Tropical Infectious Diseases: Principles, Pathogens and Practice 3rd Edition

TROPICAL INFECTIOUS DISEASES

Commissioning Editor: Sue Hodgson Development Editor: Sharon Nash Editorial Assistant: Kirsten Lowson Project Manager: Frances Affleck Design: Stewart Larking Illustration Manager: Merlyn Harvey Illustrator: Robert Britton Marketing Manager(s) (UK/USA): Richard Jones/Helena Mutak

THIRD EDITION

TROPICAL INFECTIOUS DISEASES Principles, Pathogens and Practice RICHARD L. GUERRANT MD

DAVID H. WALKER MD

PETER F. WELLER

Thomas H. Hunter Professor of International Medicine Director, Center for Global Health, Division of Infectious Diseases and International Health University of Virginia School of Medicine Charlottesville, VA, USA

Carmage and Martha Walls Distinguished University Chair in Tropical Diseases Director, Center for Biodefense and Emerging Infectious Diseases Professor and Chair, Department of Pathology University of Texas Medical Branch Galveston, TX, USA

Professor of Medicine, Harvard Medical School Professor, Immunology and Infectious Diseases Department, Harvard School of Public Health Chief, Infectious Disease Division Vice Chair of Research, Department of Medicine, Beth Israel Deaconess Medical Center Boston, MA, USA

For additional online content visit

www.expertconsult.com Edinburgh, London, New York, Oxford, Philadelphia, St Louis, Sydney, Toronto

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SAUNDERS is an imprint of Elsevier Inc. © 2011, Elsevier Inc. All rights reserved. First edition 1999 Second edition 2006 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). Chapter 41, Plague, Paul S. Mead is in the public domain apart from any borrowed figures. Chapter 60, Enterovirus Infections, Including Poliomyelitis, Mark A. Pallansch is in the public domain apart from any borrowed figures. Chapter 62, Calicivirus Infections, Gagandeep Kang, Mary K. Estes, Robert L. Atmar – Mary K. Estes retains copyright of her text and images. Chapter 90, Entomophthoramycosis, Lobomycosis, Rhinosporidiosis, and Sporotrichosis, Duane R. Hospenthal is in the public domain apart from any borrowed figures. Chapter 99, American Trypanosomiasis (Chagas Disease), Louis V. Kirchhoff is in the public domain apart from any borrowed figures. Chapter 105, Loiasis and Mansonella Infections, Amy D. Klion, Thomas B. Nutman is in the public domain apart from any borrowed figures. Chapter 128, Infectious Diseases in Modern Military Forces, Alan J. Magill, Bonnie L. Smoak, Truman W. Sharp is in the public domain apart from any borrowed figures. Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. With respect to any drug or pharmaceutical products identified, readers are advised to check the most current information provided (i) on procedures featured or (ii) by the manufacturer of each product to be administered, to verify the recommended dose or formula, the method and duration of administration, and contraindications. It is the responsibility of practitioners, relying on their own experience and knowledge of their patients, to make diagnoses, to determine dosages and the best treatment for each individual patient, and to take all appropriate safety precautions. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors assume any liability 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. Saunders British Library Cataloguing in Publication Data Tropical infectious diseases: principles, pathogens and practice. – 3rd ed.   1.  Tropical medicine.   I.  Guerrant, Richard L.  II.  Walker, David H., 1943  III.  Weller, Peter F.   616.9′883-dc22

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FOREWORD CRISIS IN THE MAINTENANCE OF GLOBAL HEALTH This third edition of the now classic Tropical Infectious Diseases: Principles, Pathogens and Practices emerges at a time of crisis in the struggle to maintain global health. Present challenges include the serious consequences of uncontrolled growth in human population; climatic extremes that create humanitarian disasters, and international spread of new, virulent, and treatment-resistant pathogens; and failures and derelictions by politicians and international organizations in the quest for a fairer distribution of resources. Some of these problems were identified in my Forewords to the first (1999) and second (2006) editions of this marvellous book but, since then, they have intensified, unabated.

AN INCREASING ROLE FOR THE SPECIALITY OF TROPICAL MEDICINE The speciality of Tropical Medicine (with Tropical Infectious Diseases as its largest component) is an unusually broad discipline. It encompasses sciences as diverse as economics, anthropology, zoology, agriculture, epidemiology, and molecular biology, equipping it to meet many of the challenges to maintaining global health. The increasing role and relevance of this approach is argued persuasively by each succeeding chapter of this book, which has been scrupulously edited by the successful team of Dick Guerrant, David Walker and Peter Weller. The third edition is resplendent in full color, a great advantage for the memorable display of clinical images, life cycles, and tables. The standardized distribution maps are a particular delight – making the lack of them for important pathogens, such as Borrelioses (with multiple agents and incomplete data as noted in Chapter 44) and Parastrongyliasis (with a broadening global range as noted in Chapter 111), all the more frustrating! – but users should beware of unwarranted certainty that a disease does not occur outside the prescribed area (e.g., malaria in Great Exuma, Bahamas occurring in travelers to March 2008). Some second-edition maps have been corrected (e.g., in Chapter 43, pinta is restored to southern Mexico and yaws to Java). The convenience and compactness of compression into a single volume is at the expense of relegating the plentiful and welcome references to a website, a cost- and space-saving inconvenience for readers of the hard copy. It is gratifying that the proportion of international authors has been increased with each new edition and that some exciting new chapters have been added. However, Karr’s apothegm “plus ça change, plus c’est la même chose” is highly apposite. What is so reassuring about the invigoratingly new and fresh presentation of this third edition of Tropical Infectious Diseases is that it continues to reemphasize, with supporting arguments and evidence, the fundamental tenets of our specialty. The following are but three examples.

