. Rhizomucor sp. Saksenaea spp. Syncephalastrum sp. Cunninghamella spp. 

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Entomophthorales Conidiobolus spp. 

Basidiobolales Basidiobolus sp. 

Dermatophytes Trichophyton spp. Microsporum spp. Epidermophyton sp. 

Opportunistic Mycoses Aspergillus spp. Fusarium spp. Geotrichum spp. Acremonium spp. Acrophialophora spp. Arthrographis sp. Beauveria sp. Chrysosporium sp. Coniochaeta spp. Nannizziopsis sp. Onychocola sp. Parengyodontium sp. Penicillium spp. Phialemonium spp. Paecilomyces spp. Purpureocillium sp. Rasamsonia spp. Sarocladium spp. (previously Acremonium) Schizophyllum sp. Talaromyces spp. Thermothelomyces sp. 

Systemic Mycoses Blastomyces spp. Coccidioides spp. Emergomyces spp. Emmonsia sp. Histoplasma capsulatum Paracoccidioides spp.

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

A

B

C

• Fig. 59.1  Rhizopus spp. showing sporangium (A) on a long sporan-

giophore (B) arising from pauciseptate hyphae. Note the characteristic rhizoids (C) at the base of the sporangiophore (×250).

The Mucorales General Characteristics The order Mucorales (Zygomycetes) characteristically produce large, ribbonlike hyphae that are irregular in diameter and contain occasional septa. Because the septa may not be apparent in some preparations, this group sometimes has been characterized as aseptate. The specific identification of these organisms is confirmed by observing the characteristic saclike fruiting structures (sporangia), which produce internally spherical, yellow or brown spores (sporangiospores) (Fig. 59.1). Each sporangium is formed at the tip of a supporting structure (sporangiophore). During maturation, the sporangium becomes fractured, and sporangiospores are released into the environment. Sporangiophores are usually connected to one another by occasionally septate hyphae called stolons, which attach at contact points where rootlike structures (rhizoids) may appear and anchor the organism to the agar surface. Identification of the Mucorales is partly based on the presence or absence of rhizoids and the position of the rhizoids in relation to the sporangiophores. 

Epidemiology and Pathogenesis Although the Mucorales (Rhizopus, Mucor, Actinomucor, Cokeromyces, Rhizomucor, Saksenaea, Apophysomyces, Lichtheimia [Absidia], Syncephalastrum, and Cunninghamella spp.) are a less common cause of infection than the aspergilli, they are an important cause of morbidity and mortality in immunocompromised patients, particularly patients with diabetes mellitus. The organisms have a worldwide distribution and are commonly found on decaying vegetable matter or old bread (bread mold) or in soil. The organism is generally acquired by inhalation or ingestion of spores or through percutaneous routes, followed by subsequent development of infection. Once established, the infection is rapidly progressive, particularly in patients with diabetes mellitus who have infections that involve the sinuses. Other immunocompromised patients who are susceptible to infection with

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Mucorales include patients with hematologic malignancies such as acute leukemia and stem cell, kidney, and liver transplant patients. Immunocompetent individuals may acquire skin infections with these fungi after traumatic injection with contaminated material. These organisms are also commonly identified as contaminants in the clinical laboratory but are also a source of nosocomial or health care–associated infections. 

Spectrum of Disease Immunocompromised patients are at greatest risk, particularly those who have uncontrolled diabetes mellitus and transplant patients who are undergoing prolonged corticosteroid, antibiotic, or cytotoxic therapy. The organisms that cause mucormycosis (an infection caused by Mucorales) have a marked propensity for vascular invasion and rapidly produce thrombosis and necrosis of tissue. One of the most common presentations is the rhinocerebral form, in which the nasal mucosa, palate, sinuses, orbit, face, and brain are involved; each shows massive necrosis with vascular invasion and infarction. Perineural invasion also occurs in mucormycoses and is a potential means of retroorbital spread (i.e., invasion into the brain). Other types of infection involve the lungs and gastrointestinal tract; some patients develop disseminated infection. The Mucorales have also caused localized skin infections in immunocompetent patients with severe burns and infections of subcutaneous tissue in patients who have undergone surgery; infection can also be a result of injury and contamination with spores or soil (Fig. 59.2). 

Laboratory Diagnosis Specimen Collection, Transport, and Processing Blood cultures are not appropriate for diagnosis of mucormycosis. Specimens from deep lesions or tissues and sterile sites should be collected rapidly and aseptically. Sufficient quantity is essential to improve the identification and recovery of the fungal isolate. Samples collected for the diagnosis of rhinocerebral forms of infection should include nasal discharge or scrapings, sinus aspirate, or a tissue specimen from a vascularized tissue. Respiratory samples may include sputum and bronchoalveolar lavage fluids. However, if these respiratory specimens result in negative results, a transbronchial or percutaneous computed tomography–guided biopsy of pulmonary lesions may be considered. These procedures pose significant risk to the patient and should be considered carefully. Separate specimens should be collected for the microbiology laboratory and the histology laboratory. Preservatives used for histologic processes, such as formalin, are inhibitory to fungal growth. Specimens should be transported in sterile containers. Tissue (biopsy specimens) should be moistened by adding a few drops of sterile saline to the container. Specimens should be transported to the laboratory within 2 hours of collection and processed immediately. Mucorales are extremely sensitive to environmental changes. See General

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C

B

A •

Fig. 59.2  Mucormycosis. (A) Orbital involvement in a cancer patient. (B) Necrotic eschar on the hard palate of a cancer patient with rhinocerebral mucormycosis. (C) Chronic nonhealing ulcer after traumatic inoculation. (Courtesy Drs. Gerald Bodey, George Viola, Saud Ahmed, and Mona Shiekh Sroujieh, The University of Texas MD Anderson Cancer Center, Houston, TX.)

Considerations for the Laboratory Diagnosis of Fungal Infections in Chapter 58. 

Direct Detection Methods The diagnosis of mucormycosis is primarily based on direct examination; nucleic acid–based testing; or recovery of the fungus from tissue, body fluids, and exudates. Stains

A mucormycosis may be diagnosed rapidly by examining tissue specimens or exudate from infected lesions in a calcofluor white and potassium hydroxide preparation. If the sample is too thick, a false negative result may occur because of insufficient dissociation of tissues. It is recommended that the negative slide be maintained overnight and reviewed again the next day. Branching, broad-diameter, predominantly nonseptate hyphae are observed (Fig. 59.3). It is important that the laboratory notify the clinician of these findings, because Mucorales grow rapidly, and vascular invasion occurs at a rapid rate.  Antigen-Protein

Antigen-protein–based assays are not used for the diagnosis of mucormycosis. In addition, beta-D-glucan testing is not useful for diagnosis.  Nucleic Acid–Based Testing

Nucleic acid testing may be performed on formalin-fixed, paraffin-embedded, fresh or frozen tissue samples. Polym­ erase chain reaction (PCR) amplification of the internal transcribed spacer, as well as seminested PCR of the 18S ribosomal ribonucleic acid (RNA)/deoxyribonucleic acid (DNA) sequence, has been used to confirm identification in samples that have been identified as histopathology positive. A real-time PCR assay has also been developed that amplifies the cytochrome b gene. Nucleic acid purification from formalin-fixed and paraffin-embedded tissues often results in poor quality of extracted DNA. Therefore, fluorescent in

• Fig. 59.3  Phase-contrast microscopy of a potassium hydroxide preparation of sputum. Note the fragmented portions (arrows) of broad, predominantly nonseptate hyphae of Rhizopus spp.

situ hybridization that does not require DNA extraction or amplification has the potential to improve the identification of the fungi. The technique uses synthetic oligonucleotides specific to the 5.8S and 18S ribosomal ribonucleic acid (rRNA) of the fungi. PCR amplification of fungal genes from the serum of high-risk patient populations has demonstrated the potential for early diagnosis of mucormycosis before the demonstration of tissue pathology.  Cultivation

Growth media containing high concentrations of carbohydrates inhibits the production of asexual fruiting bodies

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

that are required for the proper identification of the Mucorales species. It is therefore recommended that media such as potato dextrose, 2% malt, and cherry decoction (acidic) agars be used for cultivation. Growth and development of the mycelium in the Mucorales occurs within 24 to 48 hours. Subcultures should be incubated at 27°C to 30°C. The colonial morphologic features of the Mucorales allow immediate suspicion that an organism belongs to this group. Colonies characteristically produce a fluffy, white to gray or brown hyphal growth that resembles cotton candy and that diffusely covers the surface of the agar within 24 to 96 hours (Fig. 59.4). The hyphae can grow very fast and may lift the lid of the agar plate (also known as a “lid lifter”). The hyphae appear to be coarse. The entire culture dish or tube rapidly fills with loose, grayish hyphae dotted with brown or black sporangia. The different genera and species of Mucorales cannot be differentiated using colonial morphologic features. 

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• Fig. 59.4  Rhizopus colony.

Approach to Identification Mucorales are characterized by the production of branched, nonseptate, wide mycelia (10 to 20 μm). Sexual reproduction occurs by the formation of a thick walled zygospore; however, morphology of the zygospores is not generally useful for routine identification unless the species is homozygous. Asexual reproduction occurs by the formation of sporangiospores in saclike structures termed sporangiophores. The central axis of the sporangia (multispored structure) is termed the columella (singular) and a swelling of the sporangiophore below the columellae (plural) is termed the apophysis. Some species also produce rhizoids that hold the sporangiophore within the soil or growth substrate. The rhizoids are then connected to a branching root, or stolon. Rhizopus spp. have mostly unbranched sporangiophores with rhizoids that appear opposite the point where the stolon arises, at the base of the sporangiophore (Fig. 59.1). In contrast, Mucor spp. are characterized by sporangiophores that are singularly produced or branched and have a round sporangium at the tip filled with sporangiospores. They do not generally have rhizoids or stolons, which distinguishes this genus from the other genera of the Mucorales (Fig. 59.5). Lichtheimia spp. are characterized by the presence of rhizoids that originate between branched or whorled sporangiophores (Fig. 59.6) along the stolons between the rhizoids. The sporangia of Lichtheimia spp. are pyriform and have a funnel-shaped area (apophysis) at the junction of the sporangium and the sporangiophore. Usually a septum is formed in the sporangiophore just below the sporangium. Lichtheimia produce white, fast-growing wooly colonies that become grayish brown. Other genera that are encountered much less commonly in the clinical laboratory are Rhizomucor, Actinomucor, Cokeromyces, Syncephalastrum, Saksenaea, Cunninghamella, and Apophysomyces spp. and described in Table 59.1. 

Serologic Testing Serology is not useful for diagnosing mucormycosis. Patients with invasive mucormycosis develop Mucorales-specific

• Fig. 59.5  Mucor spp. showing numerous sporangia without rhizoids (×430).

A

B

•

Fig. 59.6  Lichtheimia spp. (A) showing sporangia on long sporangiophores arising from pauciseptate hyphae (B). Note that rhizoids are produced between sporangiophores and not at their bases (×250).

T cells that may be used to monitor the course of the disease but not for early diagnosis. 

Matrix-Assisted Laser Desorption Ionization Timeof-Flight Mass Spectrometry Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) has been evaluated for the identification of Mucorales. This technique has

TABLE 59.1    Morphological Differentiation of the Mucorales, Entomophthorales, and Basidiobolales

Order/ Genus

Species

Macroscopic Morphology

Microscopic Morphology

Mucorales Rhizopus spp.

R. arrhizus R. microsporus R. schipperae

Rapidly produce white cottony colonies that turn brownish to black with the production of sporangiophores. Sporangiophores are unbranched arising singly or in groups, with well-developed rhizoids at the base.

Single or clustered brown sporangiophores. Columellae are ellipsoidal, brown to grey.

Sporangiospores are angular and round to ellipsoidal.

Mucor spp.

M. circinelloides M. indicus M. irregularis M. ramosissimus M. velutinosus

Fast growing white to yellow becoming gray with time. Tall sporangiophores that are simple or branched.

Sporangiospores are hyaline and subspherical to ellipsoidal. M. circinelloides rarely produces chlamydospores, and they are absent in M. ramosissimus. Columellae are present and some species will demonstrate rhizoids, although it is not typical of the genera.

Actinomucor sp.

A. elegans

Growth is slow, colonies appear floccose and white to cream after 7 days becoming brown to beige wooly as they age.

Sporangiophores are hyaline, verticillately branched. Whorls or branches may give rise to secondary branches. Sporangiospores are spherical to ovoidal in a smooth or spiny sporangia, with a subglobose columnella present. Rhizoids and chlamydospores are produced.

Apophysomyces spp.

A. ossiformis A. trapeziformis A. variabilis

Rapid growing white to gray with sporangiophores.

Sporangiophores are smooth-walled, single, unbranched with multispored pyriform sporangia. Columellae are hemispherical, cylindrical, trapezoidal, or ellipsoidal. Apophyses are vase-, bell-, or funnel-shaped.

Cokeromyces sp.

C. recurvatus

Slow growing gray to brown.

Long, recurved, twisted stalks arise from terminal vesicles on unbranched sporangiophores. Sporangia are smooth-walled producing spherical sporangiospores. Yeastlike forms are thinto thick-walled, spherical cells that may produce a ship’s wheel appearance similar to Paracoccidioides brasiliensis when grown on brain-heart infusion or yeast extract peptone agar.

Lichtheimia spp.

L. corymbifera L. ramosa L. ornata

Fast growing white woolly colonies that become grayish brown with age.

Sporangiophores are highly branched in singles or corymbs from stolons. Rhizoids are present. Sporangia are spherical to pyriform. Columellae are hemispherical to ellipsoidal with a conical apophyses. Sporangiospores are smooth, hyaline, ellipsoidal, cylindrical, or subglobose. Irregular giant cells may be present.

Rhizomucor sp.

R miehei R. pusillus

Rapidly growing woolly, gray to brown.

Single or branched sporangiophores on aerial mycelium or stolons. Sporangia are multispored with no apophyses. Sporangiospores are round, hyaline, and smooth-walled.

Saksenaea spp.

S. erythrospora S. oblongispora S. vasiformis

White to gray colonies.

Unbranched sporangiophores with dark (melanized) rhizoids. Flask-shaped sporangia.

Syncephalastrum sp.

S. racemosum

Rapid growing white to gray, turns darker with age.

Mostly branched sporangiophores bearing globose vesicles. Smooth-walled sporangia with spherical to ovoid merospores arranged in rows. Rhizoids present.

Cunninghamella spp.

C. bertholletiae C. blakesleeana C. echinulate C. elegans

White to dark gray.

Laterally branching sporangiophores with a globose vesicle that bears a 1-spored sporangiola. Sporangiola will become a finely echinulate spherical sporangiospore.

C. coronatus C. incongruous C. lamprauges

Fast growing hyaline radially folded. Initially appear waxy and become powdery with age.

Primary conidia are spherical with prominent papilla. Villose conidia appear as the colony ages.

B. ranarum

Colonies appear as yellow, waxy with radial folds.

Primary conidiophores have swollen apices and discharge spherical conidia. Secondary conidia are pyriform with a knoblike tip. Large aseptate mycelia are produced that break into uninucleate hyphal elements.

Entomophthorales Conidiobolus spp.

Basidiobolales Basidiobolus sp.

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

demonstrated a high correlation with sequencing methods for the identification of fungal isolates (97%), indicating a potential for use as a routine identification method in the clinical laboratory. 

The Entomophthorales and Basidiobolales General Characteristics The subdivision Entomophthoromycotina contains more than 250 species distributed worldwide. However, only four species have been identified as significant in clinical samples: Order Entomophthorales; Conidiobolus coronatus, Conidiobolus lamprauges, Conidiobolus incongruus, and Order Basidiobolales: Basidiobolus ranarum. 

Epidemiology and Pathogenesis The organisms are pathogens of arthropods and animals and are primarily present in the soil, decaying vegetable material, and animal feces. Although there is a worldwide distribution, infections are more commonly identified in warm climates. Infections associated with Conidiobolus spp. have been identified in Africa, Madagascar, Mayotte, India, China, Japan, and South America. B. ranarum has been associated with infections in India, Myanmar, and Africa. Recent gastrointestinal infections with B. ranarum have been identified in the United States. Unlike mucormycosis, entomophthoromycosis occurs predominantly in immunocompetent individuals. 

Spectrum of Disease Infections from B. ranarum, basidiobolomycosis are primarily localized to subcutaneous tissue of the arms, legs, buttocks, trunk, perineum, face, or neck. Disseminated infection is rare. The infection presents as a woody, hard, painless nodule. Gastrointestinal infections have been noted. Conidiobolus spp. primarily infect the tissue around the nose and on the face. Infection is believed to be through inhalation of the spores in the nasal cavity or inoculation after trauma. After infection, swelling occurs that extends to the nose, cheeks, eyebrows, upper lip, palate, and pharynx. Rare cases of disseminated infection have occurred in immunocompromised patients. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing See General Considerations for the Laboratory Diagnosis of Fungal Infections in Chapter 58. 

Direct Detection Methods Direct examination methods should be used as previously described for the Mucorales. The Splendore-Hoeppli phenomenon (formation of asteroid bodies), the formation of eosinophilic crystals that appear radiate, starlike, asteroid, or club-shaped around a fungal infection, are associated

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with hyphae in tissue sections stained with hematoxylineosin. This is highly indicative of entomophthoromycosis but can also be observed in other bacterial, fungal, and parasitic infections. Antigen-Protein

No antigen tests are currently available.  Nucleic Acid–Based Testing

A single polymerase chain amplification assay has been developed for the diagnosis of Basidiobolus entomophthoromycosis.  Cultivation Tissue samples should be sliced or minced and cultivated on potato dextrose agar or Sabouraud agar without cycloheximide. Because of the variation in growth temperatures required for the isolation of the different organisms, cultures should be incubated at 37°C (Conidiobolus spp.) with a second culture incubated at 25°C to 30°C (Basidiobolus sp.). 

Approach to Identification B. ranarum colonies appear slightly yellow pigmented with radial folds. No aerial hyphae are present. After 7 to 10 days of growth, the culture will produce aseptate mycelia with free uninucleated hyphal elements. Sexual reproduction results in thick-walled zygospores with lateral protuberances or beaks. Primary conidiophores have swollen apices with globose spores that are forcibly discharged from the conidiophores, whereas secondary conidia appear pyriform with a knoblike adhesive tip and are passively discharged. Conidiobolus spp. is a fast-growing fungus that produces hyaline, radially folded colonies that initially appear waxy and become powdery when mycelia begin to develop. The primary conidia are spherical and have prominent papilla (small bumps). Villose (hairlike spines) conidia appear as the colony ages. C. coronatus can be differentiated from the other species based on the absence of zygospores when grown on potato dextrose agar. 

Serologic Testing No serologic tests are available for the diagnosis of entomophthoromycosis. 

The Dermatophytes General Characteristics The dermatophytes produce infections involving the superficial areas of the body, including the hair, skin, and nails (dermatomycoses). The genera Trichophyton, Microsporum, and Epidermophyton are the principal etiologic agents of the dermatomycoses. There is significant information based on molecular classification of these organisms that is not fully accepted or approved and therefore has not been majorly restructured in this edition. Table 59.2 includes an overview of the current dermatophytic genus, species,

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TABLE 59.2    Dermatophytes

Species [Proposed New Name]

Macroscopic Morphology

Microscopic Morphology

Trichophyton spp.

T. ajelloi [Arthroderma uncinatum] T. concentricum T. equinum T. erinacei T. megninii T. mentagrophytes complex T. rubrum T. schoenleinii T. simii T. soudanense T. terrestre complex T. tonsurans T. vanbreuseghemii [Arthroderma gertleri] T. verrucosum T. violaceum

Species vary from powdery to granular and cottony. Coloration is from white to yellow, or pink. Various reverse colors from white to yellow and red to brown.

Species dependent; macroconidia may or may not be present. Macroconidia are generally pencil or club shaped. Microconidia may be tear dropped or round when present.

Microsporum spp.

M. audouinii M. canis M. cookei complex [Genus Paraphyton] M. ferrugineum M. gallinae [Lophophyton gallinae] M. gypseum complex [Genus Nannizzia] M. nanum [Nannizzia nana] M. persicolor [Nannizzia persicolor] M. praecox [Nannizzia praecox] M. racemosum M. vanbreuseghemii [Lophophyton gallinae]

Generally powdery pink to buff with reverse yellow to rose or red-brown.

Macroconidia are generally smooth to rough with tapered ends (rowboat appearance). Microconidia are not present. M. nanum macroconidia appear egg-shaped or ellipsoidal.

Epidermophyton sp.

E. floccosum

Granular, tan to olive brown with reverse tan to yellow.

Macroconidia are club-shaped (beaver tail) with fewer than 6 cells. Chlamydospores may be present. Microconidia are absent.

proposed nomenclature, and general characteristics. Table 59.3 also includes those routinely isolated in the clinical laboratory. 

Epidemiology and Pathogenesis The dermatophytes break down and utilize keratin as a source of nitrogen. They usually are incapable of penetrating the subcutaneous tissue, unless the host is immunocompromised, and even then, penetration into the subcutis is rare. Species of the genus Trichophyton are capable of invading the hair, skin, and nails; Microsporum spp. involve only the hair and skin; and Epidermophyton sp. involves the skin and nails. Common species of dermatophytes recovered from clinical specimens, in order of frequency, are Trichophyton rubrum, Trichophyton mentagrophytes, Epidermophyton floccosum, Trichophyton tonsurans, Microsporum canis, and Trichophyton verrucosum. The frequency of recovery of these species may differ by geographic locale. Other geographically limited species are described elsewhere. 

Spectrum of Disease Cutaneous mycoses are perhaps the most common fungal infections of humans. They are usually referred to as tinea (Latin for “worm” or “ringworm”). The gross appearance of the lesion is an outer ring of the active, progressing infection, with central healing within the ring. These infections may be characterized by another Latin noun to designate the area of the body involved; for example, tinea corporis (ringworm of the body); tinea cruris (ringworm of the groin, or “jock itch”); tinea capitis (ringworm of the scalp and hair); tinea barbae (ringworm of the beard); tinea unguium (ringworm of the nail); and tinea pedis (ringworm of the feet, or “athlete’s foot”).

Trichophyton spp. Members of the genus Trichophyton are widely distributed and are the most important and common causes of infections of the feet and nails; they may be responsible for tinea corporis, tinea capitis, tinea unguium, and tinea barbae. They are commonly seen in adult infections, which vary

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

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TABLE 59.3    Characteristics of Dermatophytes Commonly Recovered in the Clinical Laboratory

Dermatophyte

Colonial Morphology

Growth Rate

Microscopic Identification

Microsporum audouiniia

Downy white to salmon-pink colony; reverse tan to salmon-pink.

2 weeks

Sterile hyphae; terminal chlamydoconidia, favic chandeliers, and pectinate bodies; macroconidia rarely seen (bizarre shaped if seen); microconidia rare or absent.

Microsporum canis

Colony usually membranous with feathery periphery; center of colony white to buff over orange-yellow; lemon-yellow or yellow-orange apron and reverse.

1 week

Thick-walled, spindle-shaped, multiseptate, rough-walled macroconidia, some with a curved tip; microconidia rarely seen.

Microsporum cookei complex

Velvety to granular with a wine-red reverse

1week

Thick-walled, rough-walled macroconidia with cellular compartments, no true cross walls; microconidia are teardrop-shaped.

Microsporum gallinae

Flat to velvety with a white surface with pink tinge; red reverse with diffusible pigment

1 week

Smooth to rough-walled macroconidia; thickest cell often at apex; drop-shaped microconidia

Microsporum gypseum

Cinnamon-colored, powdery colony; reverse light tan.

1 week

Thick-walled, rough, elliptical, multiseptate macroconidia; microconidia few or absent.

Epidermophyton floccosum

Center of colony tends to be folded and is khaki green; periphery is yellow; reverse yellowish-brown with observable folds.

1 week

Macroconidia: large, smooth-walled, multiseptate, clavate, and borne singly or in clusters of two or three; microconidia not formed by this species.

Trichophyton mentagrophytes complex

Different colonial types; white, granular, and fluffy varieties; occasional lightyellow periphery in younger cultures; reverse buff to reddish-brown.

7–10 days

Many round to globose microconidia, most commonly borne in grapelike clusters or laterally along the hyphae; spiral hyphae in 30% of isolates; macroconidia are thin-walled, smooth, club-shaped, and multiseptate; numerous or rare, depending on strain.

Trichophyton rubrum

Colonial types vary from white downy to pink granular; rugal folds are common; reverse yellow when colony is young, but wine/red color commonly develops with age.

2 weeks

Microconidia usually teardrop-shaped, most commonly borne along sides of the hyphae; macroconidia usually absent but when present are smooth, thin-walled, and pencilshaped.

Trichophyton schoenleiniia

Irregularly heaped, smooth, white to cream colony with radiating grooves; reverse white.

2–3 weeks

Hyphae usually sterile; many antler-type hyphae seen (favic chandeliers).

Trichophyton tonsurans

White, tan to yellow or rust, suedelike to powdery; wrinkled with heaped or sunken center; reverse yellow to tan to rust red.

7–14 days

Microconidia are teardrop- or club-shaped with flat bottoms; vary in size but usually larger than other dermatophytes; macroconidia rare (balloon forms found when present).

Trichophyton verrucosum

Glabrous to velvety white colonies; rare strains produce yellow-brown color; rugal folds with tendency to skin into agar surface.

2–3 weeks

Microconidia rare, large, and teardrop-shaped when seen; macroconidia extremely rare but form characteristic rat-tail types when seen; many chlamydoconidia seen in chains, particularly when colony is incubated at 37°C.

Trichophyton violaceuma

Port wine to deep violet colony, may be heaped or flat with waxy glabrous surface; pigment may be lost on subculture.

2–3 weeks

Branched, tortuous, sterile hyphae; chlamydoconidia commonly aligned in chains.

aThese

organisms are not commonly seen in the United States.

in their clinical manifestations. Most cosmopolitan species are anthropophilic, or “human loving”; few are zoophilic, primarily infecting animals, and one is geophilic or soil associated. The T. mentagrophytes complex includes several zoophilic and anthropophilic species.

Generally, hairs infected with Trichophyton organisms do not fluoresce under the ultraviolet (UV) light of a Woods lamp. Fungal elements must be demonstrated inside, surrounding and penetrating the hair shaft or within a skin scraping to diagnose a dermatophyte infection by direct

822 PA RT V    Mycology

examination. Confirmation requires recovery and identification of the causative organism. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing See General Considerations for the Laboratory Diagnosis of Fungal Infections in Chapter 58. 

Direct Detection Methods Stains

Calcofluor white or potassium hydroxide preparations reveal the presence of hyaline septate hyphae or arthroconidia (Figs. 58.4 and 59.7). Direct microscopic examination of infected hairs may reveal the hair shaft to be filled with masses of large arthroconidia (4 to 7 μm) in chains, characteristic of an endothrix type of invasion. In other instances, the hair shows external masses of spores that ensheathe the hair shaft; this is characteristic of the ectothrix type of hair invasion. Hairs infected with Trichophyton schoenleinii reveal hyphae and air spaces within the shaft. Evolve Procedure 59.1 describes the hair perforation test used for the differentiation of Trichophyton spp.  Antigen-Protein

Antigen-protein–based assays are not useful for the detection or identification of dermatophytes.  Nucleic Acid–Based Testing

Nucleic acid amplification assays for dermatophytes are not routine. Current traditional procedures are more cost effective for superficial infections. 

• Fig. 59.7  Calcofluor white stain of sputum showing intracellular yeast

cells of Histoplasma capsulatum (arrows). The cells are 2 to 5 μm in diameter.

Cultivation

Because the dermatophytes generally present a similar microscopic appearance in infected hair, skin, or nails, final identification typically is made by culture. A summary of the colonial and microscopic morphologic features of these fungi is presented in Table 59.2. Fig. 59.8 presents an identification schema useful to the clinical laboratory for identification of commonly encountered dermatophytes. The schema begins with the microscopic features of the dermatophytes that may be visible in the initial examination of the culture. In many instances, the primary recovery medium fails to function as well as a sporulation medium. Often the initial growth must be subcultured onto cornmeal agar or potato dextrose agar to induce sporulation. 

Approach to Identification Trichophyton spp.

Microscopically, Trichophyton organisms are characterized by smooth, club-shaped, thin-walled macroconidia with three to eight septa ranging from 4 × 8 μm to 8 × 15 μm. The macroconidia are borne singly at the terminal ends of hyphae or on short conidiophores; the microconidia (which may be described as “birds on a fence”) predominate and are usually spherical, pyriform (teardrop-shaped), or clavate (club-shaped) and 2 to 4 μm (Fig. 59.9). Only the common Trichophyton species are described here. T. rubrum and T. mentagrophytes complex are the most common species recovered in the clinical laboratory. T. rubrum is a slow-growing organism that produces a flat or heaped-up colony, generally white to reddish, with a cottony or velvety surface. The characteristic cherry-red color is best observed on the reverse side of the colony; however, this is produced only after 3 to 4 weeks of incubation. Occasional strains may lack the deep red pigmentation on primary isolation. Two types of colonies may be produced: fluffy and granular. Microconidia are uncommon in most of the fluffy strains and more common in the granular strains; they occur as small, teardrop-shaped conidia often borne laterally along the sides of the hyphae (Fig. 59.9). Macroconidia are less common, although they are sometimes found in the granular strains, where they appear as thin-walled, smooth-walled, multicelled, cigar-shaped conidia with three to eight septa. T. rubrum has no specific nutritional requirements. It does not perforate hair in vitro or produce urease. T. mentagrophytes complex produce two distinct colonial forms: the downy variety recovered from patients with tinea pedis and the granular variety recovered from lesions acquired by contact with animals. The rapidly growing colonies may appear as white to cream-colored or yellow, cottony or downy, and coarsely granular to powdery. They may produce a few spherical microconidia. The granular colonies may show evidence of red pigmentation. The reverse side of the colony is usually rose-brown, occasionally orange to deep red, and may be confused with T. rubrum. Granular colonies sporulate freely, with numerous small, spherical

Microscopic examination

Large, smooth-walled club-shaped MACROCONIDIA

CONIDIA usually not present only HYPHAE seen

Many MICROCONIDIA smooth elongated macroconidia few or absent

Large rough-walled many celled MACROCONIDIA microconidia few or absent

UREA AGAR

Colony khaki colored

Spindleshaped some with curved tip

Broadly spindle-shaped with rounded ends

Rare bizarre shaped Usually see only terminal chlamydospores

Positive in 2 days

Negative in 2 days

Potato dextrose agar

Cornmeal dextrose agar

Reverse of colony orange or yellow

Colony cinnamon color

Colony salmon color

Star-shaped powdery colonies

No growth on rice medium

Round grapelike clusters of MICROCONIDIA on cornmeal agar Few SPIRAL HYPHAE present in some cultures

Red pigment

No red pigment

Tear-shaped MICROCONIDIA along hyphae on cornmeal agar

Balloon-shaped MICROCONIDIA on cornmeal agar

Colony violet color

Colony white and wrinkled

Colony very slow growing heaped up partially submerged in medium Colony surface smooth without aerial hyphae

TRICHOPHYTON AGAR 4

ANTLER HYPHAE present

ANTLER HYPHAE sometimes present

Increased growth

Increased growth at 37°C with production of chains of CHLAMYDOSPORES with septa appearing like fission planes

TRICHOPHYTON AGAR 4

TRICHOPHYTON AGAR 4 Increased growth

Increased growth Epidermophyton floccosum

Microsporum canis

Microsporum gypseum (Nannizzia gypsea)

Microsporum audouinii

Trichophyton mentagrophytes

Trichophyton rubrum

Trichophyton tonsurans

Trichophyton violaceum

Trichophyton schoenleinii

• Fig. 59.8  Dermatophyte identification schema. (Modified from Koneman EW, Roberts GD. Practical Laboratory Mycology. 3rd ed. Baltimore: Williams & Wilkins; 1985.)

Trichophyton verrucosum

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Mycosel agar

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A

• Fig. 59.9  Trichophyton rubrum showing numerous pyriform microconidia borne singly on hyphae (×750).

microconidia in grapelike clusters and thin-walled, smoothwalled, cigar-shaped macroconidia measuring 6 × 20 μm to 8 × 50 μm, with two to five septa (Fig. 59.10). Macroconidia characteristically exhibit a definite narrow attachment to their base. Spiral hyphae may be found in one third of the isolates recovered. T. mentagrophytes complex species produce urease within 2 to 3 days after inoculation onto Christensen’s urea agar. Unlike T. rubrum, T. mentagrophytes complex perforate hair (Fig. 59.11), a feature that may be used to distinguish between the two fungi when differentiation is difficult. T. tonsurans is responsible for an epidemic form of tinea capitis that commonly occurs in children and occasionally in adults. It has displaced Microsporum audouinii as a primary cause of tinea capitis in most of the United States. The fungus causes a low-grade superficial lesion of varying severity and produces circular, scaly patches of alopecia (loss of hair). The stubs of hair remain in the epidermis of the scalp after the brittle hairs have broken off, which may give the typical “black dot” ringworm appearance. Because the infected hairs do not fluoresce under a Woods lamp, the physician should carefully search for the embedded stubs, using a bright light. Cultures of T. tonsurans develop slowly and are typically buff to brown, wrinkled, and suedelike in appearance. The colony surface shows radial folds and often develops a craterlike depression in the center with deep fissures. The reverse side of the colony is yellowish- to reddish-brown. Microscopically, numerous microconidia with flat bases are produced on the sides of hyphae. With age, the microconidia tend to become pleomorphic, are swollen to elongated, and are referred to as balloon forms (Fig. 59.12). Chlamydoconidia are abundant in old cultures; swollen and fragmented hyphal cells resembling arthroconidia may be seen. T. tonsurans grows poorly on media lacking enrichments (casein agar); however, growth is greatly enhanced by the presence of thiamine or inositol in casein agar. T. verrucosum causes a variety of lesions in cattle and in humans; it is most often seen in farmers, who acquire the infection from cattle. The lesions are found chiefly on the beard, neck, wrist, and back of the hands; they are deep,

B

• Fig. 59.10  A, Trichophyton mentagrophytes showing numerous microconidia in grapelike clusters. B, Several thin-walled macroconidia also are present (×500).

• Fig. 59.11  Hair perforation by Trichophyton mentagrophytes. Wedgeshaped areas (arrow) illustrate hair perforation (×100).

A

B

• Fig. 59.12  Trichophyton tonsurans showing numerous microconidia (A) that are borne singly or in clusters. A single macroconidium (B) (rare) is also present (×600).

pustular, and inflammatory. With pressure, short stubs of hair may be recovered from the purulent lesion. Direct examination of the hair shaft reveals sheaths of isolated chains of large spores (5 to 10 μm in diameter) surrounding the hair shaft (ectothrix) and hyphae within the hair (endothrix). Masses of these conidia may also be seen in exudate from the lesions.

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

T. verrucosum grows slowly (14 to 30 days); growth is enhanced at 35°C to 37°C and on media enriched with thiamine and inositol. T. verrucosum may be suspected when slowly growing colonies appear to embed themselves into the agar surface. Kane and Smitka1 described a medium for the early detection and identification of T. verrucosum. The ingredients for this medium are 4% casein and 0.5% yeast extract. The organism is recognized by its early hydrolysis of casein and very slow growth rate. Chains of chlamydoconidia are formed regularly at 37°C. Early detection of hydrolysis, the formation of characteristic chains of chlamydoconidia, and the restrictive slow growth rate of T. verrucosum differentiate it from T. schoenleinii, another slowly growing organism. Colonies are small, heaped, and folded; occasionally they are flat and disk-shaped. At first, they are glabrous and waxy, with a short aerial mycelium. Colonies range from gray and waxlike to bright yellow. The reverse of the colony most often is nonpigmented but may be yellow. Microscopically, chlamydoconidia in chains and antler hyphae may be the only structures observed microscopically in cultures of T. verrucosum (Fig. 58.10). Chlamydoconidia may be abundant at 35°C to 37°C (Fig. 58.16). Microconidia may be produced by some cultures if the medium is enriched with yeast extract or a vitamin (Fig. 59.13). Conidia, when present, are borne laterally from the hyphae and are large and clavate. Macroconidia are rarely formed, vary considerably in size and shape, and are referred to as “rat tail” or “string bean” in appearance. T. schoenleinii causes a severe type of infection called favus. It is characterized by the formation of yellowish cup-shaped crusts, or scutulae, on the scalp; considerable scarring of the scalp; and sometimes permanent alopecia. Infections are common among members of the same family. A distinctive invasion of the infected hair, the favic type, is demonstrated by the presence of large, inverted cones of hyphae and arthroconidia at the base of the hair follicle and branching hyphae throughout the length of the hair shaft. Longitudinal tunnels or empty spaces appear in the hair shaft where the hyphae have disintegrated. In calcofluor white or potassium hydroxide preparations, these tunnels

•

Fig. 59.13  Trichophyton verrucosum showing microconidia, which are rarely seen (×500).

825

are readily filled with fluid; air bubbles may also be seen in these tunnels. T. schoenleinii is a slowly growing organism (30 days or longer) that produces a white to light-gray colony with a waxy surface. Colonies have an irregular border consisting mostly of submerged hyphae, which tend to crack the agar. The surface of the colony is usually nonpigmented or tan, furrowed, and irregularly folded. The reverse side of the colony is usually tan or nonpigmented. Microscopically, conidia commonly are not formed. The hyphae tend to become knobby and club-shaped at the terminal ends, with the production of many short lateral and terminal branches (Fig.  59.14). Chlamydoconidia are generally numerous. All strains of T. schoenleinii may be grown in a vitamin-free medium and grow equally well at room temperature or at 35°C to 37°C. Trichophyton violaceum causes an infection of the scalp and body and is seen primarily in people living in the Mediterranean region, the Middle and Far East, and Africa. Hair invasion is of the endothrix type; the typical “black dot” type of tinea capitis is observed clinically. Direct microscopic examination of a calcofluor white or potassium hydroxide preparation of the nonfluorescing hairs shows dark, thick hairs filled with masses of arthroconidia arranged in chains, similar to those seen in T. tonsurans infections. Colonies of T. violaceum are very slow growing, beginning as cone-shaped, cream-colored, glabrous colonies. Later these become heaped up, verrucous (warty), violet to purple, and waxy in consistency. Colonies may often be described as “port wine” in color. The reverse side of the colony is purple or nonpigmented. Older cultures may develop a velvety area of mycelium and sometimes lose their pigmentation. Microscopically, microconidia and macroconidia generally are not present; only sterile, distorted hyphae and chlamydoconidia are found. In some instances, however, swollen hyphae containing cytoplasmic granules may be seen. Growth of T. violaceum is enhanced on media containing thiamine.  Microsporum spp.

Species of the genus Microsporum are immediately recognized by the presence of large (8 to 15 μm × 35 to 150 μm),

• Fig. 59.14  Trichophyton schoenleinii showing swollen hyphal tips with lateral and terminal branching (favic chandeliers). Microconidia and macroconidia are absent (×500).

826 PA RT V    Mycology

spindle-shaped, echinulate (covered with small spines), rough-walled macroconidia with thick walls (up to 4 μm) containing four or more septa (Fig. 59.15). The exception is Microsporum nanum, which characteristically produces macroconidia with two cells. Microconidia, when present, are small (3 to 7 μm) and club-shaped and are borne on the hyphae, either laterally or on short conidiophores. Cultures of Microsporum spp. develop either rapidly or slowly (5 to 14 days) and produce aerial hyphae that may be velvety, powdery, glabrous, or cottony, varying in color from whitish, to buff, to a cinnamon-brown, with varying shades on the reverse side of the colony. M. audouinii was once the most important cause of epidemic tinea capitis among school children in the United States. This organism is anthropophilic and is spread directly by means of infected hairs on hats, caps, upholstery, combs, or barber clippers. Most infections are chronic; some heal spontaneously, whereas others may persist for several years. Infected hair shafts fluoresce yellow-green under a Woods lamp. Colonies of M. audouinii generally grow more slowly than other members of the genus Microsporum (10 to 21 days), and they produce a velvety aerial mycelium that is colorless to light gray to tan. The reverse side often appears salmon-pink to reddish-brown. Colonies of M. audouinii do not usually sporulate in culture. The addition of yeast extract may stimulate growth and the production of macroconidia in some instances. Most commonly, atypical vegetative forms, such as terminal chlamydoconidia and antler and racquet hyphae, are the only clues to the identification of this organism. M. audouinii often is identified as a cause of infection by exclusion of all the other dermatophytes. M. canis is primarily a pathogen of animals (zoophilic); it is the most common cause of ringworm infection in dogs and cats in the United States. Children and adults acquire the disease through contact with infected animals, particularly puppies and kittens, although human-to-human transfer has been reported. Hairs infected with M. canis fluoresce a bright yellow-green under a Woods lamp, which is a useful tool for screening pets as possible sources of human infection. Direct examination of a calcofluor white or potassium

hydroxide preparation of infected hairs reveals small spores (2 to 3 μm) outside the hair. Culture must be performed to provide the specific identification. Colonies of M. canis grow rapidly, are granular or fluffy with a feathery border, white to buff, and characteristically have a lemon-yellow or yellow-orange fringe at the periphery. On aging, the colony becomes dense and cottony and a deeper brownish-yellow or -orange and frequently shows an area of heavy growth in the center. The reverse side of the colony is bright yellow, becoming orange- or reddish-brown with age. In rare cases, strains are recovered that show no reverse-side pigment. Microscopically, M. canis shows an abundance of large (15 to 20 μm × 60 to 125 μm), spindleshaped, multisegmented (four to eight) macroconidia with curved ends (Fig. 59.15). These are thick-walled with spiny (echinulate) projections on their surfaces. Microconidia are usually few in number, but large numbers occasionally may be seen. Microsporum gypseum complex are free-living fungi of the soil (geophilic) that only rarely causes human or animal infection and occasionally may be seen in the clinical laboratory. Infected hairs generally do not fluoresce under a Woods lamp. However, microscopic examination of the infected hairs shows them to be irregularly covered with clusters of spores (5 to 8 μm), some in chains. These arthroconidia of the ectothrix type are considerably larger than those of other Microsporum species. M. gypseum complex species grow rapidly as flat, irregularly fringed colonies with a coarse, powdery surface that appear to be buff or cinnamon colored. The underside of the colony is orange to brownish. Microscopically, macroconidia are seen in large numbers and are characteristically large, are ellipsoidal, have rounded ends, and are multisegmented (three to nine) with echinulated surfaces (Fig. 59.16). Although they are spindleshaped, these macroconidia are not as pointed at the distal ends as those of M. canis. The appearance of the colonial and microscopic morphologic features is sufficient to make the distinction between M. gypseum complex and M. canis. 

• Fig. 59.15  Large, rough-walled macroconidia of Microsporum canis

•

(×430).

Fig. 59.16  Microsporum gypseum showing ellipsoidal, multicelled macroconidia (×750).

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

•

Fig. 59.17  Epidermophyton floccosum showing numerous smooth, multiseptate, thin-walled macroconidia that appear club-shaped (×1000).

Epidermophyton sp.

E. floccosum, the only member of the genus Epidermophyton, is a common cause of tinea cruris and tinea pedis. Because this organism is susceptible to cold, specimens submitted for dermatophyte culture should not be refrigerated before culture, and cultures should not be stored at 4°C. In direct examination of skin scrapings using the calcofluor white or potassium hydroxide preparation, the fungus is seen as fine branching hyphae. E. floccosum grows slowly; the growth appears olive-green to khaki, with the periphery surrounded by a dull orange-brown. After several weeks, colonies develop a cottony white aerial mycelium that completely overgrows the colony; the mycelium is sterile and remains so even after subculture. Microscopically, numerous smooth, thin-walled, club-shaped, multiseptate (2 to 4 μm) macroconidia are seen (Fig. 59.17). They are rounded at the tip and are borne singly on a conidiophore or in groups of two or three. Microconidia are absent, spiral hyphae are rare, and chlamydoconidia are usually numerous. The absence of microconidia is useful for differentiating this organism from Trichophyton spp.; the morphology of the macroconidia (smooth, thin-walled) is useful for differentiating it from Microsporum spp. 

Serologic Testing Serology is not useful for the diagnosis of disease caused by dermatophytes. 

The Opportunistic Mycoses General Characteristics The tissue-invasive opportunistic mycoses are a group of fungal infections that occur almost exclusively in immunocompromised patients. Opportunistic fungal infections are typically identified in a host compromised by some underlying disease process, such as lymphoma, leukemia, diabetes mellitus, or another defect of the immune system. Many patients, particularly those who undergo some type of transplantation, are placed on treatment with corticosteroids,

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cytotoxic drugs, or other immunosuppressive agents to control rejection of the transplanted organ. Many fungi previously believed to be nonpathogenic are now recognized as etiologic agents of opportunistic fungal infections. Because most of the organisms known to cause infection in this group of patients are commonly encountered in the clinical laboratory as saprobes (saprophytic fungi), it may be impossible for the laboratorian to determine the clinical significance of these isolates recovered from clinical specimens. Laboratories must identify and report completely the presence of all fungi recovered, because each is a potential pathogen. Many of the organisms associated with opportunistic infections are acquired during construction, demolition, or remodeling of buildings or are hospital-acquired. Other information about the specific clinical aspects of the opportunistic fungal infections is discussed with the individual organism. 

Epidemiology and Pathogenesis Aspergillus spp. Several Aspergillus spp. are among the most commonly encountered fungi in the clinical laboratory (Table 59.4); any are potentially pathogenic in an immunocompromised host, but some species are more commonly associated with disease. There are over 350 species of Aspergillus widespread in the environment, where they colonize grain, leaves, soil, and living plants. Conidia of the aspergilli are easily dispersed into the environment, and humans become infected by inhaling them. Assessing the significance of Aspergillus organisms in a clinical specimen may be difficult. They are commonly found in cultures of respiratory secretions, skin scrapings, and other specimens. 

Pathogenesis and Spectrum of Disease Aspergillus spp. Aspergillus spp. can cause disease by ingestion of mycotoxins, traumatic inoculation, or inhalation. Aspergillus spp. are capable of causing disseminated infection, as is seen in immunocompromised patients, but also of causing a wide variety of other types of infections, including a pulmonary or sinus fungus ball, allergic bronchopulmonary aspergillosis, external otomycosis (a fungus ball of the external auditory canal), mycotic keratitis, onychomycosis (infection of the nail and nail bed), sinusitis, endocarditis, and central nervous system (CNS) infection. Most often, immunocompromised patients acquire a primary pulmonary infection that becomes rapidly progressive and may disseminate to virtually any organ. 

Fusarium spp. and Other Hyaline Septate Opportunistic Molds Molecular phylogenetic studies have indicated that organisms that were previously considered individual more accurately represent species complexes containing more than 60 different species. The most commonly isolated organisms within this group are within the Fusarium solani species complex, including F. petroliphilum, F. keratoplasticum, F. falciforme,

828 PA RT V    Mycology

TABLE 59.4    Clinically Relevant Aspergillus spp.

Species

Macroscopic Morphology

Microscopic Morphology

Seriation

A. fumigatus

Dark blue-green to gray with age; reverse variable

Smooth nonpigmented to green conidiophores short to long with a foot cell at the base; dome shaped vesicle with spherical, roughwalled conidia. Septate hyphae.

Uniseriate; columnar.

A. flavus

Yellow to dark yellow-green

Rough nonpigmented conidiophore; subglobose to globose vesicle; conidia globose (spherical) or ellipsoidal.

Uniseriate and biseriate; loosely radiate or splits into columns with age.

A. nidulansa

Dark green, buff to purplebrown; reverse red to purple

Smooth brown conidiophore; hemispherical vesicle; rough, globose conidia; cleistothecia globose and reddish-brown.

Biseriate columnar.

A. niger

Black with white margin, may have yellow surface mycelium; reverse nonpigmented to pale yellow

Smooth nonpigmented to brown conidiophore; globose vesicle; thick-walled brownish-black, rough conidia.

Biseriate radiate that splits into columns with age.

A. terreus

Tan to cinnamon-brown

Smooth, nonpigmented conidiophore; domeshaped vesicle; smooth subglobose, globose, or elliptical conidia. Single-celled conidia (aleurioconidia) may be present on submerged hyphae.

Biseriate columnar.

A. ustus

Brown-gray or olive-gray; reverse yellow to red or purple

Smooth nonpigmented to brown conidiophore; subglobose to globose vesicle; rough, globose conidia.

Biseriate; radiate to loosely columnar.

A. versicolor

Green to gray or tan with patches of pink or yellow; reverse deep red to variable

Ovate to elliptical vesicle; globose echinulate conidia.

Biseriate; radiate to loosely columnar.

aToxigenic

species.

F. solani, F. lichenicola, and F. neocosmoporiellum. The second most common group responsible for human disease is the Fusarium oxysporum species complex. Additional groups of Fusarium spp. that are clinically relevant include the Fusarium fujikuroi species complex, Fusarium incarnatum-Fusarium equiseti species complex, Fusarium chlamydosporum species complex, and the Fusarium dimerum species complex. Infection caused by Fusarium spp. and other hyaline septate monomorphic molds is becoming more common, particularly in immunocompromised patients. These organisms are common environmental microbiota and have long been known to cause mycotic keratitis after traumatic implantation into the cornea. Oftentimes infections are associated with the consumption of grains contaminated with trichothecene mycotoxins produced by F. sporotrichioides or F. poae. Disseminated fusariosis is commonly accompanied by fungemia, which is detected by routine blood culture systems. In contrast, the aspergilli are rarely recovered from blood culture, even in cases of endovascular infection. Necrotic skin lesions are common with disseminated fusariosis. Other types of infection caused by Fusarium spp. include sinusitis, wound (burn) infection, allergic fungal sinusitis, and endophthalmitis. Fusarium spp. are commonly recovered from respiratory tract secretions, skin, and other specimens from patients

who show no evidence of infection. Interpretation of culture results rests with the clinician and is often assisted by correlation with histopathology results. Geotrichum candidum is an uncommon cause of infection but has been shown to cause wound infections and oral thrush; it is an opportunistic pathogen in immunocompromised hosts. Acremonium spp. are also recognized as important pathogens in immunocompromised hosts; these have been associated with disseminated infection, fungemia, subcutaneous lesions, and esophagitis. Penicillium spp. includes more than 250 recognized species and are among the most common organisms recovered by the clinical laboratory. In North America, they are rarely associated with invasive fungal disease. However, they may be a cause of allergic bronchopulmonary penicilliosis or chronic allergic sinusitis. Talaromyces marneffei is an important and emerging pathogen in Southeast Asia and is discussed further in the section on dimorphic pathogens. Of the Purpureocillium species, Purpureocillium lilacinum appears to be the most pathogenic species and has been associated with endophthalmitis, cutaneous infections, and arthritis. Paecilomyces variotii complex includes five species with P. variotii and P. formosus being the most important pathogens, causing endocarditis, fungemia, and invasive disease. A variety of other saprobic fungi are not discussed here in detail and may be encountered in the clinical laboratory but

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

are seen less commonly. Several are included in Table 59.5. Other references are recommended for further information about identification of these organisms. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing See General Considerations for the Laboratory Diagnosis of Fungal Infections in Chapter 58. 

Direct Detection Methods Stains

Specimens submitted for direct microscopic examination containing organisms in this group demonstrate septate hyphae that usually show evidence of dichotomous branching, often of 45 degrees (Fig. 59.18). In addition, some hyphae may have rounded thick-walled cells. Although often considered to represent an Aspergillus species, these

TABLE 59.5    Other Opportunistic Fungal Organisms

Macroscopic Morphology

Microscopic Morphology

Associated with colonization in cystic fibrosis patients; keratitis, pulmonary infections, and brain abscesses.

Pale white and darken centrally to a gray or brown as the colony ages.

Unbranched, brown echinulate conidiophores, anchored by a foot-cell. Conidia are in chains and may have a distinct band.

A. kalrae

Rare opportunist that has been recovered from skin, lung, corneal ulcers, and sinusitis.

White, appear creamy, yeastlike on artificial media. Become hyphal and buff with yellow reverse.

Treelike conidiophores with lateral branches; arthroconidia.

Beauveria sp.

B. bassiana

Limited virulence in humans; has been isolated in human cases of keratitis.

Colonies are yellow to white.

Produces solitary conidia sympoidally arranged.

Chrysosporium sp.

C. zonatum

Associated with cases of pneumonia and osteomyelitis.

Colonies are yellow to white.

Produces solitary, usually singlecelled aleurioconidia that may be smooth to rough.

Coniochaeta spp.

C. mutabilis C. hoffmannii

Has been isolated in endocarditis and sinusitis in immunocompromised patients.

Colonies are white to salmon, may be moist with darkening black patches.

Phialides are short, stumpy without a basal septum. C. mutabilis forms brown chlamydospores.

Nannizziopsis sp.

N. hominis

White to yellow.

Produces solitary, usually single-celled aleurioconidia. Arthroconidia may be present.

Onychocola sp.

O. canadensis

Associated with onychomycosis.

Raised white to yellow to grayish white.

Two-celled conidia are cylindrical or swollen arthroconidia forming chains.

Parengyodontium sp.

P. album

Identified in cases of endocarditis.

Phialemonium spp.

P. obovatum

Associated with endocarditis.

Rasamsonia spp.

R. aegroticola R. eburnea R. piperina

Emerging pathogen in cystic fibrosis patients.

Genus

Species

Clinical Significance

Acrophialophora spp.

A. fusispora A. levis A. seudatica

Arthrographis sp.

Sarocladium spp. (previously Acremonium)

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Produces solitary conidia sympoidally arranged.

Cuneiform or ellipsoidal conidia.

Several cases of invasive disease have been reported.

Schizophyllum sp.

S. radiatum

Allergy related sinusitis and pulmonary disease.

Thermothelomyces sp.

T. thermophila

Known to cause fatal aortic vasculitis and associated with cerebral abscess and osteomyelitis following traumatic injection.

Produces solitary, usually singlecelled aleurioconidia.

830 PA RT V    Mycology

diagnosis followed by DNA sequencing for identification. Multiplex amplification and real-time PCR assays have been developed for the detection of systemic aspergillosis in respiratory specimens, blood, tissue, and CSF. These assays vary significantly in their performance from 60% to 100% specificity and 40% to 100% sensitivity.  Matrix-Assisted Laser Desorption Ionization Time-ofFlight Mass Spectrometry

The use of MALDI-TOF MS for the identification of fungal isolates has the potential to provide quick and accurate species identification. Numerous studies have demonstrated the utility of this technique for the identification of Aspergillus spp. and other fungal organisms. The performance of MALDI-TOF MS for the identification of Aspergillus spp. has been reported for both young and old colonies with a 98.6% correct identification.  Cultivation

•

Fig. 59.18 Papanicolaou staining of sputum shows the dichotomously branching septate hyphae (arrows) of Aspergillus fumigatus.

cannot be reliably distinguished from hyphae of Fusarium spp., Pseudallescheria boydii, or other hyaline molds.  Antigen-Protein

Antigen-protein–based assays are used to monitor patients at high risk for developing invasive fungal infections. One of these assays, the galactomannan (GM) assay, targets GM, a carbohydrate molecule with a mannose backbone that is released from the cell wall of Aspergillus spp. Aspergillus spp. are the most common source of invasive fungal infections caused by the hyaline septate molds (i.e., hyalohyphomycosis). However, the assay may yield false-positive results because of cross-reactivity with other non-Aspergillus molds, including Penicillium, Rhodotorula, Fusarium, Cryptococcus, Blastomyces, Histoplasma capsulatum, Paecilomyces, and Alternaria spp. The beta-D-glucan assay is designed to detect antigens common to all clinically important fungi. Beta-D-glucan can be detected in the serum of patients infected with systemic aspergillosis. Because the molecule is present in a variety of fungal isolates, the predictive value is not specific to infections with Aspergillus spp. It is recommended that the GM and beta-D-glucan assay be used in combination with other diagnostic tests such as nucleic acid amplification for optimal sensitivity and specificity.  Nucleic Acid–Based Tests

Nucleic acid amplification assays are not commonly performed to detect or identify the opportunistic fungi. However, a variety of both broad-range assays (those that detect all fungi) and species-specific assays have been developed and in specialized centers may be used for patient care. These panfungal PCR assays may be used for initial patient

Because aspergilli are commonly recovered, it is imperative that the organism be demonstrated in the direct microscopic examination of fresh clinical specimens or that it be recovered repeatedly from patients with a compatible clinical picture to ensure that the organism is clinically significant. Correlation with biopsy results is the best means of establishing the significance of an isolate. Most Aspergillus spp. are susceptible to cycloheximide. Specimens submitted for recovery or subculture of these species should be inoculated onto media that lack this ingredient. Aspergillus fumigatus is the most commonly recovered species from immunocompromised patients; moreover, it is the species most often seen in the clinical laboratory. Aspergillus flavus sometimes is recovered from immunocompromised patients and represents a common isolate in the clinical microbiology laboratory. Recovery of A. fumigatus or A. flavus from surveillance (nasal) cultures has been correlated with subsequent invasive aspergillosis; however, the absence of a positive nasal culture does not preclude infection. Aspergillus niger is commonly seen in the clinical laboratory, but its association with clinical disease is somewhat limited; this organism is a cause of fungus ball and otitis externa. Aspergillus terreus is a significant cause of infection in immunocompromised patients, but its frequency of recovery is much lower than that of the previously mentioned species. However, correct identification of A. terreus is important because it is innately resistant to amphotericin B. 

Approach to Identification Aspergillus spp.

A. fumigatus is a rapidly growing mold (2 to 6 days) that produces a fluffy to granular, white to blue-green colony. Mature sporulating colonies most often have a blue-green, powdery appearance. Microscopically, A. fumigatus is characterized by the presence of septate hyphae and short or long conidiophores with a characteristic “foot cell” at their base. The foot cell is T- or L-shaped at the base of the conidiophore, but it is not a separate cell. The tip of the conidiophore expands into

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

a large, dome-shaped vesicle with bottle-shaped phialides covering the upper half or two thirds of its surface. Long chains of small (2 to 3 μm in diameter), spherical, roughwalled, green conidia form a columnar mass on the vesicle (Fig. 59.19). Cultures of A. fumigatus are thermotolerant and able to withstand temperatures up to 45°C. A. flavus is a somewhat more rapidly growing species (1 to 5 days) that produces a yellow-green colony. Microscopically, vesicles are globose, and phialides are produced directly from the vesicle surface (uniseriate) or from a primary row of cells called metulae (biseriate). The phialides give rise to short chains of yellow-orange elliptical or spherical conidia that become roughened on the surface with age (Fig. 59.20). The conidiophore of A. flavus is also coarsely roughened near the vesicle. A. niger produces darkly pigmented, roughened spores macroscopically, but microscopically its hyphae are hyaline and septate, as are those of other aspergilli (i.e., it is not melanized). A. niger produces mature colonies within 2 to 6 days. Growth begins initially as a yellow colony that soon develops a black, dotted surface as conidia are produced. With age, the colony becomes jet black and powdery, but the reverse remains buff or cream colored; this occurs on any culture medium. Microscopically, A. niger shows septate hyphae, long conidiophores supporting spherical vesicles

• Fig. 59.19  Aspergillus fumigatus conidiophore and conidia (×400).

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giving rise to large metulae, and smaller phialides (biseriate), from which long chains of brown to black, rough-walled conidia are produced (Fig. 59.21). The entire surface of the vesicle is involved in sporulation. A. terreus is less commonly seen in the clinical laboratory; it produces tan colonies that resemble cinnamon. Vesicles are hemispherical, as seen microscopically, and phialides cover the entire surface and are produced from a primary row of metulae (biseriate). Phialides produce globose to elliptical conidia arranged in chains. This species produces larger cells, aleurioconidia, which are found on submerged hyphae (Fig. 59.22). 

Serologic Testing The use of serology for Aspergillus spp. has been limited to assistance in the diagnosis of chronic or allergic forms of bronchopulmonary aspergillosis and fungus ball. Serology currently has no value for the diagnosis of disseminated aspergillosis. Fusarium spp.

Colonies of Fusarium spp. grow rapidly, within 2 to 5 days, and are fluffy to cottony and may be pink, purple, yellow, green, or other colors, depending on the species. Microscopically, the hyphae are small and septate

•

Fig. 59.21  Aspergillus niger showing a larger spherical vesicle that gives rise to metulae, phialides, and conidia (×750).

C

A B

• Fig. 59.20  Aspergillus flavus showing spherical vesicles (A) that give

rise to metulae (B) and phialides (C) that produce chains of conidia (×750).

•

Fig. 59.22  Aspergillus terreus showing typical head of Aspergillus and aleurioconidia (arrow) found on submerged hyphae of this species (×500).

832 PA RT V    Mycology

and give rise to phialides producing either single-celled microconidia, usually borne in gelatinous heads similar to those seen in Acremonium spp. (Fig. 58.17) or large, multicelled macroconidia that are sickle- or boat-shaped and contain numerous septations (Fig. 59.23). Some cultures of Fusarium spp. commonly produce numerous chlamydoconidia. The most common medium used to induce sporulation is cornmeal agar. The keys to identification of Fusarium spp. are based on growth on potato dextrose agar. 

rapid growing and may appear yeastlike when initial growth is observed. Mature colonies become white to gray to rose or reddish-orange. Microscopically, small septate hyphae that produce single, unbranched, tubelike phialides are observed. Phialides give rise to clusters of elliptical, singlecelled conidia contained in a gelatinous cluster at the tip of the phialide (Fig. 58.17). The most frequently encountered species in the United States are A. kiliense and A. schlerotigenum-A. egyptiacum group. 

Geotrichum candidum

The genus Penicillium includes three subgenera: Aspergilloides, Furcatum, and Penicillium. Talaromyces is the only species (formerly Penicillium sp.) that is considered a true fungal pathogen and not an opportunist. Colonies of Penicillium spp. are most commonly shades of green or bluegreen, but pink, white, or other colors may be seen. The surface of the colonies may be velvety to powdery because of the presence of conidia. Microscopically, hyphae are hyaline and septate and produce brushlike conidiophores (i.e., penicilli). Conidiophores produce metulae from which flaskshaped phialides producing chains of conidia arise (Fig. 59.26). T. marneffei is discussed in the section on hyaline, septate, dimorphic molds. 

G. candidum often initially appears as a white to cream-colored, yeastlike colony; some isolates may appear as white, powdery molds. Hyphae are septate and produce numerous rectangular to cylindrical to barrel-shaped arthroconidia (Fig. 59.24). Arthroconidia do not alternate but are contiguous, in contrast to Coccidioides spp. (Fig. 59.25). Blastoconidia are not produced.  Acremonium spp.

The Acremonium spp. are a polyphyletic group that includes approximately 100 species. Molecular analysis is currently underway and will undoubtedly result in some additional restructuring of this group. Colonies of Acremonium spp. are

• Fig. 59.23  Fusarium spp. showing characteristic multicelled, sickleshaped macroconidia (×500).

• Fig. 59.24  Geotrichum candidum showing numerous arthroconidia. (Note that arthroconidia do not alternate with a clear (dysjunctor) cell, as in the case of Coccidioides (×430).)

Penicillium spp. and Talaromyces marneffei

•

Fig. 59.25 Mycelial form of Coccidioides spp. showing numerous thick-walled, rectangular, or barrel-shaped (arrows) alternate arthroconidia (×500).

• Fig. 59.26  Penicillium spp. showing typical brushlike conidiophores (penicilli) (×430).

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

833

Paecilomyces spp.

Colonies of Paecilomyces spp. are often velvety, tan to olivebrown, and somewhat powdery. Microscopically, Paecilomyces spp. resemble Penicillium spp. in that a penicillus is formed. However, the phialides of Paecilomyces spp. are long, delicate, and tapering (Fig. 59.27), in contrast to the more blunted phialides of Penicillium spp. The penicillus produces numerous chains of small, oval conidia that are easily dislodged. Single phialides producing chains of conidia may also be present.  Purpureocillium spp.

P. lilacinum exhibits colonies that are considered lilac in color exhibiting shades of lavender to pink. Chlamydospores are absent. The genus demonstrates a slower growth rate than Paecilomyces spp. Optimal growth temperature is 25°C to 33°C. 

•

Fig. 59.27  Paecilomyces spp. showing long, tapering, delicate phialides (arrow).

Scopulariopsis spp.

Scopulariopsis brevicaulis, S. asperula, and S. candida have been associated with onychomycosis, pulmonary infection, fungus ball, and invasive fungal disease in immunocompromised hosts. Colonies of Scopulariopsis spp. initially appear white but later become light brown and powdery. Colonies often resemble those of M. gypseum. Microscopically, a Scopulariopsis resembles a large Penicillium at first glance, because a rudimentary penicillus is produced. Annellophores produce the flask-shaped annelides, which support the lemon-shaped conidia in chains. Conidia are large, have a flat base, and are rough-walled (Fig. 59.28). The Scopulariopsis spp. include both hyaline and dematiaceous species. Scopulariopsis brumptii has been reported to have caused a brain abscess in a liver transplant patient and invasive infection in bone marrow recipients. S. candida and S. acremonium have been identified in association with invasive sinusitis. 

Serologic Testing Serology currently has no value for the diagnosis of the opportunistic disseminated fungal infections discussed. 

Systemic Mycoses As with many other groups of clinically relevant fungi, significant changes have occurred in the classification and taxonomy of these organisms. H. capsulatum is now divided into eight clades (biological taxa claiming a common ancestor), or varieties. Seven of these varieties comprise genetically and geographically distinct populations representing a species; however, the Histoplasma var. duboisii, the African containing clade, demonstrates the same mitochondrial pattern as the H. capsulatum var. capsulatum of North and South America. H. capsulatum is used throughout this section as the primary isolate of interest. The genera Blastomyces and Coccidioides have traditionally been representative of single species, Blastomyces dermatitidis and Coccidioides immitis. More recent characterization has revealed the existence of a subspecies or separate species in the genus Blastomyces, Blastomyces gilchristii.

A

B

•

Fig. 59.28  Scopulariopsis spp. showing a large penicillus (A) with echinulate conidia (B) (×430).

Polyphasic taxonomic analysis has also revealed a correlation between genotypic characteristics of these organisms and clinical phenotypic presentations. Additional species have now been proposed: B. percursus, B. parvus (formerly Emmonsia parva), B. helicus (formerly Emmonsia helica), and B. silverae. In addition, Coccidioides now comprises two species: C. immitis that includes all isolates from California and Washington State and Coccidioides posadasii, which comprises all other isolates. Phylogenetic analysis has indicated that Paracoccidioides brasiliensis can be subdivided into at least three distinct species. Several new species are proposed based on geographic regions of endemicity and include P. lutzii, P. americana, P. restrepiensis, and P. venezuelensis. The new genus Emergomyces contains many of the organisms previously included in the genus Emmonsia. Emergomyces includes Es. pasteurianus as the type species (previously Emmonsia pasteuriana), and four new species: Es. africanus, Es. orientalis, Es. canadensis, and Es. europaeus. Each species differs in yeast size and geographic distribution. Emmonsia crescens can be found in the soil and infects humans in North, Central, and South America as well as in Europe, Asia, and Africa. The distribution of disease associated with this fungi is unclear and further analysis is needed.

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General Characteristics

Coccidioides spp.

Most of the dimorphic fungi produce systemic fungal infections that may involve any of the internal organs of the body, including lymph nodes, bone, subcutaneous tissue, meninges, and skin. The dimorphic fungal pathogens most commonly encountered in North America are H. capsulatum, Blastomyces spp., and Coccidioides spp. Emmonsia spp., Paracoccidioides spp., and Emergomyces spp. are geographically distributed throughout Central and South America. Asymptomatic or subclinical infection is common with H. capsulatum and Coccidioides and may go unrecognized clinically. These infections may be detectable only by serology or after histopathologic review of tissues removed because of lesions found during a roentgenographic examination. Symptomatic infections may present signs of a mild or more severe but self-limited disease, with positive supportive evidence from cultural or immunologic findings. Patients with disseminated or progressive infection have severe symptoms, with spread of the initial disease, often from a pulmonary locus, to several distant organs. However, some cases of disseminated infection may show little in the way of signs or symptoms of disease for long periods, only to undergo exacerbation later. Immunocompromised patients most often present with disseminated infection, particularly those with advanced human immunodeficiency virus (HIV) infection (i.e., acquired immunodeficiency syndrome [AIDS]) or those receiving long-term corticosteroid therapy. The classic term systemic mycoses, used to refer to the dimorphic fungi, is somewhat misleading, because other fungi, including Cryptococcus neoformans complex, Candida albicans complex, and Aspergillus spp., may also cause disseminated systemic infections. 

Coccidioides spp. are found primarily in the desert portion of the southwestern United States and in the semiarid regions of Mexico and Central and South America. Although the geographic distribution of the organism is well defined, infection may be seen in any part of the world because of the ease of travel. The infection (coccidioidomycosis) is usually acquired by inhalation of the infective arthroconidia. The infection is not contagious; however, person-toperson spread has been reported from contaminated fomites or through an infected organ donor to a recipient. 