POVERTY, HUMAN DISASTERS, AND TROPICAL DISEASE (CHAPTERS 2, 7) x

The link between poverty, tropical developing countries, and (often classic tropical) infectious diseases is undeniable. It is partly explained

by underlying deficiencies in nutrition (Chapter 5), sanitation, and safe food and water supplies, but the role of climate and vulnerability to natural disasters has become increasingly obvious in recent earthquake, flood, and famine tragedies (Chapter 2). Nobel economics laureate Amartya Sen has emphasized the political devisiveness of human disasters, both within and between nations, a principle well illustrated by the January 2010 Haitian earthquake.1 The climate change debate continues and, in mid 2010, critical diversions of the jet stream in the upper atmosphere are likely to have caused both the devastating floods in Pakistan and north-eastern China and the heat wave in Russia. In the aftermath of natural disasters, infectious diseases become the leading cause of death. For example, in the Bengal famine of 1943–6, 80% of the 2–3 million excess deaths were attributable to endemic infections, principally malaria, other fevers and diarrhoeal diseases.1 In the response to such cataclysmic events, the experience, skills and multidisciplinary approach of Tropical Medicine professionals can be of decisive importance (Chapters 2, 7).

EMERGING PATTERNS OF TROPICAL DISEASES (CHAPTER 14) Since the publication of the second edition, the attention of WHO and national public health bodies has been largely preoccupied or distracted by the threat of an H1N1 influenza pandemic. The redistri­ bution of massive funding towards this contingency has had serious repercussions for control of other major diseases, especially the neglected tropical infectious diseases. The wisdom and propriety of WHO’s interpretation of the epidemiology and handling of this emergency is now the subject of intense enquiry.2,3 In the meantime, other less pub­ licized viral pandemics have swept across tropical regions. Chikungunya (Chapter 78) appeared in western Indian Ocean islands in early 2005 causing 100,000s of cases during which the A226V mutation was selected, allowing transmission by Aedes albopictus mosquitoes and increased virulence, and followed by spread to India (1.42 million reported cases), elsewhere in South Asia, Africa, and in rural Emilia-Romagna in Italy where autochthonous transmission was established for a while in 2007.4 Human encroachment on the natural environment with consequent ecologic disturbance and degradation has been associated with the emergence of lethal pathogens including filoviruses (Chapter 73), arenaviruses (Chapter 68), henipaviruses/paramyxoviruses (Chapter 55), rhabdo­ viruses (Chapter 79), and coronaviruses (Chapter 58). Growing evidence implicates distinctive mammalian vectors, many of which are bats whose importance for human health is becoming apparent.5

GLOBALIZATION OF TROPICAL PATHOGENS WITH THEIR RESISTANCE TO THERAPEUTIC AGENTS (CHAPTERS 12, 13, 15, 35, 96, 122) The geographic dimension, so integral to Tropical Medicine, is well illustrated by the global threat now posed by Gram-negative Enterobacteriaceae with resistance to carbapenem conferred by the

NDM-1 (New Delhi metallo-beta-lactamase) gene. A collaborative study between UK, India, and Pakistan has revealed that many patients in UK harbouring NDM-1 organisms had travel, hospital, or other links with India and Pakistan.6 NDM-1 importations from the Indian subcontinent have also been reported in other European countries, North America, and Australia. Multiply antibiotic-resistant Acinetobacter baumannii-calcoaceticus complex bacteria were imported in military and civilian casualties from field hospitals in Iraq to USA and UK where they pose an increasing risk to immunocompromised or severely ill patients in intensive care units.7 Another example would be the spread of extensively and totally resistant Mycobacterium tuberculosis.8 The breakthrough achieved in the treatment of multiply-resistant Plasmodium falciparum malaria (Chapter 96) is currently threatened by emergence of artemisinin resistance. Artesunate-resistant P. falciparum was first confirmed in Pailin on the Cambodia–Thailand border in 20089 but is now suspected in other border areas of countries along the Mekong River (Cambodia, China (Yunnan Province), the Lao People’s Democratic Republic, Myanmar, Thailand, and Vietnam) although its presence in South America and African countries is unconfirmed.10 Another wonder drug, praziquantel, which revolutionized the treatment of schistosomiasis (Chapter 122) may also be losing its efficacy against S. mansoni in Senegal, Egypt11 and Uganda. Much is happening in the world of tropical infectious diseases but both continuity and change in this fascinating field are brilliantly reflected in Guerrant, Walker, and Weller’s book, which I strongly recommend as a comprehensive, critical, up-to-date, and beautifully presented review. David A.Warrell University of Oxford, UK August 2010

REFERENCES 1. Sen A. Human Disasters. In Warrell DA, Cox TM, Firth JD, eds. Oxford Textbook of Medicine. 3rd ed. Oxford: Oxford University Press; 2010: 119. 2. British Medical Journal Correspondence: Laurell AC, Herrera JR. What happened in Mexico? BMJ. 2010;340:c3465. Zarocostas J. WHO swine flu review committee promises to probe links with drug industry. BMJ. 2010;341:c3648. 3. Lancet Infectious Diseases. Leading Edge. WHO failing in duty of transparency. Lancet Infectious Diseases. 2010;10:505. 4. Townson H, Nathan MB. Resurgence of chikungunya. Trans R Soc Trop Med Hyg. 2008 Apr;102(4):308. 5. Wong S, Lau S, Woo P, Yuen KY. Bats as a continuing source of emerging infections in humans. Rev Med Virol. 2007 Mar-Apr;17(2):67. 6. Kumarasamy KK, et al. Emergence of a new antibiotic resistance mechanism in India, Pakistan, and the UK: a molecular, biological, and epidemiological study. Lancet Infect Dis. 2010 Aug 10. [Epub ahead of print] 7. Scott P, et al. An outbreak of multidrug-resistant Acinetobacter baumanniicalcoaceticus complex infection in the US military health care system associated with military operations in Iraq. Clin Infect Dis. 2007 Jun 15;44(12):1577. 8. Gandhi NR et al. Multidrug-resistant and extensively drug-resistant tuberculosis: a threat to global control of tuberculosis. Lancet. 2010 May 22;375(9728):1830. 9. Dondorp AM et al. Artemisinin resistance: current status and scenarios for containment. Nat Rev Microbiol. 2010 Apr;8(4):272. 10. WHO. Prevention and treatment of artemisinin-resistant falciparum malaria: update for international travellers. Wkly Epidemiol Rec. 2010 May 21;85(21):195. 11. Doenhoff MJ, Cioli D, Utzinger J. Praziquantel: mechanisms of action, resistance and new derivatives for schistosomiasis. Curr Opin Infect Dis. 2008;21:659.

FOREWORD

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PREFACE In our rapidly changing world, both tropical infectious diseases and, contemporaneously, the third edition of Tropical Infectious Diseases: Principles, Pathogens and Practice have evolved substantially since the first and second editions of the book in 1999 and 2006. New agents and new threats from terrorism, increasing antimicrobial resistance, and climate and ecologic changes have created some “perfect storms,” as noted in a new Chapter 2 by Torres-Velez and Brown. Other entirely new chapters by new authors include those on Vaccines, SARS, separate Hepatitis A–E chapters, Crimean-Congo hemorrhagic fever, Tick-borne Encephalitis and Omsk Hemorrhagic Fever, Human Papillomavirus, and mucormycosis. In addition, the number of authors has increased from 231 to 271, with the number of international contributors nearly doubling from 54 to 102, representing 38 (including 15 new) countries around the world. As well as the new, topically pertinent chapters, all chapters from the previous edition have been thoroughly revised and updated, and this third edition also benefits from new, full color presentations of illustrative, clinical and scientific material throughout. Life cycle diagrams that follow the original format of human involvement in the top half and environmental stages in the bottom half, with clinically pertinent boxes annotating the human disease manifestations of infection during the patho­gens’ development in humans, are also presented in full color. Further life cycle diagrams have been included to illustrate additional microbial pathogens. The book has been condensed into a more handy single volume, while still retaining our philosophy and mission of pro­ viding useful, authoritative, scholarly, and contemporary knowledge.

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Consistent with our interests in providing a fully referenced text that is evidence-based and founded in current published knowledge, all chapters are fully referenced; the references are easily accessible, together with their abstracts, via on-line links. What has not changed is our steadfast commitment to comprehensive excellence that combines cutting edge molecular and pathophysiologic science and epidemiology with practical clinical and field experience and is reflected throughout by our outstanding authors. Our authors are the best in their fields, and we are privileged to benefit from their perspectives on the most relevant science, medicine and epidemiology of tropical infectious diseases. To them we owe a huge debt of gratitude. Finally, we thank our superbly capable and organized Development Editor, Sharon Nash. Likewise, we thank Mary Ann Winecoff, Rachel Stella, Doris Baker, and Sherrill Hebert, our Publishing Director, Sue Hodgson, and our Project Manager, Frances Affleck, whose labors have made this completely new, full color volume possible. We also acknowledge our spouses, Nancy, Margie, and Anne, who have again supported us as we have endeavored with our authors to produce an extensively updated third edition of Tropical Infectious Diseases: Principles, Pathogens and Practice. Richard L. Guerrant, MD David H. Walker, MD Peter F. Weller, MD

CONTRIBUTORS Saad H. Abdalla, MD Honorary Clinical Senior Lecturer, Department of Medicine, Imperial College London, London, UK Chapter 137

Gustavo Olszanski Acrani, PhD Research Fellow, Department of Cell Biology and Virology Research Center, University of São Paulo School of Medicine, Ribeirão Preto, SP, Brazil Chapter 58

Rakesh Aggarwal, MD DM Professor, Department of Gastroenterology, Sanjay Gandhi Postgraduate Institute of Medical Sciences, Lucknow, India Chapter 64

Ban Mishu Allos, MD Assistant Professor of Medicine and Preventive Medicine, Vanderbilt University School of Medicine, Nashville, TN, USA Chapter 19

Miriam J. Alter, PhD Robert E. Shope Professor in Infectious Disease Epidemiology, Director, Infectious Disease Epidemiology Program, Institute for Human Infections and Immunity, Professor, Department of Internal Medicine, University of Texas Medical Branch, Galveston, TX, USA Chapter 65

Jon K. Andrus, MD Deputy Director, Pan American Health Organization, Washington, DC, USA Chapter 54

Juana Angel, MD PhD Professor, Instituto de Genética Humana, Facultad de Medicina, Pontificia Universidad Javeriana, Bogotá, Colombia Chapter 61

Gregory M. Anstead, MD PhD Associate Professor, Department of Medicine, Division of Infectious Diseases, University of Texas Health Sciences Center, Director, Immunosuppression and Infectious Diseases Clinics, South Texas Veterans Healthcare System, San Antonio, TX, USA Chapters 85, 86, and 87

Eduardo Arathoon, MD Director Médico, Clínica Familiar Luis Ángel García, Asociación de Salud Integral, Guatemala City, Guatemala Chapter 85

Eurico Arruda, MD PhD Professor of Microbiology, Department of Cell Biology and Virology Research Center, University of São Paulo School of Medicine, Ribeirão Preto, SP, Brazil Chapter 58

Ray R. Arthur, PhD Director, Global Disease Detection Operations Center, Division of Global Disease Detection and Emergency Response, Center for Global Health, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 14

Robert L. Atmar, MD Professor, Departments of Medicine and Molecular Virology and Microbiology, Baylor College of Medicine, Houston, TX, USA Chapter 62

Patrick Banura, MBChB MPH Head of Department, Department of Community Health, Masaka Regional Referral Hospital, Masaka, Uganda Chapter 17

Alan G. Barbour, MD Professor, Microbiology & Molecular Genetics and Medicine, School of Medicine, University of California-Irvine, Director, Pacific-Southwest Regional Center of Excellence for Biodefense and Emerging Infections, Irvine, CA, USA Chapter 44

Alan D.T. Barrett, PhD John D. Stobo MD Distinguished Chair in Vaccinology, Director, Sealy Center for Vaccine Development, Professor, Department of Pathology, University of Texas Medical Branch, Galveston, TX, USA Chapter 10

Dan Bausch, MD MPH&TM Associate Professor, Department of Tropical Medicine, Tulane School of Public Health and Tropical Medicine, Associate Professor, Department of Medicine, Section of Infectious Diseases, Tulane University Medical Center, New Orleans, LA, USA Chapter 68

Steven L. Berk, MD Professor and Dean, School of Medicine, Texas Tech University Health Science Center, Lubbock, TX, USA Chapter 117

Pascal O. Bessong, MSc PhD Head, Department of Microbiology, Principal Investigator, AIDS Virus Research Laboratory, University of Venda, Thohoyandou, South Africa Chapter 81

Frank J. Bia, MD MPH Professor (Emeritus) Internal Medicine, Yale School of Medicine, Medical Director, AmeriCares, Stamford, CT, USA Chapter 136