Epidemiology Blastomyces spp. B. dermatitidis is uncommon as an opportunistic pathogen but may cause aggressive disease in immunocompromised individuals, producing a chronic infection that contains a mixture of suppurative and granulomatous inflammation. The disease (blastomycosis) is most commonly found in North America and extends southward from Canada to the Mississippi, Ohio, and Missouri river valleys; Mexico; and Central America. Some isolated cases have also been reported from Africa. The largest numbers of cases occur in the Mississippi, Ohio, and Missouri river valley regions. B. gilchristii isolates are primarily localized to northwestern Ontario, Wisconsin, and Minnesota. The exact ecologic niche for this organism in nature has not been determined; however, patients with blastomycosis often have a history of exposure to soil or wood, particularly near waterways. Several outbreaks have been reported and have been related to a common exposure. Blastomycosis is more common in males than in females and seems to be associated with outdoor occupations or activities. The disease also occurs in dogs. 

Emmonsia spp. Ea. crescens. are rare causes of human infection. The organism produces a self-limited, localized pulmonary infection that may appear asymptomatic. Diagnosis is generally incidental to other underlying conditions. The organism produces adiaspores, which enlarge but do not reproduce in the patient. The clinical presentation depends on the number of adiaspores inhaled. It is unclear at this time whether the organism demonstrates a species-specific geographic distribution. 

Emergomyces spp. The Emergomyces spp. appear as beige, slow growing filamentous colonies at room temperature. The conidiophores are short and unbranched, and they form at right angles to hyaline hyphae. At body temperature, they appear as small oval yeast cells. The organism is endemic in South Africa and transmitted by inhalation. Emergomyces spp. is rapidly becoming the most commonly diagnosed dimorphic fungal pathogen. 

Histoplasma capsulatum Outbreaks of histoplasmosis have been associated with activities that disperse aerosolized conidia or small hyphal fragments. Infection is acquired through inhalation of these infective structures from the environment. The severity of the disease is generally related directly to the inoculum size and the immunologic status of the host. Numerous cases of histoplasmosis have been reported in people who clean out an old chicken coop or barn that has been undisturbed for long periods and in individuals who work in or clean areas that have served as roosting places for starlings and similar birds. Spelunkers (i.e., cave explorers) are commonly exposed to the organism when it is aerosolized from bat guano in caves. An estimated 500,000 people are infected with H. capsulatum annually. The history of exposure often is impossible to document, even though histoplasmosis is perhaps one of the most common systemic fungal infections seen in the Midwest and South in the United States, including areas along the Mississippi River, the Ohio River valley, and the Appalachian Mountains. 

Paracoccidioides brasiliensis and P. lutzi Infection caused by Paracoccidioides spp. is most commonly found in South America, with the highest prevalences in Brazil, Venezuela, and Colombia. It also has been seen in

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

many other areas, including Mexico, Central America, the Caribbean, and Africa. Occasional imported cases are seen in the United States and Europe. The exact mechanism by which paracoccidioidomycosis is acquired is unclear; however, some speculate that it has a pulmonary origin and that it is acquired by inhalation of the organism from the environment. Because mucosal lesions are an integral part of the disease process, it also is speculated that the infection may be acquired through trauma to the oropharynx caused by vegetation commonly chewed by some residents of the endemic areas. The specific ecologic niche of the organism in nature is not known. 

Talaromyces marneffei T. marneffei is a dimorphic pathogenic fungus endemic to Southeast Asia, particularly the Guangxi Zhuang Autonomous Region of the People’s Republic of China. T. marneffei has been associated with the bamboo rat (Rhizomys pruinosus) and the Vietnamese bamboo rat (Rhizomys sinensis). 

Sporothrix spp. Sporothrix schenckii has been shown to be a complex of numerous species. Those involved in human infection include S. schenckii, Sporothrix brasiliensis, Sporothrix globosa, and Sporothrix luriei. Sporothrix spp. have a worldwide distribution, and their natural habitat is living or dead vegetation. Humans acquire the infection (sporotrichosis) through trauma (thorns, splinters, bites, or scratches), usually to the hand, arm, or leg. The infection is an occupational hazard for farmers, nursery workers, gardeners, florists, and miners; it is commonly known as rose gardener’s disease. Infections with S. brasiliensis have been transmitted from the bites or scratches of stray cats. Pulmonary sporotrichosis rarely occurs as a result of inhalation of spores. 

Pathogenesis and Spectrum of Disease Traditionally, the systemic mycoses have included only blastomycosis, coccidioidomycosis, histoplasmosis, and paracoccidioidomycosis. Although these fungi are morphologically dissimilar, they have one characteristic in common: dimorphism. Most of these organisms, except for Coccidioides, are thermally dimorphic. The dimorphic fungi exist in nature as the mold form, which is distinct from the parasitic or invasive form, sometimes called the tissue form. Distinct morphologic differences may be observed with the dimorphic fungi both in  vivo and in vitro, as discussed later in the chapter.

Blastomyces spp. Blastomyces spp. commonly produce an acute or chronic suppurative and granulomatous infection. Blastomycosis begins as a respiratory infection and is probably acquired by inhalation of the conidia or hyphal fragments of the organism. The infection may spread and involve secondary sites of infection in the lungs, long bones, soft tissue, and skin. 

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Coccidioides spp. Approximately 60% of patients with coccidioidomycosis are asymptomatic and have self-limited respiratory tract infections. However, the infection may become disseminated, with extension to visceral organs, meninges, bone, skin, lymph nodes, and subcutaneous tissue. Fewer than 1% of those who develop coccidioidomycosis ever become seriously ill; dissemination does occur, however, most commonly in individuals of dark-skinned races. Pregnancy also appears to predispose females to disseminated infection. This infection has been known to occur in epidemic proportions. In 1992, an epidemic occurred in northern California, with more than 4000 cases seen in Kern County near Bakersfield. People who visit endemic areas and return to a distant location may present to their local physician; therefore, the endemic mycoses should be considered in the differential diagnosis if the patient has the appropriate travel history. All laboratories should be prepared to deal with the laboratory diagnosis of coccidioidomycosis. 

Emergomyces spp. Es. pasteurianus (previously Emmonsia) does not produce adiaspores (spores that increase in size inside an animal host) in vitro on brain-heart infusion (BHI) agar incubated at 37°C like Emmonsia spp. The organism produces cells that resemble budding yeast. Emergomycosis is generally systemic and includes the appearance of widespread cutaneous lesions. 

Emmonsia spp. Nonreplicating Emmonsia spp., most notably Ea. crescens, produces 25 to 400 μm adiaspores in  vitro on BHI agar incubated at 37°C. In the natural environment the conidia are approximately 2 to 4 μm in diameter but may grow to 500 μm when inhaled into the human lung. The condition associated with inhaled conidia from Emmonsia spp. is referred to as adiaspiromycosis. The severity of the disease depends on the immunologic status of the patient as well as the inoculum size but may range from asymptomatic to fatal. Symptoms include fever, cough, dyspnea, hemoptysis, weight loss, fatigue, and possible respiratory failure. 

Histoplasma capsulatum H. capsulatum most commonly produces a chronic, granulomatous infection (histoplasmosis) that is primary and begins in the lung and eventually invades the reticuloendothelial system. Approximately 95% of cases are asymptomatic and self-limited, although chronic pulmonary infections occur. The disease can be disseminated throughout the reticuloendothelial system; the primary sites of dissemination are the lymph nodes, liver, spleen, and bone marrow. Infections of the kidneys and meninges are also possible. Resolution of disseminated infection is the rule in immunocompetent hosts, but progressive disease is more common in immunocompromised patients (e.g., patients with AIDS). Ulcerative lesions of the upper

836 PA RT V    Mycology

respiratory tract may occur in both immunocompetent and immunocompromised hosts. 

Paracoccidioides spp. Paracoccidioides produces a chronic granulomatous infection (paracoccidioidomycosis) that begins as a primary pulmonary infection. It often is asymptomatic and then disseminates to produce ulcerative lesions of the mucous membranes. Ulcerative lesions are commonly present in the nasal and oral mucosa, gingivae, and less commonly the conjunctivae. Lesions occur commonly on the face in association with oral mucous membrane infection. The lesions are characteristically ulcerative, with a serpiginous (snakelike) active border and a crusted surface. Lymph node involvement in the cervical area is common. Pulmonary infection is common, and progressive chronic pulmonary infection is found in approximately 50% of cases. In some patients, dissemination occurs to other anatomic sites, including the lymphatic system, spleen, intestines, liver, brain, meninges, and adrenal glands. 

thick-walled yeast cells 8 to 15 μm in diameter, usually with a single bud that is connected to the parent cell by a broad base (Figs. 59.29 to 59.31). A smaller form (2 to 8 μm) is seen in rare cases.  Coccidioides spp. In direct microscopic examinations of sputum or other body fluids, Coccidioides spp. appear as a nonbudding, thick-walled spherule, 20 to 200 μm in diameter, that contains either granular material or numerous small (2 to 5 μm in diameter), nonbudding endospores (Figs. 59.32 to 59.35). The endospores are freed by rupture of the spherule wall; therefore, empty and collapsed “ghost” spherules may also be present. Small, immature spherules measuring 5 to 20 μm may be confused with H. capsulatum or Blastomyces spp. Two endospores or immature spherules lying adjacent to one another may give the appearance that

Talaromyces marneffei T. marneffei commonly infects immunosuppressed individuals. The organism causes either a focal cutaneous or mucocutaneous infection, or it may produce a progressive disseminated and commonly fatal infection. Granulomatous, suppurative, and necrotizing inflammatory responses have been demonstrated. The mode of transmission and the primary source in the environment are unknown, but the bamboo rat has been implicated. 

Sporothrix spp.

•

Fig. 59.29  Blastomyces dermatitidis yeast form showing thickwalled, oval to round, single-budding, yeastlike cells (×500).

Sporothrix spp., also dimorphic fungi, are often associated with chronic subcutaneous infections. The primary lesion begins as a small, nonhealing ulcer, often of the index finger or the back of the hand. With time, the infection is characterized by the development of nodular lesions of the skin or subcutaneous tissues at the point of contact and later involves the lymphatic channels and lymph nodes that drain the region. The subcutaneous nodules ulcerate to form an infection that becomes chronic. Only rarely is the disease disseminated. Pulmonary infection may be seen in patients who inhale the spores of Sporothrix spp. 

Laboratory Diagnosis Specimen Collection, Transport, and Processing See General Considerations for the Laboratory Diagnosis of Fungal Infections in Chapter 58. 

Direct Detection Methods Stains

The microscopic morphologic features of the tissue forms, or what has been termed the parasitic forms, of the dimorphic fungi vary with the genus and are described for each. Blastomyces spp. The diagnosis of blastomycosis may easily be made when a clinical specimen is observed by direct microscopy. Blastomyces spp. appear as large, spherical,

•

Fig. 59.30 Potassium hydroxide preparation of exudate shows a large budding yeast cell with a distinct broad base (arrow) between the cells, which is characteristic of Blastomyces dermatitidis. (Phasecontrast microscopy.)

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

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• Fig. 59.34  Histologic section showing a well-developed spherule of Coccidioides spp. that is filled with endospores.

•

Fig. 59.31 Auramine-rhodamine preparation of specimen material from a bone lesion that demonstrates the characteristic broad-based budding yeast (arrow) of Blastomyces dermatitidis.

•

Fig. 59.35  Coccidioides spp. lactophenol cotton blue preparation from Sabouraud agar demonstrating arthroconidia and barrel shaped cells. (Photo courtesy Anna Hartyunyan, MLS (ASCP), Children’s Hospital, Los Angeles, CA.)

•

Fig. 59.32 Tissue form of Coccidioides spp. (i.e., the spherule). The external wall of the spherule does not stain with the silver stain, whereas the internal endospores do stain (arrowhead). Also note how the juxtaposed endospores, which have been released from a spherule that has burst, resemble budding yeast (arrow). (GMS stain; ×400.)

• Fig. 59.33  Potassium hydroxide preparation of sputum demonstrates

two spherules of Coccidioides spp. filled with endospores. When these lie adjacent to each other, they may be mistaken for Blastomyces dermatitidis. (Bright-field microscopy.)

budding yeast is present. When identification of Coccidioides is questionable, a wet preparation of the clinical specimen may be made using sterile saline, and the edges of the cover glass may be sealed with petrolatum and incubated overnight. When spherules are present, the endospores produce multiple hyphal strands.  Emergomyces spp. Emergomyces spp. can be differentiated from Emmonsia by the presence of budding yeasts and the absence of adiaspores.  Emmonsia spp. Emmonsia spp. have not been successfully cultured from human specimens. Therefore, diagnosis is dependent on the histologic appearance of a thick-walled adiaspore granuloma within the lungs. Unlike Coccidioides spp., Emmonsia spp. adiaspores do not contain endospores and are typically much larger than spherules. A recent report has indicated that PCR and DNA sequencing may be useful in diagnosing adiaspiromycosis.  Histoplasma capsulatum. Direct microscopic examination of respiratory tract specimens and other similar specimens often fails to reveal the presence of H. capsulatum. However, an astute laboratorian may detect the organism when examining Wright- or Giemsa-stained specimens of bone marrow and, in rare cases, peripheral blood.

838 PA RT V    Mycology

H. capsulatum is found intracellularly in mononuclear cells as small, round to oval yeast cells 2 to 5 μm in diameter (Fig. 59.36 and Fig. 59.7).  Paracoccidioides brasiliensis. Specimens submitted for direct microscopic examination are important for the diagnosis of paracoccidioidomycosis. Large, round or oval, multiple budding yeast cells (8 to 40 μm in diameter) are usually recognized in sputum, mucosal biopsy specimens, and other exudates. Characteristic multiple budding yeast forms resemble a “mariner’s wheel” (Fig. 59.37). The yeast cells surrounding the periphery of the parent cell range from 8 to 15 μm in diameter. Some cells may be as small as 2 to 5 μm but still exhibit multiple buds.  Talaromyces marneffei. Direct examination of infected tissues and exudates reveals that T. marneffei produces small, yeastlike cells (2 to 6 μm) that have internal cross-walls; no budding cells are produced (Fig. 59.38). Like H. capsulatum, T. marneffei may be detected in peripheral blood smears with disseminated disease.  Sporothrix spp. Exudate aspirated from unopened subcutaneous nodules or from open draining lesions often is submitted for culture and direct microscopic examination. Direct examination of this material usually has little diagnostic value, because demonstrating the rare

characteristic yeast forms is difficult. Sporothrix usually appear as small (2 to 5 μm in diameter), round to oval to cigar-shaped yeast cells (Fig. 59.39). If stained using the periodic acid-Schiff (PAS) method in histologic section, an amorphous pink material may be seen surrounding the yeast cells (Fig. 59.40).  Antigen-Protein

Immunodiffusion methods (the exoantigen test) may be used to identify isolates of these organisms based on precipitation bands of identity between specific antibodies and fungal antigen extracts. However, these assays have been largely replaced by the more rapid nucleic acid hybridization reactions and automated enzyme immunoassays. Antigen testing is available for H. capsulatum and Blastomyces in a microtiter plate double antibody sandwich EIA that can detect the antigens in urine, serum, or CSF in disseminated infections. A urinary antigen test is also available for the

• Fig. 59.36  These small, oval yeast cells that are relatively uniform in size are characteristic of Histoplasma capsulatum (×2000).

•

Fig. 59.37  Paracoccidioides brasiliensis in a bone marrow aspirate shows a yeast cell with multiple buds (arrow).

• Fig. 59.38  Talaromyces marneffei and binary fission (arrows) (×500).

•

Fig. 59.39 The deeply staining bodies in this mouse testis are the yeast forms of Sporothrix spp.

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

• Fig. 59.40  Periodic acid-Schiff (PAS) staining of exudate shows the cigar- to oval-shaped yeast cells (arrows) of Sporothrix spp.

detection of Coccidioides spp. Antigen testing is not currently available for the remaining dimorphic fungal pathogens.  Nucleic Acid Testing

Nucleic acid amplification assays are not routinely performed but are available in some reference laboratories and in research settings. A single FDA-approved assay for the detection of Coccidioides is available. Real-time or homogeneous, rapid-cycle PCR assays have been described for H. capsulatum, Blastomyces, Emmonsia, Paracoccidioides, and Coccidioides species. These assays have proven suitable for isolate identification. Reproducibility and specificity of these assays must be thoroughly evaluated for standardization before implementation in clinical laboratories. No molecular tests are available for Emergomyces spp.  Cultivation

The dimorphic fungi are regarded as slow-growing organisms, requiring 7 to 21 days for visible growth to appear at 25°C to 30°C. However, exceptions to this rule occur with some frequency. Occasionally cultures of Blastomyces and H. capsulatum are recovered in as short a time as 2 to 5 days when many organisms are present in the clinical specimen. In contrast, when a small number of colonies of Blastomyces and H. capsulatum are present, sometimes 21 to 30 days of incubation are required before they are detected. Coccidioides is consistently recovered within 3 to 5 days of incubation, but when many organisms are present, colonies may be detected within 48 hours. Cultures of P. brasiliensis are commonly recovered within 5 to 25 days, with a usual incubation period of 10 to 15 days. The growth rate, if slow, might lead the laboratorian to suspect the presence of a dimorphic fungus; however, considerable variation in the

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time for recovery exists. The exceptions to this slow growth are Coccidioides and T. marneffei, which may be recovered within 3 to 5 days. Textbooks present descriptions of the dimorphic fungi that the reader assumes are typical for each particular organism. As is true in other areas of microbiology, variation in the colonial morphologic features also occurs, depending on the strain and the type of medium used. The laboratorian must be aware of this variation and must not rely heavily on colonial morphologic features to identify members of this group of fungi. The pigmentation of colonies is sometimes helpful but also varies widely; colonies of Blastomyces and H. capsulatum are described as being fluffy white, with a change in color to tan or buff with age. Some isolates initially appear darkly pigmented, with colors ranging from gray or dark brown to red. On media containing blood enrichment, these organisms may appear heaped, wrinkled, glabrous, neutral in color, and yeastlike; often tufts of aerial hyphae project from the top of the colony. Some colonies may appear pink to red, possibly because of the adsorption of hemoglobin from the blood in the medium. Coccidioides is described as fluffy white with scattered areas of hyphae adherent to the agar surface, giving an overall “cobweb” appearance to the colony. However, numerous morphologic forms have been reported, including textures ranging from wooly to powdery and pigmentation ranging from pink-lavender or yellow to brown or buff. The definitive traditional identification method for dimorphic fungus includes observing both the mold and tissue or parasitic forms of the organism. In general, 25°C to 30°C is the optimal temperature for recovery and identification of the dimorphic fungi from clinical specimens. Temperature (35°C to 37°C), certain nutritional factors, and stimulation of growth in tissue independent of temperature are among the factors necessary to initiate the transformation of the mold form to the tissue form. Previously, Blastomyces and H. capsulatum were identified definitively by the in  vitro conversion of a mold form to the corresponding yeast form through in  vitro conversion on a blood-enriched medium incubated at 35°C to 37°C; definitive identification of Coccidioides involved conversion to the spherule form by animal inoculation. Except for Coccidioides, the conversion of dimorphic molds to the yeast form can be accomplished with some difficulty (Evolve Procedure 59.2). Some reference laboratories may still use the exoantigen test (Evolve Procedure 59.3) to identify the dimorphic pathogens. However, this test requires extended incubation before cultures may be identified. Blastomyces dermatitidis. B. dermatitidis commonly requires incubation for 5 days to 4 weeks or longer at 25°C before growth can be detected. However, it may be detected in as short a time as 2 to 3 days. On enriched culture media, the mold form develops initially as a glabrous or waxyappearing colony and is off-white to white. With age, the aerial hyphae often turn gray to brown. The waxy, yeastlike appearance is typified on media enriched with blood. Tufts of hyphae often project upward from the colonies, and this

840 PA RT V    Mycology

has been referred to as the “prickly state” of the organism. However, some isolates appear fluffy on primary recovery and remain so throughout the incubation period. On blood agar at 37°C, colonies are waxy, wrinkled, and yeastlike. Mold-to-yeast conversion usually requires 4 to 5 days. Coccidioides spp. Cultures of Coccidioides are a biohazard to laboratory workers, and strict safety precautions must be followed when cultures are examined. Mature colonies may appear within 2 to 5 days of incubation and may be present on most media, including those used in bacteriology. Laboratory workers are cautioned not to open cultures of fluffy white molds unless they are placed inside a biologic safety cabinet (BSC). Colonies of Coccidioides often appear as a delicate, cobweblike growth after 3 to 21 days of incubation. Some portions of the colony exhibit aerial hyphae, whereas in others the hyphae adhere to the agar surface. Most isolates appear fluffy white; however, colonies of varying colors have been reported, ranging from pink to yellow to purple and black. Some colonies exhibit a greenish discoloration on blood agar, and others appear yeastlike, smooth, wrinkled, and tan. Emmonsia and Emergomyces spp. Emmonsia spp. produce glabrous, colorless colonies at 25°C that produce yellow to white aerial mycelia over time. Some strains produce orange to gray mycelia. Reverse pigmentation appears gray to grayish-brown. Emergomyces spp. appears very similar at 25°C to 30°C and must be differentiated based on the microscopic production of yeastlike cells at 37°C and not adiaspores. Sporulation is enhanced on potato dextrose agar or Pablum cereal agar. The fungi produce conidia from the sides or directly on short stalks that branch from the hyphae. The hyphae may appear swollen and bear peglike structures resulting in the production of secondary conidia in a flowerlike arrangement. Histoplasma capsulatum. H. capsulatum is easily cultured from clinical specimens; however, it may be overgrown by bacteria or rapidly growing molds. A procedure that is useful for recovering H. capsulatum, B. dermatitidis, and Coccidioides spp. from contaminated specimens (e.g., sputa) uses a yeast extract/phosphate medium and a drop of concentrated ammonium hydroxide (NH4OH) placed on one side of the inoculated plate of medium. In the past, it was recommended that specimens not be kept at room temperature before culture, because H. capsulatum would not survive. The organism survives transit in the mail for as long as 16 days. However, the current recommendation is that specimens be cultured as soon as possible to ensure optimal recovery of H. capsulatum and other dimorphic fungi. H. capsulatum is usually considered a slow-growing mold at 25°C to 30°C and commonly requires 2 to 4 weeks or more for colonies to appear. However, the organism may be recovered in 5 days or less if many yeast cells are present in the clinical specimen. Isolates of H. capsulatum have been reported from blood cultures with the Isolator (Alere, Waltham, MA) within a mean time of 8 days. H. capsulatum is a white, fluffy mold that turns brown to buff with age. Some isolates ranging

from gray to red have also been reported. The organism also may produce wrinkled, moist, heaped, yeastlike colonies that are soft and cream colored, tan, or pink. Tufts of hyphae often project upward from the colonies, as described for B. dermatitidis and H. capsulatum, and cannot be differentiated using colonial morphologic features. Paracoccidioides brasiliensis. Colonies of P. brasiliensis grow very slowly (21 to 28 days) and are heaped, wrinkled, moist, and yeastlike. With age, colonies may become covered with a short aerial mycelium and turn tan to brown. The surface of colonies often is heaped with crater formations. Talaromyces marneffei. At 25°C, T. marneffei grows rapidly and produces blue-green to yellowish colonies on Sabouraud agar. A soluble, red to maroon pigment that diffuses into the agar and is often best observed by viewing the reverse of the colony is suggestive of T. marneffei. Although the growth rate and colonial morphologic features may help the laboratorian recognize the possibility of a dimorphic fungus, they should be considered in combination with the microscopic morphologic features to make the identification. T. marneffei cannot be definitively identified by morphologic features alone; thermal conversion studies or nucleic acid–based testing is needed to confirm the identification of this pathogen. Sporothrix spp. Colonies of Sporothrix spp. grow rapidly (3 to 5 days) and initially are usually small, moist, and white to cream-colored. On further incubation, these become membranous, wrinkled, and coarsely matted, with the color becoming irregularly dark brown or black and the colony becoming leathery in consistency. It is not uncommon for the clinical microbiology laboratory to mistake a young culture of Sporothrix spp. for a yeast until the microscopic features are observed. 

Approach to Identification Blastomyces dermatitidis–B. gilchristii

Microscopically, hyphae of the mold form of Blastomyces spp. are septate and delicate and measure approximately 2 μm in diameter. Commonly, ropelike strands of hyphae are seen; however, these are found with most of the dimorphic fungi. The characteristic microscopic morphologic features are single, circular to pyriform conidia produced on short conidiophores that resemble lollipops (Fig. 59.41); less commonly, the conidiophores may be elongated. The production of conidia in some isolates is minimal or absent, particularly on a medium containing blood enrichment. When incubated at 37°C, colonies of the yeast form develop within 7 days and appear waxy, wrinkled, and cream to tan. Microscopically, large, thick-walled yeast cells (8 to 15 μm in diameter) with buds attached by a broad base are seen (Fig. 59.29). Some strains may produce yeast cells as small as 2 to 5 μm, called microforms. Although these microforms may be present, a thorough search should reveal more typical yeast forms. During conversion, swollen hyphal forms and immature cells with rudimentary buds are also likely to be present. Because the conversion of Blastomyces spp. is easily accomplished, this is feasible in the clinical laboratory; however, this is the most appropriate instance in which

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

mold-to-yeast conversion should be attempted. Blastomyces may also be identified by the presence of a specific band (i.e., A band) in the exoantigen test or by nucleic acid probe testing. H. capsulatum, P. boydii, or T. rubrum may occasionally be confused microscopically with Blastomyces spp. The site of infection and the relatively slow growth rate of Blastomyces spp. and careful examination of the microscopic morphologic features usually differentiate it from these fungi. Identification can also be confirmed using the AccuProbe test (Hologic Inc., San Diego CA) for Blastomyces.  Coccidioides spp.

Microscopically, some Coccidioides cultures show small, septate hyphae that often exhibit right-angle branches and racquet forms. With age, the hyphae form arthroconidia that are characteristically rectangular to barrelshaped. The arthroconidia are larger than the hyphae from which they were produced and stain darkly with lactophenol cotton or aniline blue. The arthroconidia are separated by clear or lighter staining, nonviable cells (dysjunctor cells). These types of conidia are referred to as alternate arthroconidia (Fig. 59.25 and Fig. 59.35). Arthroconidia have been reported to range from 1.5 to 7.5 μm in width and 1.5 to 30 μm in length, whereas most are 3 to 4.5 μm in width and 3 μm in length. Variation has been reported in the shape of arthroconidia, ranging from rounded to square or rectangular to curved; however, most are barrel-shaped. Even if alternate arthroconidia are observed microscopically, definitive identification should be made using nucleic acid probe testing. If a culture is suspected of being Coccidioides, it should be sealed with tape to prevent laboratory-acquired infection. Because Coccidioides spp. are considered the most infectious of all the fungi, extreme caution should be used in handling cultures of these organisms. Safety precautions include the following: 1. If culture dishes are used, they should be handled only in a Level 3 BSC. Cultures should be sealed with tape if the specimen is suspected to contain Coccidioides spp.

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2. The use of cotton plug test tubes is discouraged, and screw-capped tubes should be used if culture tubes are preferred. All handling of cultures of Coccidioides spp. in screw-capped tubes should be performed inside a BSC. 3. All microscopic preparations for examination should be performed in a Level 3 BSC. 4. Cultures should be autoclaved as soon as final identification is made. Both species of Coccidioides were previously classified as select agents in the United States before removal from the list in 2012. Other, usually nonvirulent fungi that resemble Coccidioides microscopically may be found in the environment. Some molds, such as Malbranchea spp., also produce alternate arthroconidia, although these tend to be more rectangular; however, such species must be considered when making the identification. G. candidum and Trichosporon spp. produce hyphae that disassociate into contiguous arthroconidia; these should not be confused with Coccidioides (Fig. 59.42 and Fig. 59.24). The colonial morphologic features of older cultures of these fungi may resemble Coccidioides spp., but as noted the arthroconidia are not alternate. It is also important to remember that if confusion in identification does arise, or when occasional strains of Coccidioides that fail to sporulate are encountered, identification by exoantigen or nucleic acid testing may be performed. Identification can also be confirmed using the AccuProbe test (Hologic Inc., San Diego CA) for Coccidioides spp.  Emmonsia spp. and Emergomyces spp.

The typical mold phase for Emmonsia and Emergomyces spp. was previously described in this chapter; however, culture for conversion to the yeast phase should be grown on phytone yeast extract agar, BHI, or BHI with blood at 37°C to 40°C. Es. pasteuriana produces yeastlike cells on BHI after approximately 10 days of incubation at 37°C. Other species may require up to 14 days of incubation for the yeast phase to be identified. Some of these organisms may be morphologically indistinguishable. Multilocus gene sequencing may be necessary to fully identify clinical isolates included in these genera. 

B

A

• Fig. 59.41  The mycelial form of Blastomyces dermatitidis shows oval conidia borne laterally on branching hyphae (×1000).

• Fig. 59.42  Trichosporon spp. produce arthroconidia (A) and an occasional blastoconidium (B).

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Histoplasma capsulatum

Microscopically, the hyphae of H. capsulatum are small (approximately 2 μm in diameter) and are often intertwined to form ropelike strands. Commonly, large (8 to 14 μm in diameter) spherical or pyriform, smooth-walled macroconidia are seen in young cultures. With age, the macroconidia become roughened or tuberculate and provide enough evidence to make a tentative identification (Fig. 59.43). The macroconidia are produced either on short or long conidiospores. Some isolates produce round to pyriform, smooth microconidia (2 to 4 μm in diameter), in addition to the characteristic tuberculate macroconidia. Some isolates of H. capsulatum fail to sporulate despite numerous attempts to induce sporulation. Conversion of the mold to the yeast form is usually difficult and is not recommended. Microscopically a mixture of swollen hyphae and small budding yeast cells 2 to 5 μm in diameter should be observed. These are similar to the intracellular yeast cells seen in mononuclear cells in infected tissue. The yeast form of H. capsulatum cannot be recognized unless the corresponding mold form is present on another culture or unless the yeast is converted directly to the mold form by incubation at 25°C to 30°C after yeast cells have been observed. Nucleic acid testing is recommended as a definitive means of rapidly identifying this organism. 

is unsuccessful, the exoantigen test (Evolve Procedure 59.3) should be used to make the definitive identification of P. brasiliensis. There is no commercial DNA probe test available for the identification of P. brasiliensis.  Talaromyces marneffei

At 25°C, T. marneffei grows rapidly and produces bluegreen to yellowish colonies. A soluble red to maroon pigment, which diffuses into the agar, is highly suggestive of T. marneffei. At 37°C, conversion of mycelium to the infective, yeastlike form occurs in approximately 2 weeks. Oval, yeastlike cells (2 to 6 μm in diameter) with septa are seen; abortive, extensively branched, and highly septate hyphae may also be present (Fig. 59.38). A variety of laboratory developed nucleic acid tests have been used to identify this organism from clinical samples.  Sporothrix spp.

Microscopically, the mold form is similar to that seen with B. dermatitidis. Small hyphae (approximately 2 μm in diameter) are seen, along with numerous chlamydoconidia. Small (3 to 4 μm), delicate, globose or pyriform conidia may be seen arising from the sides of the hyphae or on very short conidiophores (Fig. 59.44). Most often cultures reveal only fine septate hyphae and numerous chlamydoconidia. After temperature-based conversion on a blood-enriched medium, the colonial morphology of the yeast form is characterized by smooth, soft-wrinkled, yeastlike colonies that are cream to tan. Microscopically, the colonies are composed of yeast cells 10 to 40 μm in diameter surrounded by narrow-necked yeast cells around the periphery, as previously described (Fig. 59.37). If in vitro conversion to the yeast form

Microscopically, hyphae are delicate (approximately 2 μm in diameter), septate, and branching. Single-celled conidia 2 to 5 μm in diameter are borne in clusters from the tips of single conidiophores (flowerette arrangement). Each conidium is attached to the conidiophore by an individual, delicate, threadlike structure (denticle) that may require examination under oil immersion to be visible. As the culture ages, single-celled, thick-walled, black-pigmented conidia may also be produced along the sides of the hyphae, simulating the arrangement of microconidia produced by T. rubrum (sleeve arrangement) (Fig. 59.45). Because of similar morphologic features, saprophytic species of the genus Sporotrichum may be confused with Sporothrix spp., and they must be differentiated. During incubation of a culture at 37°C, colonies of Sporothrix spp. transform to a soft, cream-colored to white, yeastlike appearance. Microscopically, singly or multiply budding, spherical, oval, or elongate, cigar-shaped yeast cells are observed without difficulty (Fig. 59.46). Conversion from the mold form to the yeast form is easily accomplished and usually occurs within 1 to 5 days after transfer of the culture to a medium containing blood enrichment; most isolates of Sporothrix spp. are converted to the yeast form within 12 to 48 hours at 37°C.

•

• Fig. 59.44  Mycelial form of Paracoccidioides brasiliensis shows sep-

Paracoccidioides spp.

Fig. 59.43  The mycelial form of Histoplasma capsulatum produces characteristic tuberculate macroconidia (×1000).

tate hyphae and pyriform conidia singly borne (arrow) (×430).

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

• Fig. 59.45  The mycelial form of Sporothrix spp. demonstrating pyriform to ovoid microconidia in a flowerette morphology at the tip of the conidiophore (arrow) (×750).

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Two assays, complement fixation and immunodiffusion, have been used together to detect antibodies directed toward H. capsulatum and Coccidioides spp. In the complement fixation assay, titers of 1:32 or greater indicate active infection with H. capsulatum. Titers as low as 1:2 to 1:4 have been identified in patients with coccidioidomycosis. Titers greater than 1:16 usually indicate active disease. Bands of identity form in the immunodiffusion test between known antisera, known fungal antigen, and the antibodies present in the patient’s serum. Specific bands of identity are used for serologic detection of particular fungi, whereas nonspecific bands suggest the possibility of an infection by another fungal pathogen. One or two bands of identity, the H protein and M protein bands may occur in patients with histoplasmosis. The presence of both bands indicates active infection. The presence of an M protein band may indicate early or chronic infection. Newer enzyme immunosorbent assays have been developed and demonstrate improved sensitivity and specificity for the identification of histoplasmosis and coccidioidomycosis. A latex agglutination assay has been developed for the presumptive identification of Coccidioides infections. However, the test demonstrates a high false positive rate and is not recommended for use with CSF specimens.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography • Fig. 59.46  The yeast form of Sporothrix spp. consists of cigar-shaped and oval budding cells (×500).

Sporotrichum spp. do not produce a yeast form. Molecular sequencing of the 18S rRNA or 28S rRNA along with the calmodulin gene may be required for species identification. 

Serologic Testing Fungal serology includes rapid and useful tests that may aid the diagnosis of systemic fungal infections caused by H. capsulatum, Paracoccidioides, and Coccidioides species. These tests have also been useful to study the epidemiology of these fungal infections, because even individuals with historically distant, asymptomatic, or subclinical infections often have developed an antibody response to the infecting pathogen. Unfortunately, these tests require detailed preparation and technical expertise. False-negative reactions may occur if serology specimens are drawn in immunocompromised individuals who are unable to produce an antibody response. False-positive reactions may occur because of cross reactivity with other fungi. For example, because the antigens of H. capsulatum are similar to those of B. dermatitidis, occasionally a specimen from a patient with histoplasmosis demonstrates a positive reaction for B. dermatitidis in serologic tests. Serology does not appear to be useful for the diagnosis of blastomycosis, and no tests have been developed for the diagnosis of adiaspiromycosis or emergomycosis.

Bailek R, Kern J, Herrmann T, et al.: PCR assays for identification of Coccidioides posadasii based on the nucleotide sequence of the antigen 2/proline-rich antigen, J Clin Microbiol 42:778–783, 2004. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia PA, 2020, Elsevier. Brown EM, McTaggert LR, Zhang SX, et  al.: Phylogenetic analysis reveals a cryptic species Blastomyces gilchristii, sp. nov. within the human pathogenic fungus Blastomyces dermatitidis, PloS One 8:e59237, 2013. Carey J, D’Amico R, Sutton DA, et  al.: Paecilomyces lilacinus vaginitis in an immunocompetent patient, Emerg Infect Dis 9: 1155–1158, 2003. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Croxatto A, Prod’hom G, Greub G: Applications of MALDI-TOF mass spectrometry in clinical diagnostic microbiology, FEMS Microbiol Rev 36:380–407, 2012. de Hoog G: In Atlas of clinical fungi, t.N.a.R. centraal bureau voor schimell cultures, Spain, 2001, Universita Rovira i Virgili, Utrecht and Reus. De Carolis E, Posteraro B, Lass-Flori C, et al.: Species identification of Aspergillus, Fusarium and Mucorales with direct surface analysis by matrix-assisted laser desorption ionization time-of-flight mass spectrometry, Clin Microbiol Infect 18:474–484, 2012. Fleming RV, Walsh TJ, Anaissie EJ: Emerging and less common fungal pathogens, Infect Dis Clin North Am 16:915–933, 2002. Germain G S, Summerbell R: Identifying filamentous fungi: a clinical handbook, Belmont, Calif, 1996, Star Publishing. Gomez-Munoz MT, Fernandez-Barredo S, Martinez-Diaz RA, et al.: Development of a specific polymerase chain reaction assay for the detection of Basidiobolus, Mycologia 104:585–591, 2012.

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Gutierrez-Rodero F, Moragon M, Ortiz de la Tabla V, et al.: Cutaneous hyalohyphomycosis caused by Paecilomyces lilacinus in an immunocompetent host successfully treated with itraconazole: case report and review, Eur J Clin Microbiol Infect Dis 18:814–818, 1999. Heinic GS, Greenspan D, MacPhail LA, et al.: Oral Geotrichum candidum infection associated with HIV infection: a case report, Oral Surg Oral Med Oral Pathol 73:726–728, 1992. Heys I, Taljaard J, Orth H: An Emmonsia species causing disseminated infection in South Africa, N Engl J Med 370:283–284, 2014. Hussein MR: Mucocutaneous Splendore-Hoeppli phenomenon, J Cutan Pathol 35:979–988, 2008. Kane J, Smitka C: Early detection and identification of Trichophyton verrucosumm, J Clin Microbiol 8:740–747, 1978. Kenyon C, Bonorchis K, Corcoran C, et  al.: A dimorphic fungus causing disseminated infection in South Africa, N Engl J Med 369:1416–1424, 2013. Khan ZU, Khourshee M, Makar R, et al.: Basidobolus ranarum as an etiologic agent of gastrointestinal Zygomycosis, J Clin Microbiol 39:2360–2363, 2001. Kontoyiannis DP, Wessel VC, Bodey GP, et  al.: Zygomycosis in the 1990s in a tertiary-care cancer center, Clin Infect Dis 30: 851–856, 2000. Kwon-Chung K, Bennet J. Medical mycology, Philadelphia, PA, 1992, Lea & Febiger. Martagon-Villamil J, Shrestha N, Sholtis M, et al.: Identification of Histoplasma capsulatum from culture extracts by real-time PCR, J Clin Microbiol 41:1295–1298, 2003. Meis JF, Kullberg BJ, Pruszczynski M, et  al.: Severe osteomyelitis due to the zygomycete Apophysomyces elegans, J Clin Microbiol 32:3078–3081, 1994. Modley A, Mosam A, Govender NP, et  al.: Emergomyces africanus: the mimicking fungus, Dermatopathology (Basel) 6(2): 157–162, 2019. Nucci M: Emerging moulds: Fusarium, Scedosporium, and Zygomycetes in transplant recipients, Curr Opin Infect Dis 16:607–612, 2003. Odabasi Z, Mattiuzzi G, Estey E, et  al.: Beta-D-glucan as a diagnostic adjunct for invasive fungal infections: validation, cutoff development, and performance in patients with acute myelogenous leukemia and myelodysplastic syndrome, Clin Infect Dis 39: 199–205, 2004. Ostrosky-Zeichner L, Alexander BD, Kett DH, et  al.: Multicenter clinical evaluation of the (1-3) beta-D-glucan assay as an aid to diagnosis of fungal infections in humans, Clin Infect Dis 41:654– 659, 2005.

Patel R, Gustaferro CA, Krom RA, et al.: Phaeohyphomycosis due to Scopulariopsis brumptii in a liver transplant recipient, Clin Infect Dis 19:198–200, 1994. Persaud SP, Lawton T, Burnham CD, et  al.: Comparison of urine antigen assays for the diagnosis of Histoplasma capsulatum infection, J Appl Lab Med 4(3):370–382, 2019. Pfeiffer CD, Fine JP, Safdar N: Diagnosis of invasive aspergillosis using a galactomannan assay: a meta-analysis, Clin Infect Dis 42:1417–1427, 2006. Procop GW, Cockerill III FR, Vetter EA, et al.: Performance of five agar media for recovery of fungi from isolator blood cultures, J Clin Microbiol 38:3827–3829, 2000. Schrodl W, Heydel T, Schwartze VU, et al.: Direct analysis and identification of pathogenic Lichtheimia species by matrix-assisted laser desorption ionization-time of flight analyzer mediated mass spectrometry, J Clin Microbiol 50:419–427, 2012. Schwartz IS, Govender NP, Sigler L, et al.: Emergomyces: the global rise of new dimorphic fungal pathogens, PLoS Pathog 15(9):e1007977, 2019. Skoulidis F, Morgan MS, MacLeod KM: Penicillium marneffei: a pathogen on our doorstep? J R Soc Med 97:394–396, 2004. Sun SH, Huppert M, Vukovich KR: Rapid in vitro conversion and identification of Coccidioides immitis, J Clin Microbiol 3:186–190, 1976. Torres HA, Raad II , Kontoyiannis DP: Infections caused by Fusarium species, J Chemother 15(Suppl 2):28–35, 2003. Van Burik JA, Myerson D, Schreckhise RW, Bowden RA: Panfungal PCR assay for detection of fungal infection in human blood specimens, J Clin Microbiol 36:1169–1175, 1998. Walsh TJ, Groll A, Heimenz J, et al.: Infections due to emerging and uncommon medically important fungal pathogens, Clin Microbiol Infect 10(Suppl 1):48–66, 2004. Wang SM, Shieh CC, Liu CC: Successful treatment of Paecilomyces variotii splenic abscesses: a rare complication in a previously unrecognized chronic granulomatous disease child, Diagn Microbiol Infect Dis 53:149–152, 2005. Willinger B: Laboratory diagnosis and therapy of invasive fungal infections, Curr Drug Targets 7:513–522, 2006. Woo PC, Leung SY, Ngan A, et al.: A significant number of reported Absidia corymbifera (Lichtheimia corymbifera) infections are caused by Lichtheimia ramosa (syn. Lichtheimia hongkongensis): an emerging cause of mucormycosis, Emerg Microb Infect 1:e15, 2012. Zhiyong Z, Mei K, Yanbin L: Disseminated Penicillium marneffei infection with fungemia and endobronchial disease in an AIDS patient in China, Med Princ Pract 15:235–237, 2006.

PROCEDURE 59.1

PROCEDURE 59.2

Hair Perforation Test

In Vitro Conversion of Dimorphic Molds

Method

Principle

1. Place a filter paper disk in the bottom of a sterile culture dish. 2. Cover the surface of the paper disk with sterile distilled water. 3. Add a small portion of sterilized hair(s) to the water. 4. Inoculate a portion of the colony directly onto the hair. 5. Incubate at 25°C for 10–14 days. 6. Observe the hairs regularly by placing them in a drop of water on a microscope slide. Position a coverslip and examine microscopically for conical perforations of the hair shaft (Fig. 59.11).

Dimorphic molds exist in the yeast or spherule form in infected tissue. Proof that a mold is one of the systemic dimorphic fungi can be obtained by simulating the host environment and converting the mold to the yeast form. This method is recommended for identifying Blastomyces spp., Talaromyces marneffei, and Sporothrix spp. Conversion of Coccidioides spp. to its spherule form requires special media or animal inoculation and is not recommended for clinical laboratories. Histoplasma capsulatum most often does not convert from the mold to the yeast form in vitro or does so only after extended incubation at 35°C–37°C. 

  

Method 1. Transfer a large inoculum of the mold form of the culture onto the surface of a fresh, moist slant of brainheart infusion agar containing 5%–10% sheep blood. If Blastomyces spp. are suspected, a tube of cottonseed conversion medium should be inoculated. 2. Add a few drops of sterile distilled water to provide moisture if the surface of the culture medium appears dry. 3. Leave the cap of the screw-capped tube slightly loose to allow the culture to have adequate oxygen exchange. 4. Incubate cultures at 35°C–37°C for several days; observe for the appearance of yeastlike portions of the colony. Several subcultures of any growth that appears may need to be made, because several transfers are often required to accomplish the conversion of many isolates. Cultures of Blastomyces spp., however, are usually easily converted and require 24–48 hours on cottonseed agar medium. Coccidioides spp. may be converted in vitro to the spherule form using a Converse liquid medium; however, this method is of little use to the clinical laboratory and should not be attempted. Genetic probe hybridization, DNA sequencing, species-specific polymerase chain reaction (PCR), and exoantigen detection are recommended methods of definitively identifying isolates suspected to be Coccidioides spp. Ea. crescens produces adiaspores when cultivated on Phytone Yeast Extract Agar, BHI, or BHIB at 37°C–40°C. Es. pasteurianus must be cultivated for approximately 10 days at 37°C on BHI before the production of yeastlike cells is apparent. 

Quality Control Because of the hazardous nature of stock cultures, it is not recommended that they be tested routinely. An extract of control strains can be used as a positive control. Because conversion of the dimorphic molds to the corresponding yeast or spherule forms is technically cumbersome and often involves long delays, attempts to convert the dimorphic fungi are not recommended in the routine mycology laboratory.   

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PROCEDURE 59.3

CASE STUDY 59.1

Exoantigen Test

An elderly female with diabetes visits her clinician because she has a brown, dull, discolored toenail. The clinician takes nail clippings for culture. The laboratory places the clippings on media with no cycloheximide. SAB agar reveals a rapidgrowing colony that is mature in 5 days. The colonies are initially white, become tan with age, and are velvety and powdery. The reverse is tan with a brown center. Microscopic examination reveals septate hyaline hyphae with annelides. Annelides are both solitary and in clusters that resemble a penicillus.

Principle Specific antibodies developed against particular mycelial antigens react in a gel immunodiffusion precipitin test. The mold forms of the dimorphic fungi can be identified definitively by an antigen-antibody reaction, eliminating the need for conversion to the yeast form. 

Method 1. Cover a mature fungus culture on a Sabouraud dextrose agar slant with an aqueous solution of merthiolate (1:5000 final concentration), which is allowed to remain in contact with the culture for 24 h at 25°C. The entire surface of the colony must be covered so that effective killing of the organism is ensured and solubilization of the exoantigen is maximized. 2. Filter the aqueous solution that overlays the culture through a 0.45-μm membrane filter. This should be done inside a biologic safety cabinet (BSC). 3. Concentrate 5 mL of this solution using a Minicon Macrosolute B-15 Concentrator (Millipore, Billerica, MA). The solution is concentrated 50× when testing with Histoplasma capsulatum and Blastomyces antiserum and 5× and 25× for reaction with Coccidioides antiserum. 4. Use the concentrated supernatant in the microdiffusion test. Place the supernatant in wells punched into a plate of buffered, phenolized agar adjacent to the control antigen well and test it against positive control antiserum obtained from commercial sources. 5. Allow the immunodiffusion test to react for 24 hours at 25°C. Then, observe the plate for precipitin bands of identity with the reference reagents. The sensitivity of the exoantigen test may be improved in the identification of Blastomyces by incubating the immunodiffusion plates at 37°C for 48 h; however, bands appear sharper at 25°C after 24 h. Any culture suspected of being Blastomyces should be incubated at both temperatures. 6. Coccidioides may be identified by the presence of the CF, TP, or HL antigens; H. capsulatum may be identified by the presence of H protein or M protein bands (or both); and Blastomyces may be identified by the A protein band. Detailed instructions for performing and interpreting the tests are included with the manufacturers’ package inserts. 

Quality Control Extracts from known fungi are tested each time the test is performed. Lines of identity with the unknown strain are necessary for identification.   

Data from Murray CK, Beckius ML, Green JA, et al. Use of chromogenic medium in the isolation of yeasts from clinical specimens. J Med Microbiol. 2005;54:981; and Tan GL, Peterson EM. Chromagar Candida medium for direct susceptibility testing of yeast from blood cultures. J Clin Microbiol. 2005;43:1727.

  

Questions 1. Why did the laboratory use agar without cycloheximide? 2. How would you distinguish between Talaromyces and Scopulariopsis spp.? 3. One-celled conidia are rough-walled, and spiny conidia in chains are seen. What organism is identified?   

CHAPTER 59  Hyaline Molds, Mucorales, Basidiobolales, Entomophthorales, Dermatophytes, and Opportunistic and Systemic Mycoses

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Chapter Review 1. Which test can be used to differentiate T. mentagrophytes from T. rubrum? a. Fluorescence using a Woods lamp b. In vitro hair perforation c. Red color on reverse side of colony d. Pyriform microconidia 2. Tinea capitis is caused by which dermatophyte? a.  T. tonsurans b. M. audouinii c.  M. canis d. All of the above 3.  Microsporum infection involves: a. Hair, skin, and nails b. Hair and skin c. Skin and nails d. None of the above 4. A Woods lamp is used to detect fluorescing hairs in which dermatophyte? a.  M. audouinii b. M. canis c. M. gypseum d. A and B 5. Sickle- or boat-shaped macroconidia on cornmeal agar are characteristic of: a.  Alternaria spp. b. Aspergillus spp. c.  Penicillium spp. d. Fusarium spp. 6. Which of the following produces rhizoids? a.  Mucor spp. b. Rhizopus spp. c.  Trichophyton spp. d. Microsporum spp. 7. Which dimorphic fungus is found in the Missouri River valley? a.  B. dermatitidis b. H. capsulatum c.  Coccidioides spp. d. P. brasiliensis

8. Which dimorphic fungus may be contracted by people who clean chicken coops? a.  B. dermatitidis b. H. capsulatum c.  Coccidioides spp. d. P. brasiliensis 9. Which organism is capable of producing an unusually large spore when inhaled into the human respiratory tract? a.  B. dermatitidis b. H. capsulatum c.  Coccidioides spp. d. Ea. crescens 10.  Matching: Match each term with the correct description. _____ rhizoid _____ ectothrix _____ endothrix _____ sporangia _____ tinea corporis _____ tinea cruris _____ Sporothrix schenckii _____ Trichosporon mentagrophytes _____ anthropophilic _____ zoophilic

a. inside the hair shaft b. ringworm of the body c. rose-gardener’s disease d. rootlike hypha e. infects animals f. infects humans g. athlete’s foot h. outside the hair shaft i. jock itch j. fruiting structures

60

Dematiaceous (Melanized) Molds OBJECTIVES 1. Describe the melanized fungi, including natural habitat, transmission, and diseases with signs and symptoms. 2. Identify the site where mycetomas are commonly located and the population or populations at risk of infection. 3. Compare and contrast Exophiala jeanselmei and Exophiala dermatitidis, including test methods to distinguish between the two. 4. Describe the microscopic and morphologic features of Pseudallescheria boydii, including its sexual and asexual forms. 5. Differentiate the diagnostic microscopic features of the molds included in this chapter.

Exophiala bergeri Exophiala dermatitidis Exophiala jeanselmei Exophiala oligosperma Exophiala xenobiotica Exserohilum rostratum Knufia epidermidis Lasiodiplodia theobromae Macrophomina phaseolina Neoscytalidium dimidiatum Sporothrix pallida 

Subcutaneous (Includes Mycetomas)

Alternaria alternata Aureobasidium melanogenum Cladophialophora boppii Cyphellophora europaea Cyphellophora laciniata Cyphellophora pluriseptata Curvularia spp. Hortaea werneckii Neoscytalidium dimidiatum Piedraia hortae Scopulariopsis brevicaulis Triadelphia pulvinata 

Alternaria spp. Cyphellophora suttonii Cladophialophora bantiana Curvularia spp. Diaphorthe bougainvilleicola Diaphorthe phaseolorum Exserohilum spp. Exophiala spp. Hongkongmyces pedis Knoxdaviesia dimorphospora Lasiodiplodia theobromae Lomentospora prolificans Madurella spp. Neoscytalidium dimidiatum Ochroconis mirabilis Phaeoacremonium spp. Pleurostoma richardsiae Scedosporium spp. Scopulariopsis spp. Trematosphaeria grisea Veronaea botryose 

Cutaneous and Corneal

Systemic Phaeohyphomycosis

Alternaria alternata Alternaria infectoria Bipolaris oryzae Cladophialophora boppii Cladophialophora emmonsii Cladophialophora saturnica Cladorrhinum bulbillosum Cladosporium cladosporioides Cladosporium oxysporum Curvularia lunata Curvularia senegalensis Curvularia spicifera Diaporthe longicolla Diaporthe phaseolorum Diaporthe phoenicicola

Arthrocladium fulminans Aureobasium pullulans Cladophialophora bantiana Cladophialophora modesta Curvularia spp. Exserohilum rostratum Exophiala dermatitidis Exophiala phaeomuriformis Exophiala spinifera Fonsecaea monophora Fonsecaea pedrosoi Hormonema dematioides Lasiodiplodia theobromae Lomentospora prolificans Neoscytalidium dimidiatum

SEPTATE DEMATIACEOUS MOLDS TO BE CONSIDERED Superficial Infections

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SEPTATE DEMATIACEOUS MOLDS TO BE CONSIDERED Ophiostoma piceae Phaeoacremonium parasiticum Phialophora verrucosa Rhinocladiella mackenziei Scedosporium spp. Triadelphia disseminata Verruconis gallopava 

Chromoblastomycosis Cladophialophora carrionii Cladophialophora boppii Cladophialophora samoënsis Cyphellophora ludoviensis Phialophora verrucosa Fonsecaea monophora Fonsecaea pedrosoi Fonsecaea pugnacius Rhinocladiella tropicalis Rhinocladiella aquaspersa

Epidemiology and Pathogenesis Superficial Infections Tinea nigra is a superficial skin infection caused by Hortaea werneckii, a halophilic species. It is manifested by blackish brown, macular patches on the palm of the hand or the sole of the foot. Lesions have been compared with silver nitrate staining of the skin. Black piedra is a fungal infection of the hair, scalp, and occasionally the axillary and pubic hair caused by the dematiaceous fungus Piedraia hortae. Neoscytalidium dimidiatum, a common plant pathogen, causes infections of the skin and nails that may lead to hyperkeratosis (thickening of the epidermis). These diseases occur primarily in tropical areas of the world, with cases reported from Africa, Asia, and Latin America. Phialophora spp. and Cyphellophora spp. have also been known to be the causative agents of mild skin infections and onychomycoses (M & M, Chowdhary 2015). Several species in the order Chaetothyriales are capable of causing superficial infections in humans including Cyphellophora spp., Phialophora europaea, and Knufia epidermidis. The fungi are typically associated with mild cutaneous skin infections or nail infections.

General Characteristics

Mycetoma

The dematiaceous fungi were once only characterized by the dark coloration because of their ability to produce melanin. More recently, organisms, including medically important fungi, have been classified using molecular techniques (Table 60.1). Many of the agents in this chapter cause phaeohyphomycosis or chromoblastomycosis and are known agents of superficial and subcutaneous mycoses that involve the skin and subcutaneous tissues and, less commonly, deeply invasive or disseminated disease. These organisms are ubiquitous in nature and exist as saprophytes and plant pathogens. Humans and animals serve as accidental hosts after traumatic inoculation of the organism into cutaneous and subcutaneous tissues. In the mycology laboratory, these fungal species often are initially separated by growth rate into the slowgrowing dematiaceous molds, which may require 7 to 10 days to grow, and the rapid-growing dematiaceous molds, which usually grow in less than 7 days. It has been recommended that in medical mycology the term dematiaceous only be applied to rapidly growing members of the Ploesoporales (Alternaria, Bipolaris, Curvularia, Exoserohilum, and Hongkongmyces). This has not been widely implemented and will take time for all of the changes to be apparent in clinical mycology. When nonsterile body sites are cultured, determining the significance of these organisms is very difficult. If colonies of common saprophytic molds occur near the edge of the plate and are clearly away from the inoculum, they should be considered contaminants unless additional evidence of infection is present. 

A mycetoma is a chronic granulomatous infection that usually involves the lower extremities but may occur in any part of the body. The infection is characterized by swelling, purplish discoloration, tumorlike deformities of the subcutaneous tissue, and multiple sinus tracts that drain purulent material containing yellow, white, red, or black granules called grains. The color of the granules is partly dependent on the type of infecting organism. The infection gradually progresses to involve the bone, muscle, or other contiguous tissue and ultimately requires amputation in most progressive cases. Dissemination of the organism may occur but is uncommon. Mycetomas usually are seen among people living in tropical and subtropical regions of the world whose outdoor occupations and failure to wear protective clothing predispose them to trauma. Two types of mycetomas have been described. Actinomycotic (bacterial) mycetomas are caused by the aerobic actinomycetes, including Nocardia, Actinomadura, and Streptomyces spp. (The aerobic Actinomycetes are described in detail in Chapter 18.) Eumycotic (fungal) mycetomas are caused by a heterogeneous group of fungi that have septate hyphae. Eumycotic mycetomas are subcategorized as white grain mycetomas or black grain mycetomas, a distinction determined by the pigmentation of the infecting agent’s hyphae. Some hyaline septate molds can cause mycetomas; however, the disease is covered in this section because many of the etiologic agents are dematiaceous fungi. Etiologic agents of eumycotic mycetoma to be discussed include S­ cedosporium spp. and Acremonium spp., causative agents of white grain mycetomas, and Exophiala jeanselmei, C ­urvularia spp., Cladophialophora bantiana, Trematosphaeria grisea, and Madurella spp., causative agents of black grain mycetomas.

CHAPTER 60  Dematiaceous (Melanized) Molds

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TABLE 60.1    Classification and Taxonomy of the Clinically Relevant Melanized Fungi

Order

Genera

Species

Characteristics

Botryosphaeria

Lasiodiplodia

L. theobroma

Initially spherical, thick-walled hyaline conidia that brown with age and develop a median septum.

Macrophomina

M. phaseolina

Dark sclerotic bodies and melanized mycelium.

Neoscytalidium

N. dimidiatum

Produces arthroconidia in culture.

Calosphaeriales

Pleurostoma

P. ochracea P. repens P. richardsiae

Dark hyphae with pale, tapering phialides that may be single or aggregated in dense brushes; hyaline conidia.

Capnodiales

Cladosporium

C. cladosporioides C. oxysporum C. sphaerospermum

Branching chains of single-celled or septate conidia.

Hortea

H. werneckii

Yeastlike, aseptate or septate elements.

Anthopsis

Anthopsis sp.

Ampulliform phialides that appear inverted with collarettes.

Arthrocladium

A. fulminans

Moniliform hyphae and chlamydospore-like structures.

Cladophialophora

C. bantiana C. carrionii

Long, branched conidial chains and grows at 40°C. Small conidia in branched chains.

Cyphellophora

C. europaea C. laciniata C. ludoviensis C. pluriseptica C. reptans C. suttonii

Slender, curved, transversely septate conidia.

Exophiala

E. bergeri E. dermatitidis E. jeanselmei E. oligosperma E. phaeomuriformis E. spinifera E. xenobiotica

Conidia in chains or compacted phialides. E. dermatitis is recognized by phialides without collarettes with short annellated zones.

Fonsecaea

F. compacta F. monophora F. nubica F. pedrosoi F. pugnacius

Phialophora-like phialides with collarettes and Rhinocladiella-like sympodial conidiophores.

Knufia

K. epidermidis

Produce phialides, arthroconidia, holoblastic conidia, endoconidia, and yeastlike budding cells in culture.

Phialophora

P. verrucosa

Darkened funnel-shaped collarettes.

Rhinocladiella

R. aquaspersa R. atrovirens R. basitona R. mackenziei R. similis R. tropicalis

Sympoidial conidiophores with one-celled conidia on denticles.

Veronaea

V. botryose

Sympodial rachis with denticles, single-septate conidia.

Diaporthe

D. bougainvilleicola D. longicolla D. phaseolorum D. phoenicicola

Oval to fusoid conidia or thin, curved, or bent elongated conidia.

Chaetothyriales

Diaporthales

Continued

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TABLE 60.1    Classification and Taxonomy of the Clinically Relevant Melanized Fungi—cont’d

Order

Genera

Species

Characteristics

Dothideales

Aureobasidium

A. melanogenum A. pullulans

Budding yeastlike, hyaline and melanized thickwalled hyphae that produce dark brown chlamydospores with age.

Hormonema

H. dematioides

Produce conidia in a basipetal succession.

Knoxdaviesia

K. dimorphospora

Lateral undifferentiated hyphae, epllipsoid, pale brown conidia.

Lomentospora

L. prolificans

Elongated, annellidic conidiogenous cells and obovoid conidia.

Microascus

M. brunneosporus M. cinereus M. gracilis

Dark brown to black, globose to ampulliform ascomata, with papilla. Ascospores are ellipsoid or quadrangular. Single or penicillate conidiophores with conidia in basipetal chains.

Scopulariopsis

S. asperula S. brevicaulis S. candida

Globose or pyriform, black ascomata. Annellides arranged penicillately on conidiophore being single or small groups of stalks, smooth or rough walled conidia in basipetal chains.

Scedosporium

S. apiospermum S. aurantiacum S. boydii S. dehoogii

Conidiophores with annelides. Conidia are obovoid and become brown with age. Produces laterally on hyphae or short pedicels.

Triadelphia

T. disseminate T. pulvinata

Ophiostoma

O. piceae

Subglobose, dark perithecia asci with long necks; yeastlike cells and septate hyphae.

Sporothrix

S. brasiliensis S. chilensis S. globose S. luriei S. pallida S. schenckii

Conidiophore bear clusters of thin denticles with hyaline, tear-shaped conidia. Subglobose or elongated conidia may also be present.

Alternaria

A. alternate A. infectoria

Conidia in chains with alternating septa. A. infectoria produces conidia with long apical beaks.

Bipolaris

B. australiensis B. hawaiiensis B. oryzae B. spicifera

Large ellipsoid to subcylindrical, straight conidia with a distosepta and a dark, flat basal scar.

Curvularia

C. aeria C. americana C. geniculate C. hominis C. muehlenbeckiae C. lunata C. senegalensis

Elongated conidia that are distoseptate with an asymmetrical swollen middle cell resulting in a curved appearance.

Exserohilum

E. longirostratum E. mcginnisii E. rostratum

Long, distoseptate conidia with a protruding basal hilum.

Hongkongmyces

H. pedis

Madurella

M. mycetomatis

Cladorrhinum

C. bulbillosum

Microascales

Ophiostomatales

Pleosporales

Sordariales

Aseptate conidia with intercalary conidiogenous cells with lateral phialide openings.

CHAPTER 60  Dematiaceous (Melanized) Molds

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TABLE 60.1    Classification and Taxonomy of the Clinically Relevant Melanized Fungi—cont’d

Order

Genera

Species

Characteristics

Togniniales

Phaeoacremonium

P. parasiticum Phaeoacremonium spp. (10 additional species)

Warted mycelium and slender, tubular, tapering brown phialides.

Venturiales

Ochroconis

O. mirabilis

Rust to brown olivaceous colonies that produce 1–3 septate conidia from small, open denticles on sympodial cells.

Verruconis

V. gallopava

Same as Ochroconis sp.

Madurella mycetomatis is the most common fungal agent associated with mycetoma. However, nucleic acid–based sequencing of several genes has indicated that multiple additional species exist and are associated with mycetoma. These species have likely been misidentified as M. mycetomatis in previous cases and include Madurella pseudomycetomatis, Madurella tropicana, and Madurella fahalii. Most patients with mycetomas live in tropical regions, but infections can occur in temperate zones. The most common etiologic agent of white grain mycetoma in the United States is caused by Scedosporium spp. The organisms associated with mycetoma are saprophytic and commonly found in soil, standing water, and sewage; humans acquire infections through traumatic implantation of the organism into the skin and subcutaneous tissues. 

Chromoblastomycosis Chromoblastomycosis is a chronic fungal infection acquired through traumatic inoculation of an organism, primarily into the skin and subcutaneous tissue. The infection is characterized by the development of a papule at the site of the traumatic insult that slowly enlarges to form warty or tumorlike lesions characterized as resembling cauliflower capable of spreading through the lymphatic system. Secondary infection and ulceration may occur. The lesions usually are confined to the feet and legs but may involve the head, face, neck, and other body surfaces. Histologic examination of the lesion reveals characteristic sclerotic bodies, which are copper-colored, septate cells that appear to be dividing by binary fission and resemble copper pennies. These infections cause hyperplasia of the epidermal layer of the skin, which may be mistaken for squamous cell carcinoma. Fungal brain abscess, known as cerebral chromoblastomycosis, may be caused by the dematiaceous fungi; however, it is more appropriately considered a type of phaeohyphomycosis and is discussed with that disease. Chromoblastomycosis is widely distributed, but most cases occur in tropical and subtropical areas of the world. Occasional cases are reported from temperate zones, including the United States. The infection is seen most often in areas in which agricultural workers do not wear protective clothing and suffer thorn or splinter puncture wounds.

The fungi most often associated with chromoblastomycosis include Cladophialophora carrionii, Fonsecaea monophora and pedrosoi, and Phialophora verrucosa. Additional species of Cladophialophora have also been reported as the cause of chromoblastomycosis. 

Phaeohyphomycosis Phaeohyphomycosis is a general term used to describe any infection caused by a dematiaceous organism; it includes molds; brownish, yeastlike cells; pseudohyphae; and hyphae, except those described previously. These infections may be subcutaneous, localized, or systemic, and they may be caused by a number of dematiaceous fungi. They include phaeohyphomycotic cysts, progressive soft tissue infection, brain abscess, sinusitis, endocarditis, mycotic keratitis, pulmonary infection, and systemic infection. Symptoms often include headache, neurologic manifestations, and seizures. The most common fungal isolates associated with neurologic manifestations include C. bantiana, Rhinocladiella mackenziei, Verruconis gallopava, and Exophiala dermatitidis. Alternaria, Exserohilum, Bipolaris, E. jeanselmei, Exophiala spinifera, and Curvularia spp. are also commonly associated with phaeophyomycosis. 

Pathogenesis and Spectrum of Disease The spectrum of disease caused by the dematiaceous fungi ranges from superficial infections (e.g., skin and hair) to emergent, rapidly progressive, and often fatal disease (e.g., brain abscess). The following list, which is not comprehensive, provides the common etiologic agents of diseases that may be caused by dematiaceous fungi (Table 60.2). • Mycetoma • Bacterial: Nocardia, Actinomadura, and Streptomyces spp. • White grain mycetoma: S. apiospermum complex and Acremonium and Fusarium spp. • Black grain mycetoma: Madurella spp., E. jeanselmei, and Curvularia spp. • Chromoblastomycosis: Cladophialophora, Phialophora, and Fonsecaea spp. • Phaeohyphomycosis: E. jeanselmei; E. dermatitidis; and Curvularia, Bipolaris, Alternaria, and Exserohilum spp.

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Laboratory Diagnosis

• S inusitis: Alternaria, Bipolaris, Exserohilum, and Curvularia spp. • Mycotic keratitis and endophthalmitis: E. dermatitidis, Bipolaris, and Curvularia spp. • Brain abscess: C. bantiana, E. dermatitidis, and Bipolaris spp. 

Specimen Collection, Transport, and Processing See General Considerations for the Laboratory Diagnosis of Fungal Infections in Chapter 58. 

TABLE 60.2    Common Isolated Dematiaceous Fungi

Organism

Disease

Site

Tissue Form

Chromoblastomycosis

Subcutaneous

Sclerotic bodies

Phaeohyphomycosis

Brain, subcutaneous

Septate hyphae

Verruconis gallopava

Phaeohyphomycosis

Brain, subcutaneous, lungs

Septate hyphae

Exophiala dermatitidis

Phaeohyphomycosis

Brain, eye, subcutaneous, and dissemination

Hyphal fragments and budding yeast

Pneumonial

Lungs

Hortaea jeanselmei

Mycetoma phaeomycotic cyst

Subcutaneous

Hyphal fragments and budding yeasts

Hortaea werneckii

Tinea nigra

Skin

Hyphal fragments and budding yeast

Fonsecaea spp.

Chromoblastomycosis

Subcutaneous

Sclerotic bodies

Phaeohyphomycosis

Brain

Septate hyphae

Cavitary lung disease

Lungs

Septate hyphae

Chromoblastomycosis

Subcutaneous

Sclerotic bodies

Phaeohyphomycosis

Subcutaneous

Septate hyphae

Septic arthritis

Joints

Septate hyphae

Piedraia hortae

Black piedra

Hair

Asci-containing nodules cemented to hair shafts

Madurella spp.