Tihana Bicanic, MD Research Fellow and Honorary Consultant in Infections Disease, Department of Infectious Diseases, St George’s University of London, London, UK Chapter 85

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Robert E. Black, MD MPH Edgar Berman Professor and Chair, Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, MD, USA Chapter 5

CONTRIBUTORS

Thomas P. Bleck, MD FCCM Professor of Neurological Sciences, Neurosurgery, Medicine, and Anesthesiology, and Assistant Dean, Rush Medical College, Associate Chief Medical Officer (Critical Care), Rush University Medical Center, Chicago, IL, USA Chapters 42, 79, and 136

Andrea K. Boggild, MSc MD FRCPC Staff Physician, Tropical Disease Unit, Division of Infectious Diseases, Department of Medicine, University Health Network-Toronto General Hospital, Toronto, ON, Canada Chapter 130

William Bonnez, MD Associate Professor of Medicine, Infectious Diseases Division, University of Rochester School of Medicine, Rochester, NY, USA Chapter 80

Joseph S. Bresee, MD Chief, Epidemiology and Prevention Branch, Influenza Division, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 63

Corrie Brown, DVM PhD Josiah Meigs Distinguished Professor, Department of Veterinary Pathology, College of Veterinary Medicine, University of Georgia, Athens, GA, USA Chapter 2

Lillian B. Brown, MPH PhD Research Associate Department of Epidemiology, University of North Carolina – Chapel Hill School of Medicine, Gillings School of Global Public Health, Chapel Hill, NC, USA Chapter 25

Enrico Brunetti, MD Assistant Professor of Infectious Diseases, University of Pavia, Staff Physician, Division of Infectious and Tropical Diseases, IRCCS S. Matteo Hospital Foundation, Pavia, Italy Chapter 120

Fabrizio Bruschi, MD Professor of Parasitology, Department of Experimental Pathology, MBIE, Università di Pisa, Scuola Medica, Pisa, Italy Chapter 110

Amy E. Bryant, PhD Research Scientist, Infectious Disease Section, Veterans Affairs Medical Center, Boise, ID; Affiliate Assistant Professor, Department of Medicine, University of Washington, Seattle, WA, USA Chapter 30

Carlos C. (Kent) Campbell, MD MPH Director, Malaria Control Program, PATH, Seattle, WA, USA Chapter 96

Carlos Castillo-Solorzano, MD MPH Regional Advisor on Vaccines and Immunization, Pan American Health Organization, Washington, DC, USA Chapter 54

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Martin S. Cetron, MD Director, Division of Global Migration and Quarantine, NCEZID, CDC, National Center for Emerging and Zoonotic Infectious Diseases, Atlanta, GA, USA Chapter 127

Ding-Shinn Chen, MD Distinguished Chair Professor, Division of Gastroenterology and Hepatology, Department of Internal Medicine, National Taiwan University College of Medicine, Staff Physician, Hepatitis Research Center, National Taiwan University Hospital, Taipei, Taiwan Chapter 66

Pei-Jer Chen, MD PhD Professor, Division of Gastroenterology and Hepatology, Department of Internal Medicine, National Taiwan University College of Medicine, Staff Physician, Hepatitis Research Center, National Taiwan University Hospital, Taipei, Taiwan Chapter 66

Xiang-Sheng Chen, MD PhD Professor and Vice-Director, National Center for STD Control, Nanjing, China Chapter 43

Thomas Cherian, MD Coordinator, Expanded Programme on Immunization, Department of Immunization, Vaccines and Biologicals, World Health Organization, Geneva, Switzerland Chapter 27

K.B. Chua, MBBS MMed MD PhD FRCPE FRCPath Consultant Virologist, National Public Health Laboratory, Ministry of Health Malaysia, Selangor, Malaysia Chapter 55

Myron S. Cohen, MD J. Herbert Bate Distinguished Professor, Medicine, Microbiology and Immunology and Public Health, Chief, Division of Infectious Diseases, Director, Institute of Global Health and Infectious Diseases, The University of North Carolina at Chapel Hill, Chapel Hill, NC, USA Chapters 43 and 138

Graham S. Cooke Senior Lecturer Infectious Diseases, Imperial College London, UK Honorary Associate Professor, University of KwaZulu Natal, South Africa Chapter 6

Chester R. Cooper, Jr, PhD Professor of Biological Sciences, Department of Biological Sciences, Youngstown State University, Youngstown, OH, USA Chapter 84

Edward S. Cooper, MBBS FRCP Clinical Consultant, Partnership for Child Development, Department of Infectious Disease Epidemiology, Imperial College, London, UK Chapter 114

Christina M. Coyle, MD MS Professor of Clinical Medicine, Albert Einstein College of Medicine, Director of the Tropical Medicine Clinic, Bronx Municipal Hospital, Jacobi Medical Center, Bronx, NY, USA Chapter 119

John H. Cross†, PhD Professor, Tropical Public Health, Department of Preventive Medicine and Biometrics, Uniformed Services University of the Health Sciences, Bethesda, MD, USA Chapters 111 and 123

David A.B. Dance, MB ChB MSc FRCPath Regional Microbiologist (South West), Health Protection Agency, Plymouth, Devon, UK Chapter 33

†

deceased

Chapter 136

Chandler R. Dawson, MD Professor Emeritus of Ophthalmology, Francis I. Proctor Foundation, University of California, San Francisco, San Francisco, CA, USA Chapter 46

Catherine de Martel, MD PhD Fellow, International Agency for Research on Cancer, Lyon, France Chapter 11

Ciro A. De Quadros, MD Executive Vice-President, Albert B. Sabin Vaccine Institute, Washington, DC, USA Chapter 54

Anastacio de Queiroz Sousa, MD Associate Professor of Medicine, Department of Clinical Medicine, School of Medicine, Federal University of Ceara, Director, Hospital São Jose for Infectious Diseases, Fortaleza, Ceara, Brazil Chapter 100

Nilanthi R. de Silva, MBBS MSc MD Professor of Parasitology, Faculty of Medicine, University of Kelaniya, Kelaniya, Ragama, Sri Lanka Chapter 114

Alexandre Leite de Souza, MD Infectious Diseases Specialist, Department of Medical Sciences, University of São Paulo School of Medicine (FM USP), São Paulo, Brazil Chapter 24

Christoph Dehio, PhD Professor of Molecular Microbiology, Biozentrum of the University of Basel, Basel, Switzerland Chapter 39

David J. Diemert, MD FRCP(C) Assistant Professor, Department of Microbiology, Immunology and Tropical Medicine, The George Washington University School of Medicine, Director of Clinical Trials, Sabin Vaccine Institute, Washington, DC, USA Chapter 115

Rebecca Dillingham, MD MPH Center for Global Health, Assistant Professor of Medicine, Division of Infectious Disease and International Health, University of Virginia Health System, Charlottesville, VA, USA Chapter 4

John E. Donelson, PhD Professor of Biochemistry, Carver College of Medicine, University of Iowa, Iowa City, Iowa, USA Chapter 98

J. Stephen Dumler, MD Professor, Associate Director of Clinical Microbiology, Director of Parasitology Laboratory, Division of Medical Microbiology, Department of Pathology, The Johns Hopkins University School of Medicine, The Johns Hopkins Hospital; Department of Molecular Microbiology and Immunology, The Johns Hopkins University Bloomberg School of Public Health, Baltimore, MD, USA Chapter 52

Joseph A. Duncan, MD PhD Assistant Professor of Medicine and Pharmacology, University of North Carolina School of Medicine, Department of Medicine, Division of Infectious Diseases, Chapel Hill, NC, USA Chapter 25

Herbert L. DuPont, MD Chief of Internal Medicine, St. Luke’s Episcopal Hospital, Director, Center for Infectious Diseases, The University of Texas School of Public Health, Vice Chairman, Department of Medicine, Baylor College of Medicine, Houston, TX, USA

CONTRIBUTORS

Mustapha A. Danesi, MB FRCPI FMCP FWACP Professor, Department of Medicine, College of Medicine, University of Lagos, Consultant Physician and Neurologist, Clinical Department of Medicine, Lagos University Teaching Hospital, Lagos, Lagos State, Nigeria

Chapter 129

Marlene L. Durand, MD Assistant Professor of Medicine, Harvard Medical School, Director, Infectious Disease Service, Massachusetts Eye and Ear Infirmary, Physician, Massachusetts General Hospital, Boston, MA, USA Chapter 135

Mark L. Eberhard, PhD Director, Division of Parasitic Diseases, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 107

Joshua C. Eby, MD Assistant Professor, Division of Infectious Diseases and International Health, Department of Medicine, University of Virginia School of Medicine, Charlottesville, VA, USA Chapter 31

Charles Edwards, MB BS FRCPC FACP Department Head, Department of Medicine, Queen Elizabeth Hospital, St Michael, Barbados Chapter 45

Rachel B. Eidex, PhD Chief, Refugee Health Program for Africa, Division of Global Migration and Quarantine, Centers for Disease Control and Prevention, Nairobi, Kenya Chapter 127

Jerrold J. Ellner, MD Professor of Medicine, Boston University School of Medicine, Chief, Section of Infectious Diseases, Boston Medical Center, Boston, MA, USA Chapter 35

Delia A. Enría, MD MPH Director, Instituto Nacional de Enfermedades Virales Humanas, Pergamino, Argentina Chapters 68 and 71

Onder Ergonul, MD MPH Professor of Infectious Diseases, Marmara University, School of Medicine, Istanbul, Turkey Chapter 70

Mary K. Estes, MD Professor, Departments of Molecular Virology and Microbiology and Medicine, Baylor College of Medicine, Houston, TX, USA Chapter 62

Ahmed Hassan Fahal, MBBS FRCS MD MS Professor of Surgery, Mycetoma Research Centre, University of Khartoum, Khartoum, Sudan Chapter 83

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Paul E. Farmer, MD PhD Maude and Lillian Presley Professor of Social Medicine, Chair, Department of Global Health and Social Medicine, Harvard Medical School, Chief, Division of Global Health Equity, Brigham and Women’s Hospital, Co-founder, Partners in Health, Boston, MA Chapter 4

CONTRIBUTORS

A.S.G. Faruque, MBBS MPH Scientist, International Centre for Diarrhoeal Disease Research, Bangladesh (ICDDR,B), Dhaka, Bangladesh Chapter 20

Charles Feldman, MB BCh DSc PhD Professor of Pulmonology and Chief Physician, Charlotte Maxeke Johannesburg Academic Hospital and University of the Witwatersrand, Division of Pulmonology, Department of Internal Medicine, Faculty of Health Sciences, University of the Witwatersrand Medical School, Johannesburg, South Africa Chapter 29

Heinz Feldmann, MD Chief, Laboratory of Virology, Division of Intramural Research, NIAID, NIH, Rocky Mountain Laboratories, Hamilton, MT, USA Chapter 73

Kimberley K. Fox, MD MPH Medical Epidemiologist, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 138

Silvia Franceschi, MD Head, Section of Infections, International Agency for Research on Cancer, Lyon, France Chapter 11

Manuel A. Franco, MD PhD Professor, Instituto de Genética Humana, Facultad de Medicina, Pontificia Universidad Javeriana, Bogotá, Colombia Chapter 61

Charles F. Fulhorst, DVM DrPH Professor, University of Texas Medical Branch at Galveston, Department of Pathology, Galveston, TX, USA Chapter 71

Hector H. Garcia, MD PhD Professor, Department of Microbiology, School of Sciences, Director, Center for Global Health, Universidad Peruana Cayetano Heredia, Lima, Peru Chapter 119

Robert M. Genta, MD Chief for Academic Affairs, Caris Life Sciences, Irving, Texas, Clinical Professor of Pathology and Medicine (Gastroenterology), University of Texas Southwestern Medical Center at Dallas, Staff Pathologist, Dallas Veterans Affairs Medical Center, Dallas, TX, USA Chapter 117

Robert H. Gilman, MD DTM&H Professor, International Health, Director, Institute of Tropical Medicine and Public Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, MD, USA Chapter 23

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Roger I. Glass, MD PhD Director, Fogarty International Center, National Institutes of Health, Bethesda, MD, USA Chapter 63