Mycetoma

Subcutaneous

Hyphal fragments

Phaeohyphomycosis

Subcutaneous

Septate hyphae

Sinusitis

Sinuses

Septate hyphae, possibly fungus ball

Nasal septal erosion

Nasal septum

Septate hyphae

Ulcers and onychomycosis

Skin, nails

Septate hyphae

Phaeohyphomycosis

Subcutaneous, brain, eye, bones

Septate hyphae

Sinusitis, fungus ball

Sinuses

Septate hyphae; possibly fungus ball

Sinusitis

Sinuses

Septate hyphae; possibly fungus ball

Phaeohyphomycosis

Subcutaneous, heart valves, eye, and lungs

Septate hyphae

Exserohilum spp.

Phaeohyphomycosis

Subcutaneous

Septate hyphae

Scedosporium spp. (Scedosporium apiospermum complex)

Mycetoma

Subcutaneous

Granules of hyaline hyphae

Phaeohyphomycosis

Subcutaneous, skin, joints, bones, brain, lungs

Septate, hyaline hyphae

Slow-Growing Species Cladophialophora spp.

Phialophora spp.

Rapid-Growing Species Alternaria spp.

Bipolaris spp.

Curvularia spp.

CHAPTER 60  Dematiaceous (Melanized) Molds

Direct Detection Method Stains In general, dematiaceous fungal hyphae are seen in clinical specimens by direct microscopic examination or by histopathologic examination of tissue obtained during surgery or autopsy. The dematiaceous character of the hyphae may not be appreciated if the examination is performed using calcofluor white or fluorescent microscopy alone, without observing the hyphae using traditional transmitted light microscopy. The Fontana-Masson stain, 10% silver nitrate and ammonium hydroxide, stains fungal elements brown to black in a red background. This technique improves the detection of melanin granules. The Fontana-Masson stain is useful to detect melanization that may appear as hyaline molds using light microscopy. Superficial Infections

Direct microscopic examination of a clinical specimen from a patient with tinea nigra may show dematiaceous hyphae and small budding yeast cells and/or hyphal fragments. Portions of hairs from a patient with black piedra are examined in wet mounts using potassium hydroxide (KOH) that is gently heated for nodules composed of cemented mycelium. Crushing the mature nodules reveals oval asci, containing two to eight aseptate ascospores, 19 to 55 μm long by 4 to 8 μm in diameter. The asci are spindle shaped and have a filament at each pole. Sinusitis associated with fungal infections typically reveal dense masses of pigmented, branched and septate hyphae. The hyphal elements appear as amorphous fungal balls that block the sinus cavities but are not invading the mucosal lining.  Chromoblastomycosis

The laboratory diagnosis of chromoblastomycosis is made easily. Scrapings from crusted lesions added to 10% KOH show muriform cells (aggregation of dark brown cells that resemble stones in a stonewall) or sclerotic bodies, which are rounded, brown, 4 to 10 μm in diameter, and have fission planes. They resemble copper pennies (Fig. 60.1). 

851

Mycetoma and Phaeohyphomycosis

Direct examination of clinical specimens from patients with a eumycotic mycetoma or phaeohyphomycosis demonstrates yellowish brown, septate to moniliform hyphae (string of beads), with or without budding yeast cells present. The presence of dematiaceous yeasts depends on the fungus. Dematiaceous yeasts are commonly seen in the direct examination of clinical specimens from patients with infections caused by Exophiala spp. Macroscopic examination of granules from mycetoma lesions caused by S. apiospermum complex reveal them to be white to yellow and 0.2 to 2 mm in diameter. Microscopically, the granules of S. apiospermum complex consist of loosely arranged, intertwined septate hyaline hyphae cemented together. Observation of pigmented hyphae in hematoxylin-eosin or unstained histopathologic sections is presumptive for a diagnosis of dematiaceous fungal disease. The methenamine silver stain used to detect fungal elements in tissues stains fungi black, which makes determining whether they are hyaline septate or dematiaceous septate molds impossible. Fontana-Masson stain, which stains the melanin and melanin-like pigments in the cell walls of these organisms, may be used to confirm the presence of pigmented hyphae in histologic sections. Culture of the specific etiologic agent is necessary for final confirmation. 

Serologic Testing Some serologic and skin tests may be useful for the diagnosis of allergy to dematiaceous fungi. However, serology is not useful for the diagnosis of invasive dematiaceous fungal disease. 

Nucleic Acid–Based Tests Nucleic acid amplification assays can be used for detection or identification of these fungi. Polymerase chain reaction (PCR) tests have been developed for S. apiospermum complex, and direct DNA testing is available for Madurella spp. from the actual grains. Amplification tests have been developed to detect fungal DNA in normally sterile body fluids such as cerebrospinal fluid and brain tissue in patients with suspected fungal meningitis. Nucleic acid–based sequencing of ribosomal genes may be used for the identification of fungal isolates. However, sequences should be evaluated carefully by comparison with type strains. In addition, a variety of high-resolution multigene typing systems have been used in recent years primarily for epidemiologic purposes. 

Matrix-Assisted Laser Desorption Ionization Timeof-Flight Mass Spectrometry

• Fig. 60.1  Sclerotic bodies from the tissue of a patient with chromoblas-

tomycosis (×400). (From Velasques LF, Restrepo A. Chromomycosis in the toad (Bufo marinus) and a comparison of the etiologic agent with fungi causing human chromomycosis. Sabouraudia 1975;13:1.)

Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) has been successfully used to identify clinically relevant fungal isolates including yeasts and molds. As more application of this technique becomes available in the clinical laboratory, rapid diagnosis of fungal infections will undoubtedly become

852 PA RT V    Mycology

more accurate, resulting in improved patient prognosis and treatment. 

Cultivation Although dematiaceous molds recovered in the clinical mycology laboratory may represent true pathogens, more often they represent transient microbiota, inhaled spores, or contaminants. Cultures from sterile body sites, if aseptically obtained, should not contain these molds. Cultures should be interpreted in conjunction with the results of the direct examination for fungal elements, corresponding histopathology, and discussion with the clinician to most effectively establish the diagnosis of mycotic infection caused by these organisms.

• Fig. 60.2  Yeast forms of Hortaea werneckii.

Superficial Infections

H. werneckii, the causative agent of tinea nigra, may be recovered on common fungal media but grows very slowly. Initial colonies of H. werneckii may be olive to black, shiny, and yeastlike (Fig. 60.2) and usually grow within 2 to 3 weeks. As the cultures age, colonies become filamentous, with velvety gray aerial hyphae. P. hortae, the causative agent of black piedra, is easily cultured on any fungal culture medium lacking cycloheximide. Colonies of this organism are also very slow growing, appear dark brown to black, and produce aerial mycelium. Some isolates may produce a red to brown diffusible pigment. Cyphellophora produce slender, curved, one- to threeseptate conidia. The conidia are produced on collarettes. Cultures are typically melanized without budding cells. Exophilia, which is considered a black yeast, demonstrates a high degree of morphologic variability. Colonies initially appear moist and then become wooly or velvety. The conidia are produced from narrow scars or extensions referred to as annelidic. The Exophilia are also capable of growth at 40°C and fail to assimilate nitrate. Phialophora produce phialides (flask shaped) and have no budding cells. N. dimidiatum produces a rapidly growing black arthroconidia in culture.  Mycetoma White Grain Mycetoma. Scedosporium spp. grow rapidly (5 to 10 days) on common laboratory media. Initial growth begins as a white, fluffy colony that changes in several weeks to a brownish gray (the so-called mousy gray) colony; the reverse of the colony progresses from tan to dark brown. Acremonium spp. that cause mycetomas, such as Acremonium falciforme, grow slowly and produce gray colonies.  Black Grain Mycetoma. Colonies of Madurella spp. and E. jeanselmei (Fig. 60.3) are slow growing, unlike colonies of Curvularia spp. Colonies of Madurella spp. vary from white (during the early phases of growth) to olive-brown; a brown diffusible pigment is characteristic of this fungus. Colonies of E. jeanselmei appear yeastlike and darkly pigmented (olive to black) but in time develop a velvety appearance with the production of aerial hyphae. Curvularia spp. produce a fluffy or downy, olive-gray to black colony, and growth is rapid. T. grisea forms slow-growing, velvety colonies that appear

•

Fig. 60.3  Exophiala jeanselmei on chocolate agar. (Photo courtesy Brooks Kyle Murillo-Kennedy, Houston, TX.)

smooth or radially furrowed and dark gray or olive-brown to black. The reverse side of the colonies appears black. The hyphae are septate and nonsporulating.  Chromoblastomycosis

The fungi known to cause chromoblastomycosis, Cladophialophora, Phialophora spp., and Fonsecaea spp., are all dematiaceous. These fungi are slow growing and produce heaped-up, slightly folded, darkly pigmented colonies with a gray to olive to black and velvety or suedelike appearance. The reverse side of the colonies is jet black. Microscopic examination is necessary to identify the pathogenic agent definitively.  Phaeohyphomycosis

The colonies of many of the rapidly growing dematiaceous molds are similar; identification relies on microscopic examination. The colonies of Alternaria spp. are rapidly growing, fluffy, and gray to gray-brown or gray-green. Curvularia spp. produce rapidly growing colonies that resemble those of Alternaria spp. Bipolaris spp. produce colonies that are gray-green to dark brown and slightly powdery, as do Exserohilum spp. The colonies of many of the slow-growing dematiaceous molds are also similar to one another and require identification based on microscopic morphology. E. jeanselmei and E. dermatitidis grow slowly (7 to 21 days) and

CHAPTER 60  Dematiaceous (Melanized) Molds

initially produce shiny, black, yeastlike colonies. E. dermatitidis often is mucoid and may be brown, compared with E. jeanselmei, but the two organisms are very similar in appearance. Colonies become filamentous and velvety with age because of the production of mycelium. E. spinifera produces large, stiff conidiophores. C. bantiana produces long, poorly branched conidial chains. The fungus is also capable of growth at 40°C. R. mackenziei produces pale brown conidiophores with elongated conidia on denticles (projection or peg) and may produce exophiala-like budding cells in culture. V. gallopava produces a rusty-brown to olive colony with one- to three-septate condia on small denticles. The colonial morphology of other slowly growing dematiaceous fungi (e.g., Fonsecaea spp.) was described in the previous section. 

Approach to Identification Superficial Infections H. werneckii is a dematiaceous fungus that produces yeastlike cells that may be one or two celled. Conidia produced by this organism are produced by annellophores (conidiaforming cells that produce conidia-containing transverse rings), which bear successive rings (annelides) that are difficult to see microscopically. The biophysical profile is used to differentiate this fungus from other Exophiala spp. In contrast, P. hortae usually does not sporulate on routine mycologic media but demonstrates highly septate, dematiaceous hyphae and swollen intercalary cells. 

Mycetoma The specific etiologic agent of a eumycotic mycetoma cannot be determined without culturing the organism. Culture media containing antibiotics should not be used as the sole medium for culturing clinical specimens from a mycetoma, because species of the aerobic actinomycetes are susceptible to antibacterial antibiotics and may be inhibited by these agents. White Grain Mycetoma: Scedosporium apiospermum complex and Acremonium spp.

As previously mentioned, white grain mycetoma are hyaline molds that produce septate hyphae. The features described here are useful for identification regardless of the disease process (i.e., mycetoma or hyalohyphomycosis). S. apiospermum complex is also involved in causing a variety of infections elsewhere in the body. These include infections of the nasal sinuses and septum, meningitis, arthritis, endocarditis, mycotic keratitis, external otomycosis, brain abscess, and disseminated invasive infection. Most of these more serious infections occur primarily in immunocompromised patients. S. apiospermum is the former asexual phase of Pseudoallescheria boydii. A nomenclature transition is currently in progress that would result in a single genus and species name for fungi that demonstrate multiple morphologic forms. The asexually produced conidia of S. apiospermum complex are golden brown, elliptical to pyriform and single-celled

853

and are borne singly from the tips of long or short conidiophores (annellophores) (Fig. 58.2). This anamorph (a fungus that disseminates reproductive structures without meiosis) predominates in cultures from clinical specimens. Another anamorphic form, the Graphium stage of S. apiospermum complex, is less common. It consists of clusters of conidiophores with conidia produced at the ends; it has also been referred to as coremia (Fig. 58.3). The teleomorphic form of the organism produces brown to black cleistothecia, which are pseudoparenchymatous, saclike structures containing asci and ascospores. When the latter are fully developed, the large (50 to 200 μm), thick-walled cleistothecia rupture, releasing the asci and ascospores (Fig. 58.1). Another Scedosporium species, Scedosporium prolificans, has been associated with infections other than mycetomas, such as arthritis or invasive disease in immunocompromised patients. S. prolificans differs from S. apiospermum in that it produces inflated, flask-shaped annellophores. The obsolete or previous name for S. prolificans was Scedosporium inflatum, which more accurately reflects the morphology of the conidiophore. Recognition of this organism also is important, because it is resistant to most if not all the commonly used antifungal agents. Acremonium spp. develops hyaline hyphae and produces simple, unbranched, erect conidiophores. Single-celled conidia are produced loosely or in gelatinous masses at the tip of the conidiophore (Fig. 58.17). Intercalary and terminal chlamydoconidia may also be produced.  Black Grain Mycetoma: Exophiala jeanselmei, Curvularia spp., and Madurella spp.

Sterile hyphae are produced when Madurella spp. is grown on rich fungal media. Nutritionally poor media may be used to induce sporulation. Long, tapering phialides with collarettes and sclerotia may be seen. Temperature tolerance, biochemical hydrolysis, and assimilation studies may be used to differentiate M. mycetomatis from T. grisea. (See Phaeohyphomycosis, later in the chapter, for the description of E. jeanselmei and Curvularia spp.)  Chromoblastomycosis: Cladosporium, Phialophora, and Fonsecaea spp.

The taxonomy of the organisms that cause chromoblastomycosis is complex. Their identification is based on somewhat distinct microscopic morphologic features. These are polymorphic fungi that may produce more than one type of conidiation. The genus Cladosporium includes species that produce long chains of budding, often fusiform, conidia (blastoconidia) that have a dark septal scar. The genus Phialophora includes species that produce short, flask-shaped to tubular phialides, each with a well-developed collarette. Clusters of conidia are produced by the phialides through an apical pore and often remain aggregated near the opening in a gelatinous mass. Phialophora spp. produce colonies that are wooly and olive-brown to brownish gray; some strains may appear to have concentric zones of color. Microscopically, hyphae are dematiaceous, and sporulation is

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•

Fig. 60.4  Pleurostomophora richardsiae (previously Phialophora richardsiae) showing phialides with prominent, saucerlike collarette (arrows) (×500).

common. Pleurostomophora richardsiae (previously Phialophora richardsiae) produces phialides with distinct flattened or saucerlike collarettes (Fig. 60.4). In contrast, P. verrucosa produces deeper, more cup- or flask-shaped phialides. Pleomorphic phialides may also be seen with these species; however, all produce either or both hyaline elliptical conidia or brown elliptical conidia within the phialides. The genus Fonsecaea includes organisms that exhibit a mixed type of sporulation. The genus produces a distinct Fonsecaea-type conidiophore, which somewhat resembles truncated Cladophialophora-type sporulation. It may also produce a Rhinocladiella-type sporulation, in which singlecelled conidia are produced on denticles that arise from all sides of conidiophores (sympodically). A mixture of the Fonsecaea, Rhinocladiella, and Cladophialophora types may occur, and phialides with collarettes or Phialophora-type sporulation also may be present. The diagnostic features of the Cladophialophora, Phialophora, and Fonsecaea genera can be summarized as follows: • C  ladophialophora (C. carrionii): Cladophialophora type of sporulation with long chains of elliptical conidia (2 to 3 μm × 4 to 5 μm) borne from erect, tall, branching conidiophores (Fig. 60.5). • P  hialophora spp.: P. verrucosa produces phialides, each with a distinct cup- or flask-shaped collarette (Fig. 60.6); P. richardsiae produces phialides with a flattened collarette (Fig. 60.4). Conidia are produced endogenously and occur in clusters at the tip of the phialide. • F  onsecaea spp.: Conidial heads with sympodial arrangement of conidia are seen, with primary conidia giving rise to secondary conidia (Fig. 60.7). Cladophialophora-type, Phialophora-type, and/or Rhinocladiella-type sporulation may also occur.  Phaeohyphomycosis: Alternaria, Bipolaris, Clado­ phialophora, Curvularia, Exophiala, Exserohilum, and Phialophora spp.

A useful approach to identification of the dematiaceous molds is first to determine whether single-celled or multicelled conidia are produced. If conidia are produced singly,

• Fig. 60.5  Cladophialophora spp. showing Cladophialophora type of sporulation (arrows) with chains of elliptical conidia (×430).

C B A

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Fig. 60.6  Phialophora verrucosa showing flask-shaped phialide (A) with distinct collarette (B) and conidia (C) near its tip (×750).

• Fig. 60.7  Both the Rhinocladiella and Phialophora types of sporulation may be produced by Fonsecaea pedrosoi and are demonstrated here (×430).

the laboratorian should determine whether they are produced individually or in chains (e.g., Cladophialophora spp.). In cellophane tape preparations, the chains of conidia produced by Cladophialophora spp. are easily disrupted. If multicellular conidia are produced, examining the septation within the conidium is useful. Multicellular conidia with septation in the horizontal axis of the conidium (i.e., the axis perpendicular to the longitudinal axis of the conidium)

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A • Fig. 60.9  Bipolaris spp. showing dematiaceous, multicelled conidia produced sympodically from geniculate conidiophores (×430).

B

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Fig. 60.8 (A) Alternaria spp. showing chaining multiform dematiaceous conidia with horizontal and longitudinal septa. (B) Microscopic morphology (200× magnification) of Alternaria spp. demonstrating growth within Biomed Diagnostics commercially prepared InTray fungal media. The design of the InTray media permits growth and imaging without preparation of microscopic slides, (Photo courtesy Biomed Diagnostics, Inc., White City, OR.)

are characteristic of certain organisms, such as in Bipolaris and Curvularia spp.; conidia with septation in both the longitudinal and horizontal axes of the conidium are characteristic of other fungi, such as Alternaria spp. Alternaria spp. Microscopically, hyphae are septate and golden-brown pigmented; conidiophores are simple but sometimes branched. Conidiophores bear a chain of large, brown conidia resembling a drumstick and contain both horizontal and longitudinal septa (Fig. 60.8). Observing chains of conidia sometimes is difficult, because they may be dislodged, as the culture mount is prepared.  Bipolaris spp. Hyphae are dematiaceous and septate. However, conidiophores are characteristically bent (geniculate) at the locations where conidia are attached; conidia are arranged sympodically and are oblong to fusoid. The hilum protrudes slightly (Fig. 60.9). Germ tubes are formed at one or both ends, parallel to the long axis of the conidium, when the fungus is incubated in water at 25°C for up to 24 hours (i.e., from both poles, thus the name Bipolaris).  Cladophialophora spp. Microscopically, hyphae are septate and brown. Conidiophores are long, branched, and give rise to branching chains of darkly pigmented, budding conidia. Conidia usually are single-celled and exhibit

•

Fig. 60.10  Cladophialophora spp. showing branching chains of dematiaceous blastoconidia that are easily dislodged during preparation of a microscopic mount (×430).

prominent attachment scars (dysjunctors). The cells that produce the branch points are often referred to as shield cells (Fig. 60.10). This organism commonly fails to reveal chains of conidia on wet mounts, because conidia are so easily dislodged.  Curvularia spp. Microscopically, hyphae are dematiaceous and septate. Conidiophores are geniculate (i.e., bent where conidia are attached). Conidia are arranged sympodically and are golden-brown, multicelled, and curved, with a central swollen cell (Fig. 60.11). The end cells are lighter in color than the swollen cell.  Exophiala spp. Only the Exophiala species E. jeanselmei and E. dermatitidis are considered here; although other species exist, they are recovered far less commonly in the clinical laboratory. The microscopic features of young colonies of Exophilia spp. exhibit dematiaceous, yeastlike cells (Fig. 60.12). Although these may appear to be budding, close inspection may disclose that the daughter cells are produced by annelides rather than true buds. The microscopic features of young colonies of Exophilia spp. exhibit dematiaceous, yeastlike cells. Feltlike, filamentous colonies produce dematiaceous hyphae and conidiophores that are cylindrical and have a tapered tip. Annellations may be visible at the tip, and clusters of oval to round conidia are apparent (Fig. 60.13). Potassium nitrate is

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• Fig. 60.11  Curvularia spp. showing twisted conidiophore and curved

A

conidia with a swollen central cell (arrows) (×500).

B • Fig. 60.13 (A) Exophiala jeanselmei showing elongated conidiophore •

Fig. 60.12  Exophiala dermatitidis showing dematiaceous, yeastlike cells from a young culture. These forms asexually reproduce via annelides rather than through true budding (blastoconidiation) (×500).

(annellophore) with a narrow, tapered tip (×500). (B) Exophiala dermatitidis showing elongated tubular annellophores (arrow); morphologically very similar to Exophiala jeanselmei (×500).

used by E. jeanselmei but not by E. dermatitidis. Temperature studies are also useful for differentiating the most common Exophiala species. Both E. jeanselmei and E. dermatitidis grow at 37°C, but only E. dermatitidis can grow at 40°C to 42°C.  Exserohilum spp. Hyphae are septate and dematiaceous. Conidiophores are geniculate, and conidia are produced sympodically. Conidia are elongate, are ellipsoid to fusoid, and exhibit a prominent hilum that is truncated and protruding (Fig. 60.14). The conidia are multicellular, have perpendicular septa, and usually contain five to nine septa. 

Antifungal Susceptibilities Antifungal susceptibilities for melanized fungi for most clinically relevant species are known. However, interpretive breakpoints have not been standardized. Amphotericin B and the azoles have demonstrated clinical effectiveness against infections with melanized fungi. Triazoles, posaconazole, and voriconazole have a broad spectrum of activity against most of these fungi. Occasional treatment

•

Fig. 60.14  Exserohilum spp. showing elongated multicelled conidia with prominent hila (arrows).

failure of mycetoma has been associated with the use of voriconazole.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

CHAPTER 60  Dematiaceous (Melanized) Molds

Bibliography Abd El-Bagi ME, Abdul Wahab O, Al-Thaqafi MA, et al.: Mycetoma of the hand, Saudi Med J 25:352, 2004. Ahmed AO, Van Leeuwen W, Fahal A, et al.: Mycetoma caused by Madurella mycetomatis: a neglected infectious burden, Lancet Infect Dis 4:566–574, 2004. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Chowdhary A, Perfect J, de Hoog S: Black molds and melanized yeasts pathogenic to humans, Cold Spring Harb Perspect Med 5: a019570, 2015. Gautier M, Ranque S, Normand AC, et al.: Matrix-assisted laser desorption ionization time-of-flight mass spectrometry: revolutionizing

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clinical laboratory diagnosis of mould infections, Clin Microbiol Infect 20:1366–1371, 2014. Meyer W, Guarro J: Current status of diagnosis of Scedosporium infections: what is the impact of new molecular methods? Curr Fungal Infect Rep 8:220–226, 2014. Pang KR, Wu JJ, Huang DB, et al.: Subcutaneous fungal infections, Dermatol Ther 17:523–531, 2004. Sakayama K, Kidani TT, Sugawara YY, et al.: Mycetoma of the foot: a rare case report and review of the literature, Foot Ankle Int 25:763– 767, 2004. Tintelnot K, von Hunnius P, de Hoog GS, et  al.: Systemic mycosis caused by a new Cladophialophora species, J Med Vet Mycol 33:349–354, 1995.

CASE STUDY 60.1 A 54-year-old female presents to her physician with a mildly tender subcutaneous cyst on her right forefinger. She experienced trauma to her hand while gardening 6 weeks earlier. The cyst is punctured, and the exudate is examined microscopically and cultured. Direct microscopic examination reveals yeastlike cells and brown, pigmented, branching, septate hyphae. The culture is slow growing and initially develops a shiny black, yeastlike colony that age to a filamentous, velvety texture with black reverse.

Questions 1. What genus of fungi should be considered? 2. If you saw cylindrical conidiophores that had annellation at the tips and clusters of conidia, what test or tests would you perform next to aid identification? 3. You find that this mold grows at 37°C but not at 42°C. What organism is this?

  

Chapter Review 1. Brown patches on the palm of the hand or the sole of the foot are caused by: a. Cladophialophora spp. b. Exophiala spp. c. Hortae werneckii d. Acremonium spp. 2. Adding specimen scrapings to 10% KOH to show the presence of sclerotic bodies that resemble copper pennies is useful in the diagnosis of: a Chromoblastomycosis b Phaeohyphomycosis c Mycetomas d Zygomycosis 3. Which of the following is the causative agent of black grain mycetoma? a  E. jeanselmei b Curvularia spp. c  Madurella spp. d All of the above 4. Which of the following is a fungus known to cause chromoblastomycosis? a  Curvularia spp. b Acremonium spp. c  Bipolaris spp. d Fonsecaea spp. 5. Which laboratory test may be used to differentiate E. jeanselmei from E. dermatitidis? a. Urea b. Growth at 42°C c. Esculin d. Germ tube

6. True or False _____ Dematiaceous fungi most commonly involve deeply invasive disease. _____ Most patients with mycetomas live in tropical locations. _____ Serologic assays are routinely used to diagnose infections caused by dematiaceous fungi. _____ P. boydii is resistant to amphotericin B. _____ Sclerotic bodies are present in histologic slides of lesions from chromoblastomycosis. 7. Matching: Match each term with the correct description. _____ H. werneckii a. granulomatous _____ P. hortae infection of the _____ Acremonium spp. lower extremities _____ Curvularia spp. b. conidia (with dark _____ Mycetoma septal scar) in long _____ Chromoblastomychains cosis c. tinea nigra _____ Cladophialophora d. black piedra spp. e. black grain mycetoma f. infections acquired via traumatic inoculation g. white grain mycetoma

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Atypical and Parafungal Agents OBJECTIVES 1. Describe the symptoms of Pneumocystis jirovecii infection and the cells affected by this organism. 2. List the appropriate specimen types collected for diagnosis of pneumocystis pneumonia. 3. Discuss the laboratory tests used in the diagnosis of P. jirovecii infection, including the methodology and biochemical principles. 4. List the diseases and morphologic characteristics used in diagnosing infections of the parafungi Lacazia loboi, Lagenidium spp., Pythium insidiosum, and Rhinosporidium seeberi.

GENUS AND SPECIES TO BE CONSIDERED Current Name Pneumocystis jirovecii Lacazia loboi Lagenidium spp. Pythium insidiosum Rhinosporidium seeberi

Previous Name Pneumocystis carinii

P. jirovecii originally was believed to be a trypanosome. Several factors supported the notion that P. jirovecii was a protozoan parasite; its morphology is similar to that of protozoa, and clinically it responds to antiprotozoal drugs but not to antifungal drugs in patients with pneumocystosis. Inability to maintain and propagate the organism in routine culture has further limited its characterization, although cultivation is possible under special conditions. P. jirovecii exists as three forms in its life cycle: the trophic form (trophozoite), sporozoite (precyst), and ascus (cyst), which is the diagnostic form. Although P. jirovecii has been shown to be a fungus, it differs from other fungi in various aspects. Its cell membrane contains cholesterol rather than ergosterol. The flexiblewalled trophic form is susceptible to osmotic disturbances. In addition, P. jirovecii contains only one or two copies of the small ribosomal subunit gene, whereas most other fungi contain numerous copies of this gene. Deoxyribonucleic acid (DNA) sequence analysis of the small ribosomal subunit gene in P. jirovecii has disclosed a greater sequence homology with the fungi than with the protozoa. Two independent analyses that compared the DNA sequences of P. jirovecii with those of other fungi confirmed the placement of P. jirovecii in the fungal kingdom, in the phylum Ascomycota. 

Epidemiology PNEUMOCYSTIS General Characteristics In 1999, the name of the organism that causes a pneumonia in immunocompromised humans, commonly called pneumocystis pneumonia (PCP), was changed from Pneumocystis carinii to Pneumocystis jirovecii. One of the causative organisms for the rodent form of pneumocystis is still called P. carinii; the other is Pneumocystis wakefieldiae. Although there are currently five recognized species of Pneumocystis, P. jirovecii is the only species that is known to infect humans. The organism is an opportunistic, atypical fungus that infects immunocompromised hosts and mostly manifests as PCP. However, transmission from infected patients to immunocompetent health care workers has been reported. 858

P. jirovecii has a worldwide distribution and most commonly presents as pneumonia in an immunocompromised host. Recently, PCP has been shifting toward non–human immunodeficiency virus (HIV)-infected immunosuppressed patients, such as hematologic malignancy and autoimmunity. The use of powerful immunosuppressive therapies has led to this shift. Pneumocystis is transmitted person-to-person via airborne particles. Immunocompetent individuals appear to be the reservoir for P. jirovecii, which is transmitted to immunodeficient individuals as a pathogen. Most children by age 2 to 4 years have antibodies to Pneumocystis, suggesting acquisition early in life. Vargas et al. showed that Pneumocystis DNA was present in 24 of 72 infants, as determined from nasopharyngeal specimens, and that seroconversion occurred in 85% of infants by 20 months of age.

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Since the onset of the HIV and acquired immunodeficiency syndrome (AIDS) epidemic in the 1980s, Pneumocystis has been defined as the most common opportunistic infection among those with HIV or AIDS in the United States. The introduction of highly active antiretroviral therapy (HAART) for patients with HIV has reduced the incidence of disease. However, PCP remains a significant medical problem because numerous patients with HIV do not respond to therapy, do not comply with therapy, or do not know they are infected. The results of DNA testing demonstrate the detection of P. jirovecii in immunocompetent populations as well as additional groups of patients with chronic underlying disease. 

Pathogenesis and Spectrum of Disease After P. jirovecii is inhaled, the trophic form of the pathogen is believed to adhere to type I pneumocytes (thin squamous epithelial cells of the lungs). The organisms replicate extracellularly while bathed in alveolar lining fluid. With successful replication of the organism, the alveolar spaces fill with an eosinophilic foamy material, which can be detected with hematoxylin and eosin staining. This technique does not provide direct staining of the organisms. Methenamine silver or another fungal stain may be used to identify the cyst form of the organism in lung tissue. Infection with the organism and the pathophysiologic changes described result in impaired oxygen-diffusing capacity and hypoxemia. A predominantly interstitial mononuclear inflammatory response is associated with this type of pneumonia. When first described, this pneumonia was known as interstitial plasma cell pneumonia. Symptoms of PCP include a nonproductive cough, low-grade fever, dyspnea, chest tightness, and night sweats. In patients without HIV infection, the underlying conditions most commonly seen as risk factors for this opportunistic infection are asthma, chronic obstructive pulmonary disease (COPD), cystic fibrosis, systemic lupus erythematosus (SLE), pregnancy, rheumatoid arthritis, infection with Epstein-Barr virus, ulcerative colitis, and high-dose corticosteroid therapy. During treatment with an antiretroviral medication, patients show an improvement and an increase in CD4+ cells. However, following a brief period of improvement, the patients begin to deteriorate because of an exaggerated immune response referred to as immune reconstitution inflammatory syndrome. Extrapulmonary infection has been reported in 0.6% to 3% of postmortem samples collected from patients who were diagnosed with P. jirovecii pneumonia. Extrapulmonary cysts have been identified in lymph nodes, the spleen, bone marrow, and the liver, predominantly. Additional extrapulmonary sites include the adrenal glands, gastrointestinal tract, genitourinary tract, thyroid, ear, pancreas, eyes, and skin. Multiple sites of infection typically indicate a more rapid disease progression and fatal outcome. 

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•

Fig. 61.1 Cystic forms of Pneumocystis jirovecii (arrows) stain well with methenamine silver and hematoxylin and eosin stain (×500).

Laboratory Diagnosis Specimen Collection and Transport Respiratory specimens from the deep portions of the lung, such as bronchoalveolar lavage (BAL), are best for detection of P. jirovecii. A sputum specimen submitted for direct examination should be induced sputum obtained by a trained respiratory therapist; otherwise, the rate of falsenegative results may be unacceptably high. Additional acceptable respiratory specimens include tracheal aspirates, pleural fluid, transbronchial biopsy, or cellular material from bronchial brushings. Nasopharyngeal and oropharyngeal samples have demonstrated high sensitivity and specificity for the diagnosis of PCP when used in nucleic acid–based testing methods for P. jirovecii. Collection for the diagnosis of extrapulmonary pneumocystis requires biopsy of the infected organ and histologic staining. 

Specimen Processing See the following section for specific details for specimen processing required for the various test methods. 

Direct Detection Methods Stains The diagnosis of P. jirovecii pneumonia currently is based on the clinical presentation, radiographic studies, and direct or pathologic examination of respiratory samples or biopsy material. The flexible-walled trophic forms are the predominant morphology of the organism, but these are difficult to visualize. They are somewhat discernible in Giemsa-stained material, but their pleomorphic appearance makes this form of the organism difficult to identify. Giemsa stains the nuclei of all the various life cycle stages as reddish purple with a light blue cytoplasm. A firm-walled cyst also exists, although the cysts are outnumbered by the trophozoites 10 to 1. Cysts are more easily recognized than the trophic form and may be definitively identified using a variety of stains, such as calcofluor white, methenamine silver, and immunofluorescent staining (Fig. 61.1). The cysts are spherical to

860 PA RT V    Mycology

concave, are uniform in size (4 to 7 μm in diameter), do not bud, and contain distinctive intracystic bodies. A comparison of the four most common staining methods used for P. jirovecii (i.e., Giemsa, immunofluorescent, calcofluor white, and methenamine silver) has demonstrated that immunofluorescent staining (Merifluor ­Pneumocystis; Meridian Bioscience, Cincinnati, OH), Monofluo P. jirovecii IFA (Bio-Rad, Hercules, CA), calcofluor white staining (Fungifluor; Polysciences, Warrington, PA), and methenamine silver staining (GMS) and Wright-Giemsa (Diff-Quik; Baxter Scientific, McGaw Park, IL) likely represent the best balance between sensitivity and specificity and have the best overall positive and negative predictive values. The immunofluorescent method showed greater sensitivity than the other three but a smaller negative predictive value. Therefore, if this method is used as a screening tool for the presence of Pneumocystis, a confirmatory method should be performed because of the high number of false-positive results. 

Direct Detection of (1-3)-Beta-D-Glucan The ascus (cyst) cell wall component, (1-3)-beta-D-glucan, has been used to successfully diagnose infections with P. jirovecii. Other fungi also secrete the molecule but in lower amounts. There are several commercial assays available; however, they are not all Food and Drug Administration (FDA) approved for use within the United States. The Fungitell Assay (Associates of Cape Cod, Falmouth, MA) uses patient serum for the detection of (1-3)-beta-D-glucan. Normal human serum contains low levels of (1-3)-beta-Dglucan (10 to 40 pg/mL) from the commensal yeasts that are present in the alimentary canal and gastrointestinal tract. Therefore values less than 60 pg/mL are considered negative, 60 to 79 pg/mL are indeterminate, and greater than 80 pg/ mL are positive. It is important to use additional diagnostic information and confirmatory testing in conjunction with this test, because other yeast or fungi also secrete (1-3)-betaD-glucan during infection. 

Nucleic Acid Detection A variety of nucleic acid amplification assays for P. jirovecii have been developed, including, most recently, real-time polymerase chain reaction (PCR) methods and multiplex assays. However, because of the potential for colonization of immunocompetent populations, positive nucleic acid– based testing results must be directly correlated with the patient’s history and clinical presentation. As of this writing, no tests have been FDA approved for use in the United States.

Serologic Testing Serology is useful for epidemiological purposes but not for the diagnosis. 

Cultivation P. jirovecii is very difficult to cultivate outside the lung; therefore routine culture methods are not performed. 

Approach to Identification See Direct Detection Methods. 

Treatment Trimethoprim-sulfamethoxazole (TMP-SMX) and pentamidine isethionate are the predominant agents used to treat PCP. Both drugs have significant side effects including nephrotoxicity. The use of TMP-SMX is also associated with the development of resistant strains. 

Rare Atypical and Parafungal Agents Lacazia loboi L. loboi is the causative agent of lobomycosis, a rare granulomatous zoonotic fungal infection mostly of the skin and subcutaneous tissue. The lesions and nodules are described as leprosy-like and generally appear in cooler regions of the body, indicating the pathogen does not grow well at body temperature (37°C). The organism has been found in the soil, on vegetation, and in aquatic animals in tropical and subtropical areas, especially the bottlenose dolphins. The transmission to humans is not known but theorized that infection results from traumatic entry or entry into broken skin from contaminated water. Direct dolphin to human transmission has been reported by an aquarium attendant. Human to human transmission has also been reported.

Laboratory Diagnosis and Treatment L. loboi can be seen in stained tissue biopsies; the organism itself is uncultivable in  vitro. Morphologically, the organism resembles Paracoccidioides brasiliensis (Chapter 62). Biopsied material will show yeastlike cells and an influx of inflammatory cells. The cells are uniform in size, with a thick outer membrane and appear to be chaining, connected by small tubules. The uniformity in size can help to differentiate from P. brasiliensis. Treatment of infection with L. loboi is difficult because this organism is resistant to most antifungals; therefore surgical removal is required. 

Pythium insidiosum P. insidiosum is a funguslike, aquatic oomycete organism. The organism is found in the tropical, subtropical, and temperate areas. Isolation of P. insidiosum can be performed using mycologic media such as Sabouraud dextrose agar. The organism has two phases, a more funguslike, myceliumproducing phase and a biflagellate zoospore, which is the infectious stage. Zoosporogenesis can be initiated only in water cultures in vitro. Infections are thought to be acquired by traumatic injection into the skin or the intestinal tract by traumatic lesions from contaminated aquatic environments. Infective keratitis has also been reported. Person to person transmission has not been observed. Infection with P. insidiosum includes cutaneous and subcutaneous lesions with the formation of plaques and ulcers. Orbital infections have also been reported. Vascular pythiosis can be seen

CHAPTER 61  Atypical and Parafungal Agents

more commonly in Thailand, especially in patients with thalassemia.

Laboratory Diagnosis and Treatment Serologic testing is not specific and therefore not recommended. Tissue samples are used for diagnosis and can be examined by direct microscopy. Samples may be stained with a variety of immunohistochemical stains. Stained skin and tissue samples typically demonstrate hyphal structures that are short or long, sparsely septate, tubular structures and inflammatory cells, especially eosinophils and mast cells. A 10% potassium hydroxide (KOH) preparation may also be used to visualize the hyaline hyphal elements in tissue scrapings. Culture can be used for the diagnosis of infection with P. insidiosum. Small pieces of tissue should be embedded into the mycological media and incubated at 25°C and 37°C for 24 to 48 hours. Specimens received more than 24 hours after collection should be placed in a broth tube and incubated at 37°C. Two percent Sabouraud dextrose agar or broth with or without antibiotics is the most common media used to isolate P. insidiosum. However, identification of P. insidiosum requires the development of the characteristic oogonia (sexual stage) in culture. This is extremely rare. Nucleic acid–based testing has been used to identify the organism and is recommended over culture. Identification is therefore limited by the lack of availability of the molecular test in clinical laboratories. 

Treatment P. insidiosum do not contain chitin and ergosterol similar to yeasts and fungi; therefore, the organism is resistant to antifungals. Combination antibiotics including minocycline, linezolid, and chloramphenicol have been successfully used to treat keratitis associated with P. insidiosum. 

Lagenidium spp. Lagenidium spp. is an emerging oomycete similar to P. insidiosum and the causative agent of lagenidiosis. As with P. insidiosum, Lagenidium spp. has two forms, the mycelial form and the biflagellate zoospore. Lagenidium cases have been found in wet areas. The organism is found in crabs, mosquito larvae, nematodes, and other organisms. Person-to-person transmission has not been identified. The organism causes invasive skin, subcutaneous, and arterial infections. Additional infection sites have been also noted: cornea, gastrointestinal tract, and extremities. Systemic infections have also been identified in humans and animals. 

Laboratory Diagnosis The organism can be examined using microscopy and isolated on fungal media similar to P. insidiosum. On solid media, Lagenidium spp. produces white-yellow submerged colonies. The hyphae have spherical structures, which can be seen at the end of the hyphae in liquid cultures.

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Histopathologic staining of the infective sites cannot differentiate between Lagenidium or P. insidiosum. Morphology is similar, and both initiate localized eosinophilia. The development of the zoospores and the presence of broad, aseptate hyphae in wet mounts, as well as identification by molecular methods, should be used to confirm the presence of Lagenidium spp.

Treatment Lagenidium spp. lack sterols in their cellular membranes and are therefore resistant to antifungal medications. 

Rhinosporidium seeberi R. seeberi is described as a Mesomycetozoea protistal eukaryote. This organism is similar morphologically to the parasitic form of Coccidioides. Infections are usually identified in tropical or subtropical areas. The mechanism of transmission is unknown; however, the pathogen can be found in aquatic environments, and resistant spores are present in terrestrial environments. Infection is likely acquired by exposure to resistant spores through breaks in the skin or mucous membranes. Infection results in the formation of painless polyps in the mucosal areas of the nose, eye, larynx, genitalia, and rectum.

Laboratory Diagnosis Tissue biopsies are the preferred method used for diagnosis. Wet mounts from the polyps demonstrating sporangia and endospores are usually present in cases of rhinosporidiosis. Confirmation of rhinosporidiosis disease is based on the identification of greater than 300 μm spherical sporangia with endospores and a negative fungal culture. 

Treatment R. seeberi is resistant to antifungals. Surgical removal of the infected tissue and polyps can be used, but recurrence is common.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Caroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Dick MW: Straminipilous fungi: systematics of the peronosporomycetes, including accounts of the marine straminipilous protists, the plasmodiophorids, and similar organisms, ed 1, Dordrecht, Netherlands, 2001, Kluwer Academic Publishers. Gaastra W, Lipman LJA, De Cock AWAM, et al.: Pythium insidiosum: an overview, Vet Microbiol 146:1–16, 2010. Giuintuli D, Stringer S, Stringer J: Extraordinary low number of ribosomal RNA genes in P. carinii, J Eukaryot Microbiol 41:88S, 1994. Kaplan JE, Hanson D, Dworkin MS, et al.: Epidemiology of human immunodeficiency virus–associated opportunistic infections in the United States in the era of highly active antiretroviral therapy, Clin Infect Dis 30(suppl 1):S5–S14, 2000.

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Karm MB, Mosadegh L: Extra-pulmonary Pneumocystis jirovecii infection: a case report, Braz J Infect Dis 18:681–685, 2014. Maeno S, Yoshinori O, Sunada A, et al.: Successful medical management of Pythium insidiosum keratitis using a combination of minocycline, linezolid, and chloramphenicol, Am J Ophthalmol Case Rep 15:100498, 2019. Mendoza L, Newton JC: Immunology and immunotherapy of the infections caused by Pythium insidiosum, Med Mycol 43:477–486, 2005. Otieno-Odhiambo P, Wasserman S, Hoving JC: The contribution of host cells to Pneumocystis immunity: an update, Pathogens 8:52, 2019. Procop GW, Haddad S, Quinn J, et  al.: Detection of Pneumocystis jirovecii in respiratory specimens by four staining methods, J Clin Microbiol 42:3333–3335, 2004. Putthia H, Manjuanatha BS, Astekar M, et al.: Palatal rhinosporidiosis: an unusual case report and review of the literature, J Korean Assoc Oral Maxillofac Surg 44(6):293–297, 2018.

Seyedmousavi S, Guillot J, Tolooe A, et  al.: Neglected fungal zoonoses: hidden threats to man and animals, Clin Microbiol Infect 21:416–425, 2015. Singh CA, Sakthivel P: Rhinosporidiosis, N Engl J Med 380(14):1359, 2019. Spies CFJ, Grooters AM, Lévesque CA, et al.: Molecular phylogeny and taxonomy of Lagenidium-like oomycetes pathogenic to mammals, Fungal Biol 120:931–947, 2016. Stringer JR: Pneumocystis carinii: what is it, exactly? Clin Microbiol Rev 9:489–498, 1996. Thianprasit M: Human pythiosis, Trop Dermatol 4:1, 1990. Vargas SL, Hughes WT, Santolaya ME, et  al.: Search for primary infection by Pneumocystis carinii in a cohort of normal, healthy infants, Clin Infect Dis 32:855–861, 2001. White PL, Price JS, Backx M: Therapy and management of Pneumocystis jirovecii infection, J Fungi (Basel) 4:127, 2018.

CASE STUDY 61.1 A 69-year-old male is diagnosed with follicular lymphoma and placed on a 4-week regimen of combination chemotherapy. During the fourth week of treatment, the patient develops a fever that lasts 3 days. On the third day, he is admitted to the hospital. The findings from the initial physical examination are normal except for the fever. The following laboratory test results are obtained:

White Blood Cell (WBC) count Absolute neutrophil count Serum betaD-glucan

2.5 × 109/L

Computed tomography (CT) scan of the lungs

Ground-glass opacities in the lung

1.1 × 109/L 182 pg/mL

Normal: 4.5–11.0 × 109/L Normal: 50%–60% of WBCs Normal: 100 mg/dL)

Low (50 mg/dL)

Normal or often low (>45 mg/dL)

aMust

consider cerebrospinal fluid (CSF) glucose level in relation to blood glucose level. Normally, the CSF glucose serum ratio is 0.6, or 50%–70% of the blood glucose normal value. bAbout 20%–75% of cases may have PMN leukocytosis early during infection.

level. Children older than 6 years are less likely to develop meningitis, but the risk for meningitis infection increases when the child reaches early adulthood. As previously mentioned, neonates have the highest incidence of acute meningitis, with a concomitant increased mortality rate (as high as 20%). Organisms causing disease in the newborn are different from those that affect other age groups; many of them are acquired by the newborn during passage through the mother’s vaginal vault. Neonates are likely to be infected with, in order of incidence, group B streptococci, Escherichia coli, other gram-negative bacilli, and Listeria monocytogenes; occasionally other organisms may be involved. For example, Elizabethkingia meningoseptica and Elizabethkingia anopheles are both associated with severe cases of neonatal meningitis. These organisms are normal inhabitants of water in the environment and may be transmitted from mother to fetus or acquired as a nosocomial infection. Important causes of meningitis in adults, in addition to the meningococcus in young adults, include pneumococci, L. monocytogenes, and, less commonly, Staphylococcus aureus and various gram-negative bacilli (E. coli, Klebsiella spp., Serratia marcescens, Pseudomonas aeruginosa, Acinetobacter spp., and Salmonella spp.). Meningitis caused by the latter organisms results from hematogenous seeding from various sources, including urinary tract infections. Spirochetal meningitis or neurosyphilis can be caused by dissemination of Treponema pallidum early in the infectious process. CSF abnormalities may occur in up to 9% of patients who are seronegative for syphilis and may overlap in clinical presentation, including asymptomatic, meningeal, meningovascular, parenchymatous, and gummatous. Although Naegleria fowleri is the primary cause of meningoencephalitis, additional genera may also be associated with infection, including Acanthamoeba and Balamuthia. Parastrongylus cantonensis is the most common cause of eosinophilic meningitis or encephalitis. The nematode larvae invade the CSF directly from the bloodstream and mature into adult worms migrating throughout the brain. Additional organisms

capable of causing eosinophilic meningitis include Paragonimus westermani, Gnathostoma spp. (myeloencephalitis), Baylisascaris procyonis (neural larva migrans [NLM]), and Taenia solium.  Aseptic Meningitis

Aseptic meningitis is usually viral and is characterized by an increase of lymphocytes and other mononuclear cells (pleocytosis) in the CSF; bacterial and fungal cultures are negative. This contrasts with bacterial meningitis, which is characterized by purulence and the PMN cell response in the CSF. However, viral meningitis may mimic bacterial meningitis with an increase in PMNs within the first 24 hours followed by a transition to a lymphocytic pleocytosis. Aseptic meningitis is usually self-limiting, with symptoms that may include fever, headache, stiff neck, nausea, and vomiting. In addition to the increase of lymphocytes and other mononuclear cells in the CSF, the glucose level remains normal, whereas the protein CSF level may remain normal or may be slightly elevated. Aseptic meningitis can also be a symptom for syphilis and some other spirochete diseases (e.g., leptospirosis and Lyme borreliosis). Stiff neck and CSF pleocytosis may also be associated with other disease processes, such as malignancy. Enteroviruses are currently the leading cause of aseptic meningitis. Infants and young children are the most common population susceptible to infection. Additional viral agents associated with aseptic meningitis include herpes simplex virus (HSV); varicella-zoster-virus (VZV); cytomegalovirus (CMV); Epstein-Barr virus (EBV); and human herpesviruses 6, 7, and 8. The mumps virus may contribute to aseptic meningitis cases in nonimmunized populations. 

Encephalitis/Meningoencephalitis There are over 100 different pathogens capable of causing encephalitis. Encephalitis is an acute inflammation of the

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brain parenchyma and is usually caused by direct viral invasion. Concomitant meningitis occurring with encephalitis is known as meningoencephalitis, and the cellular infiltrate present in the CSF is typically lymphocytic rather than PMNs. The host response to these CNS infections can differ somewhat from those associated with purulent or aseptic meningitis. Early during viral encephalitis, or when considerable tissue damage occurs as a part of encephalitis, the nature of the inflammatory cells found in the CSF may be no different from that associated with bacterial meningitis; cell counts, however, are typically much lower.

Viral Encephalitis Viral encephalitis, which cannot always be distinguished clinically from meningitis, is common in the warmer months. The primary agents are enteroviruses (coxsackie viruses A and B and echoviruses), mumps virus, herpes simplex virus, and arboviruses (West Nile virus, togavirus, bunyavirus, equine encephalitis, St. Louis encephalitis, and other encephalitis viruses). Herpes simplex virus is the most common virus associated with viral encephalitis in developed countries. Other viruses—such as measles, cytomegalovirus, lymphocytic choriomeningitis, Epstein-Barr virus, hepatitis, varicella-zoster virus, rabies virus, myxoviruses, and paramyxoviruses—are less commonly encountered. Any preceding viral illness and exposure history are important considerations in establishing a cause by clinical means. West Nile virus is the most common flavivirus reported in the United States with cases identified in all 48 contiguous states. Neuroinvasive infection with West Nile presents with symptoms of headache, fever, and a change in consciousness along with altered mental status. Examination of the CSF shows an increase in leukocytes with a marked increase in lymphocytes. Chemistries demonstrate an elevated protein count and normal glucose levels. Definitive diagnosis requires testing for the presence of the IgM antibody to West Nile in the serum or CSF, and because IgM does not cross the blood-brain barrier, presence of IgM antibody to West Nile in the CSF is a strong indicator for CNS infection. Polymerase chain reaction (PCR) can also be used to test for West Nile infection, but because West Nile infections have a transient and low viremia, results must be interpreted with caution. A negative result does not necessarily rule out West Nile infection. Involvement of the nervous system in patients who are infected with the human immunodeficiency virus (HIV) is common. HIV is a neurotropic (attracted to nerve cells) virus capable of entering the CNS by macrophage transport and is the cause of various neurologic syndromes. As HIV-infected individuals become progressively more immunosuppressed, the CNS becomes a target for opportunistic pathogens, such as cytomegalovirus, BK virus, and JC (John Cunningham) virus, which can produce meningitis or encephalitis. BK virus is named after the initials of the first renal transplant patient where the virus was identified in association with clinical disease. 

Parasitic Infections Parasites can cause meningoencephalitis, brain abscess (see the following discussion), or other CNS infection via two routes. A rare but devastating meningoencephalitis is caused by the free-living amoeba N. fowleri, which invades the brain via direct extension from the nasal mucosa. These organisms are acquired during swimming or diving in natural, stagnating freshwater ponds and lakes. Although N. fowleri is the primary cause of meningoencephalitis, additional genera may also be associated with infection including Balamuthia spp. and Acanthamoeba spp. (granulomatous amebic encephalitis); Sappinia pedata, is now a confirmed human parasitic pathogen linked to the development of encephalitis. Other parasites reach the brain via hematogenous spread. Toxoplasmosis, caused by an intracellular parasite that destroys brain parenchyma, is a common CNS affliction in HIV-infected patients with acquired immune deficiency syndrome (AIDS). Entamoeba histolytica and Strongyloides stercoralis have been identified in brain tissue, and the larval form of T. solium (the pork tapeworm), referred to as a cysticercus, can travel to the brain via the bloodstream and encyst within the brain tissue. Amoebic brain infection and cysticercosis cause changes in the CSF that are similar to meningitis. Parastrongylus cantonensis is the most common cause of eosinophilic meningitis. The nematode larvae invade the CSF directly from the bloodstream and mature into adult worms migrating throughout the brain. Additional organisms capable of causing eosinophilic meningitis include Gnathostoma spp., Baylisascaris procyonis, Paragonimus westermani, and T. solium. Symptoms include headache and visual disturbances. Approximately 50% of the patients’ experience vomiting and moderate fever. Approximately 10% to 20% of patients infected with Trichinella spiralis may exhibit CNS involvement with a mortality rate of 50% if untreated. Symptoms can mimic meningitis or encephalitis. 

Brain Abscess Brain abscesses (localized collections of pus in a cavity formed by the breakdown of tissue) may occasionally cause changes in the CSF and clinical symptoms similar to meningitis. Brain abscesses result from contiguous infection of the sinuses, middle ear, or mastoids (25% to 50%), hematogenously (15% to 30%), or through direct inoculation as a result of trauma or surgery (8% to 19%). Brain abscesses may rupture into the subarachnoid space, producing severe meningitis with a high mortality rate. If anaerobic organisms or viridans streptococci are recovered from CSF cultures, the diagnosis of brain abscess should be considered; however, CSF culture is typically negative in brain abscess. Patients who are immunosuppressed or who have diabetes with ketoacidosis may present with a rapid progressive fungal infection (phycomycosis) of the nasal sinuses or palatal region capable of traveling directly to the brain. The complex polymicrobial infections isolated from brain abscesses are far too extensive to list. 

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Shunt Infections Information and studies related to infections involving CSF shunts is limited. The organisms reported to be most commonly associated with infections include coagulasenegative staphylococcus, S. aureus, Propionibacterium acnes, and viridians group streptococci. A few gram-negative rods have been identified, including P. aeruginosa, Klebsiella spp., E. coli, and S. marcescens. Positive cultures are most often associated with shunt tip cultures, shunt valves, and cerebral ventricle fluid. 

Laboratory Diagnosis of Central Nervous System Infections Meningitis Except in unusual circumstances, a lumbar puncture (spinal tap) is one of the first steps in the diagnosis of a patient with suspected CNS infection, in particular, meningitis. Refer to Table 5.1 to review the procedure for collecting, transporting, and processing specimens obtained from the central nervous system.

Specimen Collection and Transport CSF is collected by aseptically inserting a needle into the subarachnoid space (lumbar puncture), at the lumbar spine region between L3, L4, or L5. Three or four tubes of CSF should be collected into sterile collection tubes that contain no additives. The tubes are numbered sequentially in the order in which they were collected along with the patient’s name. When processing the CSF collection tubes in the laboratory, tube 1 is used for chemistry studies, glucose and protein count, and immunology studies, because these tests are least affected by the presence of blood cells or bacteria introduced as a result of the spinal tap procedure; tube 2 is used for culture, allowing a larger proportion of the total fluid to be concentrated, which can facilitate the detection of infectious agents present in low numbers; tubes 3 and 4 are used for cell count and differential, because these tubes are least likely to contain cells introduced by the collection procedure. If a small capillary blood vessel is inadvertently broken during the spinal tap, blood cells picked up from this source will usually be absent from the last tube collected; comparison of counts between tubes 1 and 3 or 4 is occasionally needed if a traumatic tap is suspected or to differentiate a traumatic bloody tap from a true subarachnoid hemorrhage. In a traumatic tap, the red blood cells will be unevenly distributed among the three tubes, with the heaviest concentration of red blood cells in tube 1 and diminishing amounts in all subsequent tubes. In an intracranial hemorrhage, the red blood cells will be evenly distributed among all the tubes. If only one tube of CSF is collected, it should be submitted to microbiology first. The volume of CSF that can be collected is based on the volume available in the patient (adult versus neonate) and the opening pressure of the CSF when the needle first punctures the

999

subarachnoid space. An elevated pressure requires the CSF fluid to be withdrawn more slowly, which may prevent the collection of a larger volume. The volume of CSF is critical for detecting certain microorganisms, such as mycobacteria and fungi. A minimum of 5 to 10 mL is recommended for detecting these agents by centrifugation and subsequent culture. When the laboratory receives an inadequate volume of CSF, the physician should be consulted regarding the order of priority for laboratory tests. Processing too little specimen lowers the sensitivity of laboratory tests, which may lead to false-negative results. This is potentially more harmful to patient care than performing an additional lumbar puncture to obtain the necessary amount of sample. CSF should be hand-delivered immediately (≤15 minutes at room temperature) to the laboratory. Certain agents, such as S. pneumoniae, may not be detectable after an hour or longer. Specimens for microbiology studies should never be refrigerated; if not rapidly processed, CSF should be incubated (35°C) or left at room temperature. One exception to this rule involves CSF for viral studies. These specimens may be refrigerated for as long as 48 hours after collection or frozen at −70°C if a longer delay is anticipated until they are processed and inoculated into culture media. CSF for viral studies should never be frozen at temperatures above −70°C. CSF samples for viral identification should also not be added to transport media as they do not require antimicrobials to suppress other microorganisms. In addition, dilution using transport media may cause false-negative results. If not processed immediately, CSF specimen for hematology studies can be refrigerated, whereas the CSF for chemistry and serology can be frozen (−20°C). Information gathered from specimen analysis should be promptly relayed to the clinician, who can directly affect therapeutic outcome. Such specimens should be processed immediately upon receipt in the laboratory (STAT) and results reported to the physician as soon as possible. 

Initial Processing Initial processing of CSF for bacterial, fungal, or parasitic studies includes centrifugation of all specimens with a volume greater than 1 mL for least 15 minutes at 1500×g. CSF specimens collected for viral nucleic acid–based tests should not be centrifuged prior to nucleic acid extraction, as most viral nucleic acid will be cell associated. Specimens in which cryptococci or mycobacteria are suspected require special handling. (Discussions of techniques for culturing CSF for mycobacteria and fungi are found in Chapters 42 and 58, respectively.) If less than 1 mL of CSF is available, the specimens should be Gram stained and plated directly to blood, chocolate agar, and primary fungal isolation media, when appropriate. For bacterial culture, the supernatant is removed to a sterile tube, leaving approximately 0.5 mL of fluid. For fungal culture, the supernatant should not be removed unless a portion is required for cryptococcal antigen testing. The remaining fluid is used to suspend the sediment for visual examination or culture. Mixing of the sediment after the supernatant has been removed is critical

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for bacterial culture. Forcefully aspirating the sediment up and down into a sterile pipette several times will adequately disperse the organisms that remained adherent to the bottom of the tube after centrifugation. Laboratories that use a sterile pipette to remove portions of the sediment from underneath the supernatant will miss a significant number of positive specimens. The supernatant can be used to test for the presence of antigens, for rapid diagnostic tests (vertical flow immunochromatography), to test for N. meningitidis, or for chemistry evaluations (e.g., protein, glucose, lactate, C-reactive protein). As a safeguard, the laboratorian should keep the supernatant even if it has no immediate use. 

Cerebrospinal Fluid Laboratory Results As previously mentioned, CSF is also removed for analysis of cells, protein, and glucose. Ideally, the glucose content of the peripheral blood is determined simultaneously for comparison with CSF levels. General guidelines for the interpretation of results are shown in Table 70.2. Because the results of hematologic and chemical tests directly relate to the probability of infection, communication between the physician and the microbiology laboratory is essential. The diagnosis of acute bacterial meningitis can be excluded in patients with normal fluid parameters in almost all cases, precluding further expensive and laborintensive microbiologic processing beyond a standard smear and culture (which must be included in all cases). One exception is a patient infected with Listeria monocytogenes. Forty percent of patients with listeriosis demonstrate a positive CSF culture for isolation of the organism; however, the CSF may demonstrate normal cell counts and the Gram stain may not show any bacteria. L. monocytogenes can present as meningitis, encephalitis, abscess, or ventriculitis. Similar criteria have been used to exclude performance of smear and culture for tuberculosis and syphilis serology on CSF specimens. 

Visual Detection of Etiologic Agents After centrifugation, the resulting CSF sediment may be visually examined for the presence of cells and organisms. Stained Smear of Sediment

Gram stain must be performed on all CSF sediments. False-positive smears have resulted from inadvertent use of contaminated slides. Therefore use of alcohol-dipped and flamed or autoclaved slides is recommended. After thoroughly mixing the sediment, a heaped drop is placed on the surface of a sterile or alcohol-cleaned slide. The sediment should never be spread out on the slide surface, because this increases the difficulty of finding small numbers of microorganisms. The drop of sediment is allowed to air dry, is heat or methanol fixed, and is stained by either Gram stain (Fig. 70.3) or acridine orange. The acridine orange fluorochrome stain may allow faster examination of the slide under highpower magnification (400×) and thus a more thorough examination. The brightly fluorescing bacteria will be easily visible. All suspicious smears can be stained using the Gram

• Fig. 70.3  Gram stain of cerebrospinal fluid showing white blood cells

and many gram-positive diplococci. This specimen subsequently grew Streptococcus pneumoniae.

stain (directly over the acridine orange) to confirm the presence and morphology of organisms. Using a cytospin centrifuge to prepare slides for staining has also been found to be an excellent alternative procedure. This method for preparing smears for staining concentrates cellular material and bacterial cells up to a 1000-fold. By centrifugation, a small amount of CSF (or other body fluid) is concentrated onto a circular area of a microscopic slide (Fig. 70.4), fixed, stained, and then examined. The presence or absence of bacteria, inflammatory cells, and erythrocytes should be reported after examination. Based on demographic and clinical patient data and Gram stain morphology, the cause of most bacterial meningitis cases can be presumptively determined within the first 30 minutes after receipt of the specimen.  Wet Preparation

Amoebas are best observed by examining thoroughly mixed sediment as a wet preparation under phase-contrast microscopy. If a phase-contrast microscope is not available, observing under light microscopy with the condenser closed slightly can be used as an alternative. Amoebas are identifiable by their typical slow, methodical movement in one direction via pseudopodia. The organisms may require a little time under the warm light of the microscope before they begin to move. Organisms must be distinguished from motile macrophages, which occasionally occurs in CSF. If a wet preparation is suspicious, a trichrome stain can assist in the differentiation of amoebas from somatic cells. The pathogenic amoebas can be cultured on a lawn of K. pneumoniae or E. coli (Chapter 46).  India Ink Stain

The large polysaccharide capsule of Cryptococcus spp. allows these organisms to be visualized by the India ink stain. However, latex agglutination testing for capsular antigen is more sensitive and extremely specific. Lateral flow assays are now available for the primary screening of CSF for suspected cases of cryptococcal meningitis. Antigen testing is

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B

A

• Fig. 70.4  (A) Cytocentrifuge. (B) Device used to prepare the concentrated smears of material from body

fluid specimens such as cerebrospinal fluid by cytocentrifugation. (A, Courtesy Cytospin 2, Shandon, Inc., Pittsburgh, PA.)

recommended over the use of an India ink stain. Furthermore, strains of Cryptococcus spp. that infect patients with AIDS may not possess detectable capsules, making culture essential. To perform the India ink preparation, a drop of CSF sediment is mixed with one-third volume of India ink. The India ink can be protected against contamination by adding 0.05 mL thimerosal (Merthiolate, Sigma Chemical Co., St. Louis, MO) to the stain. After mixing the CSF and ink to make a smooth suspension, a coverslip is applied to the drop, and the preparation is examined under highpower magnification (400×) for characteristic encapsulated yeast cells, which can be confirmed by examination under oil immersion. Inexperienced microbiologists must be careful not to confuse white blood cells with yeast. The presence of encapsulated buds, smaller than the mother cell, is diagnostic. 

Direct Detection of Etiologic Agents Antigen

Commercial reagents and kits are available for the rapid detection of antigen in the CSF; the following sections review the methodologies used; for more detailed specifics, refer to Chapter 9. Bacteria. Rapid antigen detection from CSF has been largely accomplished by the techniques of latex agglutination (Chapter 9). All commercial agglutination systems use the principle of an antibody-coated particle capable of binding to specific antigen, resulting in macroscopically visible agglutination. The soluble capsular polysaccharide found in the common etiologic agents of meningitis, including the group B streptococcal polysaccharide, are well suited to serve as bridging antigens. The agglutination assays may contain either a polyclonal or monoclonal antibody or an antigen from an infectious agent. In general, the commercial systems have been developed for use with CSF, urine, or serum, although results with serum have not been as diagnostically useful as those with CSF. Soluble antigens from Streptococcus agalactiae and H. influenzae may concentrate in the urine. Urine, however, seems to produce a higher incidence of nonspecific reactions than either serum or CSF. The manufacturers’ directions must be followed for performance of antigen detection test

systems for different specimen types. Although some of the systems require pretreatment of samples (usually heating for 5 minutes), not all manufacturers recommend pretreatment. The reagents, however, may yield false positives or cross reactions if the specimen is not pretreated. Interference by rheumatoid factor and other substances, more often present in body fluids other than CSF, has also been reported. The rapid extraction of antigen procedure (REAP; Evolve Procedure 70.1) has been shown to effectively reduce a substantial portion of nonspecific and false-positive reactions, at least for tests performed with latex particle reagents. This procedure is recommended for laboratories that use commercial body fluid antigen detection kits. Some commercial systems have an extraction procedure included in the protocol. Based on the findings of several studies, only a limited number of clinically useful situations warrant bacterial antigen testing (BAT). Examples include CSF specimens from previously treated patients and Gram stain–negative CSF specimens with abnormal parameters (elevated protein, decreased glucose, or an abnormal white blood cell count). The assays are not substitutes for properly performed smears and cultures. Some of the assays demonstrate a decreased sensitivity and specificity. Considering these limitations, practice guidelines for the diagnosis and management of bacterial meningitis do not recommend routine use of BAT.  Cryptococcus spp. Reagents for the detection of the polysaccharide capsular antigen of Cryptococcus spp. are available commercially. CSF specimens that yield positive results for cryptococcal antigen should be tested with a second latex agglutination test for rheumatoid factor. The commercial test systems incorporate rheumatoid factor testing in the protocol. A positive rheumatoid factor test renders the cryptococcal latex test uninterpretable, and the results should be reported as such, unless the rheumatoid factor antibodies have been inactivated. Undiluted specimens used in latex agglutination assays or enzyme immunoassays containing large amounts of capsular antigen may yield a false-negative reaction caused by a prozone phenomenon. Patients with AIDS may have an antigen titer more than 100,000, requiring many dilutions to reach an end-point. Serial dilution protocols are useful for monitoring a patient’s response to treatment, as well as for initial diagnosis.