Jerome Goddard, PhD Associate Extension Professor of Medical and Veterinary Entomology, Mississippi State University, Department of Entomology and Plant Pathology, Mississippi State University, MS, USA Chapter 124

Eduardo Gotuzzo, MD FACP FIDSA Director, Instituto de Medicina Tropical “Alexander von Humboldt,” Universidad Peruana Cayetano Heredia, Head, Departamento de Enfermedades Infecciosas Tropicales y Dermatológicas, Hospital Nacional Cayetano Heredia, Lima, Peru Chapters 40, 129, and 141

John R. Graybill, MD Professor of Medicine (Emeritus), Department of Medicine, Division of Infectious Diseases, University of Texas Health Science Center, San Antonio, TX, USA Chapters 85, 86, and 87

Harry B. Greenberg, MD Senior Associate Dean for Research, Joseph D. Grant Professor of Medicine and Microbiology and Immunology, Stanford University School of Medicine, Stanford, CA, USA Chapter 61

Paul D. Griffiths, MBBS MD DSc FRCPath Professor of Virology, University College London Medical School, Centre for Virology, UCL Medical School, London, UK Chapter 56

Duane J. Gubler, ScD FAAAS FIDSA Professor and Director, Program on Emerging Infectious Diseases, Duke-NUS Graduate Medical School, Singapore, Asia-Pacific Institute of Tropical Medicine and Infectious Diseases, University of Hawaii, Honolulu, Hawaii, USA Chapter 75

Richard L. Guerrant, MD Thomas H. Hunter Professor of International Medicine, Director, Center for Global Health, Division of Infectious Diseases and International Health, University of Virginia School of Medicine, Charlottesville, VA, USA Chapters 1, 12, 15, and 94

Yezid Gutierrez, MD PhD Retired, Formerly Adjunct, Laboratory Medicine, Cleveland Clinic, Cleveland, OH, USA Chapter 112

Erik L. Hewlett, MD Professor of Medicine, Division of Infectious Diseases and International Health, Department of Medicine, University of Virginia School of Medicine, Charlottesville, VA, USA Chapter 31

David L. Heymann, MD Professor, Infectious Disease Epidemiology, London School of Hygiene and Tropical Medicine, Head and Senior Fellow, Chatham House Centre on Global Health Security, London, UK Chapter 7

David R. Hill, MD DTM&H FRCP FFTM Honorary Professor, London School of Hygiene and Tropical Medicine, Director, National Travel Health Network and Centre, University College London Hospitals, NHS Foundation Trust, London, UK Chapter 93

Mei-Shang Ho, MD MPH Fellow, Institute of Biomedical Sciences, Academia Sinica, Taipei, Taiwan Chapter 59

Chapter 106

Paul S. Hoffman, PhD Professor of Medicine and Microbiology, Division of Infectious Diseases and International Health, University of Virginia School of Medicine, Charlottesville, VA, USA Chapter 32

Stephen L. Hoffman, MD DTMH DSc(Hon) Chief Executive and Scientific Officer, Sanaria Inc., Rockville, MD, USA Chapter 96

Michael R. Holbrook, PhD Senior Scientist, Battelle Memorial Institute, National Institutes for Health-Integrated Research Facility, Ft. Detrick, MD, USA; Adjunct Associate Professor, Department of Pathology, and Department of Microbiology and Immunology, The University of Texas Medical Branch, Galveston, TX, USA Chapters 70 and 77

Thomas L. Holland, MD Fellow, Infectious Diseases, Duke University Medical Center, Durham, NC, USA Chapter 13

Donald R. Hopkins, MD MPH Vice President, Health Programs, The Carter Center, Atlanta, GA, USA Chapter 108

Duane R. Hospenthal, MD PhD FACP FIDSA Chief, Infectious Disease Service, Brooke Army Medical Center, Fort Sam Houston, Texas, Professor of Medicine, Edward Hébert School of Medicine, Uniformed Services University of Health Sciences, Bethesda, MD, USA Chapter 90

Peter J. Hotez, MD PhD Distinguished Research Professor and Chair, Department of Microbiology Immunology and Tropical Medicine, George Washington University, School of Medicine and Public Health, Washington, DC, USA Chapter 116

S. David Hudnall, MD Professor of Pathology and Laboratory Medicine, Director of Hematopathology, Yale University School of Medicine, New Haven, CT, USA Chapter 56

James M. Hughes, MD Professor, Division of Infectious Diseases, Department of Medicine, School of Medicine, Professor, Hubert Department of Global Health, Rollins School of Public Health, Emory University, Atlanta, GA, USA Chapter 14

Kao-Pin Hwang, MD Department of Pediatrics, Chang Gung Memorial Hospital, Kaohsiung, Taiwan Chapter 111

Raul E. Isturiz, MD FACP Senior Consultant, Centro Medico de Caracas, Department of Internal Medicine, Infectious Diseases Section, Caracas, Venezuela Chapter 141

Peter B. Jahrling, PhD Director, NIAID Integrated Research Facility, Fort Detrick, Frederick, MD, USA Chapters 57 and 73

Shahid Jameel, PhD Senior Scientist and Group Leader, Virology, International Centre for Genetic Engineering and Biotechnology (ICGEB), New Delhi, India Chapter 64

Selma M.B. Jeronimo, MD Professor, Department of Biochemistry, Bioscience Center, Universidade Federal do Rio Grande do Norte, Natal, Brazil Chapter 100

Edward C. Jones-Lopez, MD MS Assistant Professor of Medicine, Section of Infectious Diseases, Boston University School of Medicine and Boston Medical Center, Boston, MA, USA

CONTRIBUTORS

Achim M. Hoerauf, MD Professor of Microbiology and Parasitology, Director and Chair, Institute of Medical Microbiology, Immunology and Parasitology, University of Bonn Medical Center, Bonn, Germany

Chapter 35

Anna Kabanova, MD PhD student, Novartis Vaccines and Diagnostics, Siena, Italy Chapter 34

Gagandeep Kang, MD FRCPath PhD Professor, Department of Gastrointestinal Sciences, Christian Medical College, Tamil Nadu, India Chapter 62

Christopher L. Karp, MD Gunnar Esiason/Cincinnati Bell Chair, Director, Division of Molecular Immunology, Professor of Pediatrics, Cincinnati Children’s Hospital Research Foundation, University of Cincinnati College of Medicine, Cincinnati, OH, USA Chapter 139

James W. Kazura, MD Professor of International Health and Medicine, Director, Center for Global Health and Diseases, Case Western Reserve University School of Medicine, Cleveland, OH, USA Chapter 104

Peter Kern, MD DTM&H FIDSA Professor of Infectious Diseases, Head, Division of Infectious Diseases, Comprehensive Infectious Diseases Center, Ulm University Hospital and Medical Center, Ulm, Germany Chapter 120

Gerald T. Keusch, MD Professor of International Health and of Medicine, Associate Director, National Emerging Infectious Diseases Laboratories, Director, Collaborative Core, Special Assistant to the President for Global Health, Boston University, Boston, MA, USA Chapter 18

Jay S. Keystone, MD MSc (CTM) FRCPC Professor of Medicine, Tropical Disease Unit, Toronto General Hospital, Toronto, ON, Canada Chapters 126 and 140

Mehnaaz S. Khuroo, MBBS MD Consultant Pathologist, Jawahir Lal Nehru Memorial Hospital, Rainawari, Srinagar, Kashmir, J&K, India Chapter 133

Mohammad S. Khuroo, MD DM FRCP (Edin) FACP MACP Emeritus Director, Digestive Diseases Centre, Dr Khuroo’s Medical Clinic, Srinagar, Kashmir, J&K, India Chapter 133

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Ik-Sang Kim, MD PhD Professor, Department of Microbiology and Immunology, Seoul National University College of Medicine, Seoul, Korea

CONTRIBUTORS

Chapter 51

Charles H. King, MD Professor of International Health, Center for Global Health and Diseases, Case Western Reserve University School of Medicine, Cleveland, OH, USA Chapter 122

Louis V. Kirchhoff, MD MPH Professor, Division of Infectious Diseases, Department of Internal Medicine, Department of Epidemiology, University of Iowa, Staff Physician, Department of Veterans Affairs Medical Center, Iowa City, IA, USA Chapter 99

Amy D. Klion, MD Head, Eosinophil Pathology Unit, Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, Bethesda, MD, USA Chapter 105

Keith P. Klugman, MD PhD William H. Foege Professor of Global Health, Hubert Department of Global Health, Rollins School of Public Health; Professor of Epidemiology and Infectious Diseases, Emory University, Atlanta, GA, USA, Co-Director of Respiratory and Meningeal Pathogens Research Unit, Medical Research Council and University of the Witwatersrand, Johannesburg, South Africa Chapter 29

Dennis J. Kopecko, PhD Senior Research Scientist and Chief, Laboratory of Enteric and Sexually Transmitted Diseases, US Food and Drug Administration, Center for Biologics Evaluation and Research, Bethesda, MD, USA Chapter 18

Margaret Kosek, MD Assistant Scientist, Department of International Health, Johns Hopkins Bloomberg School of Public Health, Baltimore, MD, USA Chapter 5

Frederick T. Koster, MD Assistant Scientist, Respiratory Immunology and Asthma Program, Lovelace Respiratory Research Institute, Albuquerque, NM, USA Chapter 71

Phyllis E. Kozarsky, MD Professor of Medicine, Emory University School of Medicine, Atlanta, GA, Expert Consultant, Division of Global Migration and Quarantine, CDC, Atlanta, GA, USA Chapter 126

Thomas G. Ksiazek, DVM PhD Professor, Galveston National Laboratory, Department of Pathology, University of Texas Medical Branch, Galveston, TX, USA Chapter 55

Jens H. Kuhn, MD PhD MS Managing Consultant/Lead Virologist, Integrated Research Facility, Division of Clinical Research/NIAID/NIH, Fort Detrick, Frederick, MD, USA Chapter 73

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Albert J. Lastovica, PhD Extraordinary Professor of Food Microbiology, Department of Biotechnology, Faculty of Natural Sciences, University of the Western Cape, Bellville, South Africa Chapter 19

James W. LeDuc, PhD Professor, Microbiology and Immunology, Robert E. Shope MD and John S. Dunn Distinguished Chair in Global Health, Director, Galveston National Laboratory, University of Texas Medical Branch, Galveston, TX, USA Chapter 14

Peter A. Leone, MD Professor of Medicine, University of North Carolina, Medical Director, HIV/STD Prevention and Care, North Carolina Department of Health and Human Services Communicable Diseases Branch, University of North Carolina, Chapel Hill, NC, USA Chapter 25

Paul N. Levett, PhD ABMM FCCM Assistant Clinical Director, Provincial Laboratory, Saskatchewan Health, Regina, Saskatchewan, Canada Chapter 45

Michael Levin, FMed Sci FRCPCh Professor of Paediatrics and International Child Health, Department of Paediatrics, Division of Medicine, Imperial College, London, UK Chapter 6

Myron M. Levine, MD DTPH Simon and Bessie Grollman Distinguished Professor and Director, University of Maryland School of Medicine, Center for Vaccine Development, Baltimore, MD, USA Chapters 16 and 20

Aldo A.M. Lima, MD PhD Professor and Director, INCT Institute of Biomedicine and Center for Global Health, Department of Physiology and Pharmacology, School of Medicine, Federal University of Ceará, Fortaleza, Brazil Chapter 94

Gerhard Lindeque, MD FCOG(SA) FRCOG Professor of Obstetrics and Gynaecology, University of Pretoria, Pretoria, South Africa Chapter 80

David L. Longworth, MD Chair, Medical Institute, Cleveland Clinic, Cleveland, OH, USA Chapter 132

David Mabey, DM FRCP Professor of Communicable Diseases, London School of Hygiene and Tropical Medicine, London, UK Chapter 47

J. Dick Maclean†, MD FRCPC MRCP(UK) DCMT(Lond) Formerly Professor of Medicine and Director, McGill Centre for Tropical Diseases, McGill University, Senior Physician, Division of Tropical Diseases, Department of Medicine, Montreal General Hospital, Montreal, Quebec, Canada Chapter 123

Alan J. Magill, MD FACP FIDSA – COL/MC Director, Division of Experimental Therapeutics, Walter Reed Army Institute of Research, Silver Spring, MD, USA Chapter 128