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Lateral flow assays are now available (the IMMY cryptococcal LFA [Abacus ALS, Meadowbrook, Queensland, Australia]), and demonstrate superior sensitivity to latex agglutination assays. In addition, the turn-around time for results is shorter, and they can be used on urine samples and capillary blood samples.  Nucleic Acid Detection Traditional methods, including Gram stain and culture, can take several days to identify the etiological agent associated with CSF infections resulting in prolonged antimicrobial therapies and lengthened hospital stays. With the introduction of amplification technologies, such as PCR, many reports in the literature recommend the application of nucleic acid–based technologies for the diagnosis of CNS infections caused by various microorganisms. Published data indicate that nucleic acid–based assays demonstrate increased sensitivity and specificity compared with presently available techniques, particularly of CNS infections caused by herpes simplex virus and enteroviruses. PCR testing for HSV, EBV, CMV, and enterovirus in CNS infections has a sensitivity nearing 100%. The FilmArray meningitis/encephalitis panel (BioFire Diagnostics, Salt Lake City, UT) is a US Food and Drug Administration (FDA)-cleared, multiplex panel that detects 14 pathogens including 6 bacterial, 1 fungal, and 7 viral infectious agents. It is estimated that up to 50% of cases of encephalitis and 60% of cases of meningitis fail to identify an etiological agent. Delays in treatment may result in increased morbidity and mortality, and unnecessary treatment promotes the development of antibiotic resistance and increased health care costs. However, a retrospective study by Dack and colleagues indicates that the use of the BioFire FilmArray does not significantly reduce the length of stay or antimicrobial treatment associated with meningitis. This may be due to factors that include clinicians concerned with other health care–associated risks and continuing antimicrobial therapy in the presence of negative results. A significant advantage to using traditional culture methods in combination with nucleic acid–based testing is the identification of CSF infections that include more than one infectious pathogen. Although a low percentage, the FilmArray ME panel has been reported to identify several cases of coinfections when routine testing resulted in the identification of a single pathogen. Examples include bacterial and fungal pathogens such as H. influenzae and L. monocytogenes, Cryptococcus spp. and S. pneumoniae, N. meningitidis, and S. pneumoniae. Other coinfections included viral agents such as HSV-1 and HHV-6, CMV, and VZV, or combination bacterial viral infection with H. influenzae and HHV-6 or a combination fungal and viral infection with Cryptococcus spp. and HSV-1. Other advanced techniques are continually being developed, including an advanced fragment PCR based analysis capable of identifying 22 different pathogens associated with meningitis or encephalitis by F. Long et al. PCR analysis has also been successfully used to diagnose neurosyphilis. Additional multiplex laboratory developed

tests not only include bacterial, fungal, and viral agents, but others also include the detection of the free-living amoebic parasites Balamuthia mandrillaris and Acanthamoeba. The successful detection of parasites in CNS using these assays has not demonstrated sufficient clinical sensitivity and false positive results. The identification of parasitic pathogens in the CNS using nucleic acid testing requires extensive development and evaluation. Although nucleic acid–based testing does not appear to decrease length of stay or the use of antimicrobials associated with bacterial, fungal, and viral pathogens, the diagnostic sensitivity and specificity, along with the identification of pathogens that are not detected by traditional methods and the identification of rare coinfections, are significant and support the use of nucleic acid–based testing in the diagnosis and treatment of CSF infections. A variety of nucleic acid–based testing methods are now currently available. Next generation sequencing, although not widely available in routine clinical laboratories, has been used to identify pathogens in undiagnosed infectious encephalitis. Reports include the identification of astrovirus and neuroleptospirosis both in immunocompromised patients, HSV-1, HSV-2, and Brucella sp. in multiple cases of neurobrucellosis. The diagnosis of neurobrucellosis is significantly challenging as the signs and symptoms are extremely nonspecific and resemble other infectious diseases including tuberculosis, syphilis, Lyme disease, and Cryptococcus spp. infection. Traditional methods for the diagnosis of neurobrucellosis demonstrate a very low sensitivity; 28% demonstrate positive blood cultures and 15% show positive CSF cultures. These cases indicate a significant important step in the rapid and accurate diagnosis for patients of unexpected diagnoses in suspected cases of CNS infections using next generation sequencing. 

Matrix-Assisted Laser Desorption Ionization Time-of-Flight Mass Spectrometry Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) requires growth of the organism before application and spectral analysis. Studies are currently underway to determine the efficiency for the direct detection of microorganisms in CSF in cases of bacterial meningitis. A study by Bishop et  al., using fresh CSF samples, demonstrated a sensitivity of 76.2% for the identification of gram-negative bacilli postneurosurgical meningitis; however, for gram-positive cocci postsurgical and community-acquired meningitis, only a single sample was identified correctly. The interpretation of direct specimen Gram stains, followed by culture or nucleic acid–based testing, remain the gold standard for directing empirical antibiotic treatment of patients in suspected cases of meningitis, encephalitis, and other CNS infections. 

Culture Most cases of bacterial meningitis are caused by a single organism and require a limited number of culture media.

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Bacteria and Fungi

Routine bacteriologic media should include a chocolate agar plate, 5% sheep blood agar plate, and an enrichment broth, usually thioglycolate without indicator. The chocolate agar plate is needed to recover fastidious organisms, most notably H. influenzae, which are unable to grow on blood agar plates; the use of the blood agar plate aids in the recognition of S. pneumoniae. After vortexing the sediment and preparing smears, several drops of the sediment should be inoculated to each medium. Plates should be incubated at 37°C in 5% to 10% carbon dioxide (CO2) for at least 72 hours. If a CO2 incubator is not available, a candle jar or an automated environmental vacuum system can be used to create a CO2-enriched atmosphere. The broth should be incubated in air at 37°C for at least 5 to 10 days. The broth cap must be loose to allow free exchange of air. If organisms morphologically resembling anaerobic bacteria are seen on the Gram stain or if a brain abscess is suspected, an anaerobic blood agar plate may also be inoculated. These media will support the growth of almost all bacterial pathogens and several fungi. The symptoms of chronic meningitis that prompt a physician to request fungal cultures are the same as those for tuberculous meningitis. Cultures for mycobacteria are addressed in Chapter 42. For CSF fungal cultures, two drops of the well-mixed sediment should be inoculated onto Sabouraud dextrose agar or other non–blood-containing medium and brain-heart infusion with 5% sheep blood. Fungal media should be incubated in room air at 30°C for 4 weeks. If possible, two sets of media should be inoculated, with one set incubated at 30°C and the other at 35°C.  Parasites and Viruses

Conditions for the culture of free-living amoebae and viral agents are discussed in Chapters 46 and 64, respectively. The physician must notify the laboratory to culture these agents.  Brain Abscess/Biopsies Specimen Collection, Transport, and Processing. Whenever possible, biopsy specimens or aspirates from brain abscesses should be submitted to the laboratory under anaerobic conditions. Several devices are commercially available to transport biopsy specimens under anaerobic conditions. Swabs are not considered an optimum specimen, but if used to collect abscess material they should be sent in a transport device that maintains an anaerobic environment.

Biopsy specimens should be minced in sterile saline before plating and smear preparation. This processing should be kept to a minimum to reduce oxygenation. Abscess and biopsy specimens submitted for culture should be inoculated onto 5% sheep blood and chocolate agar plates. Plates should be incubated in 5% to 10% CO2 for 72 hours at 35°C. In addition, an anaerobic agar plate and broth with an anaerobic indicator, vitamin K, and hemin should be inoculated and incubated in an anaerobic environment at 35°C. Anaerobic culture plates are incubated for a minimum of 72 hours but are examined after 48 hours of incubation. Anaerobic broths should be incubated

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for a minimum of 5 days. If a fungal cause is suspected, fungal media, such as brain-heart infusion with blood and antibiotics or inhibitory mold agar, should be inoculated.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Albright RE, Christenson RH, Emlet JL, et al.: Issues in cerebrospinal fluid management: acid-fast bacillus smear and culture, Am J Clin Pathol 95:418–423, 1991. Albright RE, Graham CB, Christenson RH, et al.: Issues in cerebrospinal fluid management: CSF venereal disease research laboratory testing, Am J Clin Pathol 95:397–401, 1991. Al Masalma MA, Lonjon M, Richet H, et al.: Metagenomic analysis of brain abscesses identifies specific bacterial associations, Clin Infect Dis 54:202–210, 2011. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, PA, 2020, Elsevier. Bishop B, Geffen Y, Plaut A, et al.: The use of matrix-assisted laser desorption/ionization time-of-flight mass spectrometry for rapid bacterial identification in patients with smear-positive bacterial meningitis, Clin Microbiol Infect 24(2):171–174, 2018. Carroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Castro R, Aguas MJ, Batista T, et  al.: Detection of Treponema pallidum sp. pallidum DNA in cerebral spinal fluid by two PCR techniques, J Clin Lab Anal 30(5):628–632, 2016. Centers for Disease Control and Prevention: Epidemic/epizootic West Nile virus in the United States: guidelines for surveillance, prevention and control–3rd revision, 2003, Available at www.cdc.gov/ncidod/ dvbid/westnile/resources/wnvguidelines2003.pdf. Conen A, Walti LN, Merlo A, et  al.: Characteristics and treatment outcome of cerebrospinal fluid shunt-associated infections in adults: a retrospective analysis over an 11-year period, Clin Infect Dis 47:73–82, 2008. Culbreath K, Melanson S, Gale J, et  al.: Validation and retrospective clinical evaluation of quantitative 16S rRNA gene metagenomics sequencing assay for bacterial pathogen detection in body fluids, J Mol Diagn 21:913–923, 2019, https://doi.org/10.1016/j. jmoldx.2019.05.002. Dack K, Pankow S, Abla E: Contribution of the BioFire FilmArray meningitis/encephalitis panel. Assessing antimicrobial duration and length of stay, Kans J Med 12(1):1–3, 2019. Fan S, Ren H, Wei Y, et al.: Next-generation sequencing of the cerebral spinal fluid in the diagnosis of neurobrucellosis, Int J Infect Dis 67:20–24, 2018. Garcia LS: Diagnostic medical parasitology, ed 6, Washington DC, 2016, ASM Press. Guan H, Shen A, Lv X, et al.: Detection of virus in CSF from the cases with meningoencephalitis by next-generation sequencing, J Neurovirol 22(2):240–245, 2016. Hariharan S: BK virus nephritis after renal transplantation: a review, Kidney Int 69:655–662, 2006. Hayward RA, Shapiro MF, Oye RK: Laboratory testing on cerebrospinal fluid: a reappraisal, Lancet 1:1–4, 1987. Huang C, Morse D, Slater B, et al.: Multiple-year experience in the diagnosis of viral central nervous system infections with a panel of polymerase chain reaction assays for detection of 11 viruses, Clin Infect Dis 39:630–635, 2004.

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Korimbocus J, Scaramozzino N, Lacroix B, et al.: DNA probe array for the simultaneous identification of herpesviruses, enteroviruses, and flaviviruses, J Clin Microbiol 43:3779–3787, 2005. Liesman RM, Strasburg AP, Heitman AK, et al.: Evaluation of a commercial multiplex molecular panel for diagnosis of infectious meningitis and encephalitis, J Clin Microbiol 56(4):e01927–17, 2018. Lindsey NP, Lehman JA, Staples E, et al.: West Nile virus and other arboviral diseases-United States, 2013, Morb Mortal Wkly Rep 63(24):521–526, 2014. Long F, Kong M, Wu S, et  al.: Development and validation of an advanced fragment-analysis based assay for the detection of 22 pathogens in the cerebrospinal fluid of patients with meningitis and encephalitis, J Clin Lab Anal 33(3):e22707, 2019. Mongkolrattanothai K, Naccache SN, Bender JM, et  al.: Neurobrucellosis: unexpected answer from metagenomic nextgeneration sequencing, J Pediatric Infect Dis Soc 6(4):393–398, 2017. Onyango CO, Loparev V, Lidechi S, et al.: Evaluation of a Taqman array card for detection of central nervous system infections, J Clin Microbiol 55(7):2035–2044, 2017. Parkkinen J, Korhonen TK, Pere A, et  al.: Binding sites in the rat brain for Escherichia coli S fimbriae associated with neonatal meningitis, J Clin Invest 81:860–865, 1988. Piquet AL, Lyons JL: Infectious meningitis and encephalitis, Semin Neurol 36(4):367–372, 2016. Plaut AG: The IgA1 proteases of pathogenic bacteria, Annu Rev Microbiol 37:603–622, 1983. Poppert S, Essig A, Stoehr B, et al.: Rapid diagnosis of bacterial meningitis by real-time PCR and fluorescence in situ hybridization, J Clin Microbiol 43:3390–3397, 2005. Schwartz MN: Bacterial meningitis—a view of the past 90 years, N Engl J Med 351:1826–1828, 2004.

Segawa S, Sawai S, Murata S, et al.: Direct application of MALDITOF mass spectrometry to cerebralspinal fluid for rapid pathogen identification in a patient with bacterial meningitis, Clin Chim Acta 435:59–61, 2014. Smith LP, Hunter Jr KW, Hemming VG, et al.: Improved detection of bacterial antigens by latex agglutination after rapid extraction from body fluids, J Clin Microbiol 20:981–984, 1984. Strasinger S, DiLorenzo M: Urinalysis and body fluids, ed 5, Philadelphia, PA, 2008, FA Davis. Tarai B, Das P: FilmArray meningitis/encephalitis (ME) panel, a rapid molecular platform for diagnosis of CNS infections in a tertiary care hospital in North India: one-and-half-year review, Neurol Sci 40(1):81–88, 2019. Tunkel AR, Hartman BJ, Kaplan SL, et  al.: Practice guidelines for the management of bacterial meningitis, Clin Infect Dis 39:1267– 1284, 2004. van de Beek D, de Gan J, Spanjaard L, et al.: Clinical features and prognostic factors in adults with bacterial meningitis, N Engl J Med 351:1849–1859, 2004. Virji M, Alexandrescu C, Ferguson DJ, et al.: Variations in the expression of pili: the effect on adherence of Neisseria meningitidis to human epithelial and endothelial cells, Mol Microbiol 6:1271– 1279, 1992. Virji M, Kayhty H, Ferguson DJ, et al.: The role of pili in the interactions of pathogenic Neisseria with cultured human endothelial cells, Mol Microbiol 5:1831–1841, 1991. Wilhelm C, Ellner JJ: Chronic meningitis, Neurol Clin 4:115–141, 1986. Walsh TJ, Hayden RT, Larone DH: Larone’s medically important fungi: a guide to identification, ed 6, Washington, DC, 2018, ASM Press.

PROCEDURE 70.1

Rapid Extraction of Antigen Procedure Principle Removal of nonspecific cross-reactive material can improve the specificity of direct antigen detection particle agglutination tests. Ethylenediaminetetraacetic acid (EDTA) forms complexes with cross-reactive materials, and they are removed from the reaction mixture by centrifugation. 

Method 1. Pipette 0.05 mL fluid to be tested (cerebrospinal fluid [CSF], serum, or urine) into a 1.5-mL plastic, conical microcentrifuge tube. 2. Add 0.15 mL of 0.1 M EDTA (Sigma Chemical Co.) to the microcentrifuge tube, close the cap tightly, and vortex the tube.

3. Heat in a dry bath (available from instrument supply companies) for 3 minutes at 100°C. 4. Centrifuge the tubes for 5 minutes at 13,000×g in a tabletop microcentrifuge. Be certain that the instrument achieves the required centrifugal force. 5. Remove the supernatant with a capillary pipette and use 1 drop of this solution as the test sample in the antigen detection test, following the manufacturer’s instructions for performance of the test. 

Expected Results Nonspecific agglutination should not occur.

  

CASE STUDY 70.1 A 2-year-old girl presented at midnight to the hospital emergency department with a temperature of 104°F. She was diagnosed with bilateral otitis. She was treated with amoxicillin/ clavulanic acid and retained for observation in the hospital. During the night, the child became lethargic. She developed purpura and nuchal rigidity. A CSF sample was collected, and ceftriaxone therapy was begun. No organisms were noted on Gram stain. The next day, the laboratory reported growth of a gram-negative diplococcus.

Questions 1. What is the suspected organism in this infection, and how can the laboratory rapidly identify it? 2. Is it recommended that laboratories do susceptibility testing for N. meningitidis? 3. How can the laboratory improve the speed with which it detects this organism in CSF? 4. What measures are taken to prevent the spread of infection among health care workers who are exposed to patients with N. meningitidis?   

Chapter Review 1. Which of the following are specialized structures of the meninges that function to absorb the spinal fluid and allow it to pass into the blood? a. Leptomeninges b. Pia mater c. Arachnoid villi d. Dura mater 2. What bacteria are responsible for outbreaks of meningitis among neonates in hospital nurseries? a.  Elizabethkingia meningoseptica b. Haemophilus influenzae type b c. Group B streptococci d. Neisseria meningitides 3. All the following are pathogenic sources capable of causing brain abscesses except: a. Anaerobes b. Borrelia burgdorferi c.  Viridans streptococci d. Fungal organisms

4. When processing CSF specimens for laboratory diagnosis, the specimen appears red in some of the tubes, a sign of red blood cells and bleeding; to determine whether the blood is caused by a bloody tap or a subarachnoid hemorrhage, cell counts are done on which of the following tubes? a. Tubes 1 and 2 b. Tubes 3 and 4 c. Tubes 2 and 3 d. Tubes 1 and 4 5. Refer to the previous question. Which tube is used for chemistry and immunologic studies? a. Tube 1 b. Tube 2 c. Tube 3 d. Tube 4

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6. Culture to determine the etiologic agent causing meningitis is set up from which CSF tube? a. Tube 1 b. Tube 2 c. Tube 3 d. Tube 4 7. Which of the following organisms is a parasite that grows intracellularly, destroys brain parenchyma, and is a common CNS affliction in HIV-infected patients with AIDS? a.  Naegleria fowler b. Entamoeba histolytica c.  Taenia solium d. Toxoplasmosis 8. Which of the following causes of pediatric meningitis was significantly reduced as the result of an effective vaccination program? a. Group B Streptococcus b. Escherichia coli c.  Haemophilus influenzae type b d. Listeria monocytogenes 9. When a physician suspects Cryptococcus spp. as the etiologic agent of a CNS infection, what is the best manual way to test for it? a. India ink stain b. Cryptococcal antigen test c. Culture d. Molecular testing by PCR 10. The cerebrospinal fluid that surrounds the brain and spinal fluid functions to: a. Carry essential metabolites into the neural tissue b. Protect the central nervous system against microbial invasion by phagocytosis c.  Provide a means by which the brain monitors changes in the internal environment d. Both A and C 11.  True or False _____ In cushioning and providing buoyancy for the bulk of the brain, the effective weight of the brain is reduced by a factor of 30 by the CSF. _____  The three layers of protective membranes, which surround the brain and spinal column, are called the meninges. _____ The elderly population of patients has the highest prevalence of meningitis. _____ Encephalitis is an inflammation of the brain parenchyma and is normally caused by bacteria. _____ The first Hib vaccine was not efficacious in children younger than 18 months of age. _____ The normal CSF glucose serum ratio is 0.6, or 50% to 70% of the blood glucose normal value. _____ A component of syphilis infections is aseptic meningitis.

_____ CSF cultures from patients with brain abscesses are typically positive for anaerobes or viridians streptococci. _____ CSF is found in the subdural spaces of the brain. _____ The entire volume of CSF is exchanged every 5 to 6 hours. _____ When collecting CSF for culture studies, it is imperative to collect the correct volume of CSF. _____ When transporting CSF to the laboratory for bacterial studies, the CSF must be refrigerated and kept at a temperature of 2°C to 8°C. _____ If a physician orders viral studies on a CSF and the transport to the laboratory will be longer than 2 to 3 hours after collection, the CSF specimen must be frozen at −20°C. _____ The most sensitive method for detecting encephalitis-causing viruses in the CSF is PCR. 12.  Matching: Match each term with the correct description. _____ leptomeninges _____ dura mater _____ arachnoid membrane _____ pia mater _____ ventriculitis _____ aseptic meningitis _____ encephalitis _____ meningo-­ encephalitis _____ phycomycosis _____ spinal tap _____ cysticercus _____ choroid plexus _____ blood-brain barrier _____ meninges _____ meningitis

a. concomitant meningitis with encephalitis b. membrane covering the brain and spinal cord c. secretory cells that produce CSF d. rapidly progressive fungal infection e. self-limiting viralcaused meningitis f. larval form of Taenia solium g. defense mechanism for the CNS h. pia mater and the arachnoid membrane i. outermost membrane of the meninges j. between the dura mater and pia mater k. adheres to the outer surface of the brain and spinal cord l. purulent meningitis involving the ventricles m. infection within the subarachnoid space of the meninges n. inflammation of the brain parenchyma o. lumbar puncture

CHAPTER 70  Meningitis and Other Infections of the Central Nervous System

13.  Short Answer (1) Name the least common route of CNS infection caused by an organism. (2) Name a unique property of the HIV virus that predisposes infected individuals to viral encephalitis. (3) What is a cytospin centrifuge, and how does it assist in processing CSF specimens? (4) What is diagnostic in the smear when using the India ink stain to detect the presence of Cryptococcus spp.?

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(5) What is the purpose of the pretreatment REAP procedure? What does REAP stand for? (6) What population of patients has the highest prevalence of meningitis? Describe the associated predisposing factors. (7) What is one of the most critical steps in processing CSF for culture? (8) Explain the reasoning behind using tubes 3 and 4 in CSF collection for cell count and differential.

71

Infections of the Eyes, Ears, and Sinuses OBJECTIVES 1. Describe the anatomy of the eye, including naming the external and internal structures. 2. Name the three tissues, outer to inner, of the eyeball. 3. Differentiate normal flora of the eye and potential pathogens. 4. Describe the defense mechanisms of the eye for protection from infective agents. 5. Define the following diseases of the eye: blepharitis, hordeolum, conjunctivitis, keratitis, uveitis, and endophthalmitis. 6. List the common types of eye infections, the associated etiologic agents, and the at-risk patient population for each. 7. Define keratitis, and identify the organisms associated with the infection, the virulence factors, and the antimicrobialresistant properties for each. 8. Define endophthalmitis, explain how it is contracted, and identify the etiologic agents. 9. Explain mycotic endophthalmitis and list the risk factors that may predispose an individual to this type of infection. 10. Define a periocular infection, and list some of the associated infectious agents and the different types of clinical presentations of the infection. 11. Identify the anatomic parts of the ear, and list the structures associated with each region within the ear. 12. Define the external ear infections acute externa otitis and chronic externa otitis; list the potential pathogens. 13. Define otitis media; differentiate acute and chronic otitis media and name the most commonly encountered pathogens and the age group most often affected by this disease. 14. Explain the laboratory method used to culture the eye and the ear, including appropriate media; describe collection and transportation requirements. 15. Differentiate acute and chronic sinusitis. 16. Explain why the organisms that cause otitis media are often the same ones responsible for sinusitis. 17. List the collection methods and culture media used for cases of sinusitis. 18. Correlate signs and symptoms of infection with the results of laboratory diagnostic procedures for the identification of the etiologic agent associated with infections of the eye, ear, and sinuses.

Eyes Anatomy Eye (ocular) infections can be divided based on the area of the eye infected. The external structures of the eye—eyelids, conjunctiva, sclera, and cornea—are depicted in Fig. 71.1. The eyeball comprises three layers. From the outside in, these tissues are the sclera, choroid, and retina. The sclera is a tough, white, fibrous tissue (i.e., “white” of the eye). The anterior (toward the front) portion of the sclera is the cornea, which is transparent and has no blood vessels. A mucous membrane, called the conjunctiva, lines each eyelid and extends onto the surface of the eye itself. The choroid is the vascular layer of the eye that contains the connective tissue. The retina, the innermost layer of the eye, contains light-sensitive cells that transmit signals and images to the optical nerve. Only a small portion of the eye is exposed to the environment; about five-sixths of the eyeball is enclosed within bony orbits shaped like four-sided pyramids. The large interior space of the eyeball is divided into two sections: the anterior and posterior cavities (Fig. 71.1). The anterior cavity is filled with a clear and watery substance called aqueous humor; the posterior cavity is filled with a soft, gelatin-like substance called vitreous humor. Infections can occur in the eye’s lacrimal (pertaining to tears) system. The major components of the lacrimal apparatus include the lacrimal gland, lacrimal canaliculi (short channel), and lacrimal sac. 

Resident Microbiota Rather sparse indigenous microbiota is present in the conjunctival sac. Staphylococcus epidermidis and Lactobacillus spp. are the most common organisms; Cutibacterium acnes may also be present. Staphylococcus aureus is found in less than 30% of individuals, and Haemophilus influenzae colonizes 0.4% to 25%. Moraxella catarrhalis, various Enterobacterales, and various streptococci (Streptococcus pyogenes, Streptococcus pneumoniae, other alpha-hemolytic and

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Eyelid Eyelashes

Retina

Choroid Lens Pupil

Optic disc

Cornea Iris muscle

Anterior cavity

Optic nerve

Anterior chamber Posterior chamber

Conjunctiva

Sclera

• Fig. 71.1  Key anatomic structures of the eye. (Modified from Thibodeau GA, Patton KT. Anatomy and Physiology. 2nd ed. St Louis: Mosby; 1993.)

gamma-hemolytic forms) are found in a very small percentage of individuals. 

Diseases The eye and its associated structures are uniquely predisposed to infection by various microorganisms. The major infections of the eye are listed in Table 71.1, along with a brief description of the disease. 

Pathogenesis The eye has several defense mechanisms. The eyelashes prevent the entry of foreign material into the eye. The lids blink 15 to 20 times per minute, during which time secretions of the lacrimal glands and goblet cells wash away bacteria and foreign matter. Lysozyme and immunoglobulin A (IgA) are secreted locally and serve as part of the eye’s natural defense mechanisms. Also, the eyes themselves are enclosed within the bony orbits. The delicate intraocular structures are enveloped in a tough collagenous coat (sclera and cornea). If these barriers are broken by a penetrating injury or ulceration, infection may occur. Infection can also reach the eye via the bloodstream from another site of infection. Finally, because three of the four walls of the orbit are contiguous with the paranasal (facial) sinuses, sinus infections may extend directly to the periocular orbital structures. 

Epidemiology and Etiology of Disease Blepharitis and Hordeolum Blepharitis may appear as a bump on the eyelid that is red and swollen, resembling a pimple. Most bumps on the

eyelid are caused by an inflamed oil gland on the edge of the eyelid and are a form of hordeolum—more commonly referred to as a stye. A stye is more of an acute infection, whereas blepharitis tends to present as a chronic condition that can cause conjunctivitis, functional tear deficiency, or corneal inflammation and infection. Bacteria, viruses, and occasionally lice or Demodex mites can cause blepharitis, an infection of the eyelid surrounding the eye. Although occasionally isolated from surfaces surrounding the healthy eye, S. aureus and S. epidermidis are the most common infectious agents associated with the development of a stye and blepharitis in developed countries. Symptoms include burning, itching, the sensation of the presence of a foreign body, and crusting of the eyelids. Viruses can also cause a vesicular (blisterlike) eruption of the eyelids. Herpes simplex virus (HSV) produces vesicles on the eyelids that typically crust and heal with scarring within 2 weeks. Unfortunately, once this vesicular stage has resolved, the lesions can be confused with bacterial blepharitis. Finally, the pubic louse Phthirus pubis has a predilection for eyelash hair. The presence of this organism produces irritation, itch, and swelling of the lid margins (edges). 

Conjunctivitis Bacterial conjunctivitis, commonly referred to as “pink eye,” is the most common type of ocular infection and may be caused by allergies, bacterial, parasitic, fungal, or viral infection. The principal causes of acute conjunctivitis in a normal host are listed in Table 71.1. Age-related factors are key in the identification of the etiologic agent. Neonatal conjunctivitis (ophthalmia neonatorum) occurs within

CHAPTER 71  Infections of the Eyes, Ears, and Sinuses

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TABLE 71.1    Infections of the Eye

Infection

Description

Bacteria

Viruses

Fungi

Parasites

Blepharitis

Inflammation of the margins (edges) of the eyelids (eyelids, eyelashes, or associated pilosebaceous glands or meibomian glands); symptoms include irritation, redness, burning sensation, and occasional itching. Condition is typically bilateral.

Staphylococcus aureus, Pseudomonas aeruginosa, Capnocytophaga ochracea

Herpes simplex virus (HSV)

Malassezia furfur, Blastomyces sp.

Phthirus pubis, Leishmania donovani, Demodex mites

Conjunctivitis

Inflammation of the conjunctiva; symptoms vary according to the etiologic agent, but most patients have swelling of the conjunctiva, inflammatory exudates, and burning and itching.

Streptococcus pneumoniae, Haemophilus influenzae, S. aureus, Haemophilus spp., Chlamydia trachomatis, Neisseria gonorrhoeae, Streptococcus pyogenes, Moraxella spp., Corynebacterium spp., P. aeruginosa

Adenoviruses, HSV, varicella zoster, Epstein-Barr virus (EBV), influenza virus, paramyxovirus, rubella, human immunodeficiency virus (HIV) enterovirus, coxsackie A, variola (smallpox), SARS-CoV-2

Candida spp., Blastomyces spp., Sporothrix schenckii, Rhinosporidium seeberi

Leishmania spp., Microsporidia spp., Loa loa, Demodex (mites)

Keratitis

Inflammation of the cornea; although there are no specific clinical signs to confirm infection, most patients complain of pain and usually some decrease in vision, with or without discharge from the eye.

S. aureus, S. epidermidis, S. pneumoniae, S. pyogenes, viridans streptococci, Enterococcus faecalis, Peptostreptococcus, P. aeruginosa, Enterobacterales, Moraxella lacunata, Bacillus spp., Mycobacterium spp., spirochetes, C. trachomatis

HSV, adenoviruses, varicella zoster, vaccinia, Epstein-Barr, rubeola, enteroviruses, and coxsackie virus

Fusarium, Aspergillus spp., Candida spp., Acremonium, Alternaria, Penicillium, Bipolaris, Nosema, Vittaforma, Encephalitozoon spp.

Acanthamoeba spp., Onchocerca volvulus, Leishmania brasiliensis, Trypanosoma spp.

Keratoconjunctivitis

Infection involving both the conjunctiva and cornea; ophthalmia neonatorum is an acute conjunctivitis or keratoconjunctivitis of the newborn caused by either N. gonorrhoeae or C. trachomatis.

Refer to agents for keratitis/conjunctivitis

Refer to agents for keratitis/ conjunctivitis

Refer to agents for keratitis

Toxoplasma gondii, Toxocara

Chorioretinitis and uveitis

Inflammation of the retina and underlying choroid or the uvea; infection can result in loss of vision.

Mycobacterium tuberculosis, Treponema pallidum, Borrelia burgdorferi

Cytomegalovirus, HSV

Candida spp.

T. gondii, Toxocara, Treponema pallidum, Brucella spp.

Continued

1008 PA RT V I I     Diagnosis by Organ System

TABLE 71.1    Infections of the Eye—cont’d

Infection

Description

Bacteria

Viruses

Fungi

Parasites

Endophthalmitis

Infection of the aqueous or vitreous humor. This infection is usually caused by bacteria or fungi, is rare, develops suddenly, and progresses rapidly, often leading to blindness. Pain, especially while moving the eye, and decreased vision are prominent features.

S. aureus, S. epidermidis, S. pneumoniae, other streptococcal species, P. aeruginosa, Klebsiella pneumoniae, other gram-negative organisms, Nocardia spp.

HSV, varicella zoster

Candida spp., Aspergillus spp., Fusarium spp.

Toxocara, Onchocerca volvulus

Lacrimal infections, canaliculitis

A rare, chronic inflammation of the lacrimal canals in which the eyelid swells and there is a thick, mucopurulent discharge.

Actinomyces, Propionibacterium propionicum

Dacryocystis

Inflammation of the lacrimal sac that is accompanied by pain, swelling, and tenderness of the soft tissue in the medial canthal region.

S. pneumoniae, S. aureus, S. pyogenes, H. influenzae

Dacryoadenitis

Acute infection of the lacrimal gland; these infections are rare and can be accompanied by pain, redness, and swelling of the upper eyelid and conjunctival discharge.

S. pneumoniae, S. aureus, S. pyogenes

C. albicans, Aspergillus spp.

Note: This table is not intended to be all-inclusive for the infectious agents capable of causing eye infections.

4 weeks following birth, caused by bacterial, viral, chlamydial, or toxic reactions to chemicals. In neonates, neisserial and chlamydial infections are common and are acquired during passage through an infected vaginal canal. With the common practice of instilling antibiotic drops into the eyes of newborns in the United States, the incidence of gonococcal and chlamydial conjunctivitis has dropped dramatically. However, Chlamydia trachomatis remains responsible for one of the most important types of conjunctivitis, referred to as trachoma. Trachoma is one of the leading causes of blindness in the world, primarily in underdeveloped countries. In children, the most common causes of bacterial conjunctivitis include H. influenzae, S. pneumoniae, and perhaps S. aureus, S. pneumoniae, and Haemophilus aegyptius

have been isolated from conjunctivitis epidemics. Corynebacterium spp. colonize the lids and conjunctiva and are the overall leading cause of conjunctivitis. Inflammation of the conjunctiva is characterized by redness, itching, and discharge, and the condition is highly contagious; it can be transferred from one eye to the other by rubbing the infected eye and can be easily transferred to other individuals. Numerous other bacteria may also cause conjunctivitis. For example, diphtheritic conjunctivitis may occur in conjunction with diphtheria elsewhere in the body. Moraxella lacunata produces a localized conjunctivitis with little discharge from the eye. Distinctive clinical pictures may also occur with conjunctivitis caused by Mycobacterium tuberculosis, Francisella tularensis, Treponema pallidum, and Yersinia enterocolitica.

CHAPTER 71  Infections of the Eyes, Ears, and Sinuses

Fungi may be responsible for this type of infection as well, often in association with a foreign body that has been introduced into the eye or an underlying host immunologic problem. Fungi including Candida spp., Blastomyces spp., and Sporothrix schenckii have been associated with conjunctivitis. However, these infections are uncommon. Parasitic conjunctivitis has been associated with Leishmania spp., cryptosporidium, fly larvae, and nematodes such as Loa loa. Parasites that are known to infect the lid margin or lashes, such as lice and mites, may cause conjunctivitis as a subsequent reaction to blepharitis caused by the organism. In adults, the cause of conjunctivitis is usually viral, with adenovirus being the most common viral cause; 20% of such infections in children resulted from adenoviruses in one large US study, and 14% of infections in adult patients were caused by adenoviruses in another study. Adenovirus types 3, 4, and 7A are common. Most viral conjunctivitis is self-limited but is highly contagious, with the potential to cause major outbreaks. Worldwide, enterovirus 70 and coxsackievirus A24 are responsible for outbreaks and epidemics of acute hemorrhagic conjunctivitis. A coxsackievirus A24 variant has been reported with several outbreaks of hemorrhagic conjunctivitis in several countries. Patients can develop systemic symptoms including fever, fatigue, and limb pain; however, severe complications and death are rare. A lateral flow immunochromatographic cartridge test, the AdenoPlus (Rapid Pathogen Screening, Inc., Sarasota, FL), is available for the detection of ocular adenovirus infections. The test includes a built-in sampling pad that can be used to collect fluid by touching the eye. The assay demonstrates an 85% sensitivity and a 98% specificity, when compared with polymerase chain reaction (PCR). Negative results using the AdenoPlus should be confirmed by real-time PCR to avoid false negatives that would result in continued infection and damage to the eye. 

Keratitis Keratitis (corneal infection) may be caused by a variety of infectious agents, usually after some type of trauma to the ocular surface. Keratitis should be regarded as an emergency, because corneal perforation and loss of the eye can occur within 24 hours when organisms such as Pseudomonas aeruginosa, S. aureus, or HSV are involved. Bacteria account for 65% to 90% of corneal infections. In the United States, S. aureus, S. pneumoniae, and P. aeruginosa account for more than 80% of all bacterial corneal ulcers. Many culture-positive cases are now being recognized as polymicrobial. A toxic factor known as exopeptidase has been implicated in the pathogenesis of corneal ulcer produced by S. pneumoniae. With P. aeruginosa and Neisseria gonorrhoeae, proteolytic enzymes are responsible for the corneal destruction. Gonococcus may cause keratitis during inadequately treated conjunctivitis. Acinetobacter, which may look identical microscopically to gonococcus and is resistant to penicillin and many other antimicrobial agents, can cause corneal perforation. Many other bacteria, several viruses other than HSV, and many fungi may cause

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• Fig. 71.2  Endophthalmitis. (Courtesy Donald J. D’Amico.)

keratitis. Risk factors associated with the development of fungal keratitis caused by Candida spp. include epithelial ulceration, topical corticosteroid use, corneal transplants, and the use of soft contact lenses. Fungal keratitis may also result as a complication of trauma. Although still unusual, a previously rare etiologic agent of corneal infections has become more common in users of soft and extended-wear contact lenses. Acanthamoeba spp., free-living amoebae, can survive in improperly sterilized cleaning fluids and be introduced into the eye with the contact lens. The fungus Fusarium is an infectious disease associated with contact lens use or contact lens solutions. This genus of fungus is ubiquitous and can be found in soil and tap water and on many plants; fungal keratitis is rare but is usually associated with trauma to the eye from an object contaminated with plant matter. This infection can be serious and can lead to the loss of vision. Other bacterial and fungal causes of infection have also been traced to inadequate cleaning of lenses. Additional parasites are also associated with keratitis in different geographical regions, including the microfilariae Onchocerca volvulus, Leishmania spp., microsporidia, and trypanosomes. It is important to consider potential coinfections with other organisms, as that will decrease the effectiveness of the treatment and may result in the loss of vision. 

Endophthalmitis Surgical trauma, nonsurgical trauma (uncommonly), and hematogenous spread from distant sites of infection are the typical routes of transmission for endophthalmitis (Fig. 71.2). The infection may be limited to specific tissues within the eye or may involve all the intraocular contents. Bacteria are the most common infectious agents responsible for endophthalmitis. After surgery or trauma, evidence of the disease is usually identified within 24 to 48 hours. Postoperative infection involves primarily normal microbiota from the ocular surface. Although S. epidermidis and S. aureus are responsible for most cases of endophthalmitis after cataract removal, any bacterium, including those considered to be saprophytic,

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A

B

C

D •

Fig. 71.3  Acute retinal necrosis caused by herpes simplex. (A) External view. (B) Limited funduscopic view. (C) Funduscopic view of the normal unaffected eye. (D) View of peripheral retina demonstrating peripheral necrotizing retinitis and vasculitis (whitening and hemorrhage).

may cause endophthalmitis. In hematogenous endophthalmitis, a septic focus elsewhere is usually evident before the onset of the intraocular infection. Bacillus cereus has caused endophthalmitis in people addicted to narcotics and after transfusion with contaminated blood. Endophthalmitis associated with meningitis may involve various organisms, including H. influenzae, streptococci, and Neisseria meningitidis. Nocardia endophthalmitis may follow pulmonary infection with this organism. Mycotic infection of the eye has increased significantly since the 1980s because of the increased use of antibiotics, corticosteroids, antineoplastic chemotherapy, addictive drugs, and hyperalimentation (overeating). Fungi generally considered to be saprophytic are important causes of postoperative endophthalmitis (Table 71.1). Endogenous mycotic endophthalmitis is most often caused by Candida albicans. High-risk patients include those with diabetes or some other chronic underlying disease. Exogenous Candida spp. endophthalmitis is uncommon but may develop following surgical procedures, trauma, or keratitis. Aspergillus or Fusarium species may also be associated with traumarelated infections. Other causes of hematogenous ocular

infection include Aspergillus, Cryptococcus, Coccidioides, Sporothrix, and Blastomyces. Endophthalmitis may be a result of viral or parasitic infections. Viral causes of endophthalmitis include HSV (Fig. 71.3), varicella (herpes) zoster virus (VZV), cytomegalovirus, and measles viruses. The most common parasitic agent associated with endophthalmitis is Toxocara. Toxoplasma gondii is a well-known cause of chorioretinitis. Thirteen percent of patients with cysticercosis (Taenia solium) have ocular involvement. Onchocerca usually produces keratitis, but intraocular infection also occurs. 

Periocular Canaliculitis, one of three infections of the lacrimal apparatus (Table 71.1), is an inflammation of the lacrimal canal and is usually caused by Actinomyces or Propionibacterium propionicum. Infection of the lacrimal sac (dacryocystitis) may involve numerous bacterial and fungal agents; the major causes are listed in Table 71.1. Dacryoadenitis is an uncommon infection of the lacrimal gland characterized by pain of the upper eyelid with erythema and often involves pyogenic bacteria such as S. aureus and streptococci.

CHAPTER 71  Infections of the Eyes, Ears, and Sinuses

Chronic infections of the lacrimal gland occur in tuberculosis, syphilis, leprosy, and schistosomiasis. Acute inflammation of the gland may occur during mumps and infectious mononucleosis. Orbital cellulitis is an acute infection of the orbital contents and is most often caused by bacteria. This is a potentially serious infection because it may spread posteriorly to produce central nervous system complications. Most cases involve spread from contiguous sources such as the paranasal sinuses. In children, bloodborne bacteria, notably H. influenzae, may lead to orbital cellulitis. S. aureus is the most common etiologic agent; S. pyogenes and S. pneumoniae are also common. Anaerobes may cause cellulitis secondary to chronic sinusitis, primarily in adults. Mucormycosis of the orbit is a serious, invasive fungal infection seen particularly in patients with diabetes who have poor control of their disease, patients with acidosis from other causes, and patients with malignant disease receiving cytotoxic and immunosuppressive therapy. Aspergillus may produce a similar infection in the same settings but also can cause mild, chronic infections of the orbit. Surgical techniques involving the ocular implantation of prosthetic or donor lenses have resulted in increasing numbers of iatrogenic (resulting from the activities of a physician) infections. Isolation of C. acnes may have clinical significance in such situations, in contrast to many other sites in which it is usually considered to be a contaminant. Nontuberculous mycobacterial periocular infections have become increasingly important in patients with systemic disease. These infections are more prevalent in immunocompromised patients. 

Uveitis The uvea is the pigmented, middle layer of the eye that is between the cornea-sclera and the retina. Inflammation of the uvea is termed uveitis. Retinitis, inflammation of the retina, is also considered in this section, even though it is technically a separate structure from the uvea. There are approximately 70 to 115 cases per 100,000 individuals who are annually affected by uveitis in the United States. More than 50% of cases of uveitis are idiopathic; however, the condition may be caused by an autoimmune reaction, infection, or trauma. Infectious uveitis typically is a result of hematogenous spread of the agent of infection. The eye has a blood-eye barrier similar to the blood-brain barrier that must be breached for an infection to occur. Inflammation causes this barrier to break down, resulting in infection. The most common causes for uveitis include herpes viruses (HSV, VZV, and cytomegalovirus) and Toxoplasma spp. 

Other Infections Opportunistic infections in human immunodeficiency virus (HIV)–infected individuals can involve the eye. Systemic infections that involve the eye include cytomegalovirus, Pneumocystis jiroveci, Cryptococcus neoformans, Mycobacterium avium complex, and Candida spp. Most often the

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retina, choroid, and optic nerve are infected with these agents, resulting in significant visual morbidity (unhealthy condition) if left untreated. However, because of the widespread use of highly active antiretroviral therapy capable of assisting in immune system recovery and lowering the viral load in patients with HIV infection, the incidence of acquired immune deficiency syndrome (AIDS) and related ophthalmic infections has declined sharply. 

Laboratory Diagnosis Specimen Collection and Transport Cell cultures for the isolation of C. trachomatis have been replaced by nucleic acid–based testing methods. Cell culture is only performed in specialized laboratories for antimicrobial susceptibility. Purulent material from the surface of the lower conjunctival sac and inner canthus (angle) of the eye is collected on a sterile swab for cultures. Both eyes should be cultured separately. Chlamydial cultures are taken with a dry calcium alginate swab and placed in a 2-SP (2-sucrose phosphate) transport medium. For patients with keratitis, an ophthalmologist collects scrapings of the cornea with a heat-sterilized platinum spatula. For bacterial isolation, multiple inoculations with the spatula are made to blood agar, chocolate agar, an agar for the isolation of fungi, thioglycollate broth, and an anaerobic blood agar plate. Other special media may be used if indicated. Corneal specimens for culture of HSV and adenovirus are placed in viral transport media. Recently the collection of two corneal scrapes (one used for Gram stain and the other transported in brain-heart infusion medium and used for culture) was determined to provide a simple method for diagnosis of bacterial keratitis. Cultures of endophthalmitis specimens are inoculated with material obtained by the ophthalmologist from the anterior and posterior chambers of the eye, wound abscesses, and wound dehiscence (splitting open). Lid infection material is collected on a swab in the conventional manner. For microbiologic studies of canaliculitis, material from the lacrimal canal should be transported under anaerobic conditions. Aspiration of fluid from the orbit is contraindicated in patients with orbital cellulitis. A patient history of sinusitis in association with orbital cellulitis is an indication for obtaining an otolaryngologist’s assistance in the collection of material from the maxillary sinus by antral puncture. Blood cultures should also be obtained. Tissue biopsy is essential for the microbiologic diagnosis of mucormycosis. Because cultures are usually negative, the diagnosis is made by histologic examination. 

Direct Visual Examination All material submitted for culture should be smeared and examined directly by Gram stain or other appropriate microscopic techniques. In bacterial conjunctivitis, polymorphonuclear leukocytes predominate; in viral infection, the host cells are primarily lymphocytes and monocytes. Specimens in which Chlamydia is suspected can be stained immediately with monoclonal antibody conjugated to fluorescein for the

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detection of elementary bodies or inclusions. Using histologic stains, basophilic intracytoplasmic inclusion bodies are seen in epithelial cells. Cytologists and anatomic pathologists usually perform these tests. Direct examination of conjunctivitis specimens using histologic methods (Tzanck smear; a scraping from the lesion for collection of cells) may reveal multinucleated epithelial cells typical of herpes viral infections. However, DFA stains available for both HSV and VZV are recommended for rapid diagnosis of viral infections. In patients with keratitis, scrapings may be examined using Gram, Giemsa, periodic acid-Schiff (PAS), and methenamine silver stains. If Acanthamoeba or other amoebae are suspected, corneal scrapings or a corneal biopsy should be kept at room temperature (24°C to 28°C) and a direct wet preparation should be examined for motile trophozoites, and a trichrome stain should be added to the regimen. For this diagnosis, however, culture is by far the most sensitive detection method for the identification of the organism. In patients with endophthalmitis, the specimen is examined using Gram, Giemsa, PAS, and methenamine silver stains. When submitted in large volumes of fluid, ophthalmic specimens must be concentrated by centrifugation before additional studies are performed. 

Nucleic Acid Testing Methods In general, for nucleic acid testing methods, the manufacturer provides specific directions and/or specific collection vials or transport media. All collection, processing, and transport should follow the manufacturer’s recommended protocols. 

Other Nonculture Methods Although acute and convalescent serologic tests for viral agents might be used in the event of epidemic conjunctivitis, they typically are not performed, because the infections are self-limited. Enzyme-linked immunosorbent assay (ELISA) tests and DFA staining are available for the detection of C. trachomatis. An ELISA test of aqueous humor is available for the diagnosis of Toxocara infection. Finally, nucleic acid–based methods have replaced most of these methods and are used for the diagnosis of viral and chlamydial keratoconjunctivitis, along with other ophthalmic infections, including uveitis. 

Culture Because of the constant washing action of the tears, the number of organisms recovered from cultures of eye infections may be relatively low. Unless the clinical specimen is obviously purulent, using a relatively large inoculum and a variety of media is recommended to ensure recovery of the etiologic agent. Conjunctival scrapings placed directly onto media yield the best results. At a minimum, blood and chocolate agar plates should be inoculated and incubated under increased carbon dioxide tension (5% to 10% CO2). Because potential pathogens may be present in an eye without causing infection, it can be very helpful to culture both eyes. If a potential pathogen grows in cultures of the infected and the

uninfected eye, the organism may not be causing the infection; however, if the organism only grows in culture from the infected eye, it is most likely the causative agent. When M. lacunata is suspected, Loeffler medium may prove useful; the growth of the organism often leads to proteolysis and pitting of the medium, although nonproteolytic strains may be isolated. If diphtheritic conjunctivitis is suspected, Loeffler or cystine-tellurite medium should be used. For more serious eye infections, such as keratitis, endophthalmitis, and orbital cellulitis, a reduced anaerobic blood agar plate, a medium for the isolation of fungi, and a liquid medium such as thioglycolate broth should always be included. The diagnosis of endophthalmitis typically requires a culture of the vitreous; vitreous washings typically yield positive cultures better than more invasive techniques. Invasive techniques for the collection of vitreous include either a needle aspirate or vitrectomy. A vitrectomy is a surgical procedure that simultaneously cuts and collects some vitreous fluid. Blood cultures are also important in serious eye infections. Specimen cultures for Chlamydia and viruses should be inoculated to appropriate media from transport broth. For Chlamydia isolation, cycloheximide-treated McCoy cells should be used; for viral isolation the use of human embryonic kidney, primary monkey kidney, and Hep-2 cell lines is recommended. 

Ears Anatomy The ear is divided into three anatomic parts: the external, middle, and inner ear. Important anatomic structures are depicted in Fig. 71.4. The middle ear is part of a continuous system including the nares, nasopharynx, auditory tube, and mastoid air spaces. These structures are lined with respiratory epithelium (e.g., ciliated cells, mucus-secreting goblet cells). 

Resident Microbiota The normal microbiota within the external ear canal is rather sparse, similar to flora of the conjunctival sac. Pneumococci (S. pneumoniae), C. acnes, S. aureus, and Enterobacterales are somewhat more common. P. aeruginosa is found on occasion. Candida spp. (non–C. albicans) are also common. 

Diseases, Epidemiology, and Etiology of Disease Otitis Externa (External Ear Infections) Otitis externa is similar to skin and soft tissue infection. Two major types of external otitis exist: acute or chronic. Acute external otitis may be localized or diffuse. Acute localized disease occurs in the form of a pustule or furuncle and typically is caused by S. aureus. Erysipelas caused by group A streptococci may involve the external ear canal and the soft tissue of the ear. Acute diffuse otitis externa

CHAPTER 71  Infections of the Eyes, Ears, and Sinuses

Middle ear

External ear Pinna

Temporal bone

External auditory meatus

Tympanic membrane

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Inner ear Semicircular canals

Vestibular nerve Cochlear nerve

Acoustic nerve (VIII)

Vestibule Cochlea

Auditory ossicles (small bones)

Auditory (eustacian) tube

• Fig. 71.4  The ear. (Modified from Thibodeau GA, Patton KT. Anatomy and Physiology. 2nd ed., St Louis: Mosby; 1993.)

(swimmer’s ear) is related to maceration (softening of tissue) of the ear from swimming or hot, humid weather. Gramnegative bacilli, particularly P. aeruginosa, play an important role. A severe, hemorrhagic external otitis caused by P. aeruginosa is difficult to treat and has occasionally been related to hot tub use. Chronic otitis externa results from the irritation of drainage from the middle ear in patients with chronic, suppurative otitis media and a perforated eardrum. Malignant otitis externa is a necrotizing infection that spreads to adjacent areas of soft tissue, cartilage, and bone. If allowed to pro­ gress and spread into the central nervous system or vascular channel, a life-threatening condition may develop. P. aeruginosa, in particular, and anaerobes are commonly associated with this process. Malignant otitis media is seen in patients with diabetes who have blood vessel disease of the tissues overlying the temporal bone in which poor local perfusion of tissues results in an environment conducive for invasion by bacteria. On occasion, external otitis can extend into the cartilage of the ear, usually requiring surgical intervention. Certain viruses may infect the external auditory canal, the soft tissue of the ear, or the tympanic membrane; influenza A virus is a suspected, but not an established, cause. VZV may cause painful vesicles within the soft tissue of the ear and the ear canal. Viral agents such as influenza and bacterial agents are typically associated with acute otitis media (S. pneumoniae, H. influenzae, and M. catarrhalis). Mycoplasma pneumoniae is rarely associated with this condition. 

agents in acute disease. Group A streptococci (S. pyogenes), M. catarrhalis, S. aureus, gram-negative enteric bacilli, and anaerobes are also associated with middle ear infections. Viruses, chiefly respiratory syncytial virus (RSV), coronaviruses, enteroviruses, rhinoviruses, and influenza viruses, have been recovered from the middle ear fluid of children with acute or chronic otitis media. C. trachomatis and M. pneumoniae have occasionally been isolated from middle ear aspirates. Otitis media with effusion (fluid) is considered a chronic sequela of acute otitis media. A slowly growing organism, Alloiococcus otitis, is a pathogen that has been isolated from patients with otitis media with effusion. Chronic otitis media yields a predominantly anaerobic flora, with Peptostreptococcus spp., Bacteroides fragilis group, Prevotella melaninogenica (pigmented, anaerobic, gramnegative rods), Porphyromonas, other Prevotella spp., and Fusobacterium nucleatum as the principal pathogens; less common are S. aureus, P. aeruginosa, Proteus spp., and other gram-negative facultative bacilli. Table 71.2 summarizes the major causes of ear infections. The mastoid is a portion of the temporal bone (lower sides of the skull) containing the mastoid sinuses (cavities). Mastoiditis is a complication of chronic otitis media in which organisms find their way into the mastoid sinuses. To prevent the further spread of this infection to the central nervous system, a mastoidectomy is performed. 

Otitis Media (Middle Ear Infections)

Pathogenesis

In children (in whom otitis media is most common), pneumococci and H. influenzae are the usual etiologic

Local trauma, the presence of foreign bodies, or excessive moisture can lead to otitis externa (external ear infections).

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TABLE 71.2    Major Infectious Causes of Ear Infection

silver stains have the added efficiency of staining most bacterial and fungal organisms and several parasitic species. 

Disease

Common Causes

Culture and Nonculture Methods

Otitis externa

Acute: Staphylococcus aureus, Streptococcus pyogenes, Pseudomonas aeruginosa; other gram-negative bacilli

Ear specimens submitted for culture should be inoculated to blood, MacConkey, and chocolate agars. Anaerobic cultures should also be set up on those specimens obtained by tympanocentesis or those obtained from patients with chronic otitis media or mastoiditis. Because cultures of middle ear effusions are culture positive for only 20% to 30% of patients, conventional and nucleic acid–based test methods have been used to detect the common middle ear pathogens. 

Chronic: P. aeruginosa; anaerobes Otitis media

Acute Streptococcus pneumoniae; Haemophilus influenzae; Moraxella catarrhalis; S. pyogenes; respiratory syncytial virus; influenza virus; coronaviruses, enteroviruses, rhinoviruses Chronic: Anaerobes

Note: This table is not intended to be all-inclusive for the infectious agents capable of causing ear infections.

Sinuses Anatomy

Occasionally, an infection from the middle ear can extend by purulent drainage to the external ear. Anatomic or physiologic abnormalities of the auditory tube can predispose individuals to develop otitis media. The auditory tube is responsible for protecting the middle ear from nasopharyngeal secretions, draining secretions produced in the middle ear into the nasopharynx, and ventilating the middle ear and equilibrating air pressure with the external ear canal. If any of these functions become compromised and fluid develops in the middle ear, infection may occur. To illustrate, if a person has a viral upper respiratory infection, the auditory tube becomes inflamed and swollen. This inflammation and swelling may, in turn, compromise the auditory tube’s ventilating function, resulting in a negative, rather than a positive, pressure in the middle ear. This change in pressure can then allow for potentially pathogenic bacteria present in the nasopharynx to enter the middle ear. 

Laboratory Diagnosis Specimen Collection and Transport Although middle ear infection, or otitis media, is usually not diagnosed by culture, culture can be used for the laboratory diagnosis of external otitis; the external ear should be cleansed with a mild germicide such as 1:1000 aqueous solution of benzalkonium chloride to reduce the numbers of contaminating skin microbiota before obtaining the specimen. Material from the ear, especially that obtained after spontaneous perforation of the eardrum or by needle aspiration of middle ear fluid (tympanocentesis), should be collected by an otolaryngologist, using sterile equipment. Specimens from the mastoid are generally taken on swabs during surgery, although actual bone is preferred. Specimens should be transported anaerobically. 

Direct Visual Examination Material aspirated from the middle ear or mastoid is also examined directly for bacteria and fungi. The calcofluor white or PAS stains can reveal fungal elements. Methenamine

The sinuses, like the mastoids, are unique, air-filled cavities within the head (Fig. 71.5). The sinuses are normally sterile. These structures, as well as the eustachian tube, the middle ear, and the respiratory portion of the pharynx, are lined by respiratory epithelium. The clearance of secretions and contaminants depends on normal ciliary activity and mucous flow. 

Diseases The pathogens associated with otitis media are the same ones associated with sinusitis; bacteria from the nose and throat make their way to the inner ear and sinuses. Acute sinusitis usually develops during a cold or influenza illness and tends to be self-limited, lasting 1 to 3 weeks, and is usually more prevalent in winter and spring. Acute sinusitis is often difficult to distinguish from the primary illness. Symptoms include purulent nasal and postnasal discharge, a feeling of pressure over the sinus areas of the face, cough, and a nasal quality to the voice. Fever is sometimes present. Occasionally, acute sinusitis persists and reaches a chronic state in which bacterial colonization occurs and the condition no longer responds to antibiotic treatment. Ordinarily, surgery or drainage is required for successful management. Patients with chronic sinusitis may have acute exacerbations (flare-ups). Other complications include local extension into the orbit, skull, meninges, or brain, and development of chronic sinusitis. 

Pathogenesis Most cases of acute sinusitis are believed to be bacterial complications following a viral respiratory infection. The exact mechanisms involved are unknown. About 5% to 10% of acute maxillary sinus infections result from infection originating from a dental source. The maxillary sinuses are close to the roots of the upper teeth, providing a mechanism for dental infections to extend into the sinuses. The primary

CHAPTER 71  Infections of the Eyes, Ears, and Sinuses

Frontal sinus Ethmoid sinus Maxillary sinus

Sphenoid sinus

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have indicated that S. pneumoniae and H. influenzae are the major bacterial pathogens in adults with acute sinusitis; other species such as beta-hemolytic and alpha-hemolytic streptococci, S. aureus, and anaerobes have also been cultured but less commonly. The predominant bacterial organisms associated with chronic sinusitis include S. pneumoniae, H. influenzae, and M. catarrhalis; less commonly, isolated organisms include anaerobic streptococci, Prevotella spp., and Fusobacterium spp. Fungal pathogens such as Aspergillus, Fusarium, and C. albicans have also been identified in cases of chronic sinusitis using culture and nucleic acid–based test methods. The major causes of acute sinusitis are summarized in Table 71.3. M. catarrhalis has been isolated in chronic sinusitis in children. 

Laboratory Diagnosis •

Fig. 71.5 Location of the paranasal sinuses. (From Milliken ME, Campbell G. Essential Competencies for Patient Care. St Louis: Mosby; 1985.)

TABLE 71.3    Major Infectious Causes of Acute Sinusitis

Age Group

Common Causes

Young adults

Haemophilus influenzae, Streptococcus pneumoniae, Streptococcus pyogenes, Moraxella catarrhalis

Children

S. pneumoniae, H. influenzae, M. catarrhalis, rhinovirus

Note: This table is not intended to be all-inclusive for the infectious agents capable of causing sinusitis.

problems associated with the development of chronic sinusitis include inadequate drainage, impaired mucociliary clearance, and mucosal damage. 

Epidemiology and Etiology of Disease Although difficult to assess, the actual incidence of acute sinusitis parallels that of acute upper respiratory tract infections (i.e., being most prevalent in the fall through spring). Most studies of the microbiology of acute sinusitis are associated with maxillary sinusitis, because it is the most common type and specimen collection is available through puncture and aspiration. Acute viral sinusitis is one of the most common causes of respiratory tract infection and, in most cases, resolves without treatment. However, published estimates indicate that 0.5% to 2% of cases of acute viral sinusitis in adults are complicated by bacterial sinusitis. This scenario is even more common in children. Bacterial cultures are positive in about three-fourths of patients. Studies

In most cases, a diagnosis can be made based on physical findings, history, radiograph studies, and other imaging techniques such as magnetic resonance imaging. However, if a laboratory diagnosis is needed, an otolaryngologist collects a specimen from the maxillary sinus by puncture and aspiration or during surgery. Sinus drainage is unacceptable for smear or culture, because this material will be contaminated with aerobic and anaerobic normal respiratory microbiota; sinus washings or aspirates surgically collected are the specimens of choice. Gram-stained smears and aerobic and anaerobic cultures should be performed on each specimen. Aerobic culture media should include blood, chocolate, and MacConkey agar.

Matrix-Assisted Laser Desorption Ionization Timeof-Flight Mass Spectrometry Matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS) may be used to identify pathogens from these infections directly from pure colony isolates. The limitations of identification, however, rely on technical expertise in sample preparation and the limitations of the current database (Chapter 7).

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, PA, 2019, Elsevier. Bernardes TF, Bonfioli AA: Blepharitis, Semin Ophthalmol 25: 79–83, 2010. Carbonnelle E, Grohs P, Jacquier H, et  al.: Robustness of two MALDI-TOF mass spectrometry systems for bacterial identification, J Microbiol Methods 89:133–136, 2012. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Cramer L, Emara DM, Gadre AK: Mycoplasma an unlikely cause of bullous myringitis, Ear Nose Throat J 91:E30–E31, 2012.

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Creemers-Schild D, Gronthoud F, Spanjaard L, et al.: Fusobacterium necrophorum, an emerging pathogen of otogenic and paranasal infections? New Microbes New Infect 2:52–57, 2014. Finegold SM, Flynn MJ, Rose FV, et al.: Bacteriologic findings associated with chronic bacterial maxillary sinusitis in adults, Clin Infect Dis 35:428–433, 2002. Hendolin PH, Paulin L, Ylikoski J: Clinically applicable multiplex PCR for four middle ear pathogens, J Clin Microbiol 38:125–132, 2000. Henry CR, Flynn HW, Miller D, et al.: Infectious keratitis progressing to endophthalmitis: a 15-year study of microbiology, associated factors, and clinical outcomes, Ophthalmology 119:2443–2449, 2012. Holtz KK, Townsend KR, Furst JW, et  al.: An assessment of the AdenoPlus Point-of-Care test for diagnosing Adenoviral conjunctivitis and its effect on antimicrobial stewardship, Mayo Clin Proc Innov Qual Outcomes 1(2):170–175, 2017. Kaye SB, Rao PG, Smith G, et al.: Simplifying collection of corneal specimens in cases of suspected bacterial keratitis, J Clin Microbiol 41:3192–3197, 2003. Kim ST, Choi JH, Jeon HG, et  al.: Comparison between polymerase chain reaction and fungal culture for the detection of fungi in patients with chronic sinusitis and normal controls, Acta Otolaryngol 125:72–75, 2005.

Lynn WA, Lightman S: The eye in systemic infection, Lancet 364:1439–1450, 2004. Marciano-Cabral F, Cabral G: Acanthamoeba spp. as agents of disease in humans, Clin Microbiol Rev 16:273–307, 2003. Moorthy RS, Valluri S, Rao NA: Nontuberculous mycobacterial ocular and adnexal infections, Surv Ophthalmol 57:202–235, 2012. Palmu AA, Herva E, Savolainen H, et al.: Association of clinical signs and symptoms with bacterial findings in acute otitis media, Clin Infect Dis 38:234–242, 2004. Piccirillo JF: Clinical practice. Acute bacterial sinusitis, N Engl J Med 351:902–910, 2004. Roels P: Ocular infections of AIDS: new considerations for patients using highly active anti-retroviral therapy (HAART), Optometry 75:624–628, 2004. Sande M, Gwaltney JM: Acute community-acquired bacterial sinusitis: continuing challenges and current management, Clin Infect Dis 39:S151–S158, 2004. Skevaki CL, Galani IE, Pararas MV, et al.: Treatment of viral conjunctivitis with antiviral drugs, Drugs 71:331–347, 2011. Solomon AW, Peeling RW, Foster A, et al.: Diagnosis and assessment of trachoma, Clin Microbiol Rev 17:982–1011, 2004. Zhang L, Zhao N, Huang X, et al.: Molecular epidemiology of acute hemorrhagic conjunctivitis caused by coxsackie A type 24 variant in China, 2004-2014, Sci Rep 7:45202, 2017. Accessed October 20, 2019.

CASE STUDY 71.1 A 12-year-old boy complained of severe ear pain of 4 days duration. He was afebrile, but his tympanic membrane was erythematous and bleeding. The boy had been swimming a few days earlier in a local lake. His physician collected samples, which grew a gram-negative rod with a pleasant odor. The patient was given antibiotic eardrops and did well, with resolution of his symptoms.

Questions 1. What organism caused this infection? 2. How can this isolate be identified rapidly? 3. If the characteristic odor is lacking, what characteristics of the organism make it easy to identify?

  

Chapter Review 1. The anterior cavity of the interior space of the eyeball is filled with a clear, watery substance called: a. Vitreous humor b. Aqueous humor c. Lacrimal humor d. None of the above 2.  Which of the following organisms is the causative agent for blepharitis? a.  S. aureus b. S. epidermidis c. HSV d. All the above 3. One of the most serious types of eye infection requiring immediate medical attention because of the risk of corneal perforation is: a. Conjunctivitis b. Keratitis c. Endophthalmitis d. Canaliculitis 4. Which of the following organisms is commonly associated with corneal eye infection in soft and extendedwear contact users? a.  Acanthamoeba sp. b. Pseudomonas aeruginosa c.  Acinetobacter d. S. aureus 5. Newborns are treated with antibiotic drops immediately after birth to prevent infection by what organism? a.  C. trachomatis b. HSV c. Group B streptococci d. H. influenzae 6. The part of the eye that is a white fibrous tissue is called the: a. Cornea b. Lacrimal canaliculi c. Conjunctiva d. Sclera 7. Acute inflammation of the lacrimal gland may occur during what disease(s)? a. Syphilis and leprosy b. Tuberculosis c. Varicella zoster d. Mumps and infectious mononucleosis

8. Which of the following media is used in the culture of an eye infection if the organism suspected is M. lacunata? a. Columbia with CNA media b. Thayer-Martin media c. Loeffler medium d. Regan-Lowe media 9. All the following organisms are normally responsible for acute localized otitis media in children except: a.  S. pneumoniae b. S. aureus c.  S. pyogenes d. H. influenzae 10. The components of the lacrimal apparatus contain all the following except: a. Lacrimal gland b. Lacrimal canaliculi c. Lacrimal duct d. Lacrimal sac 11.  True or False _____ Otitis media or middle ear infection is usually not diagnosed by culture. _____ The major etiologic agent of acute sinusitis in adults is Moraxella catarrhalis. _____ The most common cause of orbital cellulitis is Staphylococcus aureus. _____ When culturing an eye specimen for infections caused by Chlamydia, cycloheximide-treated McCoy cells should be used. _____ When examining the Gram stain from a suspected case of bacterial conjunctivitis, lymphocytes and monocytes will be the predominate cells within the sample. _____ Aerobic gram-negative rods most often cause chronic otitis media. _____ PCR is often the testing method used to detect middle ear pathogens, because it is more sensitive than conventional culture. _____ Most cases of acute sinusitis are believed to be bacterial complications of the common cold. _____ In suspected cases of sinusitis, an acceptable specimen for culture is nasal drainage.

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1016.e2 PA RT V I I     Diagnosis by Organ System

12.  Matching: Match each term with the correct description. _____ bullous myringitis _____ uveitis _____ maceration _____ cornea _____ trachoma _____ exopeptidase _____ mastoiditis _____ otitis externa _____ canaliculitis _____ dacryocystitis _____ mucormycosis _____ iatrogenic _____ dacryoadenitis _____ lacrimal _____ endophthalmitis _____ blepharitis

a. swimmer’s ear b. anterior portion of the sclera; clear with no blood vessels c. infection of the iris, ciliary body, and choroid d. toxic factor produced by S. pneumoniae e. inflammation of the mastoid air cells f. resulting from the activities of a doctor g. infection of the lacrimal gland characterized by upper eyelid pain with erythema h. leading cause of blindness in underdeveloped countries caused by Chlamydia trachomatis i. infection of the internal area of the eyeball j. inflammation of the eyelids k. infection of the lacrimal sac l. softening of tissue m. invasive fungal infection of the eye orbit n. inflammation of the lacrimal canal o. painful infection of the eardrum with hemorrhagic bullae p. pertaining to tears

13.  Short Answer (1) What culture method may help a doctor determine whether an organism isolated in an eye culture is the pathogen or just normal microbiota? (2) What is the greatest risk factor associated with orbital cellulitis? (3) What type of suspected eye infection requires specimen transport to the laboratory under anaerobic conditions? (4) How does the cell culture collection of a specimen for chlamydial-suspected conjunctivitis differ from

that of conjunctivitis caused by other organisms? What testing method has replaced cell culture in most laboratories? (5) Explain the difference between acute and chronic otitis externa. (6) Differentiate acute and chronic sinusitis. (7) Describe the toxic factors exhibited by S. pneumoniae and Pseudomonas aeruginosa responsible for corneal destruction in keratitis.

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Infections of the Urinary Tract OBJECTIVES 1. Describe the anatomy and identify the structures of the male and female urinary tracts. 2. Name the organisms that colonize the urethra and are considered normal microbiota. 3. Explain how the female urinary tract anatomy may predispose females to urinary tract infections. 4. Differentiate between community-acquired urinary tract infections and hospital- and health care–associated urinary tract infections. 5. List the routes of transmission that allow bacteria to invade and cause a urinary tract infection. 6. Name the physical and chemical properties of urine that contribute to its role in the body’s defense mechanism against the bacteria capable of causing urinary tract infections. 7. Explain host and microbial factors that determine whether bacteria will be able to colonize and cause a urinary tract infection. 8. Name the properties bacteria possess that predispose them to have greater pathogenicity in causing urinary tract infections. 9. Define the five major types of urinary tract infections: pyelonephritis, cystitis, urethritis, acute urethral syndrome, and asymptomatic bacteriuria. 10. Compare complicated and uncomplicated urinary tract infections. 11. Explain the collection methods for urine specimens, including clean catch midstream urine, straight catheterized urine, a suprapubic bladder aspiration, and an indwelling catheter collection. 12. Describe the urine-screening methods available to determine bacteriuria and pyuria. 13. Explain the nitrate reductase test, the leukocyte esterase test, and the catalase test regarding their urine-screening capability. 14. Name the media required for urine cultures. 15. Explain the proper methodology for plating and interpreting a quantitative urine culture. 16. Correlate signs and symptoms with the results of laboratory diagnostic procedures for the identification of the etiologic agent associated with infections of the urinary tract.

General Considerations Anatomy The urinary tract consists of the kidneys, ureters, bladder, and urethra (Fig. 72.1). The function of the urinary tract is to make and process urine. Urine is an ultrafiltrate of blood that consists mostly of water but also contains nitrogenous wastes, sodium, potassium, chloride, and other analytes. Urine is normally a sterile fluid. Often, urinary tract infections (UTIs) are characterized as being either upper (U-UTI) or lower (L-UTI) based primarily on the anatomic location of the infection: the lower urinary tract encompasses the bladder and urethra, and the upper urinary tract encompasses the ureters and kidneys. Upper UTIs affect the ureters (ureteritis) or the renal parenchyma (pyelonephritis). Lower UTIs may affect the urethra (urethritis), the bladder (cystitis), or the prostate in males (prostatitis). Symptomatic L-UTIs and asymptomatic U-UTIs do not rule out the possibility that the infectious agent may also be affecting the upper urinary tract. The anatomy of the female urethra is of importance to the pathogenesis of UTIs. The female urethra is relatively short compared with the male urethra and lies in close proximity to the warm, moist, perirectal region, which is teeming with microorganisms. Because of the shorter urethra, bacteria can reach the bladder more easily in the female host; thus UTIs are primarily a disorder in females. In males, the incidence of UTIs increases after the age of 60 years, when the enlargement of the prostate interferes with the removal of urine from the bladder. UTIs can also be classified as uncomplicated or complicated. An uncomplicated UTI indicates that there are no structural or neurological abnormalities associated with the urinary tract. A complicated UTI indicates a history of persistent, recurring infections that may be a result of physiological factors that predispose the patient to infection. This may include previous kidney failure, obstruction, kidney transplant, immunosuppression, or urinary retention. 

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Diaphragm

Upper urinary tract

Adrenal gland Left kidney

Right kidney Ureters

Lower urinary tract

Bladder

• BOX 72.1 Resident Microbiota of the Urethra Coagulase-negative staphylococci (excluding Staphylococcus saprophyticus) Viridans and nonhemolytic streptococci Lactobacilli (adult females) Diphtheroids (Corynebacterium spp.) Nonpathogenic (saprobic) Neisseria spp. (adult women) Anaerobic cocci Propionibacterium spp. (adult patients) Commensal Mycobacterium spp. Commensal Mycoplasma spp. Yeasts (pregnant, adult females)

Urethra Male prostate

Female cervix

• Fig. 72.1  Overview of the anatomy of the urinary tract. (From Potter PH, Perry AG. Fundamentals of Nursing. St Louis: Mosby; 1985.)

Resident Microbiota of the Urinary Tract The urethra has resident microbiota that colonize its epithelium in the distal portion; these organisms are lactobacilli, corynebacteria, enterococci, and coagulase-negative staphylococci (Box 72.1). Potential pathogens, including gramnegative aerobic bacilli (primarily Enterobacterales) and occasional yeasts, are also present as transient colonizers. All areas of the urinary tract above the urethra in a healthy human were previously considered sterile. There is increasing evidence that the urinary bladder may be protected by normal microbiota. Urine is typically sterile, but noninvasive methods for collecting urine must rely on a specimen that has passed through a contaminated milieu. Therefore, quantitative cultures for the diagnosis of UTIs have been used to discriminate among contamination, colonization, and infection. 

Infections of the Urinary Tract

bacteria in urine) among females 5 through 17 years of age is 1% to 3%. The prevalence of bacteriuria in females increases gradually with time to as high as 10% to 20% in older females. In females between 20 and 40 years of age who have had UTIs, as many as 50% may become reinfected within 1 year. The association of UTIs with sexual intercourse may also contribute to this increased incidence, because sexual activity increases the chances of bacterial contamination of the female urethra. Finally, because of anatomic and hormonal changes that favor the development of UTIs, the incidence of bacteriuria increases during pregnancy. These infections can lead to serious infections in both the mother and fetus. Estrogen deficiency in postmenopausal females, resulting in a decrease in normal vaginal microbiota (in particular, lactobacilli), is associated with recurrent UTIs. UTIs are important complications of diabetes, renal disease, renal transplantation, and structural and neurologic abnormalities that interfere with urine flow. In 40% to 60% of renal transplant recipients, the urinary tract is the source for the primary occurrence of bacteremia, and in these patients, the recurrence rate is about 40%. In addition, UTIs are a leading cause of gram-negative sepsis in hospitalized patients and are the origin for about half of all health care–associated infections caused by urinary catheters. 

Epidemiology

Etiologic Agents Community-Acquired

UTIs are among the most common bacterial infections that lead patients to seek medical care. It has been estimated that more than 7 million outpatient visits, 1 million visits to the emergency department, and 100,000 hospital stays every year in the United States are a result of UTIs. Approximately 60% of all females and 5% of all males will have a UTI at some time during their lives. Of note, UTIs are also the most common hospital- and health care–associated infection. The exact prevalence of UTIs is age- and sex-dependent. During the first year of life, UTIs occur in less than 2% in males and females. The incidence of UTIs among males remains relatively low after 1 year of age and until approximately 60 years of age, when enlargement of the prostate interferes with emptying of the bladder. Extensive studies have shown that the incidence of bacteriuria (presence of

Escherichia coli is by far the most common cause of uncomplicated community-acquired UTIs. At the molecular level, E. coli, designated uropathogenic E. coli (UPEC), which causes UTIs, is sufficiently different from other types of E. coli. E. coli O25-H4 has emerged as a significant urinary tract pathogen in community-acquired infections. Other bacteria commonly isolated from patients with UTIs are Klebsiella spp., other Enterobacterales, Staphylococcus saprophyticus, and enterococci. More than 95% of uncomplicated UTIs are caused by a single bacterial species. In more complicated UTIs, particularly in recurrent infections, the relative frequency of infection caused by Proteus, Pseudomonas, Klebsiella, and Enterobacter spp. increases. In addition, community-acquired UTIs are increasingly associated with multidrug-resistant organisms such as extended beta-lactamase-resistant E. coli. 

CHAPTER 72  Infections of the Urinary Tract

Hospital- and Health Care–Associated The hospital or health care environment plays an important role in determining the organisms involved in UTIs. Hospitalized patients are most likely to be infected by antibiotic-resistant E. coli, Klebsiella spp., Proteus spp., staphylococci, enterococci, other Enterobacterales, Pseudomonas aeruginosa, Enterobacter spp., and Candida spp. The introduction of a foreign body into the urinary tract, especially one that remains in place for an extended period (e.g., Foley catheter), carries a substantial risk of infection, particularly if obstruction is present. Approximately 35% of all health care–associated infections are UTIs. Eighty percent of those infections are associated with the use of an indwelling catheter. In addition, highly antibiotic-resistant microorganisms such as extended-beta-lactamase–producing organisms (ESBL); Amp C beta-lactamase-, carbapenemase-producing Enterobacterales, and Acinetobacter spp. are increasingly identified in health care–associated UTIs. It is also not unusual to identify multiple bacterial species or infectious agents in complicated UTIs, due to repeat medical procedures and previous treatment with antibiotics. Consequently, UTI is the most common health care–associated infection in the United States, and the infected urinary tract is the most common source of bacteremia. 

Miscellaneous Other less commonly isolated agents are other gram-negative bacilli, such as Acinetobacter and Alcaligenes spp., other Pseudomonas spp., Citrobacter spp., Gardnerella vaginalis, Aerococcus urinae, and beta-hemolytic streptococci. Bacteria such as mycobacteria (predominantly in patients who are human immunodeficiency virus [HIV]-positive), Chlamydia trachomatis, Ureaplasma urealyticum, Mycoplasma hominis, Campylobacter spp., Haemophilus influenzae, Leptospira, and certain Corynebacterium spp. (e.g., C. renale) are rarely recovered from urine. In addition, Actinobaculum schaalii (vaginal and skin microbiota) and other Actinobaculum spp. may be dismissed as normal microbiota or overlooked in urine cultures because of these organisms’ slow growth rate. Because renal transplant recipients are immunosuppressed, these patients not only suffer from common uropathogens but are also susceptible to opportunistic infections with unusual pathogens. A study involving renal transplant recipients showed that for culture-negative urine, amplification of regions in bacterial 16S ribosomal ribonucleic acid (rRNA) and subsequent analysis by high-performance liquid chromatography detected the presence of several known uropathogens as well as unusual agents. For example, urine specimens from renal transplant recipients or infants may contain Listeria monocytogenes associated with a systemic infection. Salmonella spp. may be recovered during the early stages of typhoid fever; their presence should be immediately reported to the physician. If anaerobes are suspected, the physician should perform a percutaneous bladder tap unless urine can be obtained from the upper urinary tract by another means (e.g., from a nephrostomy tube). Communication by the clinician to the laboratory that such an

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agent is suspected is important for detecting such agents. In patients with “sterile pyuria,” Gram stain may reveal unusual organisms with distinctive morphology (e.g., H. influenzae, anaerobes). The presence of any organisms on smear that do not grow in culture is an important clue to the cause of the infection. The laboratory can then take the action necessary to optimize chances for recovery. As previously noted, Candida spp. may be isolated from patients with other debilitating disease (i.e., diabetes or urinary tract obstructions) and are associated with immunosuppressive therapy or immunosuppressive conditions and antibiotic treatment. Additional fungi capable of causing systemic infections that may also be identified in urine samples include Blastomyces dermatitidis, Coccidioides immitis, Cryptococcus neoformans, and Histoplasma capsulatum. The identification of a fungal isolate in a urine specimen should be carefully evaluated and reported to the attending clinician. In general, viruses and parasites are not usually considered urinary tract pathogens. However, adenovirus has been implicated as the causative agent in hemorrhagic cystitis in pediatric patients. Trichomonas vaginalis may occasionally be observed in urinary sediment, and Schistosoma haematobium can lodge in the urinary tract and release eggs into the urine. 

Pathogenesis Routes of Infection Bacteria can invade and cause a UTI via three major routes: ascending, hematogenous, and lymphatic pathways. Although the ascending route is the most common course of infection in females, ascent in association with instrumentation (e.g., urinary catheterization, cystoscopy) is the most common cause of health care–associated UTIs in both sexes. For UTIs to occur by the ascending pathway, enteric gram-negative bacteria and other microorganisms that originate in the gastrointestinal tract must be able to colonize the vaginal cavity or the periurethral area. Once these organisms gain access to the bladder, they may multiply and then pass up the ureters to the kidneys. UTIs occur more often in females, at least partially because of the short female urethra and its proximity to the anus. As previously mentioned, sexual activity can increase the chances of bacterial contamination of the female urethra. In addition, postmenopausal women are more susceptible to uropathogens due to the deficiency in estrogen and the loss of protective lactobacilli in the vaginal tract. In most hospitalized patients, UTI is preceded by urinary catheterization or other manipulation of the urinary tract. The pathogenesis of catheter-associated UTI is not fully understood. It is certain that soon after hospitalization, patients become colonized with bacteria endemic to the institution—often gram-negative aerobic and facultative bacilli carrying resistance markers. These bacteria colonize the patient’s skin, gastrointestinal tract, and mucous membranes, including the anterior urethra. With the insertion of a catheter, the bacteria may be pushed along the urethra into

1020 PA RT V I I     Diagnosis by Organ System

the bladder or, with an indwelling catheter, may migrate along the track between the catheter and the urethral mucosa, gaining access to the bladder. It is estimated that approximately 10% to 30% of catheterized patients will develop bacteriuria (presence of bacteria in urine). UTIs may also occur by the hematogenous, or bloodborne, route. Hematogenous spread usually occurs as a result of bacteremia. Any systemic infection can lead to seeding of the kidney, but certain organisms, such as Staphylococcus aureus or Salmonella spp., are particularly invasive. Although most infections involving the kidneys are acquired through the ascending route, yeast (usually Candida albicans), Mycobacterium tuberculosis, Salmonella spp., Leptospira spp., or S. aureus in the urine may indicate pyelonephritis acquired via hematogenous spread or the descending route. Hematogenous spread accounts for less than 5% of UTIs and rarely occurs with gram-negative bacilli. Finally, increased pressure on the bladder can cause lymphatic flow into the kidneys, resulting in UTI. However, evidence for the significance of this potential route is insufficient, indicating that the ascending route remains the major mechanism for the development of UTI. 

The Host-Pathogen Relationship Many individuals (females) are colonized in the vaginal or periurethral area with organisms originating from the gastrointestinal tract, yet they do not develop urinary infections. Whether an organism is able to colonize and then cause a UTI is determined in large part by a complex interplay of host and microbial factors. In most cases, the host defense mechanisms can eliminate the organisms. Urine itself is inhibitory to some of the urethral microbiota, such as anaerobes. In addition, if urine has a low pH, high or low osmolality, high urea concentration, or high organic acid content, even organisms capable of growth in the urinary tract may be inhibited. If bacteria do gain access to the bladder, the constant flushing of contaminated urine from the body either eliminates bacteria or maintains their numbers at low levels. Clearly, any interference with the act of normal voiding, such as mechanical obstruction resulting from kidney stones or strictures, will promote the development of UTI. Also, the bladder mucosal surface has antibacterial properties. If the infection is not eradicated, the site of infection remains in the superficial mucosa; deep layers of the bladder are rarely involved. In addition to the previously described host defenses, a valvelike mechanism at the junction of the ureter and bladder prevents the reflux (backward flow) of urine from the bladder to the upper urinary tract. Therefore, if the function of these valves is inhibited or compromised in any way, such as by obstruction or congenital abnormalities, urine reflux provides a direct route for organisms to reach the kidney. Hormonal changes associated with pregnancy and their effects on the urinary tract increase the chance for urine reflux to the upper urinary tract. Activation of the host immune response by uropathogens also plays a key role in fending off infection. For example,

bacterial contact with urothelial cells initiates an immune response via a variety of signaling pathways. Bacterial lipopolysaccharide (LPS; Chapter 2) activates host cells to ultimately release cytokines such as tumor necrosis factor and interferon-gamma. In addition, bacteria can activate the complement cascade, leading to the production of biologically active components such as opsonins, as well as augment the host’s adaptive immune response. Host factors that lead to host susceptibility or resistance to uropathogens have been identified. For example, a glycoprotein synthesized exclusively by epithelial cells in a specific anatomic location in the kidney, referred to as Tamm-Horsfall protein (THP) or uromodulin, serves as an antiadherence factor by binding to E. coli–expressing type 1 fimbriae (discussed later). Defensins, a group of small antimicrobial peptides, are produced by a variety of host cells such as macrophages, neutrophils, and cells in the urinary tract and attach to the bacterial cell, eventually causing the organism’s death. Although many microorganisms can cause UTIs, most cases are a result of infection by a few organisms. To illustrate, only a limited number of serogroups of E. coli (01, 02, 04, 06, 07, 08, 075, 0150, 018ab) cause a significant proportion of UTIs. Numerous investigations indicate that UPEC possesses virulence factors that enhance their ability to colonize and invade the urinary tract. Some of these virulence factors include increased adherence to vaginal and uroepithelial cells by bacterial surface structures (adhesins), pili (P [PAP] type 1), and multiple types of fimbriae; the production of alpha-hemolysin (inhibits the production of protective cytokines), cytotoxic necrotizing factor (CNF), an autotransported protease (Sat), aerobactin (iuc), and a siderophore receptor (iroN); and resistance to serum-killing activity. Also, genome sequences of UPEC strains have been determined, indicating that several potential virulence factor genes associated with the acquisition and development of UTIs are encoded on pathogenicity islands (e.g., hemolysins and E. coli P fimbriae). By definition, pathogenicity islands (Chapter 3) contain genes that are associated with virulence and are absent from avirulent (not typically found in fecal strains) or less virulent strains of the same species. UPEC strains are a major cause of community-acquired UTIs. The importance of adherence in the pathogenesis of UTIs has also been demonstrated with other species of bacteria. Once introduced into the urinary tract, Proteus strains appear to be uniquely suited to cause significant disease in the urinary tract. Data indicate that these strains can facilitate their adherence to the mucosa of kidneys. Also, Proteus is able to hydrolyze urea via urease production. The species Proteus mirabilis accounts for approximately 77% of the urinary isolates. Hydrolysis of urea results in an increase in urine pH that is directly toxic to kidney cells and stimulates the formation of kidney stones. Similar findings have been made with Klebsiella spp. S. saprophyticus also adheres better to uroepithelial cells than S. aureus or S. epidermidis. Other bacterial characteristics may be important in the pathogenesis of UTIs. Motility may be important for

CHAPTER 72  Infections of the Urinary Tract

organisms to ascend to the upper urinary tract against the flow of urine and cause pyelonephritis. Some organisms demonstrate greater production of capsular K antigen (K1, K5, and K12); this antigen protects bacteria from being phagocytized. Finally, despite numerous host defenses and even antibiotic treatments that can effectively sterilize the urine, a significant proportion of patients have recurrent UTIs. Studies show that uropathogens can invade superficial epithelial cells in the bladder and replicate, forming large foci of intracellular organisms. This invasion of bladder epithelial cells triggers the host immune response, which in turn causes the superficial cells to exfoliate within hours after infection. Although this exfoliation is considered a host defense mechanism by eliminating infected cells, intracellular organisms can reemerge from the bladder epithelial cells and invade the underlying, new superficial layer of epithelial cells, consequently persisting within the urinary tract. It has been reported that intracellular bacteria mature into numerous, large protrusions on the bladder surface they referred to as “pods.” This bacterial organization—in which the intracellular bacteria are embedded in a fibrous, polysacchariderich matrix resembling that of a biofilm—may help further explain the persistence of bladder infections despite strong host defenses. 

Types of Infection and Their Clinical Manifestations UTI encompasses a broad range of clinical entities that differ in terms of clinical presentation, degree of tissue invasion, epidemiologic setting, and requirements for antibiotic therapy. There are several types of UTIs: urethritis, ureteritis, asymptomatic bacteriuria, cystitis, the urethral syndrome, and pyelonephritis. Uncomplicated infections occur primarily in otherwise healthy females and occasionally in male infants and adolescent and adult males. Most uncomplicated infections respond readily to antibiotic agents to which the etiologic agent is susceptible. Complicated infections occur in both sexes. In general, individuals who develop complicated infections often have certain risk factors. Some of these risk factors are listed in Box 72.2. In general, complicated infections are more difficult to treat and have greater morbidity (e.g., kidney damage, bacteremia) and mortality compared with uncomplicated infections. UTIs identified in pregnant women, men, children, • BOX 72.2 Risk Factors Associated With

Complicated Urinary Tract Infections

Underlying diseases that predispose the kidney to infection (e.g., diabetes, sickle cell anemia) Kidney stones Structural or functional abnormalities of the urinary tract (e.g., a tipped bladder) Indwelling urinary catheters

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and hospitalized patients or patients in other health care– associated settings (e.g., cancer outpatient clinics) may be considered complicated infections. The organisms associated with these infections are commonly highly resistant to many antimicrobials. The clinical presentation of UTIs may vary, ranging from asymptomatic infection to pyelonephritis (infection of the kidney and its pelvis). Some UTI symptoms may be nonspecific, and the symptoms of lower UTIs may be considerably similar to those of upper UTIs.

Urethritis Symptoms associated with urethritis (infection of the urethra), dysuria (painful or difficult urination), and frequency are similar to those associated with other lower UTIs. Urethritis is a common infection. Because C. trachomatis, Neisseria gonorrhoeae, and T. vaginalis are common causes of urethritis and considered to be sexually transmitted, urethritis is discussed as a sexually transmitted disease in Chapter 73. 

Ureteritis Inflammation or infection within the ureters (ureteritis) is considered in combination with kidney infections. UTI within the ureters indicates that organisms have begun or are in the process of ascending into the kidneys and should be treated similarly to prevent further infection. 

Asymptomatic Bacteriuria Asymptomatic bacteriuria or asymptomatic UTI is the isolation of a specified quantitative count of bacteria in an appropriately collected urine specimen obtained from a person without symptoms or signs of urinary infection. Asymptomatic bacteriuria is common, but its prevalence varies widely with age, gender, and the presence of genitourinary abnormalities or underlying diseases. For example, the prevalence of bacteriuria increases with age in healthy females from as low as about 1% among school-age females to at least 20% among females 80 years of age or older living in the community, whereas bacteriuria is rare in healthy young males. Because its clinical significance was controversial (asymptomatic bacteriuria precedes UTI but does not always lead to asymptomatic infection), guidelines have been published for the diagnosis and treatment of asymptomatic bacteriuria in adults older than 18 years of age. The foundation of these guidelines rests on the premise that screening of asymptomatic subjects for bacteriuria is appropriate if bacteriuria has adverse outcomes that can be prevented by antimicrobial therapy. Thus, screening and treatment for asymptomatic bacteriuria are recommended for pregnant females (because of the risk of progression to severe symptomatic UTI and possible harm to the fetus), males undergoing transurethral resection of the prostate, and individuals undergoing urologic procedures for which mucosal bleeding is anticipated. In contrast, screening for or treatment of asymptomatic bacteriuria is not recommended for premenopausal, nonpregnant females; diabetic females;

1022 PA RT V I I     Diagnosis by Organ System

older persons living in the community; older institutionalized adults; persons with spinal cord injury; or catheterized patients while the catheter is in place. 

with acute pyelonephritis are bacteremic. Acute papillary necrosis of one or more renal pyramids (cone-shaped tissue) may occur as a complication associated with pyelonephritis. 

Cystitis

Urosepsis

Typically, patients with cystitis (infection of the bladder) complain of dysuria, frequency, and urgency (compelling need to urinate). These symptoms are a result not only of inflammation of the bladder but also of multiplication of bacteria in the urine and urethra. Often, there is tenderness and pain over the area of the bladder. In some individuals, the urine is grossly bloody. The patient may note urine cloudiness and a bad odor. Because cystitis is a localized infection, fever and other signs of systemic illness are usually not present. 

Approximately 25% of sepsis cases (severe blood infection) are a result of urosepsis, a systemic infection that may develop from community-, hospital-, or health care–associated UTIs. Urosepsis is defined as evidence of a UTI and two or more additional signs including an elevated temperature (>38°C), an elevated heart rate (>90 beats/min), an increased respiratory rate (>20 breaths/min or a PCO2 of 12,000/mm3, 10% neutrophilic band forms). Early diagnosis and treatment of UTIs are essential in preventing urosepsis. 

Acute Urethral Syndrome Another UTI is acute urethral syndrome. Patients with this syndrome are primarily young, sexually active females, who experience dysuria, frequency, and urgency but yield fewer organisms than 105 colony-forming units of bacteria per milliliter (CFU/mL) urine on culture. The criterion of greater than 105 CFU/mL of urine is highly indicative of infection in most patients with UTIs. Almost 50% of all females who seek medical attention for complaints of symptoms of acute cystitis fall into this group. Although C. trachomatis and N. gonorrhoeae urethritis, anaerobic infection, genital herpes, and vaginitis account for some cases of acute urethral syndrome, most of these females are infected with organisms identical to those that cause cystitis but in numbers less than 105 CFU/mL of urine. A cutoff of 102 CFU/ mL, rather than 105 CFU/mL, must be used for this group of patients, but concomitant pyuria (presence of eight or more leukocytes per cubic millimeter on microscopic examination of uncentrifuged urine) must also be present. Approximately 90% of these females have pyuria, an important discriminatory feature of infection. 

Pyelonephritis Pyelonephritis refers to inflammation of the kidney parenchyma, calices (cup-shaped division of the renal pelvis), and pelvis (upper end of the ureter that is located inside the kidney) and is usually caused by bacterial infection. Pyelonephritis may appear as an acute or chronic condition. Acute pyelonephritis presents with enlarged kidneys that contain surface abscesses. Chronic pyelonephritis presents with scarring on one or both kidneys and interstitial fibrosis on the pelvic wall. An inflammatory infiltrate of white blood cells, predominantly lymphocytes, is typically present. In addition, the tubules in the kidneys may either be dilated or constricted and contain colloid casts (crystalized mucous secretions). The typical clinical presentation of an upper UTI includes fever and flank (lower back) pain and, frequently, lower tract symptoms (frequency, urgency, and dysuria). Patients can also exhibit systemic signs of infection such as vomiting, diarrhea, chills, increased heart rate, and lower abdominal pain. Of significance, 40% of patients

Laboratory Diagnosis of Urinary Tract Infections As previously mentioned, because noninvasive methods for collecting urine must rely on a specimen that has passed through a contaminated milieu, quantitative cultures for the diagnosis of UTI are used to discriminate between contamination, colonization, and infection. Refer to Table 5.1 for a quick reference for collecting, transporting, and processing urinary tract specimens.

Specimen Collection Prevention of contamination by normal vaginal, perineal, and anterior urethral microbiota is the most important consideration for the collection of a clinically relevant urine specimen.

Clean-Catch Midstream Urine The least invasive and preferred routine collection procedure, the clean-catch midstream urine specimen collection, must be performed carefully for optimal results, especially with female patients. Good patient education is essential. Guidelines for proper specimen collection should be prepared on a printed card (bilingual, if necessary), with the procedure clearly described and preferably illustrated to help ensure patient compliance. The patient should be instructed to wash their hands before cleaning the periurethral area, wiping from front to back three times, each time with a clean sterile gauze pad soaked with a mild detergent to prevent contamination. Of importance, the patient should also be instructed to rinse well with two or more sponges soaked in sterile distilled water to remove the detergent, which may be bacteriostatic. Once cleansing is completed, the patient should retract the labial folds or glans penis, begin to void, and then collect a midstream urine sample. Studies show that uncleansed, first-void specimens from males are as sensitive as (but less specific than) midstream urine specimens. Sterile bags may be used for infants and children. 

CHAPTER 72  Infections of the Urinary Tract

Straight Catheterized Urine Although slightly more invasive, urinary catheterization provides a method for the collection of uncontaminated urine from the bladder in uncooperative patients or patients unable to void because of other underlying physiologic conditions. Either a physician or another trained health professional performs this procedure. Risk exists, however, that urethral organisms will be introduced into the bladder with the catheter. 

Suprapubic Bladder Aspiration After preparation of the skin, urine is withdrawn directly into a syringe through a percutaneously inserted needle during suprapubic bladder aspiration, thereby ensuring a contamination-free specimen. The bladder must be full before the procedure is performed. This collection technique may be indicated in certain clinical situations, such as pediatric practice, when urine is difficult to obtain. If good aseptic techniques are used, this procedure can be performed with little risk in premature infants, neonates, small children, and pregnant women and other adults with full bladders. 

Indwelling Catheter Patients who are housed in hospitals and long-term care facilities and those treated in other health care–associated settings such as outpatient clinics for cancer and transplant patients are more frequently required to use indwelling urinary catheters. These patients are very likely to develop bacteriuria, which predisposes them to more severe infections. Specimen collection from patients with indwelling catheters requires scrupulous aseptic technique. Health care workers who manipulate a urinary catheter in any way should wear gloves. The catheter tubing should be clamped off above the port to allow the collection of freshly voided urine. The catheter port or wall of the tubing should then be cleaned vigorously with 70% ethanol, and urine should be aspirated via a needle and syringe; the integrity of the closed drainage system must be maintained to prevent the introduction of organisms into the bladder. Specimens obtained from the collection bag are inappropriate, because organisms can multiply there, obscuring the true relative numbers. Cultures should be obtained when patients are ill; routine monitoring does not yield clinically relevant data. 

Specimen Transport Because it is an excellent supportive medium for the growth of most bacteria, urine must be immediately refrigerated or preserved. Bacterial counts in refrigerated (4°C) urine remain constant for as long as 24 hours. Urine transport tubes (BD Urine Culture Kit [Becton Dickinson Vacutainer Kits, Franklin Lakes, NJ]) containing boric acid, sodium borate, and sodium formate have been shown to preserve bacteria without refrigeration for as long as 48 hours when more than 105 CFU/mL (100,000 organisms per milliliter) are present in the initial urine specimen. The system may inhibit the growth of certain organisms, and it must be used

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with a minimum of 3 mL of urine. Boric acid products preserve bacterial viability in urine in the absence of antibiotics. For patients from whom colony counts of organisms of less than 100,000/mL might be clinically significant, plating within 2 hours of collection is recommended. The kit provides a convenient method for preserving and transporting urine from remote areas where refrigeration is not practical. 

Screening Procedures As many as 60% to 80% of all urine specimens received for culture by the acute care medical center laboratory may contain no etiologic agents of infection or contain only contaminants. Procedures developed to quickly identify those urine specimens that will be negative on culture and circumvent excessive use of media, technical staff, and the overnight incubation period are discussed in this section. A reliable screening test for the presence or absence of bacteriuria provides physicians important same-day information that a conventional urine culture may take a day or longer to provide. Many screening methods have been advocated for use in detecting bacteriuria and/or pyuria. Red blood cells or erythrocytes identified in the urine, hematuria, may also indicate UTI, but this occurs in a variety of other physiologic disorders. White blood cell casts in urine are strong evidence of pyelonephritis but can also be associated with renal disease in the absence of infection. In addition, elevated levels of protein (105 CFU/mL— extrapolated as one bacterium per microscopic field in an uncentrifuged sample). The absence of bacteria in a stained sediment from a centrifuged sample (5 minutes at 2000 rpm) indicates the probability that the specimen contains less than 104 bacteria/mL. The Gram stain should not be relied on for detecting polymorphonuclear leukocytes in urine, because leukocytes deteriorate quickly in urine that is not fresh or not adequately preserved. Many microbiologists have not adopted Gram stain examination of urine specimens because of its unreliability in detecting lower yet clinically significant numbers of organisms and because of its labor intensity. If used, urine Gram stain should be limited to patients with acute pyelonephritis, patients with invasive

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UTIs, or other patients for whom immediate information is necessary for appropriate clinical management. 

Pyuria Pyuria (10 leukocytes/mm3, using a hematocytomer from a clean-catch midstream specimen) is the hallmark of inflammation, and the presence of polymorphonuclear neutrophils (PMNs) can be detected and enumerated in uncentrifuged specimens. This method of screening urine correlates well with the number of PMNs (neutrophils) excreted per hour, the best indicator of the host’s state. Patients with more than 300,000 PMNs excreted into the urine per hour are likely to have a current infection. The standard urinalysis (usually completed in the laboratory hematology or chemistry sections) includes an examination of the centrifuged sediment of urine for enumeration of PMNs, results of which do not correlate well with either the PMN excretion rate or the presence of infection. Pyuria also can be associated with other clinical diseases, such as vaginitis, and therefore is not specific for UTIs. 

Indirect Indices Screening tests commonly detect bacteriuria or pyuria by examining for the presence of bacterial enzymes or PMN enzymes rather than the organisms or PMNs themselves. Nitrate Reductase (Griess) Test

The nitrate reductase (Griess) test looks for the presence of urinary nitrite, an indicator of UTI. Nitrate-reducing enzymes that are produced by the most common urinary tract pathogens reduce nitrate to nitrite. This test has been incorporated onto a urinary dipstick that also tests for leukocyte esterase—an enzyme produced by PMNs (discussed next). Liquid-chromatography tandem mass spectrometry (LC-MS/MS) has been used to screen urine for the presence of nitrate and nitrite for the diagnosis of UTIs. This test is not sensitive to variations in urine chemistry and has a demonstrated specificity (91%) and sensitivity (95%), which is greater than using the nitrate reductase test.  Leukocyte Esterase Test

As previously mentioned, evidence of a host response to infection is the presence of PMNs in the urine. Because inflammatory cells produce leukocyte esterase, a simple, inexpensive, and rapid method that measures this enzyme has been developed. Studies have shown that leukocyte esterase activity correlates with hemocytometer chamber counts. The nitrate reductase and leukocyte esterase tests have been incorporated into the urinary dipstick. Numerous manufacturers sell these strips commercially, and the strips are one of the most widely used enzymatic tests. Although the sensitivity of the combination strip is higher than either test alone, the sensitivity of this combination screening is not great enough to recommend its use as a standalone test in most circumstances. Of note, the leukocyte esterase test is not sensitive enough for determining pyuria in patients with acute urethral syndrome. 

Catalase

The Accutest Uriscreen (JANT Pharmacal Corp., Encino, CA) is another rapid urine-screening system based on the detection of catalase present in human somatic cells and in most bacterial species that commonly cause UTIs except for streptococci and enterococci. Approximately 1.5 to 2 mL of urine is added to a tube containing dehydrated substrate. Hydrogen peroxide is added to the urine, and the solution is mixed gently. The formation of bubbles above the liquid surface is interpreted as a positive test. Visible results are available in approximately 2 minutes. 

Automated and Semiautomated Systems Automated screening systems offer a large throughput with minimal labor and a rapid turnaround time compared with conventional cultures. Various automated or semiautomated urine-screening systems are commercially available, such as the Iris Urinalysis System (Beckman-Coulter, Inc., Brea, CA), and can analyze a urine or body fluid sample in one instrument. The instrument analyzes both the microscopic components and the urine chemistries by combining technology of both types of analyzers into one automated system. The Iris System uses a flow-imaging microscopy method to capture individual images of each particle identified in the specimen and then classifies the particle using Auto-Particle Recognition (APP) software. Siemens Medical Solutions USA, Inc. (Malvern, PA) manufactures a wide range of CLINITEK automated/ semiautomated urine analyzers that include point-of-care to high-throughput walkaway instruments. In addition, the Sysmex UN-2000 (Lincolnshire, IL) uses flow cytometry and specific fluorescent dyes for the physical and chemical identification of organisms and other particles in urine samples. Various studies have indicated that automated urinalysis instrumentation has demonstrated limitations and continue to recommend manual microscopic analysis of urine samples for the diagnosis of UTI. 

General Comments Regarding Screening Procedures In general, screening methods are insensitive at levels below 105 CFU/mL. Therefore, they are not acceptable for urine specimens collected by suprapubic aspiration, catheterization, or cystoscopy. Screening methods may also fail to detect a significant number of infections in symptomatic patients with low colony counts (102 to 103 CFU/mL), such as young, sexually active females with acute urethral syndrome. Further complicating the laboratory’s decision as to whether to adopt a screening method is whether screening results will be used to rule out infection in asymptomatic patients. Under these circumstances, testing for pyuria is essential. Therefore, given the importance of the 102 CFU/mL count and the PMN count, no screening test should be used indiscriminately. Selecting a screening method largely depends on the laboratory and the patient population being served by the laboratory. For example, there will

CHAPTER 72  Infections of the Urinary Tract

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be a cost advantage in screening urine in laboratories that receive many culture-negative specimens. On the other hand, urine from patients with symptoms of UTI plus a selected group expected to have asymptomatic bacteriuria should be cultured. For example, patients in their first trimester of pregnancy should be cultured, because these patients might appear asymptomatic but have a covert infection and become symptomatic later; UTIs in pregnant females may lead to pyelonephritis and the likelihood of a premature birth. Other situations in which patients with no symptoms of UTI might be cultured include the following: • Bacteremia of unknown source • Urinary tract obstruction • Follow-up after removal of an indwelling catheter • Follow-up of previous therapy Other factors that must be considered when selecting a rapid urine screen include accuracy, ease of test performance, reproducibility, turnaround time, and whether bacteriuria or pyuria is detected. 

Urine Culture Inoculation and Incubation of Urine Cultures Once it has been determined that a urine specimen should be cultured for isolation of the common agents of UTI, a measured amount of urine is inoculated to each of the appropriate media. The urine should be mixed thoroughly before plating. The plates can be inoculated using a calibrated loop designed to deliver a known volume, either 0.01 or 0.001 mL of urine. These loops, made of platinum, plastic, or other material, can be obtained from laboratory supply companies. The calibrated loop that delivers the larger volume of urine (0.01 mL) is recommended to detect lower numbers of organisms in certain specimens. For example, urine collected from catheterization, nephrostomies, ileal conduits, and suprapubic aspirates should be plated with the larger calibrated loop. The communication of pertinent clinical history to the laboratory is essential so that appropriate processing can be performed. The choice of media to inoculate depends on the patient population served and the microbiologist’s preference. The use of a 5% sheep blood agar plate and a MacConkey agar plate allows the detection of most gram-negative bacilli, staphylococci, streptococci, and enterococci. To save cost and somewhat streamline culture processing, many laboratories use an agar plate split in half (biplate); one side contains 5% sheep blood agar and the other half contains MacConkey or Eosin Methylene Blue agar (Fig. 72.2). In some circumstances, enterococci and other streptococci may be obscured by heavy growth of Enterobacterales. Because of this possibility, some laboratories add a selective plate for gram-positive organisms, such as Columbia colistin-nalidixic acid agar (CNA) or Enterococcosel Agar (Bile Esculin Azide Agar, Becton-Dickinson, Sparks, MD). Although some discriminatory capability may be added, cost is also added to the procedure. In addition

• Fig. 72.2  Cronobacter sakazakii isolated from a urinary sample on a 5% sheep blood agar/eosin methylene blue biplate.

to increased cost, inclusion of plated media selective for gram-positive organisms generally provides no or limited additional information. Many European laboratories use cystine-lactose electrolyte-deficient (CLED) agar. CLED agar does not contain sodium chloride, inhibiting the characteristic swarming of Proteus spp., but still supports adequate growth of most common urinary pathogens. In recent years, chromogenic media have been introduced and become commercially available from several manufacturers, allowing more specific direct detection and differentiation of urinary tract pathogens on primary plates, such as BD CHROMagar (Becton Dickson, Heidelberg, Germany). This medium uses enzymatic reactions to ­ identify E. coli and Enterococcus without additional confirmatory testing from urine specimens and provides presumptive identification of S. saprophyticus, Streptococcus agalactiae, Klebsiella-Enterobacter-Serratia, and the ProteusMorganella-Providencia groups. Before inoculation, urine is mixed thoroughly, and the top of the container is then removed. The calibrated loop is inserted vertically into the urine in a cup. Otherwise, more than the desired volume of urine will be taken up, potentially affecting the quantitative culture result (Fig. 72.3). A widely used method is described in Evolve Procedure 72.1. Once inoculated, the plates are streaked to obtain isolated colonies (Fig. 72.4). Once plated, urine cultures are incubated overnight at 35°C. Incubation for a minimum of 24 hours is typically necessary to detect uropathogens. Thus, some specimens inoculated late in the day cannot be read accurately the next morning. These cultures should either be reincubated until the next day or interpreted later in the day when a full 24-hour incubation has been completed. 

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Loop is touched to the center of the plate, from which the inoculum is spread in a line across the diameter of the plate.

• Fig. 72.3  Method for inserting a calibrated loop into urine to ensure that the proper amount of specimen adheres to the loop.

Without flaming or reentering urine, loop is drawn across the entire plate, crossing the first inoculum streak numerous times to produce isolated colonies.

• Fig. 72.4  Method for streaking with calibrated urine loop to produce isolated colonies and countable colony-forming units.

Interpretation of Urine Cultures As previously mentioned, UTIs may be completely asymptomatic, produce mild symptoms, or cause life-threatening infections. Of importance, the criteria most useful for microbiologic assessment of urine specimens is dependent not only on the type of urine submitted (e.g., voided, straight catheterization) but the clinical history of the patient (e.g., age, sex, symptoms, antibiotic therapy). One major problem in interpreting urine cultures arises because urine cultures collected by the voided technique may be contaminated with normal microbiota, including Enterobacterales. Determining what colony count represents true infection from contamination is of utmost importance and is related to the patient’s clinical presentation. Several studies have proposed the use of different cutoffs in colony counts based on clinical presentation; an example of one such set of guidelines is given in Table 72.1. Ideally, the clinician caring for the patient should provide the laboratory with enough clinical information to allow specimens from different patient populations to be identified. These specimens could then be selectively processed using the guidelines in Table 72.1. However, because microbiology laboratories frequently receive little or no clinical information about patients, questions have been raised as to whether these cutoffs are practical and realistic for routine laboratory use. Further complicating urine culture

interpretation is the increasing difficulty in distinguishing between infection and contamination as the criterion for a positive culture is lowered from 105 CFU/mL to 102 CFU/ mL. Because of these issues, many laboratories establish their own interpretative criteria for urine cultures based on the type of urine submitted (e.g., clean-catch midstream, catheterized, or surgically obtained specimens such as suprapubic aspirates). Variations in interpretative guidelines occur from one laboratory to another, but some generalities can be made; these are listed in Table 72.2. Some examples of urine culture results are shown in Fig. 72.5 to illustrate some of these interpretations. See the Evolve site for a semiquantitative procedure for the inoculation of urine cultures. In addition to the previously described guidelines, a pure culture of S. aureus is significant regardless of the number of CFUs, and antimicrobial susceptibility tests are performed. The presence of yeast in any number is reported to physicians, and pure cultures of yeast may be identified to the species level. In all urine, regardless of the extent of final workup, all isolates should be enumerated (e.g., three different organisms present at 103 CFU/mL), and those present in numbers greater than 104 CFU/mL should be described morphologically (e.g., non–lactose fermenting gram-negative rods).

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

CHAPTER 72  Infections of the Urinary Tract

TABLE 72.1    Criteria for Classification of Urinary Tract Infections by Clinical Syndrome

Category

Clinical

Laboratory Results

Acute, uncomplicated UTI in females

Dysuria, urgency, frequency, suprapubic pain No urinary symptoms in last 4 weeks before the current episode No fever or flank pain

≥10 WBC/mm3 ≥103 CFU/mL uropathogensa in CCMS urine

Acute, uncomplicated pyelonephritis

Fever, chills Flank pain on examination Other diagnoses excluded No history or clinical evidence of urologic abnormalities

≥10 WBC/mm3 ≥104 CFU/mL uropathogens in CCMS urine

Complicated UTI and UTI in males

Any combination of symptoms listed above. One or more factors associated with complicated UTIb

≥10 WBC/mm3 ≥105 CFU/mL uropathogens in CCMS urine

Asymptomatic bacteriuria: female patients

No urinary symptoms

± >10 WBC/mm3 ≥105 CFU/mL in two CCMS cultures >24 h apart

Asymptomatic bacteriuria: male patients

No urinary symptoms

± >10 WBC/mm3 ≥103 CFU/mL (suggestive) ≥105 CFU/mL (definitive) in one CCMS

aUropathogens:

Organisms that commonly cause UTIs. associated with complicated UTI include any UTI in a male patient, indwelling or intermittent urinary catheter, more than 100 mL of postvoid residual urine, obstructive uropathy, urologic abnormalities, azotemia (excess urea in the blood, even without structural abnormalities), and renal transplantation. CCMS, Clean-catch midstream urine; CFU, colony-forming unit; UTI, urinary tract infection; WBC, white blood cells. Data from Stamm WE. Criteria for the diagnosis of urinary tract infection and for the assessment of therapeutic effectiveness. Infection 20(suppl 3):S151, 1992; and Bennett J, Dolin R, Blaser M. Principles and Practice of Infectious Diseases. 8th ed. Philadelphia: Elsevier-Saunders; 2015. bFactors

TABLE 72.2    Suggested Interpretative Guidelines for Urine Cultures

Result

Specific Specimen Type/Associated Clinical Condition, if Known

≥104 CFU/mL of a single potential pathogen or for each of two potential pathogens

CCMS urine/pyelonephritis, acute cystitis, asymptomatic bacteriuria, or catheterized urines

Completea

≥103 CFU/mL of a single potential pathogen

CCMS urine/symptomatic male patients or catheterized urine or acute urethral syndrome

Complete

≥ Three organism types with no predominating organism

CCMS urine or catheterized urine

None; because of possible contamination, ask for another specimen

Either two or three organism types with predominant growth of one organism type and 105 CFU/mL of urine b. A colony bacterial count of >104 CFU/mL of urine c. A colony bacterial count of >102 CFU/mL of urine d. A colony bacterial count of >103 CFU/mL of urine 6. Which of the following tools have been used in the diagnosis of UTIs to discriminate among contamination, colonization, and infection? a. Quantitative urine cultures b. Qualitative urine cultures c. Observation of cloudy, foul-smelling urine d. Routine monitoring of nonsymptomatic patients 7. When setting up a urine culture, a calibrated loop is used that delivers a specific amount of urine to the media plate. What is that amount? a. 0.01 or 0.001 mL of urine b. 0.10 or 0.01 mL of urine c. 0.001 or 0.0001 mL of urine d. None of the above

8. True or False _____ A pure culture of S. aureus is significant regardless of the number of CFUs. _____ Screening for asymptomatic bacteriuria is recommended for nonpregnant females, diabetic females, older institutionalized subjects, patients with spinal cord injury, and catheterized patients. _____ S. aureus adheres better to uroepithelial cells than S. saprophyticus or S. epidermidis. _____ The hematogenous spread of urinary tract infections does not usually occur due to bacteremia. _____ Urinary tract infections can lead to sepsis and are the most common health care–associated infection reported from hospitals and nursing homes in the United States. _____ Salmonella spp. may be recovered from urine during the early stages of typhoid fever and should be reported immediately to the doctor. _____ Factors that may decrease bacterial contamination of urine cultures are rapid transportation to the laboratory, refrigeration of urine in cases where there is testing delay, and having female patients cleanse the labia and collect a “midstream” specimen. _____ Viruses are a common cause of urinary tract infection, and urine cultures that fail to grow on bacterial media should be inoculated into cell culture. _____ Females more frequently develop UTIs by the ascending route of bacterial invasion because of the association with instrumentation (catheterization and cystoscopy). _____ When collecting a urine specimen from patients with indwelling catheters, it is appropriate to obtain the specimen from the collection bag of the catheter. _____ The calibrated loop used to inoculate urine cultures where lower numbers of organisms are suspected is the 0.001-mL volume loop.

CHAPTER 72  Infections of the Urinary Tract

9. Matching: Match each term with the correct description. _____ reflux _____ urethra _____ ureter _____ bacteriuria _____ Foley catheter _____ hematogenous _____ pyelonephritis _____ cystitis _____ urethritis _____ pyuria _____ systemic _____ GBS _____ UPEC

a. presence of bacteria in urine b. affecting the body as a whole c. group B streptococci d. many leukocytes (white blood cells) in the urine e. tube connecting the bladder and kidney f. blood-borne g. indwelling catheter h. tube allowing urine to leave the bladder i. infection of the kidney j. uropathogenic E. coli k. inflammation of the urethra l. inflammation of the bladder m. backward flow

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10.  Short Answer (1) Explain the significance of the bacterial K-antigen. (2)  Name the single most important factor contributing to health care–associated urinary tract infections. (3)  What bacteria are normally involved in UTIs occurring by the hematogenous route? (4) What anatomic feature of the urinary tract helps to prevent urinary tract infections? (5)  Define Tamm-Horsfall protein and defensins. How do these proteins aid in fighting off infection by uropathogens? (6) Explain what the acronym “UPEC” stands for. (7) What are pathogenicity islands? (8) What might alert the microbiologist to an unusual cause of a urinary tract infection?

73

Genital Tract Infections OBJECTIVES 1. Describe the basic anatomy of the male and female reproductive systems. 2. Define the following conditions: vaginitis, cervicitis, proctitis, bartholinitis, pelvic inflammatory disease (PID), epididymitis, prostatitis, orchitis, neoplasia, urethritis, and dysuria. 3. List microorganisms that commonly are associated with vaginitis, cervicitis, and PID. 4. Describe the normal microbiota of the male and female genital tracts, and differentiate normal flora from pathogenic organisms. 5. List the media used to selectively isolate and differentiate genital tract pathogens, including modified Thayer-Martin (MTM), New York City (NYC), and colistin nalidixic agar (CNA), and the organisms capable of growth on each. 6. Compare the recommended use of various collection swabs for genital tract specimens, including the organisms inhibited by each (cotton-tipped, with or without charcoal; calcium alginate; dacron; rayon). 7. Determine specimen acceptability based on collection, transport, and diagnostic test orders for a genital tract specimen. 8. Explain the significance of gram-negative intracellular diplococci in genital specimens from both men and women. 9. Correlate signs and systems of infections with the results of laboratory diagnostic procedures for identification of the etiologic agent associated with infections of the genital tract.

General Considerations Anatomy Familiarity with the anatomic structures is important for appropriate processing of specimens from genital tract sites and interpretation of microbiologic laboratory results. The key anatomic structures for the female and male genital tract in relation to other important structures are shown in Fig. 73.1. The female reproductive system consists of two main parts: the uterus and the ovaries. The uterus produces vaginal and uterine secretions and is the location where the human fetus grows and matures during reproduction. The ovaries connect to the uterus and the fallopian tubes. The

ovaries produce the female eggs that pass through the fallopian tubes and will imbed in the uterus when fertilized by the male sperm. The uterus connects to the vaginal opening through the cervix. The male reproductive system, unlike the female, consists of several organs that are located external to the abdominal cavity. The main organs consist of the penis and the testes that produce the semen and sperm for fertilization of the female egg. The sperm is stored in a small gland coiled around the testis, the epididymis. The prostate gland surrounds the ejaculatory duct and produces semen, prostatic fluid, and seminal fluid. 

Resident Microbiota The lining of the human genital tract consists of a mucosal layer of transitional (cells capable of undergoing shape change or transitions), columnar (ciliated and longer than wide), and squamous epithelial (thin and flat) cells. Various species of commensal bacteria colonize these surfaces, causing no harm to the host except under abnormal circumstances. The colonization of the surface by resident microbiota produces a biologic barrier preventing the adherence of pathogenic organisms. Normal urethral microbiota includes coagulase-negative staphylococci and corynebacteria, as well as various anaerobes. The vulva and penis, especially the area underneath the prepuce (foreskin) of the uncircumcised male, may harbor Mycobacterium smegmatis along with other gram-positive bacteria. The microbiota of the female genital tract varies with the pH and estrogen concentration of the mucosa, which depend on the host’s age. Prepubescent and postmenopausal females primarily harbor staphylococci and corynebacteria (the same microbiota present on surface epithelium); whereas females of reproductive age may harbor large numbers of facultative bacteria, such as Enterobacterales, streptococci, and staphylococci, as well as anaerobes, such as lactobacilli, anaerobic non–spore-forming bacilli and cocci, and clostridia. Lactobacilli are the predominant organisms in secretions in normal, healthy vaginas. The lactobacilli present in vaginal secretions metabolize glucose to lactic acid, resulting in a pH of approximately 4.0. The acidic pH coupled with the organism’s ability to produce hydrogen peroxide prevents infection by exogenous sexually transmitted pathogens. Many women carry group B beta-hemolytic 1029

1030 PA RT V I I     Diagnosis by Organ System

Ovary Posterior cul-de-sac

Fallopian tube Uterus Anterior cul-de-sac Urinary bladder

Urinary bladder

Rectum

Symphysis pubis

Seminal vesicle Symphysis pubis

Cervix Fornix of vagina

Prostate gland

Ejaculatory duct

Urethra

Anus Bartholin gland

A

Urethra Vulva

Anus

Epididymis

Vagina

Testis

Glans

B • Fig. 73.1  Location of key anatomic structures of the female (A) and male (B) genital tracts in relation to other major anatomic structures.

streptococci (Streptococcus agalactiae), which may be transmitted to the neonate. Although yeasts (acquired from the gastrointestinal tract) may be transiently recovered from the female vaginal tract, they are not considered normal microbiota. 

Sexually Transmitted Diseases and Other Genital Tract Infections Genital tract infections may be classified as endogenous or exogenous. Exogenous infections may be acquired as people engage in sexual activity, and these infections are referred to as sexually transmitted diseases (STDs). In contrast, endogenous infections result from normal genital microbiota. Female genital tract infections can be divided into lower genital tract (vulva, vagina, and cervix) and upper genital tract (uterus, fallopian tubes, ovaries, and abdominal cavity) infections. Lower genital tract infections are commonly acquired by sexual or direct contact. Although the organisms that cause lower genital tract infections are not usually part of the normal genital tract microbiota, some organisms normally present in very low numbers can increase sufficiently to cause disease. Upper genital tract infections are frequently an extension of a lower tract infection in which organisms from the vagina or cervix travel into the uterine cavity and on through the endometrium to the fallopian tubes and ovaries. Similarly, an organism can spread along contiguous mucosal surfaces in the male from a lower genital tract site of infection (i.e., urethra) and cause infection in a reproductive organ, such as the epididymis. 

Genital Tract Infections Sexually Transmitted Diseases and Other Lower Genital Tract Infections Lower genital tract infections may be acquired either through sexual contact with an infected partner or through nonsexual means. These infections are some of the most common infectious diseases.

Epidemiology and Etiologic Agents STDs or sexually transmitted infections (STIs) are major public health problems in all populations and socioeconomic groups worldwide. The incidence and spread of STDs are greatly influenced by numerous factors, such as the availability of multiple sexual partners, the presence of asymptomatic infection, the frequent movement of people within populations, and increasing affluence. The number of microorganisms that can cause genital tract infections is large. These organisms are diverse, representing all four major groups of microorganisms: bacteria, viruses, fungi, and parasites. The major causes of genital tract infections are listed in Table 73.1. 

Routes of Transmission Although genital tract infections can be caused by members of the patient’s genital microbiota (endogenous infections), the overwhelming majority of lower genital tract infections are sexually transmitted. Sexually Transmitted

Chlamydia trachomatis (CT), Neisseria gonorrhoeae (GC), Trichomonas vaginalis, human immunodeficiency virus

CHAPTER 73  Genital Tract Infections

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TABLE 73.1    Major Causes of Genital Tract Infections and Sexually Transmitted Diseases

Frequency

Disease

Agent

Organism Group

More Common

Genital and anal warts (condyloma); cervical dysplasia; cancer

Human papillomavirus

Viruses

Vaginitis

Gardnerella/Mobiluncus, Trichomonas vaginalis, Candida albicans

Bacteria, parasites, fungi

Urethritis/cervicitis (also acute salpingitis, acute perihepatitis, urethritis, pharyngitis)

Neisseria gonorrhoeae, Chlamydia trachomatis, Ureaplasma urealyticum

Bacteria

Herpes genitalis (genital/skin ulcers)

Herpes simplex virus type 2 (less commonly type 1)

Viruses

AIDS

Human immunodeficiency virus (HIV)

Viruses

Hepatitis (acute and chronic infection)

Hepatitis B virus

Viruses

Lymphogranuloma venereum

C. trachomatis (L-1, L-2, L-3 serovars)

Bacteria

Granuloma inguinale

Klebsiella granulomatis (Donovania)

Bacteria

Syphilis

Treponema pallidum

Bacteria

Chancroid

Haemophilus ducreyi

Bacteria

Scabies, mites

Sarcoptes scabiei

Ectoparasites

Pediculosis pubis, “crabs” infestation

Phthirus pubis

Ectoparasites

Enteritis (homosexuals/proctitis)

Giardia duodenalis, Entamoeba histolytica, Shigella spp., Salmonella spp., Enterobius vermicularis, Campylobacter spp., Helicobacter spp.

Bacteria, parasites

Molluscum contagiosum

Poxlike virus

Viruses

Heterophile-negative mononucleosis, congenital infections

Cytomegalovirus

Viruses

Less Common

(HIV), Treponema pallidum, Ureaplasma urealyticum, Mycoplasma hominis, other mycoplasmas, herpes simplex virus (HSV), and others may be acquired during sexual activity. In addition, other agents that cause genital tract disease and may be sexually transmitted include adenovirus, hepatitis B, human T-cell lymphotropic virus (HTLV), coxsackie virus, molluscum contagiosum virus (a member of the poxvirus group), the human papillomaviruses (HPVs) of genital warts (condylomata acuminata; types 6, 11, and others) and those associated with cervical carcinoma (predominantly types 16 and 18, but numerous others are also implicated), Klebsiella granulomatis, and ectoparasites, such as scabies and lice. Some of these agents are not routinely isolated from clinical specimens. Infections with more than one agent may occur; therefore, dual or concurrent infections should always be considered. An individual’s sexual habits and practices dictate potential sites of infection. Homosexual practices and increasingly common heterosexual practices of anal-genital or oral-genital intercourse allow for transmission of a genital tract infection to other body sites, such as the pharynx or anorectic

region. In addition, these practices have required that other gastrointestinal and systemic pathogens also be considered etiologic agents of STDs. The intestinal protozoa Giardia duodenalis, Entamoeba histolytica, and Cryptosporidium spp. are significant causes of STDs, especially among homosexual populations. In the same group of patients, fecal pathogens, such as Salmonella, Shigella, Campylobacter, and Microsporidium are often transmitted sexually. Oral-genital practices may provide an opportunity for N. meningitidis to colonize and infect the genital tract. In fact, N. meningitidis is increasingly being recognized as a pathogen associated with urethritis. Outbreaks with N. meningitidis have been predominately identified in heterosexual males. Viruses shed in secretions or present in blood (cytomegalovirus [CMV]; hepatitis B, and possibly C and E; other non-A, non-B hepatitis viruses; HTLV type I [HTLV-I]; and HIV) are spread by sexual practices. Certain infections that are sexually transmitted occur on the surface epithelium of or near the lower genital tract. The major pathogens of these types of infections include HSV, Haemophilus ducreyi, and T. pallidum. 

1032 PA RT V I I     Diagnosis by Organ System

Other Routes

Organisms may also be introduced into the genital tract by instrumentation, presence of a foreign body, or chemical or immunologic processes that cause irritation and can subsequently cause infection. These infections are often a result of infection with the same organisms capable of causing skin or wound infections. Infection can also be transmitted from mother to infant either in vivo or during delivery. For example, transplacental infection may occur with syphilis, HIV, CMV, or HSV. Infection in the newborn can also be acquired during delivery by direct contact with an infectious lesion or discharge in the mother and a susceptible mucous membrane, such as the eye in the infant. STDs such as HSV, C. trachomatis, and N. gonorrhoeae may be transmitted from mother to newborn in this manner. Other organisms, such as group B streptococci, Escherichia coli, and Listeria monocytogenes originating from the mother may also be transmitted to the infant before, during, or after birth. (Infections in the fetus and newborn are discussed later in this chapter.) 

caused by common urinary tract infection isolates, adenoviruses, C. trachomatis, T. vaginalis (less frequently), and genital mycoplasmas, such as M. hominis, Mycoplasma genitalium, and U. urealyticum.  Lesions of the Skin and Mucous Membranes

Although symptoms of genital tract infections generally cause the patient to seek medical attention, a patient with an STD—especially a female—may be free of symptoms (i.e., asymptomatic). For example, gonorrhea (N. gonorrhoeae) or chlamydia (C. trachomatis) infection is usually obvious in males because of a urethral discharge, yet females with either or both infections may have either minimal symptoms or no symptoms at all. Also, the primary lesion of syphilis (chancre) can be unremarkable and go unnoticed by the patient. Therefore, the lack of symptoms does not guarantee the absence of disease. Unfortunately, these asymptomatic individuals can serve as reservoirs for infection and unknowingly spread the pathogen to other individuals. Asymptomatic infections in females caused by N. gonorrhoeae or C. trachomatis that go untreated can lead to serious sequelae, such as pelvic inflammatory disease (PID) or infertility. 

Numerous organisms can cause genital lesions that are diverse in both their appearance and their associated symptoms (Fig. 73.2) but are most often associated with STDs. The agents and their features of infection are summarized in Table 73.2. Some of these infections, such as genital herpes (caused by HSV) or genital warts (caused by HPVs and discussed in Chapter 65), are common; whereas others, such as lymphogranuloma venereum and granuloma inguinale, are uncommon in the United States. Genital skin and mucous membrane infections are often polymicrobial, making the diagnosis difficult. In addition, the characteristics of the lesions may vary from one type of infectious process to another for the same organisms. For example, specific HPV genotypes infect mucosal cells in the cervix and anus. The virus can cause a progressive spectrum of abnormalities classified as low-grade and high-grade squamous intraepithelial neoplasia (the process of rapid cell growth that is faster than normal and continues to grow—i.e., a tumor) and in some cases, progress to invasive cervical or anal cancer. The patient’s history of behavior and other relevant clinical information are important when attempting to identify the infectious agent associated with the lesion. For example, recurrent genital lesions and periods of dysesthesias (pain when touched) with intermittent outbreaks suggests infection with HSV 1 or 2. The virus remains inactive within the nerve ganglia during asymptomatic periods and then reemerges after an illness or other physiologic stress placed on the host. A patient’s medication history may also attribute to the eruption of a genital lesion. An individual who has recently completed antibiotic therapy for an unrelated condition who has sexual contact with a partner infected with Candida albicans may be more susceptible to infection. In addition, patients who have underlying autoimmune diseases, such as Crohn disease or HIV infection, are more susceptible to other infections and the development of genital lesions. 

Dysuria

Vaginitis

Clinical Manifestations Clinical manifestations of lower genital tract infections are as varied and diverse as the etiologies. Asymptomatic

Although a common presenting symptom associated with urinary tract infection, dysuria (painful urination) can also result from an STD caused by organisms such as N. gonorrhoeae, C. trachomatis, and HSV.  Urethral Discharge

The presence of an inflammatory exudate at the tip of the urethral meatus is generally observed in males; the symptoms of urethral infection in females are not commonly localized. Most males complain of discomfort at the penile tip as well as dysuria. Urethritis (swelling and irritation of the urethra) may be gonococcal, caused by N. gonorrhoeae, or nongonococcal. Nongonococcal urethritis (NGU) can be

Inflammation of the vaginal mucosa, called vaginitis, is a common clinical syndrome accounting for approximately 10 million office visits each year. Females who present with vaginal symptoms often complain of an abnormal discharge and additional symptoms, such as an offensive odor or itching. Vulvitis, local irritation of external genitalia, may be associated with vaginitis. The three most common causes of vaginitis in premenopausal females are vaginal candidiasis, bacterial vaginosis (BV) (group B streptococci, E. coli, and enterococci), and trichomoniasis. C. albicans causes about 80% to 90% of the cases of vaginal candidiasis; other species of Candida account for the remaining cases. Yeast can be carried vaginally in small

CHAPTER 73  Genital Tract Infections

A

B

C

D

1033

• Fig. 73.2  Genital lesions of the skin and mucous membranes that are sexually transmitted. (A) Genital

herpes showing vesicular lesions. (B) Typical chancre of primary syphilis. (C) Early chancroid lesion of the penis. (D) Condyloma acuminatum. (From Farrar WE, Wood MJ, Innes JA, et al. Infectious Diseases Text and Color Atlas. 2nd ed. London: Gower Medical Publishing; 1992.)

numbers and produce no symptoms. Most patients experiencing candidiasis complain of perivaginal itching, often with little or no discharge. Irritating symptoms such as erythema are also associated with candidiasis. Discharge is classically thick and “cheesy” in appearance (Fig. 73.3). Nonalbicans infections are more commonly associated with infections because of the increased use of topical antifungal genital creams and ointments. Vaginal infection with T. vaginalis, a protozoan parasite, produces a profuse, slightly offensive, yellow-green discharge; patients commonly complain of itching. Additional symptoms may include dysuria and dyspareunia (persistent genital pain during intercourse). Some patients will pre­ sent with a strawberry-appearing vaginal mucosa because of capillary dilation. The pH of the vaginal secretions will also typically increase to higher than 4.5, and numerous leukocytes and motile trophozoites may be present. About 25% of females carrying trichomonads are asymptomatic. In addition to vaginitis caused by these two organisms, there is a third type referred to as bacterial vaginosis (BV). Initially, BV was thought to be associated with Gardnerella vaginalis infection, but G. vaginalis was isolated from 40% of women without vaginitis. Therefore, the presence of G.

vaginalis should not be considered diagnostic for BV. BV is polymicrobial in etiology, involving G. vaginalis and other facultative and anaerobic organisms. The exact mechanism for the onset of BV is unknown, although it appears to be associated with a reduction in lactobacilli and hydrogen peroxide production, a rise in the vaginal pH, and the overgrowth of BV-associated organisms. Synergistic activity of various anaerobic organisms, including Prevotella spp., Porphyromonas spp., Bacteroides spp., Peptostreptococcus spp., Mobiluncus spp. (curved, motile rods), and Mycoplasma spp., as well as G. vaginalis, seems to contribute to the pathology of BV. BV is characterized by perivaginal irritation that is considerably milder than trichomoniasis or candidiasis and is usually associated with a foul-smelling discharge, often described as having a “fishy” odor. This odor is a result of products of bacterial metabolism (polyamines) being volatilized by vaginal fluids. Some patients also complain of abdominal discomfort. Dysuria and dyspareunia are rare. It appears that BV and trichomoniasis frequently coexist. Because BV can recur in the absence of sexual re-exposure and other settings (e.g., non–sexually active females or virgins), BV is not exclusively sexually transmitted. BV also increases a female’s risk of acquiring HIV, is associated with

1034 PA RT V I I     Diagnosis by Organ System

TABLE 73.2    Summary of Common Causes of Genital Lesions of the Skin and Mucous Membranes

Agent

Disease

Lesion

Major Associated Symptoms

Herpes simplex virus

Genital herpes

Papules, vesicles (blisters), pustules, or ulcers

Multiple lesions that are usually painful and tender, can recur (Fig. 73.2A).

Treponema pallidum

Primary syphilis

Genital ulcer (chancre)

Usually a single lesion, painless; lesion has even edges, represents the first of three stages of syphilis (Fig. 73.2B).

Haemophilus ducreyi

Chancroid

Papule that becomes pustular and ulcerates (chancroid); multiple ulcers may develop

Ulcer is deeply invasive, tender, painful, and purulent in appearance; edges of lesion are ragged (Fig. 73.2C).

Chlamydia trachomatis serotype L1, L2, and L3

Lymphogranuloma venereum

Small ulcer or vesicle that heals spontaneously without leaving a scar

After lesion heals, painful, swollen lymph nodes (lymphadenopathy) develop 2–6 weeks later; fever and chills; severe lymphatic obstruction and lymphedema can develop.

Klebsiella

Granuloma inguinale

Single or multiple subcutaneous nodules

Indolent and chronic course; nodules enlarge granulomatis and erode through the skin, producing a deep red, sharply defined ulcer that is painless.

Human papillomavirus

Condylomata acuminate (primary genotypes 6 and 11)

Genital warts

Warts have a cauliflower-like appearance; usually multiple lesions that can be flat or elevated; usually asymptomatic apart from physical presence (Fig. 73.2D).

Condylomata planum (primary genotypes 16, 18, 31, 33)

Flat, genital warts

Cervical warts that must be visualized by using a magnifying lens after the application of acetic acid (called colposcopy); infections can cause neoplasias that, in some cases, can progress to cervical cancer.

•

Fig. 73.3 Vulvovaginal candidiasis. Visible adherent white patches with surrounding erythema on the cervical mucosa.

increased complications in pregnancy, and may be involved in the pathogenesis of PID. Although uncommon, there are other infectious causes of vaginitis. Three are briefly mentioned here because Gram stain of vaginal secretions may be helpful. The clinical syndrome referred to as desquamate inflammatory vaginitis resembles a bacterial vaginitis. The syndrome manifests in premenopausal patients with a diffuse, exudative vaginitis with massive vaginal cell exfoliation, purulent vaginal discharge, and an occasional vaginal and cervical spotted rash.

Laboratory findings include an elevated pH (>5.0) of vaginal secretions. Also, numerous polymorphonuclear cells, an increased number of parabasal cells, the absence of gram-positive bacilli, and their replacement by occasional ­gram-positive cocci may be observed on direct Gram stain (Fig. 73.4). Basal cells appear because of the extensive exfoliation of epithelial cells. Symptoms associated with another disorder, lactobacillosis, resemble those of candidiasis and often follow antifungal therapy. Gram stain or wet mount typically reveals many very long lactobacilli. These predominately anaerobic lactobacilli are 40 to 75 μm in length and are significantly longer than the average normal microbiota lactobacillus (5 to 15 μm). Finally, preexisting lesions caused by other diseases may become secondarily infected with a mixed anaerobic microbiota of fusobacteria and spirochetes. This is referred to as fusiform-spirochete disease; this infection can progress rapidly. Gram stain examination reveals inflammatory cells in conjunction with gram-negative fusiform bacterial morphotypes and spirochetes.  Cervicitis

Polymorphonuclear neutrophils (PMNs) are normally present in the endocervix; however, an abnormally increased number of PMNs may be associated with cervicitis (inflammation of the cervix). Therefore, a purulent discharge from the endocervix can be observed in some cases of cervicitis. The endocervix is the site from which N. gonorrhoeae is

CHAPTER 73  Genital Tract Infections

PB

1035

PMN

PB

SEC

PMN

A

B •

Fig. 73.4  Gram stain of vaginal secretions from a patient with desquamate inflammatory vaginitis. (A) Numerous polymorphonuclear cells (PMNs), a squamous epithelial cell (SEC), a parabasal cell (PB), and the absence of lactobacilli are observed. (B) Numerous PMNs, several PBs, and the absence of lactobacilli are observed.

most frequently isolated in females with gonococcal infections. In patients presenting with cervicitis, C. trachomatis and M. genitalium can also be isolated; chlamydia has not been associated with cervicitis. Patients are often infected with both N. gonorrhoeae and C. trachomatis. Because most females with cervicitis caused by gonococci or chlamydia are asymptomatic, and cervical abnormalities are either subtle or absent in these patients, an appropriate laboratory diagnosis to detect these organisms must be performed. HSV and HPV can also infect the cervix. In females with herpes cervicitis, the cervix is friable (bleeds easily) and may have ulcers. Affected patients may also have lower abdominal pain.  Anorectal Lesions

As previously mentioned, because of the homosexual practice and increasingly common heterosexual practice of analgenital intercourse, sites of infection in addition to those in the genital tract must be considered. The anorectum and pharynx are commonly infected with the classic STDs, including anal warts and cancer caused by HPV, as well as other viruses and parasites. Patients with symptoms of proctitis (inflammation of the rectum) caused by N. gonorrhoeae or C. trachomatis complain of itching, mucopurulent anal discharge, anal pain, bleeding, and tenesmus (painful straining during a bowel movement). Anorectal infection caused by HSV is associated with severe anal pain, rectal discharge, tenesmus, and systemic signs and symptoms, such as fever, chills, and headaches. In HIV-infected individuals and other immunocompromised patients, these infections tend to last longer, are more severe, and are more difficult to treat compared with infection in immunocompetent individuals. Anorectal lesions are common in HIV-infected patients and include anal condylomata, anal abscesses, and ulcers. Anal abscesses and ulcers can be caused by various organisms, including CMV, Mycobacterium avium complex, HSV, Campylobacter spp., and Shigella, as well as traditional etiologic agents of STDs. 

Bartholinitis

In adult females, the Bartholin gland is a 1-cm mucus-producing gland on each side of the vaginal orifice. Each gland has a 2-cm duct that opens on the inner surface of the labia minora. If infected, this duct can become blocked and result in a Bartholin gland abscess (Bartholinitis). Although N. gonorrhoeae and C. trachomatis can cause infection, anaerobic and polymicrobic infections originating from normal genital microbiota are more common. 

Infections of the Reproductive Organs and Other Upper Genital Tract Infections Besides the lower genital tract, infections can occur in the reproductive organs of both males and females.

Females Infection of the female reproductive organs (i.e., uterus, fallopian tubes, ovaries, and even the abdominal cavity) can occur. The organisms spread as they ascend from lower tract sites of infection. Organisms may also be introduced to the reproductive organs by surgery, instrumentation, or during childbirth. Pelvic Inflammatory Disease

PID is an infection that results when cervical microorganisms travel upward to the endometrium (inner membrane of the uterus), fallopian tubes, and other pelvic structures. This infection can produce one or more of the following inflammatory conditions: endometritis, salpingitis (inflammation of the fallopian tubes), localized or generalized peritonitis, or abscesses involving the fallopian tubes or ovaries. Patients with PID often have intermittent abdominal pain and tenderness, vaginal discharge, dysuria, and possibly systemic symptoms, such as fever, weight loss, and headache. Serious complications, such as permanent scarring of the fallopian tubes and infertility, can arise if PID is untreated.

1036 PA RT V I I     Diagnosis by Organ System

TABLE 73.3    Common Etiologic Agents of Prenatal and Neonatal Infections

Time of Infectiona

Route of Infection

Common Agents

Prenatal

Transplacental

Bacteria: Listeria monocytogenes, Treponema pallidum, Borrelia burgdorferi Viruses: cytomegalovirus (CMV), rubella, HIV, parvovirus B19, enteroviruses Parasites: Toxoplasma gondii, Plasmodium spp.

Ascending

Bacteria: group B streptococci, Escherichia coli, L. monocytogenes, Chlamydia trachomatis, genital mycoplasmas Viruses: CMV, herpes simplex virus (HSV)

Natal

Passing through the birth canal

Bacteria: group B streptococci, E. coli, L. monocytogenes, N. gonorrhoeae, C. trachomatis Viruses: CMV, HSV, enteroviruses, hepatitis B virus, human immunodeficiency virus (HIV)

Postnatal

All the aforementioned routes, from the nursery environment, or from maternal contact (e.g., breastfeeding)

All agents listed previously and various organisms from the nursery environment, including gram-negative bacteria and viruses, such as respiratory syncytial virus

aSome

newborns develop infections during the first 4 weeks of postnatal life. Infections may be delayed manifestations of earlier prenatal (before birth), natal, or postnatal (after birth) acquisition of pathogens.

Infection with N. gonorrhoeae or C. trachomatis in the lower genital tract can lead to PID if a female is not adequately treated. Other organisms, such as anaerobes, gramnegative rods, streptococci, and mycoplasmas, may ascend through the cervix, particularly after parturition (childbirth), dilation of the cervix, or abortion. The presence of an intrauterine device (IUD) is associated with a slightly higher rate of PID. Such infections caused by Actinomyces have been associated with the use of IUDs.  Infections After Gynecologic Surgery

After gynecologic surgery, such as a vaginal hysterectomy, patients frequently develop postoperative infections including pelvic cellulitis or abscesses. The major pathogens include normal human microbiota: aerobic gram-positive cocci, gram-negative bacilli, anaerobes such as Peptostreptococcus spp., and genital mycoplasmas.  Infections Associated With Pregnancy

Infections can also occur in females during pregnancy (prenatal) or after the birth (postpartum) of a child. These infections may, in turn, be transmitted to the infant; they are not only capable of compromising the mother’s health but also the health of the developing fetus or neonate. While developing within the uterus, the fetus is protected from most environmental factors, including infectious agents. The human immune system does not become fully competent until several months after birth. Immunoglobulins that cross the placental barrier, primarily immunoglobulin G (IgG), protect the newborn from many infections until the infant begins to produce immunoglobulins of his or her own in response to antigenic stimuli. This unique environmental niche, however, does expose the vulnerable fetus to pathogens present in the mother.

• BOX 73.1 Organisms Commonly Isolated in

Chorioamnionitis

Anaerobic bacteria Genital mycoplasmas Group B streptococci Escherichia coli

Prenatal infections (those that occur any time before birth) may be acquired hematogenously or by ascending genital tract routes from mother to infant. If the mother has a bloodstream infection, organisms can reach and cross the placenta, with possible spread of infection to the developing fetus. Organisms that can cross the placenta are listed in Table 73.3. Alternatively, organisms can also infect the fetus by the ascending route from the vagina through torn or ruptured fetal membranes. Chorioamnionitis is an infection of the uterus and its contents during pregnancy. This infection is commonly acquired when organisms spread from the vagina or cervix after premature or prolonged rupture of the membranes, or during labor. Organisms that are commonly isolated from amniotic fluid are listed in Box 73.1. Other maternal infections associated with adverse pregnancy outcomes that are not generally sexually transmitted include parvovirus B19, rubella, and L. monocytogenes. 

Males Infections in male reproductive organs can also occur and include epididymitis, prostatitis, and orchitis (testicular swelling). Epididymitis, an inflammation of the epididymis, is commonly seen in sexually active men. Patients complain of fever and pain and swelling of the testicle. Patients may also present with dysuria and a urethral discharge. There

CHAPTER 73  Genital Tract Infections

are two general types of epididymitis: nonspecific bacterial epididymitis caused by aerobic gram-negative rods, enterococci, or Pseudomonas spp.; and sexually transmitted epididymitis most commonly associated with N. gonorrhoeae and C. trachomatis. However, bacterial epididymitis is typically associated with an underlying genitourinary abnormality that requires surgery or urethral catheterization. Infections may also be caused by enteric bacteria and coagulasenegative staphylococci in males over 35 years of age and in homosexual males; these infections are often associated with obstruction by the prostate gland. Prostatitis is a term to clinically describe adult male patients who have perineal, lower back, or lower abdominal pain, urinary discomfort, or ejaculatory complaints. Prostatitis is caused by both infectious and noninfectious means. Bacteria can cause an acute or chronic prostatitis. Patients with acute bacterial prostatitis have dysuria and urinary frequency—symptoms typically associated with lower urinary tract infection. These patients frequently have systemic signs of illness, such as fever. Chronic bacterial prostatitis is an important cause of persistent bacteriuria in males leading to recurrent bacterial urinary tract infections. The common causes of these infections are similar to the bacterial causes of lower urinary tract infections, such as E. coli, Pseudomonas spp., and other enteric organisms. Finally, inflammation of the testicles, orchitis, is uncommon and generally acquired by the blood-borne dissemination of viruses. Mumps is associated with most cases. Patients exhibit testicular pain and swelling after infection. Infections range from mild to severe. In addition, epididymo-orchitis may occur after infection of the epididymis. Organisms typically isolated from bacterial orchitis include staphylococci, streptococci, E. coli, Klebsiella pneumoniae, and Pseudomonas aeruginosa. Gonorrhea

Gonorrhea is a common STI caused by the bacterium N. gonorrhoeae. The infection may be spread by direct contact with secretions within the mouth, vagina, penis, or perianal region. The organism reproduces in warm, moist areas of the body including the urethra of males and females, fallopian tubes, uterus, and cervix. Symptoms occur 2 to 5 days after infection in females. Males may not display symptoms for up to 1 month after infection. Symptoms in females include a vaginal discharge, pain and frequency on urination, sore throat, abdominal pain, fever, and painful sexual intercourse. Males experience pain and frequency during urination, a penile discharge, red or swollen urethra, and tenderness in the testes. The characteristics of the urethral discharge may vary from cloudy to clear and is therefore an unreliable indicator for gonococcal urethritis in males. Gonorrhea can be directly diagnosed by Gram staining a sample of urethral discharge, cervical specimens, or joint fluids. N. gonorrhoeae is a gram-negative diplococcus with a characteristic kidney bean shape on Gram stain. The detection of intracellular diplococci in male secretions

1037

is diagnostic for N. gonorrhoeae. Extracellular diplococci in females is an indication of normal genital microbiota; however, intracellular diplococci indicate the presence of pathogenic organisms. Definitive diagnosis in females must include confirmation by culture. Infection with N. gonorrhoeae can lead to increased complications, including PID and gonorrheal ophthalmia neonatorum (eye infections) in newborns. N. meningitidis has been increasingly associated with urethritis. Nongonococcal urethritis (NGU) is most commonly associated with C. trachomatis infection. Additional organisms that may be isolated from specimens in cases of NGU include U. urealyticum and M. genitalium.  Syphilis

Syphilis is an STD that is caused by the bacterium T. pallidum. The organism is transmitted from person to person through direct contact with infected lesions on the external genital area, vagina, anus, or rectum. Syphilis may also be transmitted from mother to baby during pregnancy. Many individuals can be infected and remain asymptomatic for years, making the control of this disease difficult. The disease is characterized by three stages: primary, secondary, and tertiary (also referred to as late or latent syphilis). Direct diagnosis may be accomplished by dark-field microscopy of material from an infectious lesion. However, serology provides a more accurate and reliable method for diagnosis. (See Chapter 45 for a more detailed description of the disease and laboratory diagnosis.) 

Laboratory Diagnosis of Genital Tract Infections Lower Genital Tract Infections Urethritis, Cervicitis, and Vaginitis Specimen Collection

This discussion focuses on those specimens submitted for culture or direct examination. Procedures for the collection and transport of specimens for detection of agents by other noncultural methods (e.g., detection of infectious agents using nucleic acid–based tests) should be followed according to the respective manufacturer’s instructions. (Refer to Table 5.1 for a review of collection, transport, and processing of genital tract specimens.) Urethral. Urethral discharge may occur in both males and females infected with pathogens, such as N. gonorrhoeae, N. meningitidis, and T. vaginalis. The presence of infection is more likely to be asymptomatic in females, because the discharge is usually less profuse and may be masked by normal vaginal secretions. U. urealyticum can also be isolated from male urethral discharge. A small urogenital swab designed expressly for collection of such specimens should be used. These swabs are made of cotton or rayon, treated with charcoal to adsorb material toxic to gonococci, and wrapped tightly over one end of a thin wire shaft. Cotton- or rayon-tipped swabs on a thin

1038 PA RT V I I     Diagnosis by Organ System

wire may also be used to collect specimens for isolation of mycoplasmas and chlamydiae. Calcium alginate swabs are generally more toxic for HSV, gonococci, chlamydia, and mycoplasmas than treated cotton swabs. Because dacron swabs are least toxic, they are recommended for viral specimens. Dacron-tipped swabs on plastic shafts are also acceptable for chlamydiae and genital mycoplasmas. To obtain a urethral specimen, a swab is inserted approximately 2 cm into the urethra and rotated gently before withdrawing. Because chlamydiae are intracellular pathogens, it is important to remove epithelial cells (with the swab) from the urethral mucosa. Separate swabs for cultivation of gonococci, chlamydiae, and ureaplasma are required. When profuse urethral discharge is present, particularly in males, the discharge may be collected externally without inserting a sampling device into the urethra. However, a urethral swab for chlamydiae must be collected on males. A few drops of first-voided urine have also been used successfully to detect gonococci in males. Because T. vaginalis may be present in urethral discharge, material for culture should be collected by swab, as described, and another specimen collected on a swab and placed into a tube containing 0.5 mL of sterile physiologic saline. This specimen should be delivered to the laboratory immediately. Direct wet mounts and cultures for T. vaginalis can be performed from this second specimen. Commercial media for culture of Trichomonas are available. The first few drops of voided urine make a suitable specimen for recovery of Trichomonas from infected males, if it is inoculated into culture media immediately. Alternatively, material may be smeared onto a slide for a fluorescent antibody stain. Plastic envelopes for direct examination and subsequent culture are also available (InPouch TV, BIOMED, White City, OR); sensitivity of this system is superior to other available methods, and organism viability is maintained up to 48 hours. In addition, several other techniques are available, including enzyme immunoassay, latex agglutination tests, and the Affirm VPIII probe (Becton Dickinson, Cockeysville, MD). Nucleic acid–based test methods are the most sensitive for the detection and identification of T. vaginalis directly in clinical specimens. N. gonorrhoeae may be detected from clinical specimens using a variety of nucleic acid–based methods using a DNA probe that hybridizes to organismal ribosomal ribonucleic acid (rRNA). Fully automated systems for complete sample processing are available that reduce technical time. These tests are rapidly evolving and widely used for quick detection of N. gonorrhoeae from vaginal, urethral, thin-prep, and urine specimens. N. meningitidis should be considered when a urethral Gram stain indicates the presence of gramnegative intracellular diplococci and a nonculture based test, such as nucleic acid amplification test (NAAT), is negative for the presence of N. gonorrhoeae.  Cervical and Vaginal. Organisms that cause purulent vaginal discharge (vaginitis) include T. vaginalis, gonococci, Candida spp. and, rarely, beta-hemolytic streptococci. The same organisms that cause purulent infections in the urethra

may also infect the epithelial cells in the cervical opening (os), as can HSV. Mucus is removed by gently rubbing the area with a cotton ball. The urethral swab is inserted into the cervical canal and rotated and moved from side to side for 30 seconds before removal. Swabs are handled as previously described for urethral swabs for isolation of Trichomonas and gonococci. Chlamydiae cause a mucopurulent cervicitis with discharge. Endocervical specimens are obtained after the cervix has been exposed with a speculum, which allows visualization of vaginal and cervical architecture, and after ectocervical mucus has been adequately removed. The speculum is moistened with warm water, because many lubricants contain antibacterial agents. Because normal vaginal secretions contain great quantities of bacteria, care must be taken to prevent or minimize contamination of swabs for culture by contact with these secretions. A small, nylon-bristled cytology brush, or Cytobrush, may be used to ensure that cellular material is collected. Collection may result in patient discomfort and bleeding. In addition to cervical specimens, which are particularly useful for isolating herpes, gonococci, mycoplasmas, and chlamydiae, vaginal discharge specimens may be collected. Organisms likely to cause vaginal discharge include Trichomonas, yeast, and the agents of BV. Swabs for diagnosis of BV are dipped into the fluid that collects in the posterior fornix of the vagina. Genital tract infections caused by sexually transmitted agents in children (preadolescents) are most often the result of sexual abuse. Because of medico-legal implications, the laboratory should treat specimens from such patients with extreme care, carefully identifying and documenting all isolates. Although nucleic acid–based testing methods are available for the identification of organisms associated with sexual abuse cases, culture remains the preferred method of detection for C. trachomatis and N. gonorrhoeae in medico-legal cases. In addition, cultivation of the isolate may be required to link the specific isolate to the perpetrator using epidemiologic studies. It is important to follow the current rules and regulations of each state for the collection and processing of isolates in these situations. Because it is impossible to exclude contamination with vaginal microbiota, obtaining swabs of Bartholin gland exudate is not recommended. Infected Bartholin glands should be aspirated with needle and syringe after careful skin preparation, and cultures should be evaluated for anaerobes and aerobes.  Transport. Swabs collected for isolation of gonococci may be transported to the laboratory in modified Stuart’s or Amie’s charcoal transport media and held at room temperature until inoculated to culture media. Good recovery of gonococci is possible if swabs are cultured within 12 hours of collection. Material that must be held longer than 12 hours should be inoculated directly to one of the commercial systems designed for recovery of gonococci, described later in this chapter.

CHAPTER 73  Genital Tract Infections

• Fig. 73.5  Gram-negative intracellular diplococci, which are diagnostic

for gonorrhea in urethral discharge and presumptive for gonorrhea in vaginal discharge.

Swabs for isolation of chlamydiae and mycoplasmas are transported in specific transport media containing antibiotics and other essential components. Specimens for chlamydia culture should be transported on ice. (Specimens transported at room temperature should be inoculated within 15 minutes of collection.) Specimens can be stored at 4°C for up to 24 hours. If culture inoculation will be delayed more than 24 hours, specimens should be quickfrozen in a dry ice and 95% ethanol bath and stored at −70°C until cultured. If collected and transported in specific transport media, specimens for genital mycoplasma culture may be transported on ice or at room temperature. If not in genital mycoplasma transport media, specimens should be transported on ice to suppress the growth of contaminating microbiota.  Direct Microscopic Examination

In addition to culture, urethral discharge may be examined by Gram stain for the presence of gram-negative intracellular diplococci (Fig. 73.5), usually indicative of gonorrhea in males. After inoculation to culture media, the swab is rolled over the surface of a glass slide, covering an area of at least 1 cm2. Specimens collected from within the urethra may contain small cuboidal epithelial cells with a large nucleus. Numerous PMNs, more than four per oil immersion field, will also be visible in acute urethritis. If the Gram stain is characteristic of normal skin or genital microbiota, cultures of urethral discharge need not be performed. Urethral smears from females may also be examined. If extracellular organisms resembling N. gonorrhoeae are seen, the microbiologist should continue to examine the smear for intracellular diplococci. Presumptive diagnosis can be useful when decisions are to be made regarding immediate therapy, but confirmatory cultures or an alternative nonculture method should always be performed on specimens from females. Some strains of N. gonorrhoeae are sensitive to the amount of vancomycin present in selective media. If suspicious organisms seen on smear fail to grow in culture, reculture on chocolate agar without antibiotics may be warranted and N. meningitidis should also be considered. In addition, a Gram

1039

•

Fig. 73.6 Clue cells in vaginal discharge suggestive of bacterial vaginosis.

stain that contains numerous PMNs without intracellular gram-negative diplococci may also be suggestive of NGU. Fluorescein-conjugated monoclonal antibody reagents are sensitive and specific for visualization of the inclusions of C. trachomatis in cell cultures or elementary bodies in urethral and cervical specimens containing cells. Reagents for direct staining of specimens are available commercially in complete collection and test systems, but the increased technologist time required limits the usefulness of this method for laboratories that receive many specimens, except as a confirmatory test for other antigen detection systems with borderline results. In some studies, the sensitivity of visual detection of chlamydia with these reagents has been similar to that of culture. False-positive results should not occur if at least 10 morphologically compatible fluorescing elementary bodies are seen on the smear. No direct visual methods exist for detection of mycoplasmas, but nucleic acid–based assays are the most sensitive for diagnosis of these fastidious organisms. Direct microscopic examination of a wet preparation of vaginal discharge provides the simplest rapid diagnostic test for T. vaginalis and can be examined immediately. Motile trophozoites of Trichomonas can be visualized in a routine wet preparation in two thirds of cases or a direct fluorescent antibody (DFA) stain, Merifluor (chlamydia) (Meridian Diagnostics, Cincinnati, OH) may be used. Positive findings on a wet mount are diagnostic for trichomoniasis, but results are often negative in males. Budding cells and pseudohyphae of yeast can also be easily identified in wet preparations by adding 10% potassium hydroxide (KOH) to a separate preparation, thereby dissolving host cell protein and enhancing the visibility of fungal elements. BV, characterized by a foul-smelling discharge, can be diagnosed microscopically or clinically. The discharge is primarily sloughed epithelial cells, many of which are completely covered by tiny, gram-variable rods, and coccobacilli. These cells are called clue cells (Fig. 73.6). The absence of inflammatory cells in the vaginal discharge is another sign of BV. Although G. vaginalis has been historically associated with the syndrome and can be cultured on a human blood bilayer plate, culture is not recommended for diagnosis of BV. A clinical diagnosis of BV is dependent on the presence

1040 PA RT V I I     Diagnosis by Organ System

of three or more of the following criteria: homogeneous, gray discharge; clue cells seen on wet mount or Gram stain; a pH higher than 4.5; and an amine or fishy odor elicited by the addition of a drop of 10% KOH to the discharge on a slide or on the speculum. BV may be differentiated from other vaginal infection by Gram stain (Fig. 73.7). A grading system for Gram stains of vaginal discharge has been developed (see Evolve Procedure 73.1). This system is based on the presence or absence of certain bacterial morphologies. Typically, in patients with BV, lactobacilli are either absent or few in number, whereas curved, gram-variable rods (Mobiluncus spp.) or G. vaginalis and Bacteroides morphotypes predominate. The Gram stain is more sensitive and specific than either the wet mount for detection of clue cells or culture for G. vaginalis, and the smear can be saved and reexamined later.  Culture

Samples for isolation of gonococci may be inoculated directly to culture media, obviating the need for transport medium. Commercially produced systems have been developed for this purpose, and many clinicians inoculate standard plates directly if convenient access to an incubator is available. Modified Thayer-Martin medium is most often used, although New York City (NYC) medium has the added advantage of supporting the growth of mycoplasmas and gonococci. The specimen swab is rolled across the agar with constant turning to expose all surfaces to the medium. Specimens must be inoculated to additional media for isolation of yeast, streptococci, and mycoplasmas. Yeast grows well on Columbia agar base with 5% sheep blood and colistin and nalidixic acid (CNA), although more selective media are available. Most yeast and streptococci also grow on standard blood agar; thus, adding special fungal media, such as Sabouraud brain-heart infusion agar (SABHI), is unwarranted. A specimen from the lower vagina followed by the rectum using the same swab at 35 to 37 weeks’ gestation, reliably

predicts the presence of group B streptococci at delivery. The swab should be transported to the laboratory in a nonnutritive transport medium, such as Amie’s or Stuart’s, without charcoal and then inoculated into a recommended selective broth medium, such as Todd-Hewitt broth supplemented either with gentamicin and nalidixic acid or with CNA referred to as LIM broth. Selective enrichment broths are subcultured to agar the next day to isolate and identify group B streptococci. In addition, the presence of group B streptococci in urine in any concentration from a pregnant female is a marker for heavy genital tract colonization. Any quantity of group B streptococci in urine from pregnant women should be worked up in the laboratory (Chapter 72). T. vaginalis may be cultured in Diamond’s medium (available commercially) or plastic envelopes inoculated with discharge material. Culture techniques are most sensitive. A commercially available biphasic genital mycoplasma culture system (Mycotrim-GU, Irvine Scientific, Santa Ana, CA) can be used to culture Mycoplasma spp. and U. urealyticum, although commercially prepared media are not as sensitive as fresh media. M. genitalium may not grow on commercial media because of the presence of thallium acetate.  Nonculture Methods Various nonculture methods may be used to diagnose genital tract diseases, including serology, latex agglutination, nucleic acid hybridization and amplification assays, and enzyme immunoassays. Most assays detect a single or possibly two genital tract pathogens and are commercially available. These methods are described in more detail in chapters relating to individual pathogens. As previously discussed, BV involves several organisms. Although the Gram stain offers high sensitivity and specificity, it is not immediately available. Currently, commercial laboratory tests are available to aid in the diagnosis of BV, but not all are available in the United States; a test for sialidase (OSOM BVBLUE, Sekisui Diagnostics, Framingham, MA) in conjunction with measuring pH has been reported to be

A B A B

A

B • Fig. 73.7  (A) Predominance of lactobacilli in Gram stain from healthy vagina. (B) Absence of lactobacilli and presence of Gardnerella vaginalis (A arrows) and Mobiluncus spp. (B arrows) morphologies.

CHAPTER 73  Genital Tract Infections

a rapid, highly sensitive, and specific means to diagnose BV. Sialidases are secreted from anaerobic gram-negative rods, such as Bacteroides and Prevotella as well as Gardnerella, and play a role in bacterial nutrition, cellular interactions, and immune response evasion, which in turn improves the ability of bacteria to adhere to, invade, and destroy mucosal tissue. A hybridization assay (Affirm VP III Microbial Identification Test; Becton Dickinson Microbiology Systems, Burlington, NC) is commercially available to diagnose BV, as well as genital tract infections caused by Candida spp. and T. vaginalis. Once the appropriate reagents and specimen are added to special trays, the entire hybridization assays are then performed using instrumentation (Fig. 73.8). Evaluations indicate this system is sensitive and specific. In addition to microscopic examination, NAAT should be performed on vaginal, cervical, or urine specimens when cervicitis is suspected. A variety of assays are available for the detection of associated pathogens, such as the automated Aptima Combo 2 Assay (Fig. 73.9). This assay is a transcription-mediated amplification (TMA) test that utilizes target capture for the in  vitro qualitative detection and differentiation of rRNA from C. trachomatis and/or N. gonorrhoeae using the Panther System (Hologic) as a testing platform. 

Genital Skin and Mucous Membrane Lesions External genital lesions are usually either vesicular or ulcerative. Causes of lesions can be determined by physical examination, histologic or cytologic examination, and microscopic examination or culture of exudate. Gram staining is typically not useful for the identification of organisms and the evaluation of genital lesions because of the presence of contaminating normal microbiota. Vesicles in the genital area are almost always attributable to viruses, and herpes simplex is the most common cause. Epithelial cells from the base of a vesicle may be spread onto the surface of a slide (Tzanck smear) and examined for the typical multinucleated giant cells of HSV by staining with

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Wright-Giemsa stain or immunofluorescent antibody stains for viral antigens. NAATs are recommended for HSV testing that provide excellent sensitivity. Additionally, or alternatively, the material may be transported for culture of the virus as outlined in Evolve Procedure 73.2. Specimens positive for HSV will typically demonstrate cytopathic effect (changes in cell morphology) within 48 hours. Cultures have been largely replaced by NAATs. Several commercial fluorescein-conjugated monoclonal and polyclonal antibodies directed against herpetic antigens of either type 1 or 2 are available. When fluorescent antibody–stained lesion material containing enough cells is viewed under ultraviolet light, the diagnosis can be made in 70% to 90% of patients. Laboratories that routinely process genital material for herpes should be using immunofluorescent staining reagents when a rapid answer is desired; otherwise, culture, which is generally positive in 2 days. Nonfluorescent markers, such as biotin-avidin-horseradish peroxidase or alkaline phosphatase, have also been conjugated to these specific antibodies, often allowing for earlier detection of herpes-infected cells in tissue culture monolayers. Serologic assays are currently available to distinguish HSV 1 and HSV 2 and have been modified that allow the use of these tests in a clinic setting. Lesions caused by HPV are typically characterized using the Papanicolaou smear (Pap) or a biopsy. These methods lack specificity, and, therefore, positive smears should be confirmed with a nucleic acid–based test such as hybrid capture or polymerase chain reaction. Material from lesions suggestive of syphilis should be examined by dark-field or fluorescent microscopy. Darkfield microscopy is not useful for the differentiation of pathogenic from nonpathogenic treponemes. A two-step serologic test using the rapid plasma reagin (RPR), Venereal Disease Research Laboratory (VDRL), or unheated serum reagin (USR) test—followed by a confirmatory test—is the recommended procedure for the diagnosis of syphilis.

• Fig. 73.8  Affirm VP III Microbial Identification Test used to differentiate the three major causes of vaginitis/

bacterial vaginosis from a single sample within 1 hour. (Courtesy Becton Dickinson Microbiology Systems. Affirm is a trademark of Becton Dickinson and Co.)

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Lysis releases nucleic acids Target RNA Hybridization of specific capture oligonucleotides with target RNA

Capture oligonucleotides

Capture of target RNA by magnetic particles

Isothermal amplification with reverse transcriptase and RNA polymerase

Reverse transcriptase

Reverse transcriptase

RNA

cDNA

dsDNA RNA polymerase RNA

RNA amplicon Fluorophore Detection with single stranded nucleic acid torches tagged with fluorophore and quencher

Quencher

+ Nucleic acid torch

• Fig. 73.9  The Aptima Combo 2 Assay is a transcription-mediated amplification (TMA) test that utilizes tar-

get capture for the in vitro qualitative detection and differentiation of ribosomal RNA (rRNA) from Chlamydia trachomatis (CT) and/or Neisseria gonorrhoeae (GC) using the Panther System (Hologic) as a testing platform. NAAT, Nucleic acid amplification test. (Adapted from teaching materials, courtesy Jim Flanigan, American Society for Clinical Laboratory Science.)

(These procedures are described in Chapter 45.) NAAT have also been developed for the detection of T. pallidum. All lesions suspected of infectious etiology may be Gram stained in addition to the procedures described. The smear of lesion material from a patient with chancroid may show many small, pleomorphic, slender, gramnegative rods and coccobacilli arranged in chains and groups referred to as a “school of fish,” characteristics of H. ducreyi. However, culture has been shown to be more sensitive for diagnosis of this agent. Material collected on cotton or dacron swabs may be transported in modified Stuart’s medium. Specimens should be inoculated to culture media within 1 hour of collection. A special agar, consisting of Mueller-Hinton–based chocolate agar enriched with 1% IsoVitaleX (Becton-Dickinson, Franklin Lakes, NJ) and vancomycin (3 mg/mL), has yielded good isolation if cultures are incubated in 5% to 7% carbon dioxide in a moist atmosphere, such as a candle jar. H. ducreyi grows best at 33°C. NAAT have also been developed for the detection of H ducreyi. Granuloma inguinale (K. granulomatis) (Fig. 73.8) is diagnosed by staining a crushed preparation of a small piece of biopsy tissue obtained from the edge of the base of the ulcer with Wright’s or Giemsa stain and finding characteristic Donovan bodies (bipolar staining rods within

macrophages). No acceptable media for isolation of K. granulomatis are available. 

Buboes Buboes, swollen lymph glands in the inguinal (pelvic) region, are often evidence of a genital tract infection. Buboes are common in patients with primary syphilis, genital herpes, lymphogranuloma venereum, and chancroid. Patients with acquired immune deficiency syndrome (AIDS) may show generalized lymphadenopathy. Other diseases that are not sexually transmitted, such as plague, tularemia, and lymphoma, can also produce buboes. Material from buboes may be aspirated for microscopic examination and culture. Isolation by cell culture or the identification of C. trachomatis using nucleic acid–based test methods is considered diagnostic. 

Infections of the Reproductive Organs Pelvic Inflammatory Disease PID is often caused by the same organisms that cause cervicitis or by organisms that make up the normal microbiota of the vaginal mucosa. Diagnosis is often made based on signs and symptoms. Because of the profuse normal microbiota of the vaginal tract, specimens must be collected in such a way as to prevent vaginal microbiota contamination. Aspirated

CHAPTER 73  Genital Tract Infections

material collected by needle and syringe represents the best specimen. If this cannot be obtained at the time of surgery or laparoscopy, collection of intrauterine contents using a protected suction curetting device or double-lumen sampling device inserted through the cervix is also acceptable. Culdocentesis (aspiration of fluid in the cul-de-sac), after decontamination of the vagina by povidone-iodine, is satisfactory but rarely practiced today. Aspirated material should be placed into an anaerobic transport container. The presence of mixed anaerobic microbiota, gonococci, or both, can be rapidly detected from a Gram stain. Direct examination with fluorescent monoclonal antibody stain may also detect chlamydiae. All specimens should be inoculated to media that allow the recovery of anaerobic, facultative, and aerobic bacteria, gonococci, fungi, mycoplasmas, and chlamydiae. All material collected from normally sterile body sites in the genital tract should be inoculated to chocolate agar and placed into a suitable broth, such as thioglycollate, in addition to the other types of media noted. If only specimens obtained on routine swabs inserted through the cervix are available, cultures should be performed for detection of gonococci and chlamydiae. 

Miscellaneous Infections Infections of the male prostate, epididymis, and testes are usually bacterial. Uropathogens, such as E. coli, P. aeruginosa, and Enterococcus spp. cause more than 60% of acute bacterial prostatitis. In younger males, chlamydia and N. gonorrhoeae predominates as the cause of sexually transmitted epididymitis and possibly of prostatitis. Orchitis, or inflammation of the testes, can be caused by the same uropathogens as the other conditions as well as viral infections. The mumps virus is the cause of most cases of viral orchitis. Urine or discharge collected via the urethra is the specimen of choice unless an abscess is drained surgically or by needle and syringe. The first few milliliters of voided urine may be collected before and after prostatic massage to try to pinpoint the anatomic site of the infection. Cultures are inoculated to support the growth of anaerobic, facultative, and aerobic bacteria, as well as gonococci. 

Infections of Neonates and Human Products of Conception Suspected infections acquired by the fetus because of a maternal infection that crosses the placenta (congenital infection) can be diagnosed culturally or serologically in the newborn. Because maternal IgG crosses the placenta, serologic tests are often difficult to interpret (Chapter 9). Nucleic acid testing for viruses included in the Herpesviridae are more widely used and are more sensitive than cell culture and antigen detection. Although HSV, varicella-zoster virus (VZV), enteroviruses, and CMV can be cultured easily—as can most bacterial agents—rubella and parvovirus B19 are more difficult to culture. Nasal and urine specimens offer the greatest yield for viral isolation, although blood,

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cerebrospinal fluid, and material from a lesion can also be productive. Systemic neonatal herpes without lesions may be difficult to diagnose unless tissue biopsy material is examined, because the viruses may not be present in cerebrospinal fluid or blood. Bacteria and fungi can be isolated from lesions, blood, and other normally sterile sites. Determining the presence of fetal immunoglobulin M (IgM) directed against the agent in question establishes the serologic diagnosis of congenital infection. Until recently, ultracentrifugation was required for separation of IgM from IgG, the only definitive means of preventing false-positive results caused by maternal IgG or fetal rheumatoid factor. Ion-exchange chromatography columns, antihuman IgG, and bacterial proteins that bind to IgG, are commercially available for removing cross-reactive IgG and rheumatoid factor to obtain more homogeneous IgM for differentiation of fetal antibody. Indirect fluorescent antibody and enzymelinked immunosorbent assay (ELISA) test systems are commercially available to detect IgM against T. gondii, rubella, CMV, HSV, and VZV. Interference by rheumatoid factor is still a consideration in most commercial IgM test systems (Chapter 9). The ability to detect viral inclusions in tissue, conjunctiva scrapings, and vesicular lesions, traditionally performed with Giemsa stain, has been improved because of the availability of monoclonal and polyclonal fluorescent antibody reagents, which are described in the chapters that discuss individual agents. Infections that infants can acquire as they pass through an infected birth canal or that are related to difficult labor, premature birth, premature rupture of the membranes, or other events include the following: • HSV and CMV • Gonorrhea • Group B streptococcal sepsis • Chlamydial conjunctivitis and pneumonia • E  . coli or other neonatal meningitis In the laboratory, these infections are diagnosed by direct detection, including NAATS or culturing for the agents when possible, or by performing serologic tests. The appropriate specimens (e.g., cerebrospinal fluid, serum, pus, tracheal aspirate) should be examined and inoculated immediately. Routine body surface cultures of infants in intensive care have not been shown to be helpful for predicting subsequent disease. The use of viral cell culture methods has been replaced in most laboratories by NAAT methods. Finally, certain infectious agents are known to cause fetal infection and even abortion. For example, L. monocytogenes, the causative agent of mild flulike symptoms in the mother, can cause extensive disease and abortion of the fetus if infection occurs late in the pregnancy. Therefore, isolation of the organism from the placenta and from tissues of the fetus is important.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

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Bibliography Anderson MR, Klink K, Cohrssen A: Evaluation of vaginal complaints, JAMA 291:1368–1379, 2004. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, PA, 2020, Elsevier. Carroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Clarridge JE, Shawar R, Simon B: Haemophilus ducreyi and chancroid: practical aspects for the clinical microbiology laboratory, Clin Microbiol Newsl 12:137–141, 1990. Creatsas G, Deligeoroglou E: Microbial ecology of the lower genital tract in women with sexually transmitted diseases, J Med Microbiol 61:1347–1351, 2012. Curry A, Williams T, Penny ML: Pelvic inflammatory disease: diagnosis, management, and prevention, Am Fam Physician 100(6):357– 364, 2019. Fredricks DN, Fiedler TL, Marrazzo JM: Molecular identification of bacteria associated with bacterial vaginosis, N Engl J Med 353:1899–1911, 2005. Hammerschlag MR, Guillen CD: Medical and legal implications of testing for sexually transmitted infections in children, Clin Microbiol Rev 23:493–506, 2010. Hillier SL, Krohn MA, Rabe LK, et al.: Normal vaginal flora, H2O2producing lactobacilli and bacterial vaginosis in pregnant women, Clin Infect Dis 16(Suppl 4):S273–S281, 1993.

Johnson RE, Newhall WJ, Rapp JR, et al.: Screening tests to detect Chlamydia trachomatis and Neisseria gonorrhoeae infections—2002, MMWR 51(RR-15):1–38, 2002. Kellogg JA, Seiple JW, Klinedinst JL, et  al.: Comparison of cytobrushes with swabs for recovery of endocervical cells and for Chlamydiazyme detection of Chlamydia trachomatis, J Clin Microbiol 30:2988–2990, 1992. Nugent RP, Krohn MA, Hillier SL: Reliability of diagnosing bacterial vaginosis is improved by a standardized method of Gram stain interpretation, J Clin Microbiol 29:297–301, 1991. Schmid GP, Faur YC, Valu JA, et  al.: Enhanced recovery of Haemophilus ducreyi from clinical specimens by incubation at 33°C versus 35°C, J Clin Microbiol 33:3257–3259, 1995. Schrag S, Gorwitz R, Fultz-Butts K, et  al.: Prevention of perinatal group B streptococcal disease, revised guidelines from CDC, MMWR Recomm Rep 51(RR-11):1–22, 2002. Sobel JD: What’s new in bacterial vaginosis and trichomoniasis? Infect Dis Clin North Am 19:387–406, 2005. Wilson J: Managing recurrent bacterial vaginosis, Sex Transm Infect 80:8–11, 2004. Wood JC, Lu RM, Peterson EM, et al.: Evaluation of mycotrim-GU for isolation of Mycoplasma species and Ureaplasma urealyticum, J Clin Microbiol 22:789–792, 1985.

PROCEDURE 73.1

PROCEDURE 73.2

Preparing and Scoring Vaginal Gram Stains for Bacterial Vaginosis

Collection of Material From Suspected Herpetic Lesions

Organism Morphotype Lactobacillus-like (parallelsided, gram-positive rods)

Mobiluncus-like (curved, gram-negative rods) Gardnerella/Bacteroides-like (tiny, gram-variable coccobacilli and rounded, pleomorphic, gram-negative rods with vacuoles) Score 0–3 4–6 7–10

Number/Oil Immersion Field >30 5–30 1–4 5 30 5–30 1–4 65 years), immunosuppression or other severe underlying GI disease, use of proton pump inhibitors, and exposure to a health care setting. No single laboratory test will establish the diagnosis unequivocally. Two major types of tests are available for routine use: culture for direct detection of the organism and detection of cytotoxin (toxin A, B, or both) by cell culture cytotoxicity neutralization or EIA. In addition, most laboratories are using nucleic acid–based testing methods. Some of the commercially available nucleic acid–based assays include BD Gene Ohm (BD Diagnostics, La Jolla, CA), Cepheid Xpert (Cepheid, Sunnyvale, CA), FilmArray (BioFire Diagnostics Inc., Salt Lake City, UT), Roche LightCycler (Roche Applied Science), and ProGastro (Hologic, San Diego, CA). Nucleic acid–based testing demonstrates high sensitivity and specificity for the diagnosis of C. difficile–associated diarrhea. Diagnosis of a CDI should include a method to identify the organism and a method to assess the toxin status including immunoassay for either toxin A or B, cell culture cytotoxicity, or a nucleic acid method for the detection of toxin B. This prevents an incorrect diagnosis of clinical disease and administration of unnecessary medications or antibiotics in a patient colonized with a nontoxigenic strain.

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A

B

C

D

E

F •

Fig. 74.7 Colonies of a lactose-positive organism growing on xylose-lysine deoxycholate (XLD) agar (A) and Hektoen enteric (HE) agar (B). Colonies of Salmonella enteritidis (lactose-negative) growing on XLD (C) and HE agar (D). (Note how both agars detect H2S production.) Colonies of Shigella (lactose-negative) growing on XLD (E) and HE agar (F).

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, PA, 2020, Elsevier-Saunders. Buvens G, Pierard D: Low prevalence of STEC autotransporter contributing to biofilm formation (Sab) in verocytotoxin-producing E. coli isolates of humans and raw meats, Eur J Clin Microbiol Infect Dis 31:1463–1465, 2012. Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM.

Gavin PJ, Thomson RB: Diagnosis of enterohemorrhagic Escherichia coli infection by detection of Shiga toxins, Clin Microbiol Newsl 26:49, 2004. Goldenberg SD, Cliff PR, French GL: Glutamate dehydrogenase for laboratory diagnosis of Clostridium difficile infection, J Clin Microbiol 48:3050–3051, 2010. Herwaldt BL, Beach MJ: The return of Cyclospora in 1997: another outbreak of cyclosporiasis in North America associated with imported raspberries, Ann Intern Med 130:210–220, 1999. Kaye SA, Obrig TG: Pathogenesis of E. coli hemolytic-uremic syndrome, Clin Microbiol Newsl 18:49, 1996. Kehl SC: Role of the laboratory in the diagnosis of entero-hemorrhagic Escherichia coli infections, J Clin Microbiol 40:2711–2715, 2002. Kellner T, Parsons B, Chui L, et al.: Comparative evaluation of enteric bacterial culture and a molecular multiplex syndromic panel in

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children with acute gastroenteritis, J Clin Microbiol 57(6):e00205– e00219, 2019. https://doi.org/10.1128/JCM.00205-19. Loo VG, Poirier L, Miller MA, et  al.: A predominantly clonal multiinstitutional outbreak of Clostridium difficile-associated diarrhea with high morbidity and mortality, N Engl J Med 353:2442–2449, 2005. MacKenzie AM, Orrbine E, Hyde L, et  al.: Performance of the ImmunoCard STAT! E. coli O157:H7 test for detection of Escherichia coli O157:H7 in stools, J Clin Microbiol 38:1866– 1868, 2000. Marder EP, Cieslak PR, Cronquist AB, et al.: Incidence and trends of infections with pathogens transmitted commonly through food and the effect of increasing use of culture-independent diagnostic tests on surveillance-foodborne diseases active surveillance network, 10 U.S. Sites, 2013-2016, MMWR Morb Mortal Wkly Rep 66(15):397–403, 2017. McDonald LC, Killgore GE, Thompson A, et al.: An epidemic, toxin gene-variant of Clostridium difficile, N Engl J Med 353:2433– 2441, 2005. Novicki TJ, Daly JA, Mottice SL, et  al.: Comparison of sorbitol MacConkey agar and a two-step method which utilizes enzymelinked immunosorbent assay toxin testing and a chromogenic agar to detect and isolate enterohemorrhagic Escherichia coli, J Clin Microbiol 38:547–551, 2000.

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O’Horo JC, Jones A, Sternke M, et al.: Molecular techniques for diagnosis of Clostridium difficile infection: systematic review and metaanalysis, Mayo Clin Proc 87:643–651, 2012. Redondo N, Carroll A, McNamara E: Molecular characterization of Campylobacter causing human clinical infection using wholegenome sequencing: virulence, antimicrobial resistance and phylogeny in Ireland, PloS One 14(7):e0219088, 2019. https://doi. org/10.1371/journal.pone.0219088. Sansonetti PJ: Genetic and molecular basis of epithelial cell invasion by Shigella species, Rev Infect Dis 13(Suppl 4):S285–S292, 1991. Schmidt H, Hensel M: Pathogenicity islands in bacterial pathogenesis, Clin Microbiol Rev 17:14–56, 2004. Voth DE, Ballard JD: Clostridium difficile toxins: mechanisms of action and role in disease, Clin Microbiol Rev 18:247–263, 2005. Vuik F, Dicksved J, Lam SY, et al.: Composition of the mucosa-associated microbiota along the entire gastrointestinal tract of human individuals, United European Gastroenterol J 7(7):897–907, 2019. Wilkins TD, Bartlett JG: Clostridium difficile testing: after 20 years, still challenging, J Clin Microbiol 41:531–534, 2003. Zollner-Schwetz I, Hogenauer C, Joainig M, et al.: Role of Klebsiella oxytoca in antibiotic-associated diarrhea, Clin Infect Dis 47:e74– e78, 2008.

CASE STUDY 74.1 A 30-year-old male developed diarrhea with severe abdominal cramping 3 days after eating in a local restaurant. He became febrile and weak and went to his physician. A stool specimen was collected and immediately hand-carried to the laboratory. Numerous white blood cells and bacteria were seen in a wet mount, but the bacteria were noteworthy in that they were nonmotile. The patient was treated with ciprofloxacin. A culture was performed that grew a non–lactose-fermenting gramnegative rod that was identified as Shigella sonnei.

Questions 1. What agent of diarrhea becomes nonviable in a stool that has not been cultured within 30 minutes of collection? 2. If the culture is going to be delayed in transit, what preservative and storage temperature is best for preservation of the specimen? 3 After culturing the organism, what traditional method is used to identify Shigella to the species level, and what is the importance of such identifications?   

CASE STUDY 74.2 A 52-year-old female on immune-suppressive therapy for rheumatoid arthritis presents with a 2-day history of severe, watery diarrhea. This is accompanied by chills and reported fever of 101°F. She has no complaints of nausea or myalgia, but she does have severe abdominal cramps and loss of appetite. She had recently visited her niece, who had just purchased a pet turtle. She reports no other significant travel history or ill contacts and is treated with antidiarrheal medications as an outpatient. On the third day of illness she returns in a worsening condition with the following physical examination and laboratory results: • Vitals: Temp 102°F, respiration 20, BP 90/56, pulse 98 • Examination is essentially unremarkable with exception of diffusely tender abdomen; there is no rash or adenopathy, and the patient appears dehydrated • WBC 16K (reference range 5–10 × 109/L) with 82% granulocytes • Hgb = 14.5 • Liver function tests: amylase, lipase are all normal

• S  odium is 144 (reference range 135–145 mEq/L) • K = 3.0 (reference range 3.6–5.0 mEq/L) She is admitted to the hospital for rehydration, potassium replacement, and additional investigation. The initial diagnosis is sepsis syndrome. Because diarrhea is the major feature of her illness, a stool sample is submitted for culture and fecal WBC count. The patient rapidly convalesced by the second day of admission on empiric triple antibiotic therapy. While the patient was hospitalized, her niece became ill with severe diarrhea and was successfully treated as an outpatient.

Questions 1. What is the likely agent of infection in this case? 2. Did the administration of antibiotics improve or worsen the patient’s condition? Explain your answer. 3. Why did the niece recover so quickly, whereas the initial patient suffered a much more severe disease including dehydration and sepsis?   

Chapter Review 1. Self-limiting food poisoning is often a result of ingestion of: a.  C. botulinum spores b. Contaminated hamburger containing E. coli c. Salads or other foods contaminated with S. aureus d. V. cholerae–contaminated water 2. What does enterotoxic diarrheal disease produce? a. Bloody mucus–containing diarrhea b. Profuse watery stool c. Increased fecal WBC counts d. Bloody diarrhea 3. Repression of normal microbiota by the intake of antimicrobials often results in a GI infection with: a. Enteropathogenic E. coli b. P. aeruginosa c.  C. difficile d. Candida albicans

4. All the following protect the GI tract from infection except: a. Peristaltic movement b. Resident bacterial microbiota c. Mucus d. Alkaline pH 5. A patient presented with headache and malaise and is slightly emaciated. The patient indicates that she has had watery diarrhea for approximately 24 hours. The infection is most likely: a. A result of infection with Shigella sp. b. Infection in the small intestine c. Infection in the large intestine d. A result of infection with E. coli 6. Several tests are available for the detection of enterotoxin, including which of the following? a. Immunodiffusion, serology, culture b. Immunodiffusion, latex agglutination, or molecular (gene identification) 1063.e1

1063.e2 PA RT V I I     Diagnosis by Organ System

c. PCR, immunodiffusion, culture d. Culture followed by serology 7. Which organism is associated with thrombotic thrombocytopenia purpura? a.  S. dysenteriae b. C. difficile c. STEC d. V. cholerae 8. Which of the following is a flagellated protozoan that attaches to the intestine via a ventral sucker? a.  Cryptosporidium b. Cystoisospora spp. c.  Strongyloides stercoralis d. G. duodenalis 9. A patient presents to the emergency room with fever and bloody, mucus-laden diarrhea. The patient is complaining of abdominal cramping. What is the likely agent of infection? a.  Salmonella b. E. coli c.  Shigella d. C. difficile 10.  True or False _____ E. coli O157:H7, Shigella, and C. difficile can withstand the exposure to gastric acids. _____ Enterotoxigenic diarrhea is always the result of an infection with a GI pathogen.

_____ Failure to isolate any enteric pathogens in a stool is diagnostic for no evidence of infection. _____ Stool specimens contain lots of debris, including plant material, metabolic by products, and bacteria, therefore negating the necessity for transport media. 11.  Matching: Match each term with the correct description. _____ C. botulinum _____ Salmonella _____ C. jejuni _____ C. difficile _____ ETEC _____ C. perfringens _____ G. duodenalis _____ esophagitis _____ V. cholerae _____ B. cereus



a. health care– associated b. rice c. honey d. eggs e. microaerophilic f. subunits A and B g. heat-labile toxin h. meats and gravies i. traveler’s diarrhea j. HIV

12.  Short Answer (1) Describe a typical protocol for the media requirements used for culture identification of enteric pathogens. Why is it still important to culture enteric pathogens if molecular testing is used for identification? (2) Describe the proper laboratory diagnosis for C. difficile–associated diarrhea.

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Skin, Soft Tissue, and Wound Infections OBJECTIVES 1. Identify the layers of the skin and describe the function of the skin in host defense, including the physical and chemical properties. 2. List the organisms that colonize the skin and are considered normal microbiota. 3. Describe the mechanisms that allow bacteria to invade and cause skin and soft tissue infections. 4. Define each of the following manifestations of skin infection: Macule Papule Nodule Pustule Vesicle Bulla Scales Ulcer 5. Characterize each of the following types of infection, and describe the associated laboratory diagnosis: Folliculitis Furuncle Carbuncle Erysipelas Erythrasma Erysipeloid Impetigo Cellulitis Dermatophytoses Necrotizing fasciitis Myositis 6. List an organism that commonly causes the following types of infection, and describe the associated laboratory diagnosis: Postoperative Bite Burn 7. Define sinus tract, including organisms associated with this condition. 8. Identify the pathogens most commonly associated with infections in patients with diabetes mellitus. 9. Describe and evaluate a specimen submitted for culture from the following types of infection: ulcers, nodules or abscess, pyoderma or cellulitis, vesicles or bullae, sinus tracts, fistula, burns, postsurgical wounds, and bite infections. 10. Correlate patient signs and symptoms with laboratory results to identify the etiologic agent associated with a skin, soft tissue, or wound infection. 1064

General Considerations The skin serves as a barrier between the internal organs and the external environment. Skin is subjected to frequent trauma and therefore is at risk of infection. In addition, manifestations visible on the surface of the skin can provide clues for the identification of an internal systemic disease.

Anatomy of the Skin The skin is divided into two distinct layers: the epidermis (the outermost layer), and the dermis. The subcutaneous tissue beneath the dermis connects the skin to underlying structures (Fig. 75.1). The epidermis is made up of stratified squamous epithelium. Hair follicles, sebaceous glands (oil-producing), and sweat glands open to the skin surface through the epidermis. The dermis is composed of dense connective tissue rich in blood and nerve endings, and some hair follicles and sebaceous glands originate here. The subcutaneous tissue contains loose connective tissue and is rich in fat. Deeper hair follicles and sweat glands originate in this layer. Below the subcutaneous layer are thin fascial membranes (sheets or bands of fibrous tissue) covering muscles, ligaments, and other connective tissue. 

Function of the Skin The skin is the body’s largest and thinnest organ. It forms a self-repairing and protective boundary between the body’s internal environment and the external environment. Skin plays a crucial role in the control of body temperature, excretion of water and salts, synthesis of important chemicals and hormones, and as a sensory organ. The skin has an important protective function because of the composition of the outermost layer of the epidermis, which is composed of cells containing keratin, a water-repellent protein. The skin’s normal microbiota, pH, and chemical defenses (high salt and acidic environment) also help prevent colonization by many pathogens. Examples of resident microbiota are listed in Box 75.1, although variation can occur among people. 

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Hair

Epidermis

Dermis

Sebaceous gland

Sweat gland

Hair follicle Subcutaneous tissue

Fascia Muscle

• Fig. 75.1  Diagram of the skin. • BOX 75.1 Normal Microbiota of the Skin Diphtheroids Staphylococcus epidermidis Other coagulase-negative staphylococci Cutibacterium acnes

Prevalence, Etiology, and Pathogenesis Approximately 15% of all patients who seek medical attention have either some skin disease or a skin lesion, many of which are infectious. Various bacteria, fungi, parasites, and viruses may be involved. These infections can include one or several causative agents. Because of the diversity of etiologic agents and the potential complexity of these infections, only the most common infections involving the skin and subcutaneous tissues will be addressed. Skin infections can arise from the invasion of certain organisms from the external environment through breaks in the skin or from organisms that reach the skin through the blood as part of a systemic disease. In some infections, such as staphylococcal scalded-skin syndrome, toxins produced by the bacteria cause skin lesions. In others, lesions are a result of the host’s immune response to microbial antigens. Because of the diversity of etiologic agents, clinicians will often rely on the appearance of skin lesions for diagnostic clues to determine the required laboratory testing. The physical characteristics of the lesions can indicate the need for smear, culture, biopsy, or surgical procedures. Some of the terms most commonly used to describe manifestations

of skin infections are provided in Table 75.1. Fig. 75.2 shows examples of some skin lesions. 

Skin and Soft Tissue Infections Infections of the Epidermis and Dermis Numerous infections of the skin may occur. Several of the most common are discussed here.

Infections in or Around Hair Follicles Folliculitis, furuncles, and carbuncles are localized abscesses either in or around hair follicles. These infections are distinguished from one another based on size and the extent of involvement in subcutaneous tissues. Table 75.2 summarizes each infection’s respective clinical features. For the most part, these infections are precipitated by blockage of the hair follicle with skin oils (sebum) or because of minor trauma resulting from friction, such as that caused by clothes rubbing against the skin. Staphylococcus aureus is the most common etiologic agent for all three infections. Members of the Enterobacterales, Malassezia furfur, and Candida spp. may also cause folliculitis. Outbreaks of folliculitis caused by Pseudomonas aeruginosa have been reported to be associated with the use of whirlpools, swimming pools, and hot tubs. 

Infections in the Keratinized Layer of the Epidermis Because of their ability to utilize the keratin in the epidermal cells, the dermatophyte fungi are significant and well-suited pathogens for infection. Unlike the previously discussed infections, dermatophytes do not invade the deeper layers of

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TABLE 75.1    Manifestations of Skin Infections

Term

Description

Possible Etiologic Agents (Infections)

Macule

A circumscribed (limited), flat discoloration of the skin

Dermatophytes Treponema pallidum (secondary syphilis) Viruses, such as enteroviruses (exanthems rashes)

Papule

An elevated, solid lesion ≤5 mm in diameter

Human papillomavirus types 3 and 10 (flat warts) Pox virus (molluscum contagiosum) Sarcoptes scabiei (scabies) Staphylococcus aureus, Pseudomonas aeruginosa, etc. (folliculitis)

Nodule

A raised, solid lesion >5 mm in diameter

Corynebacterium diphtheriae Sporothrix schenckii Miscellaneous fungi (subcutaneous mycoses) Mycobacterium marinum Nocardia spp. S. aureus (furuncle)

Pustule

A circumscribed, raised, pus-filled (leukocytes and fluid) lesion

Candida spp. Dermatophytes Herpes simplex virus Neisseria gonorrhoeae (gonorrhea) S. aureus (folliculitis) S. aureus or group A streptococci (impetigo) Varicella-zoster virus (chickenpox)

Vesicle

A circumscribed, raised, fluid-filled (blisterlike) lesion ≤5 mm in diameter

Herpes simplex virus Varicella-zoster virus (chickenpox and shingles)

Bulla

A circumscribed, raised, fluid-filled lesion >5 mm in diameter

Clostridial species (necrotizing gas gangrene) Herpes simplex virus Other gram-negative bacilli S. aureus (bullous impetigo and scalded-skin syndrome) Vibrio vulnificus and other Vibrio spp.

Scales

Dry, horny, platelike lesions

Dermatophytes (tinea)

Ulcer

A lesion with loss of epidermis and dermis

Bacillus anthracis (cutaneous anthrax) Bowel microbiota (decubiti) Haemophilus ducreyi (chancroid) T. pallidum (chancre of primary syphilis)

Adapted from Lazar AJF. Robbins Basic Pathology. 8th ed. St Louis: Saunders; 2007.

skin. Because keratin is also present in hair and nails, these fungi may also cause superficial infections at these sites (see Chapter 60 for more information). 

Bacillus anthracis, Corynebacterium diphtheriae, Mycobacterium marinum, Nocardia spp., and Sporothrix schenckii. 

Infections in the Deeper Layers of the Epidermis and Dermis

Infections of the Subcutaneous Tissues

Most infections in the deeper layers of the epidermis and dermis result from the inoculation of microorganisms by traumatic breaks in the skin. These superficial skin infections usually do not require surgical intervention. Table 75.3 summarizes these infections. In most instances, these infections resolve with local care and only occasionally require antimicrobial therapy. Cutaneous ulcers usually involve a loss of epidermal and part of the dermal tissues. In contrast, nodules are inflammatory foci in which the epidermal and dermal layers remain largely intact. Various bacteria and fungi can cause ulcerative or nodular skin lesions after direct traumatic inoculation. Examples of these causative agents include

Infections of the subcutaneous tissues may manifest as abscesses, ulcers, or boils. The most common etiologic agent of subcutaneous abscesses in healthy individuals is S. aureus. Many subcutaneous abscesses are polymicrobial. To a large degree, the organisms isolated from these abscesses depend on the site of infection. For example, anaerobes are commonly isolated from abscesses of the perineal, inguinal, and buttock area, whereas nonperineal infection is commonly caused by a polymicrobial infection containing facultative organisms. Progressive synergistic gangrene, or Meleney ulcer, is a slowly progressive infection of the subcutaneous tissue that usually begins as an ulcer after trauma or surgery. The infection leads to subcutaneous necrosis and enlargement of a

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A

B

C

D •

Fig. 75.2  (A) Viral maculopapular rash. (B) Furuncle. (C) Folliculitis caused by Staphylococcus aureus showing numerous pustules. (D) Desquamation (shedding or scaling) of the skin resulting from scarlet fever caused by group A streptococci. (A and D, From Habif TB. Clinical Dermatology: A Color Guide to Diagnosis and Therapy. 3rd ed. St Louis: Mosby; 1996.)

TABLE 75.2    Infections Involving Hair Follicles

Infection

Skin Manifestations

Folliculitis (minor infection of hair follicles)

Papules or pustules that are pierced by a hair and surrounded with redness

Furuncle (boil)

Abscess that begins as a red nodule in a hair follicle that ultimately becomes painful and full of pus

Carbuncle

Furuncles that coalesce and spread more deeply to the dermis and subcutaneous tissues; they usually have multiple sites, which drain to the skin surface (sinuses)

visible ulcer. This is a true polymicrobial infection in which microaerophilic streptococci grow synergistically with S. aureus. The infection may also include other facultative or anaerobic organisms. In many instances, infections of the epidermis and dermis extend deeper and become subcutaneous infections and may even reach the fascia or muscle. For example, erysipelas (Fig. 75.3) can develop into subcutaneous cellulitis and eventually necrotizing fasciitis. Similarly, folliculitis can readily develop into a subcutaneous abscess or carbuncle that can extend to the fascia. Cellulitis can also extend to the subcutaneous tissues (Fig. 75.4). Anaerobic cellulitis is associated with the production of large amounts of gas by organisms that may be present in the subcutaneous tissue. This type of infection is most often located in the extremities and is particularly common among patients with

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TABLE 75.3    Infections of the Epidermal and Dermal Layers of the Skin

Infection

Key Features of Infection

Etiologies

Other Comments

Erysipelas

Primarily involves the dermis and most superficial parts of the subcutaneous tissue; lesions are painful, red, swollen, and indurated; patients are febrile, and regional lymphadenopathy (swollen glands) is often present; lesion has a marked, well-demarcated, raised border (Fig. 75.3)

Group A streptococci (Streptococcus pyogenes [sometimes groups B, C, or G streptococci])

Infants, children, and elderly individuals are most affected; primarily a clinical diagnosis

Erythrasma

Chronic infection of the keratinized layer of the epidermis; lesions are dry, scaly, itchy, and reddish brown

Corynebacterium minutissimum— possible cause

Common in diabetics; resembles dermatophyte infection

Erysipeloid

Purplish-red, nonvesiculated skin lesion with an irregular, raised border; the lesions itch and burn; fever and other systemic symptoms are uncommon

Erysipelothrix rhusiopathiae

Uncommon; considered an occupational disease

Impetigo

Erythematous (red) lesions that may be bullous (less common) or nonbullous

Nonbullous—group A streptococci (S. pyogenes) Bullous—Staphylococcus aureus

Cellulitis

Diffuse, spreading infection involving the deeper layers of the dermis; lesions are ill-defined, flat, painful, red, and swollen; patients have fever, chills, and regional lymphadenopathy (Fig. 75.4)

Commonly: Group A streptococci and other streptococci, S. aureus Less common: Aeromonas spp., Vibrio spp., and Haemophilus influenzae (typically affects young children)

Dermatophytoses

Superficial fungal infections of the skin and its appendages (i.e., ringworm, athlete’s foot, jock itch, and infections of nails and hair)

Epidermophyton, Microsporum, and Trichophyton spp.

Hidradenitis

Chronic infection of obstructed apocrine (sweat) glands in the axillar, genital, or perianal areas with intermittent discharge of often foul-smelling pus

S. aureus, Streptococcus anginosus group, anaerobic streptococci, and Bacteroides spp.

Infected pilonidal tuft cyst or hairs

Pain and swelling; redness

Anaerobes, including Bacteroides fragilis group, Prevotella, Fusobacterium, anaerobic gram-positive cocci, and Clostridium spp.

Primarily a clinical diagnosis

• Fig. 75.3  Erysipelas caused by group A streptococci. •

Fig. 75.4 Cellulitis. (From Farrar WE, Wood MJ, Innes JA, et  al. Infectious Diseases: Text and Color Atlas. 2nd ed. London: MosbyWolfe; 1992.)

CHAPTER 75  Skin, Soft Tissue, and Wound Infections

diabetes. The infection may involve the neck, abdominal wall, perineum, connective tissue, or other areas. Anaerobic cellulitis may also occur as a postoperative condition. The onset and spread of the lesion are usually slow, and patients may not immediately show obvious systemic effects. The causative agents in deep tissue infections are almost always a mixture of aerobic, facultative, and anaerobic organisms. Common aerobic, facultative organisms include Escherichia coli, alpha-hemolytic and nonhemolytic streptococci, and S. aureus. However, group A streptococci and other members of the Enterobacterales may be encountered as well. The anaerobes are typically found in greater numbers and in more varieties and include Peptostreptococcus spp., Bacillus fragilis group strains, Prevotella spp., Porphyromonas spp., other anaerobic gram-negative bacilli, and clostridia. Bacteremia is not usually present. 

Infections of the Muscle Fascia and Muscles There are several uncommon, yet serious or potentially serious, forms of deep and often extensive soft tissue and skin infections.

Necrotizing Fasciitis Necrotizing fasciitis is a serious infection that is relatively uncommon. The basic pathology involves infection of the fascia overlying the muscles, often with involvement of the overlying soft tissue. At the fascial level, no barrier exists to prevent the spread of infection, so fasciitis may extend widely and rapidly to involve large areas of the body in a short amount of time. There are three distinct types of necrotizing fasciitis based on the bacteriological agents involved in the infection. Type I is polymicrobial and typically involves Bacteroides or Peptostreptococcus in combination with one or more facultative organism, such as group A streptococci or Enterobacterales. Type II is generally a monomicrobic infection with group A streptococci but may be associated with another species, often S. aureus. Type III is a result of infection with marine gram-negative pathogens, such as Aeromonas hydrophilia or Vibrio spp. Necrotizing fasciitis is most often acute and can affect any area of the body. 

Progressive Bacterial Synergistic Gangrene Progressive bacterial synergistic gangrene is usually a chronic necrotic condition of the skin most often encountered as a postoperative complication, particularly after abdominal or thoracic surgery or other medical procedures and devices, such as a colostomy (opening that connects the colon to the abdominal wall). The lesions may be extensive and, with involvement of the abdominal wall, may lead to evisceration (extrusion of the internal organs). As the name implies, this is a synergistic polymicrobial infection with microaerophilic streptococci and S. aureus. Other organisms may also be present, including anaerobic streptococci, Proteus spp., other gram-negative bacilli, or other facultative and anaerobic bacteria. This type of infection is uncommon. Cultures should be taken from the advancing outer edge

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of the lesion (not the central portion of the wound). This prevents missing cultivation of microaerophilic streptococci that may be involved in the infection. 

Myositis Myositis (inflammation of muscle) can be caused by a variety of organisms. The nature of the pathologic process is variable, sometimes involving extensive necrosis of muscle or focal collections of suppuration (pus) in muscle (pyomyositis). This may occur because of a penetrating wound, vascular insufficiency in an extremity, or another contiguous infection. The most common cause of acute bacterial myositis from hematogenous spread is S. aureus. Categories of bacterial myositis include pyomyositis, psoas abscess (pus in the iliopsoas muscle compartment), S. aureus myositis, group A streptococcal necrotizing myositis, group B streptococcal myositis, clostridial gas gangrene, and nonclostridial myositis. Serious vascular problems resulting from loss of blood supply may lead to death of muscle tissue, leading to a secondary infection (vascular gangrene). Organisms that cause myositis or other muscle pathology are listed in Box 75.2. 

Wound Infections Besides skin and soft tissue infections, wound infections occur primarily from breaks in the skin because of complications associated with surgery, trauma, and bites, or from diseases that interrupt the mucosal or skin surface.

• BOX 75.2 Organisms Producing Myositis or Other

Muscle Pathology

Clostridium perfringens Clostridium novyi Clostridium septicum Clostridium histolyticum Clostridium sordellii Clostridium sporogenes Paraclostridium bifermentans (previously Clostridium) Bacillus spp. Aeromonas spp. Peptostreptococcus spp. Microaerobic streptococci Bacteroides spp. Enterobacterales Staphylococcus aureus Group A streptococci (Streptococcus pyogenes) Pseudomonas spp. Vibrio vulnificus Mycobacterium tuberculosis Salmonella enterica serotype Typhi Legionella spp. Rickettsia spp. Viruses Trichinella spp. Taenia solium Toxoplasma gondii

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Microbial biofilms have also increased the length and chronicity of wound infections.

Postoperative Infections Surgical site infections are among the most common health care–associated infections. Sources of surgical wound infections can include the patient’s normal microbiota or organisms present in the hospital environment. These organisms are introduced to the patient by medical procedures or underlying disease or trauma (e.g., burns). The nature of the infecting organism depends on the patient’s underlying condition and the location of the medical treatment or procedure. The most common organism involved in postoperative infections is S. aureus. Surgical procedures in the colorectal or other lower gastrointestinal areas have the highest incidence of postoperative infections because of the presence of intestinal bacteria. These infections are most likely to be caused by enteric gram-negative bacteria, anaerobes, and enterococci. Principal pathogens are listed in Box 75.3. 

Bites Bite wounds can be the result of human teeth, as well as a large variety of domestic or wild animals. The infectious agents associated with the bite may come from the environment, the victim’s normal skin microbiota, and the oral flora of the biter. This makes the variety of potential infectious agents quite diverse. Human bite (Fig. 75.5) infections can be attributed to occlusion bites or closed-fist injuries. Not surprisingly, the most commonly isolated organisms are normal oral microbiota. Most frequently isolated are viridans streptococci (particularly Streptococcus anginosus), S. aureus, and Eikenella corrodens. Common anaerobes isolated include Prevotella, Fusobacterium, Veillonella, and Peptostreptococcus species. These infections are usually polymicrobial and contain both aerobic and anaerobic organisms.

Animal bites account for approximately 1% of all emergency room visits in the United States. The most common animal bites are from domestic cats and dogs. Bites from these animals (Fig. 75.6) usually are infected with organisms commonly found in the animal’s oral and nasal fluids. The most commonly isolated aerobes are Pasteurella, Streptococcus, Neisseria, and Staphylococcus species. The most commonly isolated anaerobes are Fusobacterium, Bacteroides, and Porphyromonas species. Similar to human bites, animal bite wound infections are usually polymicrobial and include aerobes and anaerobes. Other far less common animal bites may also become infected. Rat bite infections are usually caused by Streptobacillus moniliformis. Snakebites may become infected with A. hydrophilia. 

Burns Burns are a significant cause of mortality due to hospitalassociated infections; 42% to 65% of deaths in burn patients are associated with infection. Infected burn wounds may be associated with many organisms, causing significant mortality, and may interfere with the success of skin grafts. Bacteria cause 70% of burn wound infections, 20% to 25% are caused

• BOX 75.3 Organisms Encountered in

Postoperative Wound Infections

Staphylococcus aureus Coagulase-negative staphylococci Streptococcus pyogenes Streptococcus anginosus group streptococci (Streptococcus anginosus, Streptococcus constellatus, Streptococcus intermedius) Microaerophilic streptococci Enterococci Proteus, Morganella, and Providencia spp. Other Enterobacterales Escherichia coli Pseudomonas spp. Candida spp. Bacteroides spp. Prevotella and Porphyromonas spp. Fusobacterium spp. Clostridium spp. Peptostreptococcus spp. Non–spore-forming, anaerobic, gram-positive bacilli

• Fig. 75.5  Human bite infection.

• Fig. 75.6  Animal bite infection caused by Pasteurella spp.

CHAPTER 75  Skin, Soft Tissue, and Wound Infections

by fungi, and anaerobic organisms and viruses cause 5% to 10%. Burn wound infections can commonly be identified as four types: impetigo; surgical infections; cellulitis; bacteremia and invasive, systemic infections. Factors that contribute to the development of infection include loss of the skin barrier, coagulated proteins and other microbial nutrients, loss of vascularity of the wound, dehydration of surrounding tissue, and the inflammatory response of the patient’s immune system. Gram-positive organs tend to be isolated from early infections, whereas the incidence of infection with gramnegative organisms increases with the length of hospitalization. The organisms isolated most often from burns include S. aureus, P. aeruginosa, enterococci, Enterobacter spp., and E. coli. Other organisms, such as fungi (e.g., Candida spp., Aspergillus niger, Fusarium spp., and Mucor spp.) and viruses may also be involved in invasive burn infections. The risk of infections with multidrug-resistant organisms also increases with the length of stay for burn patients. These organisms include P. aeruginosa, Acinetobacter baumannii, Stenotrophomonas maltophilia, and S. aureus. 

Special Circumstances Regarding Skin and Soft Tissue Infections In addition to the infections previously discussed, other circumstances can cause skin and underlying soft tissue to become infected. Some of these infections are associated with an immunocompromised host; others are manifestations of systemic infection. 

Infections Related to Vascular and Neurologic Problems Frequently, a patient with infections associated with vascular or neurologic problems has diabetes mellitus. These patients have a high risk of developing infections, especially in their lower extremities. The excess glucose present in their blood can result in impaired microvascular circulation and peripheral motor neuropathy, leading to an increased risk of infection. Any skin-damaging injury or surgery greatly increases that risk. In addition, because of the complications associated with impaired circulation and neuropathy, the infected tissue of a diabetic patient does not heal as rapidly as that of a healthy individual. An estimated 25% of adults with diabetes will develop a foot infection, and the risk increases with age. Foot infections can lead to amputations and greatly increased mortality. Periodically, an acute cellulitis and lymphangitis may be associated with chronic, low-grade infection, thereby making control of the patient’s diabetes difficult. Peripheral vascular disease unrelated to diabetes may also predispose a patient to skin and soft tissue infections, but usually these infections are easier to manage because there is no associated neuropathy. Foot infections in diabetic patients can accelerate dramatically, producing devastating consequences without proper treatment. Therefore, appropriate techniques used to obtain a microbiologic sample are critical. Culture of aspirated fluid or pus, not surface swabbing, is more likely to

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• Fig. 75.7  Sacral decubitus ulcer.

yield a causative agent—particularly if taken from a deep pocket within the wound. In addition, culture of debrided infected tissue improves the diagnosis of these infections. The most common bacteria isolated from mild to moderate diabetic foot infections include S. aureus, group B streptococci, members of the Enterobacterales, and anaerobes. More severe infections are usually polymicrobial. Extension of the infection into the underlying bone produces a difficult-to-manage osteomyelitis. Definitive diagnosis of this infection requires a specimen of bone obtained during open or percutaneous biopsy. Venous insufficiency may also predispose an individual to infection, again primarily in the lower extremities (often in the calf or lower leg rather than the foot). Infections related to poor blood supply often involve S. aureus and group A streptococci. Those with open ulcers may become colonized with Enterobacterales and P. aeruginosa. Anaerobes are also frequently involved in these infections, as a result of the blood supply creating anaerobic conditions. Anaerobes that may be involved include Bacteroides fragilis group, Prevotella, Porphyromonas, Peptostreptococcus and, less commonly, Clostridium species. Another common type of infection in this general category, especially in the elderly or chronically ill, bedridden patient, is infected decubitus ulcer (pressure sore; Fig. 75.7). Anaerobic conditions are present in the lesions as a result of tissue necrosis. Most of these lesions are located near the anus or on the lower extremities. Because these patients are relatively helpless and have limited mobility, the ulcers may become contaminated with gastrointestinal bacteria, leading to chronic infection. These conditions contribute to further death of tissue and extension of the decubitus ulcer. Bacteremia is a possible complication; B. fragilis group is often involved, as are clostridia and other enteric bacteria. Health care–associated pathogens, such as S. aureus and P. aeruginosa, may also be recovered. 

Sinus Tracts and Fistulas Sometimes, a deep-seated infection will develop a channel, called a sinus tract, to the skin surface. The sinus tract will

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A

B

• Fig. 75.8  Actinomycosis. (A) Note “lumpy jaw.” (B) Side view. Note sinuses in skin of face and neck.

drain fluid and pus onto the skin. The infections involved are often chronic and may include osteomyelitis. The organisms commonly involved in sinus tract formation with an underlying osteomyelitis include S. aureus, various mem­ bers of the Enterobacterales, P. aeruginosa, anaerobic gramnegative bacilli, and anaerobic gram-positive cocci. In the case of actinomycosis (Fig. 75.8), with or without bone involvement, the organisms involved include Actinomyces spp., Aggregatibacter actinomycetemcomitans, Propionibacterium propionicum, Prevotella or Porphyromonas species, and other non–spore-forming anaerobes. Chronic draining sinuses may also be found in patients with tuberculosis and atypical mycobacterial infection, Nocardia infection, and infections associated with implanted foreign bodies. Curetting or biopsy specimens from the debrided, cleansed sinus should be used for culture. Abnormal channels connecting epithelial surfaces, either between two internal organs or between an organ and the skin epithelium, are known as fistulas. Infections associated with fistulas often pose insurmountable problems in terms of collecting a meaningful specimen because the organ involved may contain indigenous microbiota. Examples include perirectal fistulas from the small bowel to the skin associated with Crohn disease or chronic intraabdominal infection. When the bowel is involved, cultures for specific organisms, such as mycobacteria or Actinomyces, are useful. An attempt should always be made to rule out specific associated underlying causes, such as tuberculosis, actinomycosis, and malignancy. A biopsy should be performed in these situations. 

Systemic Infections and Skin Manifestations Cutaneous manifestations of systemic infections, such as bacteremia or endocarditis, may be important clues for the clinician. These represent an opportunity for direct detection or culture for the presence of an organism. For example, a scraping of petechiae (tiny red hemorrhagic spots in

the skin) from patients with meningococcemia may demonstrate the presence of gram-negative diplococci. In other patients, the skin lesion may represent a metastatic infection. In Vibrio vulnificus sepsis, dramatic-appearing cutaneous ulcers with necrotizing vasculitis or bullae may be seen. In some patients, skin lesions may represent a noninfectious complication of a local or systemic infection, such as scarlet fever or toxic shock syndrome. Various organisms involved in systemic infections capable of producing cutaneous lesions are listed in Box 75.4. 

Laboratory Diagnostic Procedures Infections of the Epidermis and Dermis For many of the infections of the epidermis and dermis, such as impetigo, folliculitis, cellulitis, and erysipelas, diagnosis is generally based on clinical observations. Table 75.3 provides the key features and etiologic agents of these infections.

Erysipeloid In erysipeloid, usually the Gram stain or culture of superficial wound drainage is negative. However, culture of a fullthickness skin biopsy taken at the margin of the lesion can confirm the clinical diagnosis. 

Superficial Mycoses and Erythrasma If a dermatophyte infection is suspected, the lesion is cleaned, and scrapings are obtained from the active border of the lesion. These scrapings should be treated with 10% potassium hydroxide and examined for the presence of hyphae. The specimen may also be cultured if necessary (Chapter 59). A Woods lamp examination of the skin lesions for tinea versicolor may show golden-yellow fluorescence. Erythrasma, which is caused by infection with Corynebacterium minutissimum, can be diagnosed by making smears from the lesion revealing gram-positive pleomorphic

CHAPTER 75  Skin, Soft Tissue, and Wound Infections

• BOX 75.4 Organisms Involved in Systemic

Infection With Cutaneous Lesions

Viridans streptococci Staphylococcus aureus Enterococci Group A and other beta-hemolytic streptococci Neisseria gonorrhoeae Neisseria meningitidis Haemophilus influenzae Pseudomonas aeruginosa Pseudomonas spp. Listeria monocytogenes Vibrio vulnificus Salmonella enterica serotype Typhi Mycobacterium tuberculosis Mycobacterium leprae Treponema pallidum Leptospira spp. Streptobacillus moniliformis Bartonella bacilliformis Bartonella henselae Rickettsia spp. Candida spp. Cryptococcus spp. Blastomyces dermatitidis Coccidioides immitis Histoplasma capsulatum

bacilli. If necessary, skin scrapings may be cultured in media containing serum. A Woods lamp examination of the skin lesions may reveal a coral red fluorescence resulting from the production of porphyrin by C. minutissimum. 

Erysipelas and Cellulitis As previously mentioned, diagnosis of erysipelas and cellulitis can generally be made based on clinical observation. Swab specimens from bullae, pustules, or ulcers may be cultured. Culturing of needle aspiration or punch biopsy specimens is not recommended and rarely informative. Blood cultures are also typically negative. 

Vesicles and Bullae These fluid-filled lesions characteristically involve specific organisms (Table 75.1). Material in a blisterlike lesion may vary from serous (resembling serum) fluid to serosanguineous (composed of serum and blood) fluid or hemorrhagic (bloody) fluid. Large bullae may permit withdrawal of fluid by needle and syringe aspiration. Specimens from tiny vesicles may need to be collected with a swab. The clinician can usually anticipate whether the lesion is viral or bacterial and may even suspect an organism. Specimens should be submitted for viral or bacterial culture based on the clinical presentation. Bullous lesions are often associated with sepsis, requiring the collection of blood for nucleic acid testing or cultures. Gas gangrene caused by Clostridium perfringens and other clostridia is characterized by bronzed skin with bullous lesions. Gram stain of the fluid from the lesions typically reveals gram-positive bacilli. 

1073

Infections of the Subcutaneous Tissue Proper collection and transport of specimens are important factors in the laboratory diagnosis of all infections. Specimen collection for the diagnosis of subcutaneous tissue infection is particularly difficult, because many of these lesions are open and readily colonized by nosocomial pathogens that may not be involved in the systemic underlying infection. The most reliable specimens for determining the etiology of ulcers and nodules are those obtained from the base of the ulcer or nodule after removal of overlying debris, or by surgical biopsy of deep tissues, avoiding contact with the superficial layers of the lesion. A Gram stain of the specimen should be performed, and material aerobically cultured on blood and MacConkey agar. If fungi, Nocardia spp., or mycobacterial infection is suspected, appropriate fungal media or mycobacterial media should be used. These culture methods are addressed in greater detail in Chapters 42 and 59, respectively. Similar challenges exist in collecting material for culture from sinus tracts. The material should be obtained from the deepest portion of the sinus tract. If systemic symptoms, such as fever, are present, blood cultures should also be collected. A Gram stain should be routinely performed. Cultures should be inoculated to recover both facultative and anaerobic bacteria in the same manner as for surgical wounds. Nucleic acid–based assays may be used to directly identify organisms, such as S. aureus, in wound infections. In addition, when deep tissue or bone infections are suspected and yield negative culture results, molecular assays may aid in the identification of the infectious agent. 

Infections of the Muscle Fascia and Muscles Blood cultures should always be collected from patients with significant myonecrosis. Transport of material (tissue is recommended, followed by purulent material, and then a swab) should be under anaerobic conditions. Gram stains should be routinely performed. Cultures should be inoculated to recover both facultative and anaerobic bacteria in the same manner as for surgical wounds. 

Wound Infections Postoperative Because anaerobic bacteria are involved in many of these infections, specimen collection should be completed carefully to avoid indigenous microbiota. Specimen transport in anaerobic conditions is essential. Unusual organisms associated with postsurgical wound infections, such as Mycoplasma hominis, Mycobacterium chelonae, Mycobacterium fortuitum, fungi, and even Legionella spp., should not be overlooked. A Gram-stained smear of material submitted for culture should be examined. Exudates from superficial wounds should routinely be inoculated to blood, Mac­ Conkey, and Colistin-nalidixic acid (CNA) agars, as well as an enrichment broth. Material from deep wounds should be inoculated onto media for both anaerobic and aerobic

1074 PA RT V I I     Diagnosis by Organ System

cultures. More detailed information regarding the processing of specimens for anaerobic cultures is presented in Chapters 40 and 41. 

Bites Bite wound infections usually involve relatively small lesions and minimal exudate. A swab specimen for aerobic culture and one in anaerobic transport media should be collected. Surrounding skin should be thoroughly disinfected before the specimen is obtained. The best material for culture is purulent exudate aspirated from the depth of the wound or samples obtained during surgery involving incision and drainage or debridement (removal of all dead and necrotic tissue). Gram-stained smears should be prepared and examined. For aerobic cultures, a minimum of blood, MacConkey, and chocolate agar should be inoculated. 

Burns For many burn patients, diagnosis of infection is based on clinical symptoms, signs, and examination of the burn wound. When possible, cultures should be performed on any purulent wound exudates, and blood cultures should also be collected. Surface specimens should be collected with a moistened sterile swab using a minimal amount of pressure. Sometimes a quantitative or semiquantitative culture (Evolve Procedure 75.1) of a tissue biopsy specimen is used for infection surveillance, or to identify the most prevalent organism in a polymicrobial infection. This type of culture is reported in colony-forming units (CFUs) per gram of tissue, with a result of 105 CFUs/g or more indicative of a potentially serious infection.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Bailey E, Kroshinsky D: Cellulitis: diagnosis and management, Dermatol Ther 24:229–239, 2011. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, PA, 2020, Elsevier-Saunders.

Capoor MR, Sarabahi S, Tiwari VK, et al.: Fungal infections in burns: diagnosis and management, Indian J Plast Surg 43:S37–S42, 2010. Carroll KC, Pfaller MA: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM Press. Crum-Cianflone NF: Bacterial, fungal, parasitic, and viral myositis, Clin Microbiol Rev 21:473–494, 2008. Humphreys H: Preventing and controlling the risk of post-operative surgical-site infections, Eur Infect Dis 2:110–112, 2008. Hurlow JJ, Humphreys GJ, Bowling FL, et al.: Diabetic foot infection: a critical complication, Int Wound J 15(5):814–821, 2018. Hurt JB, Maday KR: Management and treatment of animal bites, JAAPA 4:27–31, 2018. Lachiewicz AM, Hauck CG, Weber DJ, et  al.: Bacterial infections after burn injuries: impact of multidrug resistance, Clin Infect Dis 65(12):2130–2136, 2017. Lazar AJF: The skin. In: Robbins basic pathology, ed 8, St. Louis, MO, 2007, Saunders. Levy PY, Fenollar F: The role of molecular diagnostics in implantassociated bone and joint infection, Clin Microbiol Infect 18:1168– 1175, 2012. Lipsky BA: Medical treatment of diabetic foot infections, Clin Infect Dis 39(Suppl 2):S104–S114, 2004. Mayhall CG: The epidemiology of burn wound infections: then and now, Clin Infect Dis 37:543–550, 2003. Mena KD, Gerba CP: Risk assessment of Pseudomonas aeruginosa in water, Rev Environ Contam Toxicol 201:71–115, 2006. Murphy E: Microbiology of animal bites, Clin Microbiol Newsl 30:47–50, 2008. Oehler RL, Velez AP, Mizrachi M, et al.: Bite-related and septic syndromes caused by cats and dogs, Lancet Infect Dis 9:439–447, 2009. Polavarapu N, Ogilvie MP, Panthaki ZJ: Microbiology of burn wound infections, J Craniofac Surg 19:899–902, 2008. Salkind AR, Rao KC: Antibiotic prophylaxis to prevent surgical site infections, Am Fam Physician 83:585–590, 2011. Sankar RU, Biswas R, Raja S, et al.: Brain abscess and cervical lymphadenitis due to Paraclostridium bifermentans: a report of two cases, Anaerobe 51:8–11, 2018. Talan AD, Abrahamian FM, Moran GJ, et al.: Clinical presentation and bacteriologic analysis of infected human bites in patients presenting to emergency departments, Clin Infect Dis 37:1481–1489, 2003. Williams DT, Hilton JR, Harding KG: Diagnosing foot infection in diabetes, Clin Infect Dis 39(Suppl 2):S83–S86, 2004.

PROCEDURE 75.1 Semiquantitative Bacteriologic Culture of Tissue

Principle The degree or extent of bacterial wound contamination is directly related to the risk of wound sepsis. Because of this relationship, physicians use the results of a quantitative culture (the number of colony-forming units [CFUs] per gram of the eschar biopsy) in their management of severely burned patients. 

Method 1. Cut a piece of tissue, measuring several cubic millimeters, aseptically onto a small, preweighed, sterile urine cup. 2. Determine the weight of the tissue by subtracting the weight of the aluminum foil from the total weight. 3. Place the specimen and 2 mL of sterile nutrient broth in a sterile tissue grinder; macerate the specimen. 4. Inoculate 0.1 mL of sample to a blood agar plate, in duplicate, and an anaerobic blood agar plate (if indicated), in duplicate. In addition, inoculate 0.01 mL of sample using a calibrated loop to a blood agar plate, in duplicate. Spread the inoculum on the plates with a sterile glass spreading rod or a loop.

5. Incubate plates in 5% to 10% carbon dioxide overnight and count the colonies of bacteria on the plates that contain 30 to 300 CFUs. If more than 300 colonies are obtained on both plated dilutions, the factor 300 is used as N for calculations and the result is considered greater than the value. 6. Calculate the number of CFUs per gram of tissue with the following formula: Number of CFUs counted × Reciprocal of volume of homogenate inoculated (10-1 or 10-2) × 2 (volume of diluent used for tissue homogenization) ÷ weight of tissue For example, for a tissue that weighed 0.002 g, 68 CFUs were observed on the plate that received the 10–2 dilution of suspension: 68 × 102 × 2 0. 002

=

136 × 102 2 × 10−3

= 6.8 × 106 CFU / g

  

Modified from a method published by Buchanan K, Heimbach DM, Minshew BH, et al. Comparison of quantitative and semiquantitative culture techniques for burn biopsy. J Clin Microbiol. 1986;23:258.

  

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1074.e2 PA RT V I I     Diagnosis by Organ System

CASE STUDY 75.1 A 59-year-old female presented with swelling and pain in an injured left hand and enlarged lymph nodes in the axial region of her left arm. Her hand was injured when she stopped on her way home from work to rescue a small kitten that had wandered into the road. When she attempted to pick up the frightened kitten, it scratched her left hand. A specimen for culture was obtained by aspiration from the wound. A Gram stain showed small, gram-negative bacilli. The culture showed no growth on blood and MacConkey agar and sparse growth on chocolate agar.

Questions 1. The isolate was oxidase negative and catalase negative. Given this information, what is the suspected causative bacterium? 2. What would be the expected results if urease and nitrate reductase tests were performed? 3. What serology tests could be performed to identify this bacterium?

  

CASE STUDY 75.2 A 45-year-old female presented to the clinic with a complaint of a painful skin lesion on her back. This developed spontaneously 24 hours earlier. One month prior, she had a similar lesion on her thigh that was treated successfully with a sulfa drug but was not cultured. Evaluation revealed no fever, and she appeared to be in mild distress. Laboratory results revealed the following: CBC: WBC 12K, HGB 13K, HCT 39, PLTS 325K. The abscess was drained. A Gram stain of the thick, cream-colored purulent drainage revealed gram-positive cocci in clusters. The lesion was thoroughly irrigated, and the patient

was prescribed oral sulfa antimicrobial therapy. The lesion resolved, and the woman returned 2 months later with another lesion.

Questions 1. Given the patient’s clinical presentation and previous complaint of a lesion on her thigh, what would be the suspected infecting organism? 2. Because of the recurring nature of her infection, how would the physician determine whether the patient was colonized with the organism?   

CASE STUDY 75.3 A 50-year-old male presented with extreme pain in his left thigh. He had an elevated measurement of creatine phosphokinase (CPK), an enzyme found predominantly in the heart, brain, and skeletal muscle. When the CPK is elevated, it usually indicates injury or stress to one or more of these areas. His heart muscle CPK fraction was normal. A biopsy specimen was collected by fine-needle aspiration. Gram stain showed gram-positive bacilli with subterminal spores. The aerobic culture was sterile, but the anaerobic culture grew a pure culture of bacteria with the same Gram stain morphology as seen in the direct smear. The

A

colony had irregular edges like a medusa head; a film of growth swarmed over the entire plate in 24 hours (Fig. 75.9).

Questions 1. The isolate was indole-negative. Given this information, what is the genus and species of these bacteria? 2. What does the positive test for CPK indicate in this patient? 3. Infections with Clostridium septicum are an indication of what underlying diseases?

B • Fig. 75.9  Demonstration of the swarming film of growth of Clostridium septicum at 24 hours (A) and Clostridium sporogenes (B).

  

CHAPTER 75  Skin, Soft Tissue, and Wound Infections

1074.e3

Chapter Review 1. Primary functions of the skin include all the following except: a. Assists in maintaining body temperature b. Hydrates the body by absorbing fluid c. Provides a protective outer barrier d. Acts as a sensory organ e. Acts as a site for chemical synthesis 2. Which chemical properties of the skin assist in the prevention of infection? a. pH, NaCl, oil b. NaCl, pH, sebum, bacteria c. pH, bacteria d. Sebum, NaCl 3. What factor enhances the development of an infection after a burn? a. Minimal loss of skin b. Dehydration and loss of tissue c. Increased vascularity d. Lack of inflammation 4. A patient presented to the physician with a wound to the right index finger. The patient indicated his pet rat had bit him. Upon cultivation of the wound, a gram-positive catalase-positive rod was isolated. What is the most likely organism? a. S. aureus b. P. multocida c. S. moniliformis d. C. perfringens 5. The technologist received a swab specimen from a perirectal fistula. The specimen appears contaminated with feces. What should the technologist do? a. Reject the specimen b. Culture the specimen c. Culture the specimen for both aerobic and anaerobic organisms d. Culture the specimen for both aerobic and anaerobic organisms and notify the physician of possible specimen contamination

6. Matching: Match each term with the correct description. _____ cellulitis _____ erythrasma _____ folliculitis _____ macule _____ pustule _____ scales _____ necrotizing fasciitis _____ impetigo _____ erysipelas _____ epidermis _____ papule _____ vesicle _____ ulcer _____ myositis _____ erysipeloid _____ carbuncle _____ dermis _____ nodule _____ bulla _____ dermatophytoses _____ subcutaneous _____ furuncle

a. limited to superficial tissue b. B. anthracis c. blisterlike d. solid elevated lesion e. dense connective tissue f. abscess g. indurated lesions h. purple, irregular, raised i. group A streptococcus j. C. minutissimum k. bullous, large blisters l. hair surrounded by redness m. facial membranes n. slightly elevated solid lesion o. necrotizing gas gangrene p. muscle fascia q. inflammation of the muscle r. platelike s. circumscribed purulent lesion t. flat discoloration u. outer squamous epithelium v. subcutaneous

7. Short Answer (1)  What organisms are commonly associated with folliculitis? (2) In what ways are infections in the subcutaneous tissues usually manifested? (3) What are the various ways that wound infections can occur? (4) Why are patients with diabetes mellitus prone to foot infections?

76

Normally Sterile Body Fluids, Bone and Bone Marrow, and Solid Tissues OBJECTIVES 1. Describe the five main cavities of the human body; also name the membranes associated with these cavities and state the function of these membranes. 2. Define each of the following body cavity fluids and explain the diagnostic culture methods for each: pleural fluid, pericardial fluid, peritoneal fluid, joint fluid, and dialysis fluid. 3. Define parietal and visceral pleura. 4. Define cellulitis; name the etiologic agents of this illness and explain the associated risk factors for the development of disease. 5. Define pleural effusion; explain the difference between exudative pleural effusion and transudative pleural effusion. 6. Explain when a pleural effusion becomes an empyema and what medical condition contributes to the development of an empyema? 7. Define pericarditis and myocarditis; explain the physical conditions that may contribute to the accumulation of pericardial fluid. 8. Define peritonitis; differentiate between primary and secondary peritonitis. 9. Name the etiologic agents most commonly isolated from primary peritonitis cases in children, adults, sexually active females, and immunocompromised patients. 10. Define osteomyelitis; explain how this infection is transmitted, the diagnostic method, and the organisms most commonly responsible for this type of infection. 11. Explain the process for culturing organisms from the following specimens: bone, tissue, and bone marrow. 12. Correlate patient signs and symptoms with laboratory results to identify the etiologic agent associated with the body fluid, bone and bone marrow, and other solid tissue infection.

T

he human body is divided into five main body cavities: cranial, spinal, thoracic, abdominal, and pelvic. Each cavity is lined with membranes, and within the body wall and these membranes, or between the membranes and organs, are small spaces filled with minute amounts of

fluid. The purpose of this fluid is to bathe the organs and membranes, reducing friction between organs. Bacteria, fungi, viruses, or parasites can invade any body tissue or sterile body fluid site. Although from different areas of the body, all specimens discussed in this chapter are considered normally sterile. Therefore, even one colony of a potentially pathogenic microorganism may be significant. (Refer to Table 5.1 for a quick guide regarding collection, transport, and processing of specimens from sterile body sites.)

Specimens From Sterile Body Sites Fluids In response to infection, fluid may accumulate in any body cavity. Infected solid tissue often presents as cellulitis or with abscess formation. Areas of the body from which fluids are typically sent for microbiologic studies (in addition to blood and cerebrospinal fluid [Chapters 67 and 70]) are listed in Table 76.1. In general, peritoneal, pleural, and pericardial fluids may be cultured in aerobic and anaerobic blood culture bottles. This method should not be used if a polymicrobial infection is suspected, as some organisms can overgrow fastidious or other slow-growing organisms resulting in incomplete identification of the pathogens.

Pleural Fluid Lining the entire thoracic cavity (Chapter 68) of the body is a serous membrane called the parietal pleura. Covering the outer surface of the lung is another membrane called the visceral pleura (Fig. 76.1). Within the pleural space between the lung and chest wall is a small amount of fluid called pleural fluid that lubricates the surfaces of the pleura (the membranes surrounding the lungs and lining of the chest cavity). Normally, equilibrium exists among the pleural membranes, but in certain disease states, such as cardiac, hepatic, or renal disease, excess amounts of this fluid can be produced and accumulates in the pleural space; this is known as a pleural effusion. Pleural effusions can either 1075

1076 PA RT V I I     Diagnosis by Organ System

be exudative or transudative. Exudative pleural effusions are caused by inflammation, infection, and cancer, whereas transudative effusions result from systemic changes, such as congestive heart failure. Normal pleural fluid contains few or no cells and has a consistency similar to serum, but with a lower protein count. Pleural fluid containing numerous white blood cells is indicative of infection. Pleural fluid specimens are collected by thoracentesis, a procedure in which a needle is inserted through the chest wall into the pleural space and the excess fluid aspirated. This fluid is then submitted to the laboratory as thoracentesis fluid, pleural fluid, or empyema fluid. The fluid, or effusion, can then be analyzed for cell count, total protein, glucose, lactate dehydrogenase, amylase, cytology, nucleic acid testing, and culture. The total protein and glucose results determine whether the effusion is transudate or exudate. The patient’s serum or plasma glucose level is needed to compare with the results indicated in the body fluid. Several characteristics can be used to determine whether a fluid is a transudate or exudate (Table 76.2). TABLE   Microbiology Laboratory Body Fluid 76.1  Collection Sites

Body Area

Fluid Name(s)

Thorax

Thoracentesis or pleural or empyema fluid

Abdominal cavity

Paracentesis or ascitic or peritoneal fluid

Joint

Synovial fluid

Pericardium

Pericardial fluid

When effusions are extremely purulent (i.e., full of pus), the effusion is referred to as an empyema. Empyema often arises as a complication of pneumonia, but other infections near the lung (e.g., subdiaphragmatic infection) may seed TABLE 76.2    Pleural Fluid Effusion Characteristics

Transudate

Exudate

Appearance

Clear

Cloudy

Specific gravity

1.015

Total protein

3.0 mg/dL

LD fluid/serum ratio

0.6

Cholesterol

60 mg/dL

Cholesterol fluid/ serum ratio

0.3

Bilirubin fluid/ serum ratio

0.6

Total protein fluid/ serum ratio

0.6

White blood cells

100/mL is usually indicative of infection), the number of organisms is usually too low for detection on Gram stain of the peritoneal fluid sediment unless a concentrating technique is used; fungi are more readily detected. Many recent studies show that improved sensitivity can be achieved by using automated blood culture systems in which 10 mL of fluid is inoculated into culture bottles. Rapid detection of pathogens has also been successful using polymerase chain reaction (PCR) and 16s rRNA sequencing. The sensitivity associated with 16s rRNA sequencing varies significantly due to the inability to distinguish organisms due to high genetic similarity and should be used in conjunction with culture. Most infections originate from the patient’s normal skin microbiota; Staphylococcus epidermidis and S. aureus are the most common etiologic agents, followed by streptococci, aerobic or facultative gram-negative bacilli, Candida spp., Corynebacterium spp., and others. The oxygen content of peritoneal dialysate is usually too high for the development of anaerobic infection. Among the gram-negative bacilli isolated, Pseudomonas spp., Acinetobacter spp., and the Enterobacterales are commonly observed. 

Pericardial Fluid The heart and contiguous major blood vessels are surrounded by the pericardium, a protective tissue. The area between the epicardium—which is the membrane surrounding the heart muscle—and the pericardium is called the pericardial space, and normally contains 15 to 20 mL of clear fluid. If an infectious agent is present within the fluid, the pericardium may become distended and tight, and eventually tamponade (interference with cardiac function and circulation) can ensue. Up to 500 mL of fluid can accumulate during infection, which may seriously complicate cardiac function. Agents of pericarditis (inflammation of the pericardium) are usually viruses, especially coxsackie virus. Bacterial pericarditis usually occurs during a severe systemic infection. Parasites, certain fungi, and noninfectious causes are also associated with this disease. Myocarditis (inflammation of the heart muscle itself ) may accompany or follow pericarditis. The pathogenesis of disease involves the host inflammatory response contributing to fluid buildup, as well as cell and tissue damage. Common causes of myocarditis include viral infections with coxsackie virus, echoviruses, or adenovirus. The most common etiologic agents of pericarditis and myocarditis are listed in Box 76.1. Other bacteria, fungi, and parasitic agents have been recovered from pericardial effusions. Patients who develop pericarditis resulting from agents other than viruses are often immunocompromised or suffering from a chronic disease. An example is infective endocarditis, in which a myocardial abscess develops and then ruptures into the pericardial space. 

CHAPTER 76  Normally Sterile Body Fluids, Bone and Bone Marrow, and Solid Tissues

• BOX 76.1 Common Etiologic Agents of

• BOX 76.2 Most Commonly Encountered Etiologic

Viruses

Bacterial

Enteroviruses (primary coxsackie A and B and, less commonly, echoviruses) Adenoviruses Influenza viruses 

Staphylococcus aureus Beta-hemolytic streptococci Streptococci (other) Haemophilus influenzae Haemophilus spp. (other) Bacteroides spp. Fusobacterium spp. Neisseria gonorrhoeae Pseudomonas spp. Salmonella spp. Pasteurella multocida Moraxella osloensis Kingella kingae Moraxella catarrhalis Capnocytophaga spp. Corynebacterium spp. Clostridium spp.

Pericarditis and Myocarditis

Bacteria (Relatively Uncommon) Mycoplasma pneumoniae Chlamydia trachomatis Mycobacterium tuberculosis Staphylococcus aureus Streptococcus pneumoniae Enterobacterales and other gram-negative bacilli 

Fungi (Relatively Uncommon) Coccidioides immitis Aspergillus spp. Candida spp. Cryptococcus spp. Histoplasma capsulatum 

1079

Agents of Infectious Arthritis

Finegoldia spp. Eikenella corrodens Actinomyces spp. Mycobacterium spp. Mycoplasma spp. Ureaplasma urealyticum Borrelia burgdorferi 

Fungal Candida spp. Cryptococcus spp. Coccidioides immitis Sporothrix schenckii 

Viral Hepatitis viruses Rubella Other viruses (rarely)

Parasites (Relatively Uncommon) Entamoeba histolytica Toxoplasma gondii

Joint Fluid Arthritis is an inflammation in a joint space. Infectious arthritis may involve any joint in the body. Infection of the joint usually occurs secondary to hematogenous spread of bacteria or, less often, fungi as a direct extension of infection of the bone. It may also occur after injection of material, especially corticosteroids, into joints or after insertion of prosthetic material (e.g., total hip replacement). Although infectious arthritis usually occurs at a single site (monoarticular), a preexisting bacteremia or fungemia may seed more than one joint to establish polyarticular infection, particularly when multiple joints are diseased, such as in rheumatoid arthritis. In bacterial arthritis, the knees and hips are the most commonly affected joints in all age groups. In addition to active infections associated with viable microorganisms within the joint, sterile, self-limited arthritis caused by antigen-antibody interactions may follow an episode of infection, such as meningococcal meningitis. When an etiologic agent cannot be isolated from an inflamed joint fluid specimen, either the absence of viable agents or inadequate transport or culturing procedures may be the cause. For example, even under the best circumstances, Borrelia burgdorferi is isolated from the joints of fewer than 20% of patients with Lyme disease. Nonspecific test results, such as increased white blood cell count, decreased glucose, or elevated protein, may indicate that an infectious agent is present but is inconclusive.

Overall, S. aureus is the most common etiologic agent of septic arthritis, accounting for approximately 70% of infections. In adults younger than 30 years of age, however, N. gonorrhoeae is commonly isolated. Haemophilus influenzae has been the most common agent of bacteremia in children younger than 2 years of age; consequently, it has been the most common cause of infectious arthritis in these patients, followed by S. aureus. The widespread use of H. influenzae type B vaccine has contributed to a change in this pattern. Streptococci, including groups A (Streptococcus pyogenes) and B (Streptococcus agalactiae), pneumococci, and viridans streptococci, are prominent among bacterial agents associated with infectious arthritis in patients of all ages. Among anaerobic bacteria, Bacteroides, including B. fragilis, may be recovered as well as Fusobacterium necrophorum, which usually involves more than one joint during sepsis. Among people living in certain endemic areas of the United States and Europe, infectious arthritis is a prominent feature associated with Lyme disease. Chronic monoarticular arthritis is commonly caused by mycobacteria, Nocardia asteroides, and fungi. Some of the more common etiologic agents of infectious arthritis are listed in Box 76.2. These agents act to stimulate a host inflammatory response, which is initially responsible for the pathology of the infection. Arthritis is also a symptom associated with infectious diseases caused by certain agents, such as Neisseria meningitidis, group A streptococci (rheumatic fever), and Streptobacillus moniliformis, in which the agent cannot be recovered from joint fluid. Presumably, antigen-antibody complexes formed during active infection accumulate in a joint, initiating an inflammatory response that is responsible for the ensuing damage.

1080 PA RT V I I     Diagnosis by Organ System

Infections in prosthetic joints are usually associated with somewhat different etiologic agents than those in natural joints. After insertion of the prosthesis, organisms that gained access during the surgical procedure slowly multiply until they reach a critical mass and produce a host response. This may occur long after the initial surgery; approximately half of all prosthetic joint infections occur more than 1 year after surgery. Normal skin bacteria are the most ­common etiologic agents, with S. epidermidis, other coagulasenegative staphylococci, Corynebacterium spp., and Cutibacterium spp. being the most prevalent. However, S. aureus is also a major pathogen in this infectious disease. Alternatively, organisms may reach joints during hematogenous spread from distant, infected sites. Arthritis caused by viral agents usually occurs simultaneously with the systemic illness and may be the result of direct entry to the joint or a host immune-mediated response. Viral arthritis is associated with Parvovirus B19, Alphaviruses such as Chikungunya, rubella, hepatitis viruses B and C, HIV-1, and HTLV-1. Diagnosis of joint infections requires an aspiration of joint fluid for culture and microscopic examination. Inoculating the fluid directly into blood culture bottles may prevent the fluid from clotting. Some of the fluid may be Gram-stained and inoculated onto blood, as well as chocolate and anaerobic media. The use of AFB (acid-fast bacteria) and fungal media must also be considered. 

Bone Bone Marrow Aspiration or Biopsy Diagnosis of diseases, including brucellosis, histoplasmosis, blastomycosis, tuberculosis and leishmaniasis, can sometimes be made by detection of the organisms in bone marrow. Brucella spp. can be isolated on culture media, as can fungi, but parasitic agents must be visualized in smears or sections made from bone marrow material. Bone marrow aspirates are not likely to assist in the identification of most bacterial diseases. Many transplant centers will submit bone marrow aspirates in lysis centrifugation tubes or sterile containers for bacterial culture. If sterile containers are received, the sample should be placed in a blood culture bottle and incubated in the automated instrument. Many of the etiologic agents associated with disseminated infections in patients with human immunodeficiency virus (HIV) may be visualized or isolated from bone marrow. Some of these organisms include cytomegalovirus, Cryptococcus neoformans, and Mycobacterium avium complex. 

Bone Biopsy A small piece of infected bone is occasionally sent to the microbiology laboratory to identify the etiologic agent of osteomyelitis (infection of bone). Patients develop osteomyelitis from hematogenous spread of an infectious agent, invasion of bone tissue from an adjacent site (e.g., joint

infection, dental infection), breakdown of tissue caused by trauma or surgery, or lack of adequate circulation followed by colonization of a skin ulceration with microorganisms. Once established, infections in bone may progress toward chronicity, particularly if blood supply is insufficient in the affected area. S. aureus, seeded during bacteremia, is the most common etiologic agent of osteomyelitis among patients of all age groups. The toxins and enzymes produced by this bacterium, as well as its ability to adhere to smooth surfaces by expressing high-affinity adhesions to components of the bone matrix and producing a protective glycocalyx coating, contribute to the organism’s pathogenicity. Osteomyelitis in younger patients is often associated with a single agent. Such infections are usually of hematogenous origin. Other organisms recovered from hematogenously acquired osteomyelitis include coagulase negative staphylococci, Finegoldia, Salmonella spp., Haemophilus spp., Enterobacterales, Pseudomonas spp., F. necrophorum, and various fungi. S. aureus or P. aeruginosa is often recovered from patients with drug addictions. Parasites or viruses are rarely, if ever, etiologic agents of osteomyelitis. Bone biopsies from infections that have spread to a bone from a contiguous source or that are associated with poor circulation, especially in patients with diabetes, are likely to yield multiple isolates. Gram-negative bacilli are increasingly common among hospitalized patients; a break in the skin (surgery or intravenous line) may precede establishment of gram-negative osteomyelitis. Breaks in skin from other causes, such as a bite wound or trauma, also may be the initial event leading to underlying bone infection. For example, a human bite may lead to infection with Eikenella corrodens, whereas an animal bite may result in Pasteurella multocida osteomyelitis. Poor oral hygiene may lead to osteomyelitis of the jaw with Actinomyces spp., Capnocytophaga spp., and other oral microbiota, particularly anaerobes. Pigmented Prevotella and Porphyromonas, Fusobacterium, and Finegoldia spp. are often involved. A pelvic infection in females may result in a mixed aerobic and anaerobic osteomyelitis of the pubic bone. Patients with neuropathy (pathologic changes in the peripheral nervous system) in the extremities—notably patients with diabetes, who may have poor circulation, may experience an unrecognized or notable trauma. They develop ulcers on the feet that do not heal, become infected, and may eventually progress to involve underlying bone. These infections are usually polymicrobial, involving anaerobic and aerobic bacteria. Prevotella or Porphyromonas, other gram-negative anaerobes, including the B. fragilis group, Finegoldia spp., S. aureus, and group A and other streptococci are common agents. Nucleic acid–based testing, such as PCR, is useful in determining the infectious organism associated with the patient’s condition and can be used for rapid diagnosis in conjunction with traditional culture. 

CHAPTER 76  Normally Sterile Body Fluids, Bone and Bone Marrow, and Solid Tissues

• BOX 76.3 Infectious Agents in Tissue Requiring

Special Media

Actinomyces spp. Brucella spp. Legionella spp. Bartonella henselae (cat-scratch disease bacilli) Systemic fungi Mycoplasma spp. Mycobacterium spp. Viruses

Solid Tissues Pieces of tissue are removed from patients during surgical or needle biopsy procedures or may be collected at autopsy. Any agent of infection may cause disease in tissue, and laboratory practices should be adequate to recover bacteria, fungi, and viruses and detect the presence of parasites. Fastidious organisms (e.g., Brucella spp.) and agents of chronic disease (e.g., systemic fungi and mycobacteria) may require special media and long incubation periods for isolation. Some agents requiring special supportive or selective media are listed in Box 76.3. In addition, some organisms may be visualized using histopathology, such as Treponema pallidum, Klebsiella granulomatous, or Spirillum minus. DNA sequencing and other nucleic acid–based methods are useful in the detection of organisms that cause genital ulcers or organisms that are found in complex biofilms. 

Laboratory Diagnostic Procedures Specimen Collection and Transport Requirements for the collection and transport of specimens from sterile body sites vary because of the numerous types of specimens that can be collected and submitted to the laboratory for testing. For all specimens, the recommended procedures and transport systems for molecular detect varies and manufacturer directions should be followed accordingly.

Fluids and Aspirates Most specimens (pleural, peritoneal, pericardial, and synovial fluids) are collected by aspiration with a needle and syringe. Collecting pericardial fluid is not without risk to the patient, because the sample is collected from the cavity immediately adjacent to the heart. Collection is performed by needle aspiration with electrocardiographic monitoring or as a surgical procedure. Laboratory personnel should be alerted in advance of the procedure, ensuring that the appropriate media, tissue culture media, and stain procedures are available immediately. Body fluids from sterile sites should be transported to the laboratory in a sterile tube or airtight vial. Between 1 and 5 mL of specimen is adequate for isolation of most bacteria,

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but the larger the specimen, the better, particularly for isolation of Mycobacterium tuberculosis and fungi; at least 5 mL should be submitted for recovery of these organisms. Ten milliliters of fluid are recommended for the diagnosis of peritonitis. Anaerobic transport vials are available from several sources. These vials are prepared in an oxygen-free atmosphere and are sealed with a rubber septum or short stopper through which the fluid is injected. Transportation of fluid in a syringe capped with a sterile rubber stopper is not recommended. Most clinically significant anaerobic bacteria survive adequately in aerobic transport containers (e.g., sterile, screw-capped tubes) for short periods if the specimen is purulent and of adequate volume. However, collection in anaerobic transport media is recommended, and procedures vary in different laboratories. Specimens received in anaerobic transport vials should be inoculated to routine aerobic (an enriched broth, blood, chocolate, and sometimes MacConkey agar plates) and anaerobic media as quickly as possible. Specimens for recovery of fungi or mycobacteria may be transported in sterile, screwcapped tubes. At least 5 to 10 mL of fluid are required for adequate recovery of small numbers of organisms. If gonococci or chlamydia are suspected, additional aliquots should be sent to the laboratory for smears and appropriate cultures. Percutaneous catheters are placed during many surgical procedures to prevent the accumulation of exudate and blood at the operative site. Often, the laboratory receives drainage fluids from these catheters for culture when signs and symptoms suggest infection. However, culture of such fluid is potentially misleading when the fluid becomes contaminated within the catheter or collection device, or when the fluid does not originate from a site of the infection. Direct aspiration of potentially infected fluid collections, rather than catheter drainage fluid, should be submitted for culture for the assessment of deep tissue infections in patients. With respect to pericardial, pleural, synovial, and peritoneal fluids, the inoculation of blood culture broth bottles at the bedside or in the laboratory may be beneficial. An additional specimen should be submitted to the laboratory for a Gram stain. The specimen in the blood culture bottle is processed as a blood culture, facilitating the recovery of small numbers of organisms and diluting out the effects of antibiotics. Citrate or sodium polyanetholesulfonate (SPS) may be used as an anticoagulant. Specimens collected by percutaneous needle aspiration (paracentesis) or at the time of surgery should be inoculated into aerobic and anaerobic blood culture bottles as soon as possible. Fluid from patients receiving CAPD can be submitted to the laboratory in a sterile tube, urine cup, or the original bag. The bag is entered with a sterile needle and syringe to withdraw fluid for culture. Fluid should be directly inoculated into blood culture bottles (20 mL recommended [10 mL in each of two culture bottles]). Numerous studies

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•

Fig. 76.3  Mincing a piece of tissue for culture using sterile forceps and scissors. Note: Perform this procedure in a biosafety cabinet.

indicate that in addition to blood culture bottles, an adult Isolator tube is a sensitive and specific method for culture. 

Bone Bone marrow is typically aspirated from the interstitium of the iliac crest. Usually, this material is not processed for routine bacteria as previously indicated, because blood cultures are equally useful, and false-positive cultures for skin bacteria (S. epidermidis) are common. Some laboratories report good recovery from bone marrow material injected into a pediatric Isolator tube (ISOLATOR 1.5 mL, Alere, Waltham, MA) as a collection and transport device. The lytic agents within the Isolator tube are believed to lyse cellular components, presumably freeing intracellular bacteria for enhanced recovery. Bone removed at surgery or by percutaneous biopsy is sent to the laboratory in a sterile container. 

Tissue Tissue specimens are obtained after careful preparation of the skin. It is critical that biopsy specimens be collected aseptically and submitted to the microbiology laboratory in a sterile container; a wide-mouthed, screw-capped bottle or plastic container is recommended. Anaerobic organisms survive within infected tissue long enough to be recovered from culture. A small amount of sterile, nonbacteriostatic saline may be added to keep the specimen moist. Because homogenizing with a tissue grinder can destroy some organisms by the shearing forces generated during grinding, it is often best to use sterile scissors and forceps to mince larger tissue specimens into small pieces suitable for culturing (Fig. 76.3). Legionella spp. may be inhibited by saline. A section of lung should be submitted without saline for Legionella isolation. If anaerobic organisms are of concern, a small amount of tissue can be placed into a loosely capped, wide-mouthed plastic tube and sealed into an anaerobic pouch system, which also seals in enough moisture for survival of organisms in tissue until the specimen is plated. The surgeon should take responsibility for seeing that a second specimen is submitted to anatomic pathology for histologic studies.

Formaldehyde-fixed tissue is not useful for recovery of viable microorganisms, although some organisms can be recovered after very short periods of time. Material from draining sinus tracts should include a portion of the tract’s wall obtained by deep curettage. Tissue from infective endocarditis should contain a portion of the valve and vegetation if the patient is undergoing valve replacement. In some cases, contaminated material may be submitted for microbiologic examination. Specimens, such as tonsils or autopsy tissue, may be surface cauterized with a heated spatula or blanched by immersing in boiling water for 5 to 10 seconds to reduce surface contamination. The specimen may then be dissected with sterile instruments to permit culturing of the specimen’s center, which will not be affected by the heating. Alternatively, larger tissues may be cut in half with sterile scissors or a blade, and the interior portion cultured for microbes. Because surgical specimens are obtained at great risk and expense to the patient, and because supplementary specimens cannot be obtained easily, it is important that the laboratory save a portion of the original tissue (if enough material is available) in a small amount of sterile broth in the refrigerator and at −70°C (or, if necessary, at −20°C) for at least 4 weeks in case additional studies are indicated. If the entire tissue must be ground up for culture, a small amount of the suspension should be placed into a sterile tube and refrigerated. 

Specimen Processing, Direct Examination, and Culture Fluids and Aspirates Techniques for laboratory processing of sterile body fluids are similar except for those previously discussed that are directly inoculated into blood culture bottles. Clear fluids may be concentrated by centrifugation or filtration, whereas purulent material can be inoculated directly to media. A body fluid received in the laboratory that is already clotted must be homogenized to release trapped bacteria and minced or cut to release fungal cells. Either processing of such specimens in a motorized tissue homogenizer or grinding them manually in a mortar and pestle or glass tissue grinder allows better recovery of bacteria. Hand grinding is often preferred, because motorized grinding can generate considerable heat and thereby kill microorganisms in the specimen. Grinding may lyse fungal elements; therefore, it is not recommended with specimens processed for fungi. Small amounts of whole material from a clot should be aseptically cut with a scalpel and placed directly onto media for isolation of fungi. All fluids should be processed for direct microscopic examination. In general, if one organism is seen per oil immersion field, at least 105 organisms per milliliter of specimen are present. Often only a few organisms are present in normally sterile body fluids. Organisms must be concentrated in body fluids. For microscopic examination, cytocentrifugation (Fig. 70.4) should be used to prepare

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Gram-stained smears, because organisms can be further concentrated up to 1000-fold. Body fluids should be concentrated by either filtration or high-speed centrifugation. Once the sample is concentrated, the supernatant is aseptically decanted or aspirated with a sterile pipette, leaving approximately 1 mL of liquid in which to thoroughly mix the sediment. Vigorous vortexing or drawing the sediment up and down into a pipette several times is required to adequately suspend the sediment. This procedure should be carried out in a biologic safety cabinet. The suspension is used to inoculate media. Direct potassium hydroxide (KOH) or calcofluor white preparations for fungi, and acid-fast stain for mycobacteria, can also be performed. (See Chapter 6 for detailed descriptions related to the preparation of smears for staining procedures.) Specimens for fungi should be examined by direct wet preparation or by preparing a separate smear for periodic acid-Schiff (PAS) staining in addition to Gram stain. Either 10% KOH or calcofluor white is recommended for visualization of fungal elements from a wet preparation. In addition to hyphal forms, material from the thoracic cavity may contain spherules of Coccidioides or budding yeast cells. Lysis of leukocytes before concentration of CAPD effluents can significantly enhance recovery of organisms. Filtration of CAPD fluid through a 0.45-mm pore membrane filter allows a greater volume of fluid to be processed and usually yields better results. Because the numbers of infecting organisms may be low (less than one organism per 10 mL of fluid), a large quantity of fluid must be processed. Sediment obtained from at least 50 mL of fluid has been recommended. If the specimen is filtered, the filter should be cut aseptically into three pieces: one of which is placed on chocolate agar for incubation in 5% carbon dioxide, one on MacConkey agar, and the other on a blood agar plate for anaerobic incubation. If fluids have been concentrated by centrifugation, the resulting sediment should be inoculated to an enrichment broth, blood, and chocolate agars. Because these specimens are from normally sterile sites, selective media are inadvisable, because they may inhibit the growth of some organisms. Appropriate procedures for the isolation of anaerobes, mycobacteria, fungi, Chlamydia spp., and viruses should be used when such cultures are clinically indicated. 

or material taken from a draining sinus leading to an area of osteomyelitis, may not reflect the actual etiologic agent of the underlying osteomyelitis. Cultures of bone samples obtained during wound debridement surgery appear to be more useful for directing antibiotic therapy for better clinical outcome. Diagnosis of prosthetic (artificial) joint infections is often difficult. Unfortunately, there is no universally accepted definition for the diagnosis of infection in the absence of microbiologic evidence, because clinical symptoms such as pain do not differentiate infection from mechanical joint failure. There is no standardized approach to the laboratory diagnosis of these infections, and published data are conflicting. Further complicating the diagnosis is that the most common bacteria causing prosthesis infections are common skin contaminants such as coagulase-negative staphylococci. Some studies have reported that culture is relatively insensitive, possibly because of the organisms residing in biofilms, whereas PCR assays were able to detect most pathogens associated with prosthetic joint infections. Five or six operative bone specimens should be submitted for culture and three or more should yield the isolation and identification of the same organism for a definite diagnosis of infection. However, a study using PCR and culture using multiple media types and prolonged incubation found that appropriate culture was adequate to exclude bacterial infection in hip prostheses, and PCR did not enhance diagnostic sensitivity for infection. Normal bone is difficult to break up; however, most infected bone is soft and necrotic. Grinding the specimen in a mortar and pestle may break off some pieces. Small shavings from the most necrotic-looking areas of the bone specimen may sometimes be scraped off aseptically and inoculated onto media. Pieces should be placed directly into media for recovery of fungi. Small bits of bone can be ground with sterile broth to form a suspension for bacteriologic and mycobacterial cultures. If anaerobes are to be recovered, all manipulations are best performed in an anaerobic chamber. If such an environment is unavailable, microbiologists should work quickly within a biosafety cabinet to inoculate prereduced anaerobic plates and broth with material from the bone. 

Bone

Tissue should be manipulated in a laminar flow biologic safety cabinet. Processing tissue within an anaerobic chamber is even better. The microbiologist should cut through the infected area (which is often discolored) with a sterile scalpel blade. Half of the specimen can be used for fungal cultures and the other half for bacterial cultures. Both types of microbial agents should be considered in all tissue specimens. Some samples should also be sent to surgical pathology for histologic examination. Specimens should be processed for viruses or acid-fast bacilli when requested. Material that is to be cultured for parasites should be finely minced or teased before inoculation into broth. Direct examination of stained tissue for parasites is often

Clotted bone marrow aspirates or biopsies must be homogenized or ground to release trapped microorganisms. Specimens are inoculated to the same media as for other sterile body fluids. A special medium for enhancement of growth of Brucella spp. and incubation in 10% carbon dioxide may be needed. A portion of the specimen may be inoculated directly to fungal media. Sections are also made from biopsy material (bone) for fixation, staining, and examination (usually by anatomic pathologists) for the presence of mycobacterial, fungal, or parasitic agents. With respect to obtaining specimens from patients suspected of having osteomyelitis, cultures taken from open wound sites above infected bone,

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performed in the anatomic pathology laboratory. Imprint cultures of tissues may yield bacteriologic results identical to homogenates and may help differentiate microbial infection within the tissue’s center from surface colonization (growth only at the edge). Additional media can be inoculated for incubation at lower temperatures, which may facilitate recovery of certain systemic fungi and mycobacteria. Tissue may also be inoculated to tissue culture cells for isolation of viruses. Brain, lung, spinal fluid, and blood are generally good specimens for viral isolation. Tissue may be examined by immunofluorescence for the presence of herpes simplex virus, varicella-zoster virus, cytomegalovirus, or rabies viral particles. Lung tissue should be examined by direct fluorescent antibody test for Legionella spp. The tissues of all fetuses, premature infants, and babies who have died of an infectious process should be cultured for Listeria. Specimens of the brain, spinal fluid, blood, liver, and spleen are most likely to contain the organism.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Ahmadi SH, Neela V, Hamat RA, et al.: Rapid detection and identification of pathogens in patients with continuous ambulatory peritoneal dialysis (CAPD) associated peritonitis by 16s rRNA gene sequencing, Trop Biomed 30(4):602–607, 2013. Atkins BL, Athanasou N, Deeks JJ, et al.: Prospective evaluation of criteria for microbiological diagnosis of prosthetic-joint infection at revision arthroplasty, J Clin Microbiol 36:2932–2939, 1998. Bennett J, Dolin R, Blaser M: Principles and practice of infectious diseases, ed 9, Philadelphia, PA, 2020, Elsevier. Bourbeau P, Riley J, Heiter BJ, et al.: Use of the BacT/Alert Blood culture system for culture of sterile body fluids other than blood, J Clin Microbiol 36:3273–3277, 1998.

Carroll KC, Pfaller MA, Landry ML, et al.: Manual of clinical microbiology, ed 12, Washington, DC, 2019, ASM. Chapin-Robertson K, Dahlberg SE, Edberg SC: Clinical and laboratory analyses of cytospin-prepared Gram stains for recovery and diagnosis of bacteria from sterile body fluids, J Clin Microbiol 30:377–380, 1992. Everts RJ, Heneghan JP, Adholla PO, et  al.: Validity of cultures of fluid collected through drainage catheters versus those obtained by direct aspiration, J Clin Microbiol 39:66–68, 2001. Ince A, Rupp J, Frommelt L, et al.: Is “aseptic” loosening of the prosthetic cup after total hip replacement due to nonculturable bacterial pathogens in patients with low-grade infection? Clin Infect Dis 39:1599–1603, 2004. Khatri G, Wagner DK, Sohnle PG: Effect of bone biopsy in guiding antimicrobial therapy for osteomyelitis complicating open wounds, Am J Med Sci 321:367–371, 2001. Kim SH, Jeong HS, Kim YH, et al.: Evaluation of DNA extraction methods and their clinical application for direct detection of causative bacteria in continuous ambulatory peritoneal dialysis culture fluids from patients with peritonitis by using broad-range PCR, Ann Lab Med 32(2):119–125, 2012. Levy PY, Fenollar F: The role of molecular diagnostics in implantassociated bone and joint infection, Clin Microbiol Infect 18:1168– 1175, 2012. National Kidney and Urologic Diseases: Kidney disease statistics for the United States, 2009. Runyon B, Antillon MR, Akriviadis EA, et al.: Bedside inoculation of blood culture bottles with ascitic fluid is superior to delayed inoculation in the detection of spontaneous bacterial peritonitis, J Clin Microbiol 28:2811–2812, 1990. Teitelbaum I, Burkart J: Peritoneal dialysis, Am J Kidney Dis 42:1082– 1096, 2003. Von Essen R, Holtta A: Improved method of isolating bacteria from joint fluids by the use of blood culture bottle, Ann Rheum Dis 45:454–457, 1986.

CASE STUDY 76.1 A 79-year-old female had left-knee arthroplasty to replace an arthritic joint with a prosthetic joint. The surgery was complicated and lasted more than 1 h. She received antibiotics at the time of the surgery but not postoperatively, in accordance with usual protocols for this type of surgery. She did well postoperatively and went home in 7 days. After several weeks, she complained of low-grade fevers and pain in the joint. Her physician aspirated 50 mL of fluid from the knee and inoculated 10 mL into each of both aerobic and anaerobic blood culture bottles. Some fluid was also sent to the laboratory, where numerous white blood cells were found but no organisms were seen on Gram stain. The anaerobic blood culture bottle turned positive at 48 h. A gram-positive coccus was identified. The aerobic bottle remained negative.

Questions 1. What is the likely genus of the organism in the blood culture bottle? 2. What is the likely source of the infection in this patient? 3. The physician wanted the laboratory to be sure that the organism was not isolated because of poor technique in collection of the joint fluid specimen. Because the diagnosis of a septic joint means that the patient must have more surgery and long-term therapy, the physician wanted to be certain of the diagnosis. How can the laboratory ascertain that the organism caused the infection?

  

Chapter Review 1. When performing diagnostic testing on ascitic fluid, which of the following findings is expected? a.  Increased number of inflammatory cells and an elevated protein level b. Decreased number of inflammatory cells and an elevated specific gravity c. Increased number of inflammatory cells and a low specific gravity d. Increased number of inflammatory cells and a low protein level 2. Which of the following groups of organisms are the leading cause of myocarditis? a.  E. coli, the B. fragilis group, and enterococci b. Mycobacteria spp. and Candida spp. c. Coxsackie virus, adenovirus, and echovirus d. Influenza A and parainfluenza 1 and 3 3.  Which of the following organisms are commonly responsible for peritoneal infection among young, sexually active females? a.  S. pneumoniae and group A streptococci b. N. gonorrhoeae and C. trachomatis c. Enterobacterales and other gram-negative bacilli d. Candida albicans and Candida krusei 4. What type of fluid would be collected from the thorax body cavity? a. Thoracentesis or pleural or empyema fluid b. Paracentesis or ascitic or peritoneal fluid c. Synovial fluid d. Pericardial fluid 5.  What type of fluid would be collected from the abdominal cavity? a. Thoracentesis or pleural or empyema fluid b. Paracentesis or ascitic or peritoneal fluid c. Synovial fluid d. Pericardial fluid 6. Most cases of ascites are caused by what category of disease? a. Congestive heart failure b. Kidney disease

c. Pancreatic disease d. Liver disease 7. Poor dental hygiene can lead to osteomyelitis of the jaw; what organism is most often responsible for this type of infection? a.  Actinomyces spp. b. S. aureus c.  Prevotella spp. d. Fusobacterium spp. 8.  When processing lung tissue for diagnostic culture evaluation, which of the following suspect organisms is inhibited by saline? a.  H. influenzae b. Legionella pneumophila c.  S. pneumoniae d. K. pneumoniae 9. Which of the following pairs of organisms is most commonly the cause of peritonitis for patients receiving CAPD therapy? a. Anaerobic bacilli and gram-negative bacteria b. S. aureus and S. pyogenes c.  S. epidermidis and S. aureus d. Actinomyces and Bacteroides melaninogenicus 10.  Which of the following types of organisms play a prominent role in intraabdominal infection of secondary peritonitis? a. Gram-positive cocci b. Gram-negative bacilli c. Anaerobic bacteria d. Beta-hemolytic streptococci 11. When examining fluids by direct microscopic examination, if one organism is seen per oil immersion field, how many organisms per milliliter of specimen are present? a. 55 b. 75 c. 105 d. 155 1084.e1

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12. When testing tissue from fetuses, premature infants, and babies who have died of an infectious process, which of the following organisms must be cultured for? a.  Cytomegalovirus b. Bordetella pertussis c.  Legionella pneumophila d. Listeria monocytogenes 13.  True or False _____ One colony of a potentially pathogenic microorganism may be significant when culturing fluid from normally sterile body sites. _____ In bacterial arthritis, the knee and hip are the most commonly affected joints in all age groups. _____ Secondary peritonitis is rare and results when infection spreads from the blood and lymph nodes. _____ Exudative pleural effusions are caused by systemic body changes, such as congestive heart failure. _____ Normal pleural fluid may contain as many as 300 white blood cells per milliliter. _____ Pericarditis is often caused by parasites. _____ B. burgdorferi is commonly isolated from the synovial fluid of patients with Lyme disease and joint infections. _____ To culture bone for fungal studies, small shavings or pieces of the bone should be inoculated directly into the media. _____ The total protein and lactate dehydrogenase measurements of pleural fluid are used for the differentiation of transudates from exudates. _____ Parasites or viruses are rarely etiologic agents of osteomyelitis. 14.  Matching: Match each term with the correct description. _____ paracentesis _____ monoarticular _____ tamponade _____ osteomyelitis _____ pericarditis _____ visceral pleura _____ parietal pleura _____ CAPD _____ peritoneum _____ ascites _____ synovial fluid _____ peritoneal fluid

a. excess amounts of pleural fluid b. purulent exudative pleural effusions c. serous membrane lining walls of abdominal and pelvic cavity and outer coat of organs d. CAPD e. increased amounts of fluid in the peritoneal cavity f. inflammation of the heart muscle g. percutaneous needle aspiration

_____ peritonitis _____ thoracentesis _____ effusion _____ empyema _____ myocarditis _____ arthritis _____ homeostasis

h. inflammation of the peritoneum i. involving one joint j. inflammation in a joint space k. inflammation of the bone l. inflammation of the pericardium m. membrane covering the lungs n. physiologic balance o. collection of fluid from the thoracic cavity by needle aspiration p. interference with cardiac function q. membrane lining the entire thoracic cavity r. joint fluid s. fluid of the peritoneum that acts as a lubricant between the bowel, inner abdominal, and pelvic side walls

15.  Short Answer (1)  Explain why a pediatric Isolator tube can be used as a collection and transport device for bone marrow specimens. (2)  Explain the difference between exudative and transudative pleural effusions. (3)  What is Fitz-Hugh–Curtis syndrome? (4)  What laboratory analysis is typically performed on pleural fluid? (5)  What factors are believed to contribute to the pathogenicity of S. aureus in osteomyelitis? (6)  Explain how patients develop osteomyelitis. (7)  Explain how body fluids are processed for culture in the laboratory. (8)  Why is hand grinding of tissue or clotted body fluids preferred over using a motorized tissue homogenizer? (9)  Explain why prosthetic joint infections are associated with different etiologic agents from those in natural joints.

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Quality in the Clinical Microbiology Laboratory OBJECTIVES 1. Distinguish between the terms total quality management (TQM), continuous quality improvement (CQI), performance improvement (PI), quality assurance (QA), LEAN, Six Sigma, and individualized quality control program (IQCP). 2. Describe the quality program associated with the microbiology laboratory. 3. Identify acceptable guidelines for specimen collection and transport and give examples of unacceptable specimens. 4. State the purpose of the Standard Operating Procedure Manual. 5. Explain the requirements for laboratory personnel, use of reference laboratories, and elements of patient reports. 6. Define proficiency testing (PT) and outline the necessary steps to achieve successful results. 7. Design a log to check performance for instruments and media used in the microbiology laboratory. 8. Explain the requirements for antimicrobial susceptibility tests (ASTs). 9. Compare the maintenance of reference-quality control stocks in bacteriology, mycology, mycobacteriology, virology, and parasitology. 10. Outline a QA program for the microbiology laboratory to include all phases of infectious disease diagnosis and differentiate between external and in-house QA audit programs. 11. Describe daily monitoring activities by microbiologists and supervisors that result in providing quality care to the patient population.

S

ince the publication of the report “To Err is Human” by the Institute of Medicine, the endeavor for a safer and a more efficient health care delivery system has been in full force. The issue of quality in the medical laboratory has evolved over more than four decades after the publication of the recommendations for quality control (QC) in 1965. Just as microbial taxonomy has changed over the years, the approach to quality has evolved as well. QC is now seen as only one part of the total laboratory quality program. Quality also includes total quality management (TQM), continuous quality improvement (CQI) or performance improvement (PI), individualized quality

control program (IQCP), and quality assurance (QA). TQM, CQI, and PI are umbrella terms, encompassing the entire institution’s quality program. TQM evolved as an activity to improve patient care by having the laboratory monitor its work to detect deficiencies and subsequently correct them. CQI, IQCP, and PI went a step further by seeking to improve patient care by placing the emphasis on preventing mistakes; CQI, IQCP, and PI advocate continuous training to guard against having to correct deficiencies. IQCP, however, goes one more step further, and allows a laboratory to determine an IQCP that may result in running QC less frequently, as long as the risk assessment demonstrates accuracy and supports the validity of the process. The LEAN methodology concentrates on eliminating redundant motion, recognizing waste, and identifying what creates value from the client’s perspective. The main objective for the medical laboratory is to deliver quality patient results at the lowest cost, within the shortest time frame, while maintaining client satisfaction. It involves five principles: value, value stream, flow, pull, and continuous improvement. The first principle is to define the value in the process from the client’s perspective, which is what the patient knowingly pays for the attributes of service. Next, the value stream is identified for each process providing that value, challenging the wasted steps, and eliminating them. The next part involves ensuring the service flows continuously through the remaining value-added step. Then, it is all pulled together by introducing a continuous flow of events between all steps of the process where continuous flow is possible. The last principle is continuous improvement through management working toward perfection on an ongoing basis, so the number of steps and time is constantly under scrutiny. The scope of resources and the information needed to provide the service to the client needs to be monitored also. These principles can increase quality, throughput, capacity, and efficiency while decreasing cost, inventory, space, and lead time. Ultimately it will provide better patient care within the clinical laboratory. Six Sigma is a relatively new concept compared with TQM. Six Sigma originated in 1986 from Motorola’s drive to reduce defects by minimizing variation in processes through 1085

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metrics measurement. The process focuses on CQIs for achieving near perfection by restricting the number of possible defects to fewer than 3.4 defects per million. Six Sigma projects follow a project methodology inspired by Deming’s Plan-Do-Study-Act Cycle. The methodology is composed of five phases bearing the acronym DMAIC: define, measure, analyze, improve, control. The purpose is to aid in making precise measurements, identifying exact problems, and providing measurable solutions. When implemented correctly, Six Sigma can help organizations reduce operational costs by focusing on reducing defects, minimizing turnaround time, and trimming costs. The main difference between TQM and Six Sigma is the approach. TQM tries to improve quality by ensuring conformance to internal requirements, whereas Six Sigma focuses on improving quality by reducing the number of defects and impurities. Six Sigma is also fact-based, data-driven, and results-oriented, providing quantifiable and measurable bottom-line results, linked to strategy and related to customer requirements. QC is associated with the internal activities that ensure diagnostic test accuracy. QA is associated with the external activities that ensure positive patient outcomes. Positive patient outcomes in the microbiology laboratory include: • Reduced length of stay • Reduced cost of stay • Reduced turnaround time for diagnosis of infection • Appropriate antimicrobial therapy • Customer (physician or patient) satisfaction CQI and PI, through well-thought-out programs of QC and QA, are part of the requirements for laboratory accreditation under Clinical Laboratory Improvement Amendments (CLIA, 1988). IQPC provides a mechanism for the laboratory to review the preanalytical, analytical, and postanalytical phases of testing and create an individual QC process unique to the laboratory’s testing and environment. An IQPC must include a risk assessment, QC, and quality assessment plan. An IQPC must assess the following: • Specimen • Test system • Reagents • Environment • Testing personnel CLIA does not require any specific types of tools or assessment be utilized if the laboratory demonstrates that the QC plan meets CLIA requirements. The medical laboratory director remains responsible for ensuring that the plan meets CLIA guidelines.

Quality Program Laboratory quality is defined as accuracy, reliability, and timeliness of reported test results. The laboratory results must be as accurate as possible, all aspects of the laboratory operations must be reliable, and reporting must be timely to be useful in a clinical or public health setting. Each laboratory must establish and maintain written policies and

procedures that implement and monitor quality systems for all phases of the total testing process (preanalytical, analytical, and postanalytical), as well as general laboratory systems. The medical laboratory director is primarily responsible for the QC and QA programs. However, all laboratory personnel must actively participate in both programs. Federal guidelines (CLIA, 1988) are considered minimum standards and are superseded by higher standards imposed by individual states or private certifying agencies, such as the College of American Pathologists (CAP) or The Joint Commission (TJC). Using a set of standards established by the United States military for the manufacture and production of equipment, the International Organization for Standardization (ISO) established standards for industrial manufacturing. The ISO 9000 documents provide guidance for quality in manufacturing and service industries and can be broadly applied to many other kinds of organizations. ISO 9001:2000 addresses general quality management system requirements and applies to laboratories. There are two ISO standards that are specific to laboratories: • ISO 15189:2007. Medical laboratories—requirements for quality and competence. • ISO/IEC 17025:2005. General requirements for the competence of testing and calibration laboratories. Another important international standards organization for laboratories is the Clinical and Laboratory Standards Institute, or CLSI. CLSI uses a consensus process involving many stakeholders for developing standards. CLSI has two documents that are very important in the clinical laboratory: • QMS14-A: Quality Management System: Leadership and Management Roles and Responsibilities; Approved Guideline. • QMS01-A4: Quality Management System: A Model for Laboratory Services; Approved Guideline—Fourth Edition. The basic elements of a QC program are described in the following sections. 

Specimen Collection and Transport The laboratory is responsible for providing written policies and procedures that ensure positive identification and optimum integrity of a patient’s specimen, from the time of collection or receipt of the specimen through completion of testing and reporting of results. These guidelines and instructions should be available to health care providers for use when specimens are collected. The written collection instructions should be in detail and include the following: • Test purpose and limitations • Patient selection criteria • Timing of specimen collection (e.g., before antimicrobials are administered) • Optimal specimen collection sites • Approved specimen collection methods • Specimen transport medium criteria

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• S pecimen transport time and temperature • Specimen holding instructions if it cannot be transported immediately (e.g., hold at 4°C for 24 hours) • Minimum acceptable volume requirements where applicable • Availability of test (onsite or sent to reference laboratory) • Turnaround time • Result reporting procedures The collection instructions should include information on how a requisition should be filled out electronically or by hand, and the laboratory must include a statement indicating that the requisition must be filled out entirely. In addition to standard information, such as patient name, hospital or laboratory number, and ordering physician, other critical information includes (1) whether the patient is receiving antimicrobial therapy, (2) the suspect agent or syndrome, (3) immunization history (if applicable), and (4) travel history when certain microorganisms or parasites are suspected. The laboratory should also establish criteria for unacceptable specimens. Examples of unacceptable specimens include the following: • Unlabeled, mislabeled, or incompletely labeled specimens • Quantity not sufficient for testing (QNS) • Use of an improper transport medium, such as stool for ova and parasites not submitted in preservative(s) • Use of improper swab, such as use of wooden shaft or calcium alginate tip for viruses • Inappropriate handling of specimen with respect to temperature, timing, or storage requirements • Improper collection site for test requested, such as stool for respiratory syncytial virus • Specimen leakage from transport container • Sera excessively hemolyzed, lipemic, or contaminated with bacteria The rejected specimen will be logged electronically or manually. A rejection of specimen report will be sent to the ordering clinician. The rejected specimens will not be returned. Sometimes, specimens not meeting the requirements may be accepted by the laboratory if the specimen is irretrievable or it has been acquired through an invasive procedure. If this happens, approval by the ordering physician and laboratory director must be secured with a disclaimer on the final report, indicating that the specimen was not collected properly, and the results should be interpreted with caution. 

should be available in the work areas. It is the definitive laboratory reference and is used often for questions related to individual tests. Any obsolete procedure should be dated when removed from the SOPM and retained for at least 2 years. 

Standard Operating Procedure Manual

Patient Reports

The requirement for a Standard Operating Procedure Manual (SOPM) is considered part of the QC program. The SOPM should define test performance, tolerance limits, reagent preparation, required QC, result reporting, and references. The SOPM should be written in the format of CLSI and must be reviewed and signed annually or biannually by the medical laboratory director who appears on the CLIA certificate; in addition, all changes must be approved and dated by the laboratory director. The SOPM

There should be an established system for supervisory review of all laboratory reports. This review involves checking the specimen workup to verify that the correct conclusions were reached, and no clerical errors were made in reporting results. Reports should be released only to individuals authorized by law to receive them (physicians and various midlevel practitioners). Clinicians should be notified about “panic values” immediately. Panic values are potentially lifethreatening results; for example, positive Gram stain for

Personnel It is the medical laboratory director’s responsibility to employ sufficient qualified personnel for the volume and complexity of the work performed. For example, published studies regarding staffing of virology laboratories suggest one technologist per 500 to 1000 specimens per year. Technical on-the-job training must be documented, and the employee’s competency must be assessed twice in the first year and annually thereafter. Continuing education programs should be provided, and verification of attendance should be maintained in the employee’s personnel file. CLIA has improved the regulations associated with personnel competency (CLIA subpart K:493.1235). Laboratory employee competency assessment must include the following: (1) Direct observation of test performance, to include patient preparation (if applicable), specimen handling, processing, and testing; (2) monitoring the recording and reporting of test results; (3) review of intermediate test results or work sheets, QC results, patient results, and preventative maintenance records; (4) direct observation of performance or instrument maintenance and function checks; (5) assessment of test performance through testing previously analyzed specimens and internal blind testing of samples or external patient samples; and (6) assessment of problem-solving skills. These competency assessments must be documented and completed by qualified personnel. 

Reference Laboratories Not all testing can be completed in one facility. A laboratory test that cannot be performed in-house and needs to be sent somewhere else is considered a reference laboratory test. The reference laboratory is a separate entity from the facility that collects and sends the specimen. It must be accredited or licensed. The referral laboratory’s name, address, and licensure numbers should be included on the patient’s final report. 

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cerebrospinal fluid (CSF) or a positive blood culture. Reference ranges must be included on the report where appropriate. All patient records should be maintained for at least 2 years. However, maintaining records for at least 10 years may be needed to support medical necessity in the event of a postpayment billing audit by the Centers for Medicare and Medicaid Services. 

Proficiency Testing Proficiency testing (PT) is a QA measure used to monitor the laboratory’s analytical performance compared with its peers and reference standards. It provides an external validation tool and objective evidence of the laboratory competence for patients, as well as accrediting and oversight agencies. Laboratories are required to participate in a PT program for each analyte (test) for which a program is available; the laboratory must maintain an average score of 80% to maintain licensure in any subspecialty area. If a new regulated analyte is added, the medical laboratory must contact the PT agency to add that analyte as soon as possible. Laboratories must remain enrolled in the same PT program for one year before changing to a different program. The federal government no longer maintains a PT program, but some states, such as New York—as well as several private accrediting agencies, such as CAP—send out “blind unknowns.” These unknowns are to be treated exactly as patient specimens, from accessioning into the laboratory computer or manual logbook through workup and reporting of results. The testing personnel and medical laboratory director are required to sign a statement when the PT is completed attesting to the fact that the specimen was handled exactly like a patient specimen. In this way, PT specimens establish the accuracy and reproducibility of a laboratory’s day-to-day performance. The laboratory’s procedures, reagents, equipment, and personnel are all checked in the process. Furthermore, errors on PT help point out deficiencies, and the subsequent education of the staff can lead to overall improvements in laboratory quality. When scores (evaluations) come back, critiques accompanying them should be discussed with the entire technical staff. Evidence of corrective action in the event of problems should be documented, including changes in procedures, retraining of personnel, or the purchase of alternative media and reagents. Some laboratories have a system of internal PT in addition to those received from external agencies. When external audit is not available for a testing method, laboratories are required by law to set up an internal program to revalidate the test, at least semiannually. Internal PT samples can be set up by (1) seeding a simulated specimen and labeling it as an autopsy specimen so that no one panics if a pathogen is recovered, (2) splitting a routine specimen for workup by two different technologists, or (3) sending part of a specimen to a reference laboratory to compare and confirm the laboratory’s result. 

Performance Checks Instruments Instruments and equipment logs should contain the following information: • Instrument name, serial number, and date of implementation in the laboratory. • Procedure and periodicity of function checks with at least the frequency specified by the manufacturer; function checks must be within the manufacturer’s established limits before patient testing is conducted. • Acceptable performance ranges. • Instrument function failures, including specific details of steps taken to correct the problems (corrective action). • Date and time of service requests and response. • Maintenance records as defined and with at least the minimum frequency specified by the manufacturer. In addition, with the advances in technology, some instruments have on-board controls that are intended to be the same procedure as using external QC material. If the material represents a similar matrix to the patient samples, and the material goes through the exact same testing procedure, the on-board controls may be used to satisfy the previous need for external QC. It is the medical laboratory director’s responsibility to ensure that the material used meets all regulatory requirements. Instrument maintenance records should be retained in the laboratory for the life of the instrument. Specific guidelines regarding the periodicity of testing for autoclaves, biologic safety cabinets, centrifuges, incubators, microscopes, refrigerators, freezers, water baths, heat blocks, and other microbiology laboratory equipment can be found in a few of the references listed at the end of this chapter. 

Commercially Prepared Media Exempt From Quality Control The CLSI Subcommittee on Media Quality Control collected data over several years regarding the incidence of QC failures of commonly used microbiology media. Based on its findings, the subcommittee published a list of media that did not require retesting in the user’s laboratory if purchased from a manufacturer who follows CLSI guidelines. The laboratory must inspect each shipment for cracked media or Petri dishes, hemolysis, freezing, unequal filling, excessive bubbles, clarity, and visible contamination. The manufacturer must supply written assurance that CLSI standards were followed; this verification must be maintained along with the laboratory’s QC protocol. 

User-Prepared and Nonexempt, Commercially Prepared Media QC forms for user-prepared media should contain the amount prepared, the source of each ingredient, the lot number, the sterilization method, the preparation date,

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the expiration date (usually 1 month for agar plates and 6 months for tubed media), and the name of the preparer. Both user-prepared and nonexempt, commercially prepared media should be checked for proper color, consistency, depth, smoothness, hemolysis, excessive bubbles, and contamination. A representative sample of the lot should be tested for sterility; 5% of any lot is tested when a batch of 100 or fewer units is received, and a maximum of 10 units are tested in larger batches. A batch is any one shipment of a product with the same lot number; if a separate shipment of the same lot number of a product is received, then it is considered a different batch and needs to be tested separately. Sterility is examined by incubating the medium for 48 hours under the environmental conditions and temperature routine used within the laboratory. Both user-prepared and nonexempt, commercially prepared media should also be tested with QC organisms of known physiologic and biochemical properties. Tables listing specific organisms to test for various media can be found in a few of the references listed at the end of this chapter. 

Antimicrobial Susceptibility Tests The goal of QC testing of antimicrobial susceptibility tests (ASTs) is to ensure the precision and accuracy of both the supplies and the microbiologists performing the test. The laboratory must check each lot number and shipment of antimicrobial agent(s) before, or concurrent with, initial use, using approved control organisms. Criteria regarding frequency of testing are the same regardless of the methodology, such as minimum inhibitory concentration (MIC) broth dilution or Kirby-Bauer (Chapter 11). Each new shipment of microdilution trays or Mueller-Hinton plates should be tested with CLSI-approved American Type Culture Collection (ATCC [Rockville, MD]) reference strains. Reference strains for MIC testing are selected for genetic stability and give MICs within the midrange of each antimicrobial agent tested. Reference strains for Kirby-Bauer testing have clearly defined mean diameters for the respective zone of inhibition for each antimicrobial tested. ATCC numbers of reference strains are different for various AST methods. QC MICs and zone diameters are annually updated and published by the CLSI Subcommittee on Antimicrobial Susceptibility Testing. New tables should be obtained from the CLSI regularly. Each susceptibility test system must also be tested with use (usually daily) for 20 consecutive days. If three or fewer MICs or zone of inhibition diameters per drug-reference strain combination are outside the reference range during the 20-day testing period, laboratories may switch to weekly QC testing. Thereafter, aberrant results obtained during the weekly testing must be vigorously investigated. If a source of error, such as contamination, incorrect reference strain used, or incorrect atmosphere of incubation is found, QC testing may simply be repeated. However, if no source of error is uncovered, 5 consecutive days of retesting must be

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performed. If accuracy and precision are again acceptable, weekly QC testing may resume; if the problem drug/organism combinations are still outside the reference ranges, 20 days of consecutive testing must be reinitiated before weekly testing can be reinstated. Under no circumstances should any drug/organism combination be reported for a patient isolate if QC testing has failed.

Stains and Reagents Containers of stains and reagents should be labeled as to contents, concentration, storage requirements, date prepared (or received), date placed in service (commonly called the “date opened”), expiration date, source (commercial manufacturer or user prepared), and lot number. All stains and reagents should be stored according to the manufacturer’s recommendations and tested with positive and negative controls before use. Tables listing specific organisms to test for various stains or reagents can be found in several references at the end of this chapter. Outdated materials or reagents that fail QC even after retesting with fresh organisms should be discarded immediately. Patient specimens should not be tested using the lot number in question until the problem is resolved; in the case of a repeat failure, an alternative method should be used, or the patient’s specimen should be sent to a reference laboratory. 

Antisera The lot number, date received, condition received, and expiration date must be recorded for all shipments of antisera. In addition, the antisera should be dated when opened. New lots must be tested concurrently with previous lots, and testing must include positive and negative controls. Periodicity of testing thereafter should follow the requirements of agencies that inspect an individual laboratory and may include with use, monthly, or semiannual checks. 

Kits Kits that have been approved by the US Food and Drug Administration (FDA) need to be tested as specified in the manufacturer’s package insert. Each shipment of kits must be tested even if it is the same lot number as a previously tested lot, because temperature changes during shipment may affect the performance. Components of reagent kits of different lot numbers must not be interchanged unless otherwise specified by the manufacturer. 

Maintenance of Quality Control Records All QC results should be recorded and, when applicable, must include a review of the effectiveness of corrective actions taken to resolve problems, revision of policies and procedures necessary to prevent recurrence of problems, and discussion with appropriate staff. If temperature is adjusted or a biochemical test is repeated, the new readings

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within the tolerance limits should be listed. In many laboratories, the supervisor reviews and initials all forms weekly; the director then reviews each one monthly. QC records should be maintained for at least 2 years, except those on equipment, which must be saved for the life of each instrument. 

Maintenance of Reference Quality Control Stocks Stock organisms may be obtained from the ATCC, commercial vendors, or PT programs; well-defined clinical isolates may also be used. The laboratory should have enough organisms on hand to cover the full range of testing of all necessary materials, such as media, kits, and reagents.

Bacteriology Nonfastidious, aerobic bacterial organisms can be saved up to 1 year on trypticase soy agar (TSA) slants. Long-term storage (less than 1 year) of aerobes or anaerobes can be accomplished either by lyophilization (freeze drying) or freezing at −70°C. Frozen, nonfastidious organisms should be thawed, re-isolated, and refrozen every 5 years; fastidious organisms should be thawed, re-isolated, and refrozen every 3 years. Stock isolates may be maintained by freezing them in 10% skim milk; trypticase soy broth (TSB) with 15% glycerol; 10% horse blood in sterile, screw-cap vials; or the Microbank commercially available system (Pro Lab Diagnostics, Round Rock, TX). 

Mycology Yeasts may be treated as nonfastidious bacterial organisms for maintaining stock cultures. Molds can be stored on potato dextrose agar (PDA) slants at 4°C for 6 months to 1 year. For longer-term storage, PDA slants may be overlaid with sterile mineral oil and stored at room temperature. Alternatively, sterile water can be added to an actively sporulating culture on PDA, the conidia (spores) can be teased apart to dislodge them from the agar surface, and the water can then be dispensed to sterile, screw-top vials. These vials should be capped tightly and stored at room temperature. 

Mycobacteriology Acid-fast bacilli (AFB) may be kept on Lowenstein-Jenson (LJ) agar slants at 4°C for up to 1 year. They may also be frozen at −70°C in 7H9 broth with glycerol. 

Virology Viruses may be stored indefinitely at −70°C in a solution containing a cryoprotectant, such as 10% dimethyl sulfoxide (DMSO) or fetal bovine serum. 

Parasitology Slides and photographs must be available for QC purposes. Trichrome and other permanent slides may be purchased from commercial vendors. Clinical slides may be preserved indefinitely by adding a drop of Permount and a coverslip. Clinical slides prepared in-house should be inspected periodically, because the preservation solution may deteriorate and crack over time. 

Quality Assurance Program Because QA is the method by which the overall process of infectious disease diagnosis is reviewed, any of the steps involved in the diagnosis of an infectious disease may be studied. These steps include the following phases: • Preanalytical Phase • Ordering of test by the clinician • Processing of test by the clinician • Processing of test request by the clerical staff • Collection of specimen by health care providers or patients • Transport of specimen to the laboratory • Initial processing of specimen in the laboratory, including specimen accessioning • Analytical Phase • Examination and workup of culture by the microbiologist • Interpretation of specimen results by the microbiologist • Postanalytical Phase • Formulation of a written or printed report by the microbiologist • Communication of the microbiologist’s conclusions to the clinician in written or printed format • Interpretation of report by the clinician • Institution of appropriate therapy by the clinician Analytical testing (the work completed in the microbiology laboratory) is now seen as only one part of a continuing spectrum of steps that begins when the physician orders the test and ends when he or she receives the results and treats the patient. QA audits are planned and conducted by examining the three phases of testing. The goal is to look at the proficiency with which the patient is served by the whole facility, including the laboratory. The outcome is to look at the consequences to the patient based on the work that was performed. QA audits involve an analysis of how the system works and how it can be improved. 

Types of Quality Assurance Audits One way to conduct a QA audit is for the laboratory to subscribe to an external interlaboratory QA program. Topics to be audited are selected nationally. Data are then collected for a specified period and then returned to the program provider for analysis. A summary is compiled and returned to the institution with a comparison

CHAPTER 77  Quality in the Clinical Microbiology Laboratory

with other facilities of similar size and scope of service. That way, an individual facility can compare its results with those of its peers, a process called benchmarking. QA audits in microbiology may include areas such as (1) blood culture use, (2) health care–associated infections, (3) cumulative susceptibility results, (4) antibiotic usage, (5) turnaround time for CSF Gram stains, (6) viral hepatitis test use, (7) laboratory diagnosis of tuberculosis, (8) blood culture contamination rates, (9) appropriateness of the ordering of stools for microbiology testing, and (10) sputum quality. Other laboratory-wide audits are also applicable to the microbiology laboratory, including error reporting, quality of reference laboratories, and effects of laboratory computer downtime. A facility that does not subscribe to an external program may select topics for audits through suggestions from the medical, nursing, or pharmacy staff; complaints from the medical or nursing staff; or deficiencies and/or observations noted in the laboratory. Physicians may suggest an audit to measure the transcription accuracy of their orders by nursing unit clerical personnel. Nursing administrators may suggest an audit of contaminated urine cultures to assess the compliance of the nursing staff in instructing patients about proper urine culture collection techniques. Pharmacists may notice improper antibiotic use by the clinical staff—for example, a patient was not placed on the appropriate therapy after the pathogen was reported or the patient remains on antibiotic therapy to which his or her organism is resistant after the susceptibility report has been charted. Complaints from the medical or nursing staff can involve failure of the laboratory to conduct all the tests requested on the requisition, performance of the wrong test, or an unexpected delay in turnaround time of test results. All complaints to the laboratory must be documented. Corrective action and follow-up with the laboratory, medical, and nursing staff must also be documented. Deficiencies or problems in the laboratory’s performance should also be documented. If, for example, the laboratory notices a dramatic and unseasonable rise in the number of positive respiratory syncytial virus (RSV) direct antigen tests in the summer, and the problem is traced back to a QC problem that a new employee did not recognize, a QA audit might be indicated to study the outcome of the patients, including inappropriate treatment for RSV and failure to institute treatment for the true causative agent. Alternatively, microbiologists may notice they are receiving many ova and parasite (O&P) examinations and stool cultures on patients hospitalized for more than 3 days. Because current cost containment guidelines suggest that this is inappropriate, the microbiology laboratory personnel could undertake a study to determine the percentage of positive results and the number of patients who tested positive for Clostridiodes difficile cytotoxin, which is the more likely cause of diarrhea in patients hospitalized for more than 3 days. If the audit showed that none of the stool cultures or O&P examinations tested positive and no stools were analyzed for

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C. difficile cytotoxin, then these findings would be presented to the medical staff. Some months after the medical staff inservice, the number of stool culture and O&P requests on patients hospitalized longer than 3 days would be reevaluated in the hope of a dramatic decrease in numbers of inappropriate tests. 

Conducting a Quality Assurance Audit Box 77.1 is an example of how an in-house QA audit may be conducted. 

Continuous Daily Monitoring Daily activities of microbiologists and supervisory personnel ensure that patients get the best quality care. These activities include (1) comparing results of morphotypes seen on direct examinations with what grows on the culture to ensure that all organisms have been recovered, (2) checking antimicrobial susceptibility reports to verify that profiles match those expected from a species, and (3) studying culture and susceptibility reports for clusters of patients with unusual infections or multidrug-resistant organisms. These and many other processes result in continual improvement to all test systems, ultimately resulting in quality patient care.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Anderson NL, Noble MA, Weissfeld AS, et al.: Quality systems in the clinical microbiology laboratory. In Sewell DL, coordinating, editors: Cumitech 3B, Washington, DC, 2005, ASM Press. Centers for Medicare and Medicaid Services. CLIA IQCP, What is an IQCP? Accessed July 6, 2016. Available at: https://www.cms.gov/Regulationsand-Guidance/Legislation/CLIA/Downloads/CLIAbrochure13. pdf. Clinical and Laboratory Standards Institute: Quality management system: leadership and management roles and responsibilities; approved guideline QMS14-A, Wayne, PA, 2015, Clinical and Laboratory Standards Institute. Clinical and Laboratory Standards Institute: Quality management system: a model for laboratory services; approved guideline QMS01-A4 — fourth edition, Wayne, PA, 2015, Clinical and Laboratory Standards Institute. Clinical and Laboratory Standards Institute: Methods for dilution antimicrobial susceptibility tests for bacteria that grow aerobically; approved standard M7-A10, Wayne, PA, 2015, Clinical and Laboratory Standards Institute. Clinical and Laboratory Standards Institute: Performance standards for antimicrobial disk susceptibility tests; approved standard M2-A12, Wayne, PA, 2015, Committee for Clinical and Laboratory Standards Institute. Clinical Laboratory Improvement Amendments (CLIA) Regulations. Subpart K. 2011;493:1235.

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• BOX 77.1 Quality Assurance Audit on STATa Turnaround Times

Background After a complaint regarding turnaround time for STAT RSV direct antigen tests one winter, the microbiology laboratory at General Hospital has decided to audit its turnaround time. The medical staff indicated that it would like to turn the test around in 2.5 h (150 min) from the time of collection to the time the physician is notified; the medical staff feels that this will ensure the maximum patient benefits. 

Study Design All RSV requests for direct antigen testing were evaluated for a 3-month period to determine whether laboratory personnel were meeting this turnaround time. 

Results REPORTS GIVEN IN 5 μm in diameter) that cannot travel more than 3 feet (pertussis) • Airborne contact—for example, inhalation of droplets (>5 μm) that can travel large distances on air currents (tuberculosis) • Vector-borne contact—for example, disease spread by vectors, such as mosquitoes (malaria) or rats (rat-bite fever); this mode of transmission is rare in hospitals within developed countries Once the reservoir is known, the infection control practitioner can implement control measures, such as reeducation regarding hand washing (in the case of spread by HCWs) or hyperchlorination of air-conditioning cooling towers such as in the case of legionellosis (caused by the bacterium Legionella pneumophila). 

Role of the Microbiology Laboratory The microbiology laboratory supplies the data on organism identification and antimicrobial susceptibility profiles that the infection control practitioner reviews daily for evidence of HAI. Thus, the laboratory personnel must be able to detect potential microbial pathogens and then accurately identify them to species level and perform appropriate susceptibility testing. The microbiology laboratory staff should also monitor multidrug-resistant organisms by tabulating data on antimicrobial susceptibilities of common isolates and studying trends indicating emerging resistance. Significant or abnormal findings or trends in susceptibility profiles of isolates should be immediately reported to the infection control practitioner. If an outbreak is suspected, the laboratory works in tandem with the infection control committee by (1) saving all isolates, (2) culturing possible reservoirs (patients, personnel, or the environment), and (3) performing typing of strains to establish relatedness between isolates of the same species. Microbiology laboratories are also obligated by law to report certain isolates or syndromes to public health authorities. It is the responsibility of the laboratory to be familiar with the reporting requirements within the service area of the laboratory. 

Characterizing Strains Involved in an Outbreak The ideal system for typing microbial strains involved in outbreaks should be standardized, reproducible, sensitive, stable, readily available, inexpensive, applicable to a wide range of microorganisms, and field tested in other epidemiologic investigations. Standardization of methods is

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important so that results from one institution can be compared with those from another, facilitating a larger investigation if deemed necessary. Although no such perfect system is currently available, several methods are used to aid in typing epidemic strains. There are two major ways to type strains using either phenotypic traits or molecular typing methods. Classic phenotypic techniques include biotyping (analyzing unique biologic or biochemical characteristics), the use of antibiograms (analyzing antimicrobial susceptibility patterns), and serotyping (serologic typing of bacterial or viral antigens, such as bacterial cell wall [O] antigens). Bacteriocin typing, which examines an organism’s susceptibility to bacterial peptides (proteins), and bacteriophage typing, which examines the ability of bacteriophages (viruses capable of infecting and lysing bacterial cells) to attack certain bacterial strains, have been useful for typing Pseudomonas aeruginosa and S. aureus, respectively; these techniques, however, are not widely available. Genotypic, or molecular, methods have largely replaced phenotypic methods as a means of confirming the relatedness of strains involved in an outbreak. Plasmid analysis and restriction endonuclease analysis of chromosomal DNA are widely used. Plasmids are extrachromosomal pieces of genetic material (nucleic acids) that self-replicate (reproduce) in tandem with the bacterial genome and are passed to newly replicated organisms during cell division. Plasmids may also be transferred from one bacterial cell to another by conjugation or transduction (Chapter 2). Plasmid analysis has often been used to explain the occurrence of unusual or multiple-antibiotic resistance patterns. It has been shown that plasmids or R factors (resistance genes carried on plasmids) can facilitate outbreaks when a specific plasmid is transmitted from one genus of bacteria to another. Plasmid profiles, patterns created when plasmids are separated based on molecular weight by agarose gel electrophoresis, can also be used to characterize the similarity of bacterial strains. Relatedness of strains is based on the number and size of plasmids, with strains from identical sources showing identical plasmid profiles. Plasmids themselves or chromosomal deoxyribonucleic acid (DNA) may also be typed by means of restriction endonuclease digestion patterns. Restriction enzymes recognize specific nucleotide sequences in DNA and produce double-stranded cleavages that break the DNA into smaller fragments. The fragments of various sizes are separated using gel electrophoresis based on molecular weight. The specific recognition sequence and cleavage site have been defined for a great many of these enzymes. Modifications of the basic restriction endonuclease technique have been developed to reduce the number of bands generated to fewer than 20 to make the gels easier to interpret. These include pulsed-field gel electrophoresis (PFGE) and hybridization of ribosomal ribonucleic acid (RNA) with short fragments of DNA. Plasmid restriction digests have been used to type S. aureus and coagulase-negative staphylococci, and PFGE is the preferred method for typing enterococci, enteric gram-negative rods, and other gram-negative rods.

Other molecular methods, such as polymerase chain reaction (PCR), are used in conjunction with these methods for strain typing. In addition to strain typing, susceptibility to some antibiotics can be predicted by PCR and sequencing of specific genetic loci within the bacterial genome. Advances in next generation sequencing (NGS), which allows a laboratory to sequence an entire microorganism genome in less than an hour, has the potential to revolutionize epidemiological studies. This will undoubtedly improve the speed associated with the identification of antibioticresistant bacterial infections or confirm results observed in culture-based assays. Molecular methods (nucleic acid–based tests) are discussed in more detail in Chapter 8. 

Preventing Health Care–Associated Infections The CDC published guidelines in the 1970s specifying isolation precautions in hospitals. Techniques for isolation precautions included (1) HCWs washing their hands between caring for different patients; (2) isolation of infected patients in private rooms or placing patients with the same clinical syndrome in semiprivate rooms if private rooms are not available; (3) wearing of masks, gowns, and gloves when caring for infected patients; (4) bagging of contaminated articles, such as bed linens, before removing them from the room; (5) processing of all isolation rooms after the patient is discharged; and (6) placement of cards on the patient’s door specifying the type of isolation and instructions for visitors and HCWs. Categories of isolation were also established that included (1) strict isolation for highly contagious diseases such as chickenpox, pneumonic plague, and Lassa fever; (2) respiratory isolation for diseases such as measles, H. influenzae, or Neisseria meningitidis; (3) enteric precautions for diseases such as amoebic dysentery, Salmonella, and Shigella; (4) contact isolation for patients infected with multidrug-resistant bacteria; (5) acid-fast bacilli (AFB) isolation for persons suspected of having a Mycobacterium tuberculosis infection; (6) drainage and secretion precautions for persons with conjunctivitis and burns; and (7) blood and body fluid precautions for individuals with acquired immune deficiency syndrome (AIDS). Over time, a system of disease-specific precautions was added to the categoryspecific ones, and hospitals were given the option of using one of the two systems. Disease-specific precautions were more cost-effective, in that only those precautions specifically necessary were used to interrupt the transmission of a single disease. In 1996, the CDC developed a new system of standard precautions synthesizing the features of universal precautions (Chapter 4) and body substance isolation. Standard precautions are used in the care of all patients and apply to blood; all body fluids, secretions, and excretions except sweat, regardless of whether they contain visible blood; nonintact skin; and mucous membranes.

CHAPTER 78  Infection Control

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• BOX 78.1 Infection Control Measures for

with one of the viral hemorrhagic fevers (Ebola, Marburg, and others). 

• H  ealth care workers (HCWs) should wash hands frequently using a plain soap except in special circumstances—for example, preoperatively or after handling dressings from patients on contact isolation. • HCWs should wear gloves when touching blood, body fluids, secretions, excretions, and contaminated items. • HCWs should wear a mask, gown, eye protection, or face shield as appropriate. • Each hospital should ensure that it has adequate procedures for routine care, cleaning, and disinfection of environmental surfaces, beds, bed rails, and bedside equipment. • Hospitals should handle, transport, and launder used linen soiled with blood, body fluids, secretions, and excretions in a manner that prevents skin and mucous membrane exposure and contamination of clothing, and that prevents the transfer of microorganisms to other patients or the environment. • HCWs should take care to prevent injuries when using needles, scalpels, and other sharp instruments or devices. • HCWs should use equipment, such as mouthpieces and resuscitation bags, instead of mouth-to-mouth resuscitation. • HCWs should refrain from handling patient care equipment if they have exudative lesions or weeping dermatitis. • Hospitals should place incontinent or nonhygienic patients in a private room. • Hospitals should ensure that reusable equipment is properly sterilized. • Hospitals should ensure that single-use items are discarded properly.

Surveillance Methods

Standard Precautions

  

Modified from Healthcare Infection Control Practices Advisory Committee (HICPAC), 2007.

In addition, transmission-based precautions are used for patients known (or suspected) to be infected with pathogens spread by airborne or droplet transmission or by contact with dry skin or fomites. Box 78.1 lists infection control measures for standard precautions. Table 78.1 lists the infectious agents or syndromes along with the respective infection control measures for each transmission-based precaution. Many infection control practitioners find these guidelines substantially less cumbersome to implement than the old category- and disease-specific measures. Health care facilities, however, may modify these guidelines to fit their individual situations as long as their number of HAIs remains low. Some of the potential agents of bioterrorism can be transmitted person to person (smallpox, pneumonic plague, and viral hemorrhagic fevers including Ebola) and some cannot (anthrax). The ones that can be easily transmitted have specific transmission-based precautions—that is, airborne precautions for smallpox, droplet precautions for patients with pneumonic plague, and contact precautions for individuals

Most routine environmental cultures in a health care facility are now considered to be of little use and should not be performed unless there are specific epidemiologic implications. The decision to perform these cultures should be determined by the infection control team. However, certain surveillance cultures are still performed as a method of limiting outbreaks. These include culturing cooling towers or hot water sources for Legionella spp.; culturing water and dialysis fluids; endotoxin testing; culturing blood bank products, especially platelets; and surveillance cultures for VRE, MRSA or oxacillin-resistant S. aureus, and VRSA using rectal and oropharyngeal swabs. Physical rehabilitation centers often culture hydrotherapy equipment (whirlpools) quarterly to verify that cleaning methods are adequate; some centers perform such routine cultures more frequently. Routine surveillance of air handlers, food utensils, food equipment surfaces, and respiratory therapy equipment is no longer recommended; neither is monitoring infant formulas prepared in-house nor items purchased as sterile. A better approach is for the infection control team to monitor patients for the development of an HAI that might be related to the use of contaminated commercial products. In the event of an outbreak or an incident related to suspected contamination, a microbiologic study would be indicated. However, most often, such infections are caused by inuse contamination, rather than contamination during the manufacturing process. Suspect lots of fluid and catheter trays should be saved, and the US Food and Drug Administration (FDA) should be notified if contamination of an unopened product is suspected. Although some institutions still require preemployment stool cultures and ova and parasite examinations on food handlers, most now recognize that this is of limited value. It is much more important for food handlers to submit specimens for these tests if they develop diarrhea. Similarly, most hospitals no longer screen personnel routinely for nasal carriage of S. aureus. Although a significant percentage of the general population, including hospital personnel, are known to carry this organism, most individuals rarely shed enough organism to pose a hazard, and there is no simple way to predict which nasal carriers will disseminate staphylococci. All steam and dry-heat sterilizers and ethylene oxide gas sterilizers should be checked at least once weekly with a liquid spore suspension test vial to verify complete sterilization. Health care facilities that perform bone marrow transplantation or treat hematologic malignancies may also conduct surveillance cultures of severely immunocompromised patients who occupy laminar flow rooms. In these instances, the isolation of specific organisms may have predictive value for subsequent systemic infection. Air sampling for fungi during construction is also indicated, especially if patients

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TABLE 78.1    Transmission-Based Precautions

Type of Precaution

Specific Etiologic Agents or Syndromes

Infection Control Measures

Airborne

Measles Varicella Tuberculosis Smallpox

Place patient in private room that has monitored negative air pressure, 6–12 air changes per hour, and appropriate discharge of air outdoors or monitored HEPA filtration of room air before air is circulated to other areas of the hospital or cohorting of patients—that is, placing patients with the same infection in the same room, if private rooms are not available. Health care workers (HCWs) should wear respiratory protection when entering rooms of patients with known or suspected tuberculosis and, if not immune, for patients with measles or varicella as well. Transport patients out of their room only after placement of a surgical mask.

Droplet

Invasive Haemophilus influenzae type b infection, including meningitis, pneumonia, epiglottitis, and sepsis Invasive Neisseria meningitidis infection, including meningitis, pneumonia, and sepsis Diphtheria (pharyngeal) Mycoplasma pneumoniae Pertussis Pneumonic plague Streptococcal pharyngitis, pneumonia, or scarlet fever in infants and young children Adenovirus, influenza virus Mumps Parvovirus B19 Rubella

Place patient in private room without special air handling or ventilation or cohort patients. HCWs should wear mask when working within 3 feet of patient. Transfer patients out of their room only after placement of a surgical mask.

Contact

Gastrointestinal, respiratory, skin, or wound infections, or colonization with multidrug-resistant bacteria Clostridiodes difficile For diapered or incontinent patients: Escherichia coli O157:H7, Shigella, hepatitis A virus, or rotavirus Respiratory syncytial virus, parainfluenza virus, and enterovirus infections in infants and young children Skin infections such as diphtheria (cutaneous), herpes simplex virus (neonatal or mucocutaneous), impetigo, major abscesses, cellulitis, or decubiti, pediculosis (lice infestation), scabies (mite infestation), staphylococci furunculosis (boils) in infants and young children, zoster (disseminated or in the immunocompromised host) Viral hemorrhagic infections (Ebola, Lassa, or Marburg)

Place patient in a private room without special air handling or ventilation or cohort patients. HCWs should wear gloves when entering the patient’s room. HCWs should wash hands with a special antimicrobial agent or a waterless antiseptic agent. HCWs should wear a mask and eye protection during activities that are likely to generate splashes of blood, body fluids, secretions, or excretions. HCWs should wear a gown during procedures likely to generate splashes. HCWs should ensure reusable equipment is properly sterilized. HCWs should ensure that single-use items are properly discarded.

Modified from Healthcare Infection Control Practices Advisory Committee (HICPAC), 2007.

are immunocompromised and are being treated near the construction site. The US Pharmacopeia published requirements for monitoring of sterile compounding in hospital pharmacies. The laminar flow hoods, biologic safety cabinets, clean rooms, and donning areas must be monitored weekly or monthly

so that intravenous or intrathecal products and drugs used in the operating room are made (compounded) under sterile conditions.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

CHAPTER 78  Infection Control

Bibliography American Recovery and Reinvestment Act of 2009, Public Law 111-5, 42 U.S.C. § 241(a). Banerjee SN, Emori TG, Culver DH, et  al.: The national nosocomial infection surveillance system: secular trends in nosocomial primary bloodstream infections in the United States, 1980-1989, Am J Med 91(Suppl 3B):86S, 1991. Centers for Disease Control and Prevention: Public health focus: surveillance, prevention and control of nosocomial infections, Morb Mortal Wkly Rep 41:783–787, 1992. Centers for Disease Control and Prevention: Healthcare-Associated Infections (HAIs): state-based HAI prevention, 2015, Available at http://www.cdc.gov/HAI/state-based/. Accessed 16 November 2018. Centers for Disease Control and Prevention: Healthcare facility HAI reporting requirements to CMS via NHSN: current or proposed requirements, 2014, Available at: http://www.cdc.gov/nhsn/PDFs/ CMS/CMS-Reporting-Requirements.pdf. Accessed 16 November 2018. Centers for Disease Control and Prevention: National Healthcare Safety Network (NHSN), 2015, Available at: http://www.cdc.gov /nhsn/. Accessed 16 November 2018. Centers for Disease Control and Prevention: National Healthcare Safety Network (NHSN). About NHSN, 2015, Available at: http:// www.cdc.gov/nhsn/about.html. Accessed 16 November 2018. Centers for Disease Control and Prevention: National and state healthcare associated infections: progress report, 2015, Available at: http:/ /www.cdc.gov/HAI/pdfs/progress-report/hai-progress-report.pdf. Accessed 16 November 2018. Coffin SE, Zaoutis TE: Healthcare-associated infections. In Long SS, Pickering LK, Prober CG, editors: Principles and practice of pediatric infectious diseases, ed 3, New York, NY, 2008, Churchill Livingstone. Craven DE, Chroneou A, Zias N, Hjalmarson KI: Ventilatorassociated tracheobronchitis: the impact of targeted antibiotic therapy on patient outcomes, Chest 135:521–528, 2009. Craven DE, Steger KA, Barber TW: Preventing nosocomial pneumonia: state of the art and perspectives for the 1990s, Am J Med 91(Suppl 3B):44S–53S, 1991. Edwards JR, Peterson KD, Andrus ML, et al.: The National Healthcare Safety Network (NHSN) report, data summary for 2006 through 2007, issued November 2008, Am J Infect Control 36:609–626, 2008. Emori TG, Gaynes RP: An overview of nosocomial infections, including the role of the microbiology laboratory, Clin Microbiol Rev 6:428–442, 1993. Garibaldi RA, Cushing D, Lerer T: Risk factors for postoperative infection, Am J Med 91(Suppl 3B):158S–163S, 1991. Garner JS, Favero MS: Guideline for hand washing and hospital environmental control, 1985, Atlanta, 1985, Centers for Disease Control, pp PB85–923404. Garner JS, Simmons BP: CDC guideline for isolation precautions in hospitals, Atlanta, 1983, Centers for Disease Control, pp PB85–923401. Gastmeier P, Geffers C, Brandt C, et al.: Effectiveness of a nationwide nosocomial infection surveillance system for reducing nosocomial infections, J Hosp Infect 64:16–22, 2006. Griffith JT, Rohde RE: Ebola: implications for the clinical laboratory, Clin Lab Sci 1–6, 2014, December, Available at: http://www.asc ls.org/images/publications/journals/Ebola_virus_manuscript.pdf. Accessed 16 November 2018.

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Guidelines for the management of adults with hospital-acquired: ventilator-associated, and healthcare-associated pneumonia, Am J Respir Crit Care Med 171:388–416, 2005. Guidelines for the Prevention of Intravascular Catheter-Related Infections. Centers for Disease Control and Prevention. Available at: www.c dc.gov/mmwr/PDF/rr/rr5110.pdf.Accessed November 16, 2018. Horan TC, Andrus M, Dudeck MA: CDC/NHSN surveillance definition of healthcare-associated infection and criteria for specific types of infections in the acute care setting, Am J Infect Control 36:309–332, 2008. Hospital Infection Control Practices Advisory Committee: Guideline for infection control in healthcare personnel, Am J Infect Control 26:289, 1998. Hospital Infection Control Practices Advisory Committee: Guideline for isolation precaution in hospitals, Infect Control Hosp Epidemiol 17:53–80, 1996. Hospital Infection Control Practices Advisory Committee: Guideline for prevention of intravascular device-related infections, Am J Infect Control 24:262–277, 1996. Hospital Infection Control Practices Advisory Committee: Guideline for prevention of nosocomial pneumonia, Atlanta, 1994, Centers for Disease Control and Prevention, pp PB95–176970. Hospital Infection Control Practices Advisory Committee: Guideline for prevention of surgical site infection, Infect Control Hosp Epidemiol 20:247–280, 1999. Hospital Infection Control Practices Advisory Committee: Recommendations for preventing the spread of vancomycin resistance, Infect Control Hosp Epidemiol 16:105–113, 1995. Hospital Infections Program, National Center for Infectious Diseases, Centers for Disease Control and Prevention: Public health focus: surveillance, prevention, and control of nosocomial infections, Morb Mortal Wkly Rep 41(42):783–787, 1992. Javis WR: Infection control and changing health-care delivery systems, Emerg Infect Dis 7:170–173, 2001. Jewett JF, Reid DE, Safon LE, et  al.: Childbed fever: a continuing entity, J Am Med Assoc 206:344–350, 1968. Klevens RM, Edwards JR, Richards CL, et al.: Estimating healthcareassociated infections in US hospitals, 2002, Public Health Rep 122:160–166, 2007. Marchetti A, Rossier R: Economic burden of healthcare-associated infections in US acute care hospitals: societal perspective, J Med Econ 16:1399–1404, 2013. McGowan Jr JE, Weinstein RA: The role of the laboratory in control of nosocomial infection. In Bennett JV, Brachman PS, editors: Hospital infections, ed 3, Boston, 1992, Little, Brown. Miller JM, Bell M: Epidemiologic and infection control microbiology. In Isenberg HD, editor: Clinical microbiology procedures handbook, ed 2, Washington, DC, 2004, ASM Press. Mitchell-Hogan A, Rohde RE, Tille P, et al.: The changing role of the healthcare environment, Clin Lab Sci 29(1):44–48, 2016. Nichols RL: Surgical wound infection, Am J Med 91(Suppl 3B):54S– 64S, 1991. Reagan J, Rohde RE, Mitchell-Hogan A, et al.: The legal landscape: HAI public reporting in the United States, Clin Lab Sci 29(1):39– 43, 2016. Rohde RE, Felkner M, Reagan J, et al.: Healthcare-associated infections (HAI): the perfect storm has arrived!, Clin Lab Sci 29(1):28– 31, 2016. Rubin R: Hospital-acquired conditions declining, J Am Med Assoc 320(4):331, 2018, https://doi.org/10.1001/jama.2018.10151. Rutala WR, Weber DJ: The Healthcare Infection Control Practices Advisory Committee (HICPAC): guideline for disinfection and

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sterilization in healthcare facilities, 2008, Available at: http://ww w.cdc.gov/hicpac/pdf/guidelines/Disinfection_Nov_2008.pdf. Accessed 16 November 2018. Siegel JD, Rhinehart E, Jackson M, Chiarello L. The Healthcare Infection Control Practices Advisory Committee. 2007 Guideline for Isolation Precautions: Preventing Transmission of Infectious Agents in Healthcare Settings. Centers for Disease Control and Prevention. Available at: www.cdc.gov/ncidod/dhqp/pdf/guidelines/Isolation2 007.pdf. Accessed November 16, 2018. Stamm WE: Catheter-associated urinary tract infections: epidemiology, pathogenesis, and prevention, Am J Med 91(Suppl 3B):65S– 71S, 1991. Tille P, Rohde RE, Felkner M, et al.: The perfect storm: emerging trends and pathogens in healthcare, Clin Lab Sci 29(1):32–38, 2016. U.S. Department of Health and Human Services Office of Disease Prevention and Health Promotion: National action plan to prevent

healthcare-associated infections: road map to elimination, 2015, Available at: http://www.health.gov/hai/prevent_hai.asp#hai_p lan. Accessed 16 November 2018. US Pharmacopeial Convention, Inc: Pharmaceutical compounding— sterile preparations. In United States pharmacopeia, vol. 27. Rockville, MD, 2004, US Pharmacopeial Convention, Inc, (Suppl 1. Wenzel RP, editor: Prevention and control of nosocomial infections, ed 3, Baltimore, 1997, Williams & Wilkins. Wenzel RP, Edmond MB: The impact of hospital-acquired bloodstream infections, Emerg Infect Dis 7:174–177, 2001. Wong ES, Hooton TM. Guideline for Prevention of CatheterAssociated Urinary Tract Infections. Centers for Disease Control and Prevention. Available at: www.cdc.gov/ncidod/dhqp/gl_catheter_ assoc.html. Accessed November 16, 2018.

Chapter Review 1.  How do HAIs differ from community-acquired infections? a. HAIs are acquired after patients are admitted to the medical facility. b. HAIs are contracted outside of a health care setting. c. HAIs are present on admission. d.  Organisms of HAI are usually more antibiotic sensitive. 2. Which of the following factors could determine the likelihood that a given patient would acquire a health care–associated infection? a. Susceptibility of the patient to the infection b. The virulence of the infecting organism c. The nature of the patient’s exposure to the infecting organism d. All the above 3. What is the most common type of health care–associated infection? a. Urinary tract infection b. Pneumonia c. Surgical site infection d. Bloodstream infection 4. The emergence of antibiotic-resistant microorganisms can be mainly attributed to: a. The size of the medical facility b. The use and overuse of antibiotics c. The patient’s blood type d. Geographic location 5. Within an infection control committee, who is primarily responsible for collecting and analyzing surveillance data, monitoring patient care practices, and participating in epidemiologic investigations? a. The microbiologist b. The infection control practitioner c. The epidemiologist d. The pharmacist 6. The spread of MRSA from patient to patient on the hands of HCWs is an example of which mode of transmission for microorganisms? a. Airborne contact b. Direct contact c. Indirect contact d. Vector-borne contact

7. In the event of an infectious outbreak, the microbiology laboratory works in tandem with the infection control committee by fulfilling all the following duties except: a. Saving all isolates b. Culturing possible reservoirs (patients, personnel, or the environment) c. Performing typing of strains to establish relatedness between isolates of the same species d. Monitoring the number of missed days by employees because of sickness 8. In characterizing strains involved in an outbreak, what are the two major ways to type strains? a. Classic phenotypic techniques b. Molecular typing methods c. Both A and B d. None of the above 9. Many isolation techniques can be implemented in a health care facility; however, which of the following techniques remains the cornerstone of the modern infection control programs? a. HCWs washing hands between patients b. Segregation of infected patients in private rooms c. Wearing of masks, gowns, and gloves when caring for infected patients d. Bagging of contaminated articles such as bed linens 10. Which of the following statements is true about surveillance cultures? a. Routine environmental cultures in a medical facility are now considered to be of great value. b. Certain surveillance cultures are still performed as a method of limiting outbreaks, such as culturing cooling towers or hot water sources for Legionella spp. c.  Routine surveillance of air handlers, food utensils, and food equipment surfaces is strongly recommended. d. Preemployment stool cultures and ova and parasite examinations on all food handlers should be a requirement.

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79

Sentinel Laboratory Response to Bioterrorism OBJECTIVES 1. Define and give examples of a bio crime. 2. Distinguish between covert and overt assaults. 3. Define and give examples of select agents. 4. Site two laws that govern the possession of select agents. 5. List the government agencies that must be notified, by registration, before a laboratory may possess a select agent. 6. State the components of a biosecurity plan. 7. Summarize the standard operating procedures required for laboratories that maintain select agents. 8. Diagram and give a brief description of the Laboratory Response Network. 9. Identify responsibility of each hierarchical level through which an agent passes in its process of being defined as an agent of bioterrorism. 10. Outline the steps microbiology laboratories must follow if a select agent is isolated from a clinical specimen. 11. Explain the requirements for operation as a sentinel laboratory. 12. Name the government agencies responsible for the investigation and management of a bioterrorism event. 13. Given microbiologic characteristics, identify the most likely suspect agent of terror.

General Considerations Bioterrorism may have started centuries ago; however, lack of proper documentation made it difficult for historians and microbiologists to differentiate natural epidemics from alleged biological attacks. The practice of clinical microbiology changed significantly after Bacillus anthracis was intentionally released into the United States postal system in October 2001. Before this release there were a few events in which microorganisms were used to intentionally harm the civilian population in the United States. The first incident, in 1984, was a large community outbreak of salmonellosis caused by the intentional contamination of restaurant salad bars in The Dalles, Oregon. In this incident, a cult leader, Baghwan Sri Rajneesh, set out to influence the outcome of a local election by incapacitating voters. Cultures of Salmonella enterica serotype Typhi were grown at a laboratory within the cult’s compound. Ultimately, 751 individuals fell ill; luckily there were no deaths. Another religious cult, Aum Shinrikyo, known for

the famous sarin gas attack in Tokyo in 1995, was developing biological weapons using Clostridium botulinum and B. anthracis, but with no proof of effectiveness. In 1996, an outbreak among laboratory workers took place and was caused by a microbiology technologist in Dallas, Texas, purposely contaminating muffins and donuts with Shigella dysenteriae type 2. Forty-five laboratory workers developed gastroenteritis; four individuals were hospitalized. The event in October 2001 stunned the country. Although there had previously been sporadic instances of suspicious letters, those events proved to be hoaxes. This outbreak resulted from the delivery of weaponized anthrax spores in mailed letters or packages; ultimately there were 11 cases of inhalational anthrax and 11 of cutaneous disease. Five individuals died. The attacks prompted institutions to implement or modify bioterrorism readiness plans. The US government also reviewed the public health response and identified areas for improvement.

Bio Crime A bioterrorism event, also known as a bio crime, is an intentional assault on a person, or group of people, using a pathogen or toxin. The assault may be overt or covert. An overt attack is announced. The letters sent to Senators Daschle and Leahy in 2001 are examples of an overt event; a note inside each envelope announced that the individual opening it had been exposed to B. anthracis spores. A covert attack is unannounced; the recipient receives no indication that a threat is present. The package sent to the journalist at American Media, Inc., is an example of a covert event; an environmental investigation of his office uncovered the anthrax spores after his death and the illness of a coworker. 

Government Laws and Regulations The bombings at the World Trade Center in 1993 and the federal building in Oklahoma City in 1995 led Congress to pass the Antiterrorism and Effective Death Penalty Act of 1996. Section 511 (d) restricts the possession and use of materials capable of producing catastrophic damage in the hands of terrorists by requiring their registration. A companion law, the Uniting and Strengthening America by Providing Appropriate Tools Required to Intercept 1103

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and Obstruct Terrorism (USA PATRIOT) Act of 2001, prohibits any person to knowingly possess any biologic agent, toxin, or delivery system of a type or in a quantity that, under the circumstances, is not reasonably justified by prophylactic, protective, bona fide research, or other peaceful purpose. Later, the Public Health Security and Bioterrorism Preparedness and Response Act of 2002 required institutions to notify the Department of Health and Human Services (DHHS) or the United States Department of Agriculture (USDA) of the possession of specific pathogens or toxins called select agents. Therefore, clinical laboratories possessing any select agents must register with the Centers for Disease Control and Prevention (CDC), a branch of the DHHS. Violation of any of these statutes carries criminal penalties. The pathogens and toxins classified as select agents are listed in Box 79.1. The list is updated as needed. Bioterrorism agents are divided into three categories: A, B, or C. Category A agents are considered those presenting the highest risk to public health and national security, because they are easily disseminated or transmitted from person to person and have high mortality rates. Category A includes pathogens such as B. anthracis and Yersinia pestis. Category B agents are moderately easy to disseminate and have moderate to low mortality rates. This category includes Brucella spp. and Clostridium perfringens toxin. Category C contains emerging pathogens that could be engineered for mass spread in the future. Additional information may be found in Appendix F of the fifth edition of the CDC and the National Institutes of Health (NIH) manual Biosafety in Microbiological and Biomedical Laboratories (BMBL). The publication contains national guidelines to promote the safety and health of workers in biological and medical laboratories.

Biosecurity Biosecurity is the latest issue of concern for microbiology laboratory directors and managers. Laboratories must conduct a risk assessment and threat analysis to write a security plan. This plan must include physical security (e.g., electronic card key access and locked freezers and refrigerators), and data system (laboratory information system) security and security policies for personnel. Most hospital clinical laboratories have decided not to store any select agents. Some commercial laboratories, on the other hand, store select agents for use as positive controls for comparison with suspect samples. These laboratories must write standard operating procedures (SOPs) for (1) the access of select agents; (2) specimen accountability; (3) the receipt of select agents into the laboratory; (4) the transfer or shipping of select agents from the laboratory; (5) the reporting of incidents, injuries, and breaches of security; and (6) an emergency response plan if security is breached or the isolate is unintentionally released during an accident. They must also register the agents with the CDC.

• BOX 79.1 List of Select Agentsa

Viruses Crimean-Congo hemorrhagic fever virus Eastern equine encephalitis virus Ebola viruses Hendra virus Herpesvirus 1 (Herpes B virus) Lassa fever virus Marburg virus Monkeypox virus Nipah virus Reconstructed 1918 influenza virus Rift Valley fever virus South American hemorrhagic fever viruses (Junin, Machupo, Sabia, Flexal, Guanarito) Tick-borne encephalitis complex viruses Variola major virus (smallpox virus) Variola minor virus (Alastrim) Venezuelan equine encephalitis virus 

Bacteria Bacillus anthracis Brucella abortus, Brucella melitensis, Brucella suis Burkholderia (Pseudomonas) mallei Burkholderia (Pseudomonas) pseudomallei Clostridium botulinum Francisella tularensis Yersinia pestis Rickettsiae Coxiella burnetii Rickettsia prowazekii Rickettsia rickettsii 

Toxins Abrin Botulinum toxins Clostridium perfringens epsilon toxin Conotoxins Diacetoxyscirpenol Ricin Saxitoxin Shiga-like ribosome inactivating proteins Shigatoxin Staphylococcal enterotoxins T-2 toxin Tetrodotoxin   

aHHS

and USDA Select Agents and Toxins, 7 CFR Part 331, 9 CFR Part 121, and 42 CFR Part 73.

Each clinical laboratory should have a bioterrorism response plan. The plan should include policies and procedures to be enacted when a suspicious isolate cannot be ruled out as a biothreat agent. If a laboratory has any questions about isolating, identifying, or submitting an organism that may be an agent of bioterrorism, laboratory personnel should call the state public health laboratory. The select agent must be either sent to a public health laboratory or destroyed within 7 days of identification. If the agent is autoclaved, its destruction must be documented using Animal and Plant Health Inspection Service (APHIS)/CDC Form 4, available at https://www.selectagents.gov/forms.html .

CHAPTER 79  Sentinel Laboratory Response to Bioterrorism

National Laboratories

Reference Laboratories

Sentinel Laboratories

• Fig. 79.1  Laboratory network for biologic terrorism.

1105

Most of these laboratories are hospital-based clinical institutions and commercial diagnostic laboratories. Reference laboratories possess the required reagents and technology to perform confirmatory testing on pathogens. According to the CDC, reference laboratories are made up of more than 150 state and local public health, military, international, veterinary, agriculture, food, and water testing laboratories. In addition to laboratories in the United States, facilities located in Australia, Canada, the United Kingdom, Mexico, and South Korea serve as reference laboratories abroad. Confirmed bioterrorism agents are sent to a national laboratory. National laboratories, such as those at the CDC, US Army Medical Research Institute for Infectious Diseases, or the Naval Medical Research Center, are responsible for the definitive characterization of agents (Fig. 79.1).

Laboratory Response Network

Role of the Sentinel Laboratory

Laboratory testing and communication between clinical and public health laboratories are critical when responding to a bioterrorism event. To address this issue, the CDC, in partnership with the Association of Public Health Laboratories and the Federal Bureau of Investigation (FBI), established the Laboratory Response Network (LRN). The LRN is a threetier system. Sentinel (formerly level A) laboratories receive patient samples, rule out pathogens, and transfer suspicious specimens to reference laboratories. According to the CDC, the LRN works with the American Society for Microbiology (ASM) and state public health laboratory directors to ensure that private and commercial laboratories are part of the LRN. There are an estimated 2500 private and commercial laboratories serving in the sentinel capacity in the United States.

The main role of sentinel microbiology laboratories is to determine whether a targeted agent is suspected in a human specimen. Detection and recognition of a possible bioterrorism event depend on the following: • A laboratory having an active microbial surveillance and monitoring program • Vigilant technologists looking for a disease that (1) does not occur naturally in a geographic region (e.g., plague in New York City); (2) is transmitted by an aerosol route of infection; and (3) is a single case of disease caused by an unusual agent (e.g., Burkholderia mallei) • Good communication with infection control practitioners, infectious disease physicians, and local or regional public health laboratories

TABLE a 79.1    Algorithm for Sentinel Laboratories for Likely Bioterrorism Agents

Agent

Sentinel Laboratory Procedures

Comments

Bacillus anthracis

Colony: large, nonhemolytic, stands up like a beaten egg (Fig. 79.2) Gram stain: large, gram-positive rods (Fig. 79.3) Catalase: positive Motility: nonmotile Optional: use of the Red Line Alert Test (Tetracore, Inc.), cleared by the Food and Drug Administration, to rule out B. anthracis (see Chapter 15 for a more complete discussion of this test)

May be mistaken for Bacillus megaterium

Brucella spp.

Colony: small, nonhemolytic Gram stain: lightly staining tiny gram-negative coccobacilli Oxidase: positive Urease: positive Motility: nonmotile

May be mistaken for Haemophilus or Francisella

Francisella tularensis

Colony: pinpoint growth after 48 h Gram stain: pleomorphic, minute, faintly staining gram-negative coccobacilli Oxidase: negative Urease: negative Beta-lactamase: positive

May be mistaken for Haemophilus or Actinobacillus

Continued

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TABLE Algorithm for Sentinel Laboratories for Likely Bioterrorism Agentsa—cont’d 79.1

Agent

Sentinel Laboratory Procedures

Comments

Yersinia pestis

Colony: pinpoint growth on blood agar after 24 h Gram stain: gram-negative rods exhibiting bipolar staining Catalase: positive Oxidase: negative Urease: negative Indole: negative

Rapid systems may misidentify as Shigella spp., H2S-negative Salmonella spp., Acinetobacter spp., and Yersinia pseudotuberculosis

Clostridium botulinum

None

Send all specimens to reference laboratory; patient must get antitoxin immediately

Smallpox and hemorrhagic fever viruses

None

Smallpox can be mistaken for herpes virus if inoculated into routine tissue culture cells

aSee

individual chapters for a more detailed discussion of each organism.

• Fig. 79.2  Colony of Bacillus anthracis.

• Fig. 79.3  Gram stain of Bacillus anthracis.

Sentinel laboratories must have a class II biologic safety cabinet, have copies of level A protocols containing the algorithms for ruling out suspicious microorganisms (Table 79.1), and participate in an applicable proficiency testing program such as the College of American Pathologist’s Laboratory Preparedness Survey. Because sentinel laboratories rule out and refer microorganisms, proper knowledge of appropriate packaging and shipping is critical (Chapter 4); all specimens must be classified as infectious. Sentinel laboratories should never accept nonhuman specimens such as those from animals or the environment. Such specimens should be submitted directly to the nearest reference laboratory. Rapid communication between LRN sentinel members and their reference public health laboratories is essential. Each sentinel laboratory must know how to contact public health officials 24 hours/day. Sentinel laboratories, however, do not make the determination that a bioterrorist event has occurred and do not notify law enforcement. The FBI has primary responsibility when a bioterrorism event occurs as outlined in Presidential Decision Directive 39. A bioterrorist event is first and foremost a criminal investigation. The Federal Emergency Management Agency (FEMA) has the lead role in consequence management. FEMA receives assistance from the Department of Defense (DOD), Department of Energy (DOE), USDA, Department of Transportation (DOT), DHHS, and Environmental Protection Agency (EPA). FEMA, for example, calls for the deployment of the National Pharmaceutical Stockpile by the CDC so victims may be appropriately treated. Early recognition is the key to saving lives, and sentinel laboratorians are on the front lines in the fight against bioterrorism. Because sentinel laboratories are charged with ruling out possible bioterrorism agents and referring suspicious isolates to reference laboratories for confirmatory testing, each sentinel laboratory’s bioterrorism response plan must include a telephone number and contact information for the reference laboratory.

CHAPTER 79  Sentinel Laboratory Response to Bioterrorism

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• BOX 79.2 Infectious Agents, Toxins, and Procedures for Which Sentinel Level Clinical Microbiology

Laboratory Guidelines Are Provideda

Anthrax (Bacillus anthracis) Brucella spp. Botulinum toxin BT readiness plan Burkholderia spp. Coxiella burnetii aComplete

Novel influenza viruses Packing and shipping Plague (Yersinia pestis) Smallpox Staphylococcal enterotoxin B Tularemia (Francisella tularensis)   

guidelines available at https://www.asm.org/index.php/science-skills-in-the-lab/sentinel-guidelines

Catalase (+)

Oxidase (+)

Motile

Oxidase ( -)

Nonmotile

Urease (+)

Oxidase (+/-)

Urease (-)

Brucella spp.

Burkholderia pseudomallei

MAC (+)

Large gram-positive rods

Small gram-negative coccobacilli Burkholderia mallei

Nonmotile

Burkholderia pseudomallei

Burkholderia mallei

MAC (NG)

Yersinia pestis

Bacillus anthracis

Motile

Francisella tularensis

• Fig. 79.4  Algorithm for the differentiation of bioterrorism agents.

A sentinel laboratory’s key responsibility is to be familiar with likely agents involved in a bio crime; it must have SOPs to accomplish this task. To standardize the process nationwide, ASM has compiled a series of guidelines entitled “Sentinel level clinical laboratory guidelines for suspected agents of bioterrorism and emerging infectious diseases.” Guidelines for individual bacteria listed in Box 79.2 may be accessed on the ASM website at https://www.asm.org/ index.php/science-skills-in-the-lab/sentinel-guidelines. Algorithms for the identification of likely bioterrorism agents are provided in Table 79.1 and Fig. 79.4.

Visit the Evolve site for a complete list of procedures, review questions, and case studies.

Bibliography Barras V, Greub G: History of biological warfare and bioterrorism, Clin Microbiol Infect 20(6):497–502, 2014. Centers for Disease Control and Prevention: Laboratory security and emergency response guidance for laboratories working with select agents, MMWR (Morb Mortal Wkly Rep) 51(RR19):1–6, 2002.

Hawley RJ, Eitzen Jr EM: Biological weapons—a primer for micro­ biologists, Annu Rev Microbiol 55:235–253, 2001. Jernigan JA, Stephens DS, Ashford DA, et  al.: Bioterrorism-related inhalational anthrax: the first 10 cases reported in the United States, Emerg Infect Dis 7:933–944, 2001. Jorgensen J, Pfaller M, Carroll K, et al.: Manual of clinical microbiology, ed 11, Washington, DC, 2015, ASM Press. Klietmann WF, Ruoff KL: Bioterrorism: implications for the clinical microbiologist, Clin Microbiol Rev 14:364–381, 2001. Kolavic SA, Kimura A, Simons SL, et al.: An outbreak of Shigella dysenteriae type 2 among laboratory workers due to intentional food contamination, J Am Med Assoc 278:396–398, 1997. Morse SA: Bioterrorism: laboratory security, Lab Med 32:303, 2001. Sewell DL: Laboratory safety practices associated with potential agents of biocrime or bioterrorism, J Clin Microbiol 41:2801–2809, 2003. Snyder JW: Role of the hospital-based microbiology laboratory in preparation for and response to a bioterrorism event, J Clin Microbiol 41:1–4, 2003. Torok TJ, Tauxe RV, Wise RP, et al.: A large community outbreak of salmonellosis caused by intentional contamination of restaurant salad bars, J Am Med Assoc 278:389–395, 1997. US Department of Health and Human Services/CDC and National Institutes of Health: In Chosewood LC, Wilson DE, editors: Biosafety in microbiological and biomedical laboratories (BMBL), ed 5, Washington, DC, 2009, US Department of Health and Human Services.

CASE STUDY 79.1 From October 4 to November 2, 2001, the first 10 confirmed cases of inhalational anthrax caused by intentional release of B. anthracis were identified in the United States. Epidemiologic investigation indicated that the outbreak, in the District of Columbia, Florida, New Jersey, and New York, resulted from intentional delivery of B. anthracis spores through mailed letters or packages. This case study describes the clinical presentation and course of these cases of bioterrorism-related inhalational anthrax. The median age of patients was 56 years (range 43–73 years), 70% were male, and except for one, all were known or believed to have processed, handled, or received letters containing B. anthracis spores. The median incubation period from the time of exposure to onset of symptoms, when known (n = 6), was 4 days (range 4–6 days). Symptoms at initial presentation included fever or chills (n = 10), sweats (n = 7), fatigue or malaise (n = 10), minimal or nonproductive cough (n = 9), dyspnea (n = 8), and nausea or vomiting (n = 9). The median white blood cell count was 9.8 × 103/mm3 (range 7.5–13.3), often with increased neutrophils and band forms. Nine patients had elevated serum transaminase levels, and

six were hypoxic. All 10 patients had abnormal chest x-rays; abnormalities included infiltrates (n = 7), pleural effusion (n = 8), and mediastinal widening (n = 7). Computed tomography of the chest was performed on eight patients, and mediastinal lymphadenopathy was present in seven. With multidrug antibiotic regimens and supportive care, survival of patients (60%) was markedly higher (