Ismael Maguilnik, MD Professor of Internal Medicine, Gastroenterology, Federal University of Rio Grande do Sul, Brazil, Head of Endoscopy, Department of Clinical Hospital, Federal University of Rio Grande do Sul, Brazil Chapter 117

†

deceased

Chapter 39

James H. Maguire, MD MPH Professor of Medicine, Brigham and Women’s Hospital, Boston, MA, USA Chapters 97, 101, and 103

Siddhartha Mahanty, MBBS MPH Staff Clinician, Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA Chapters 123 and 139

Shinji Makino, DVM PhD Edgar and Mary Frances Monteith Distinguished Professorship in Viral Genetics, Professor, Department of Microbiology and Immunology, The University of Texas Medical Branch, Galveston, TX, USA Chapter 69

Christian W. Mandl, MD PhD Associate Professor of Virology, Department of Virology, Medical University of Vienna, Vienna, Austria, Vice President, Novartis Vaccines and Diagnostics, Cambridge, MA, USA Chapter 77

Thomas J. Marrie, MD Dean, Faculty of Medicine, Dalhousie University, Halifax, Nova Scotia, Canada Chapters 32, 48, and 53

Barry J. Marshall, AC FRACP FAA FRS Clinical Professor of Microbiology, The University of Western Australia, Faculty of Life and Physical Sciences, School of Biomedical Biomolecular and Chemical Sciences, Discipline of Microbiology and Immunology, Crawley, WA, Australia Chapter 23

Gregory J. Martin, MD Captain, Medical Corps, US Navy, National Naval Medical Center, Associate Professor of Medicine and Preventive Medicine, Uniformed Services University, Bethesda, MD, USA Chapter 134

Tadahiko Matsumoto, MD DMSc Assistant Director of Clinical Medicine and Consultant Dermatologist, Yamada Institute of Health and Medicine, Tokyo, Japan, Clinical Professor, Department of Dermatology, Juntendo University, Tokyo, Japan, Clinical Professor, Department of Dermatology, Kurume University, Kurume, Japan Chapter 84

Steven D. Mawhorter, MD DTM&H Staff Physician and Director, International Traveler’s Health Clinic, Department of Infectious Diseases, Cleveland Clinic, Associate Professor of Medicine, Cleveland Clinic Lerner College of Medicine of Case Western Reserve University, Cleveland, OH, USA Chapter 132

James S. McCarthy, MD Professor of Medicine and Senior Consultant in Infectious Diseases, Royal Brisbane and Women’s Hospital, Queensland Institute of Medical Research, University of Queensland, Brisbane, Australia Chapters 109 and 113

Michael R. McGinnis, PhD Fungus Testing Lab, Department of Pathology, University of Texas Health Science Center at San Antonio, Professor Emeritus, Department of Pathology, University of Texas Medical Branch, Galveston, TX, USA Chapters 82 and 89

Paul S. Mead, MD MPH Chief, Epidemiology and Surveillance Activity, Bacterial Diseases Branch, Division of Vector-borne Infectious Diseases, Centers for Disease Control and Prevention, Fort Collins, CO, USA Chapter 41

Wayne M. Meyers, MD PhD DSc Visiting Scientist, Department of Environmental and Infectious Disease Sciences, Armed Forces Institute of Pathology, Washington, DC, USA

CONTRIBUTORS

Ciro Maguiña, MD Director, Adjunto del IMT-AVH-UPCH, Departamento de Medicina, Universidad Peruana Cayetano Heredia, Médico Infectólogo Tropicalista, Departamento de Medicina Tropical, Hospital Nacional Cayetano Heredia, Lima, Peru

Chapters 36 and 37

Robert F. Miller, MB BS FRCP Reader in Clinical Infection, University College London, London, UK Chapter 91

Samuel I. Miller, MD Professor of Medicine, Microbiology, Genome Sciences, Adjunct Professor of Immunology, University of Washington School of Medicine, Seattle, WA, USA Chapter 17

James N. Mills, PhD Chief, Medical Ecology Unit, Division of Viral and Rickettsial Diseases, National Center for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 68

Thomas P. Monath, MD Partner, Kleiner Perkins Caufield and Byers, Harvard, MA, USA Chapter 74

Christopher C. Moore, MD Assistant Professor, Division of Infectious Diseases and International Health, Department of Medicine, University of Virginia, Charlottesville, VA, USA Chapter 17

Thomas A. Moore, MD FACP Clinical Professor, Department of Medicine, University of Kansas School of Medicine, Wichita, KS, USA Chapters 109 and 113

J.C. Morrill, DVM PhD Visiting Professor, Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, TX, USA Chapter 69

J. Glenn Morris, Jr, MD MPH&TM Director, Emerging Pathogens Institute, Professor of Medicine (Infectious Diseases), College of Medicine, Adjunct Professor, College of Public Health and Health Professions, University of Florida, Gainesville, FL, USA Chapter 21

Megan Murray, MD Associate Professor of Epidemiology, Department of Epidemiology, Harvard School of Public Health, Boston, MA, USA Chapter 3

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K. Darwin Murrell Adjunct Professor, USUHS School of Medicine, Department of Preventive Medicine and Biometrics, Bethesda, MD, USA, Honorary Professor, Faculty of Life Science, University of Copenhagen, Denmark

Ynés R. Ortega, PhD MPH Associate Professor, Center for Food Safety, University of Georgia, Griffin, GA, USA Chapter 95

Chapter 110

CONTRIBUTORS

G. Balakrish Nair, PhD Director, National Institute of Cholera and Enteric Diseases, Beliaghata, Kolkata, India Chapter 21

Theodore E. Nash, MD Head, Gastrointestinal Parasites Section, Laboratory of Parasitic Diseases, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, USA Chapter 93

Barnett R. Nathan, MD Assistant Professor, Department of Neurology, University of Virginia, Charlottesville, VA, USA Chapter 42

Ricardo Negroni, MD Honorary Professor of Microbiology and Parasitology, University of Buenos Aires School of Medicine, Consultant, Francisco Javier Muñiz Hospital, Buenos Aires, Argentina Chapter 86

Anne Nicholson-Weller, MD Professor of Medicine, Harvard Medical School, Divisions of Allergy/ Inflammation and Infectious Diseases, Department of Medicine, Beth Israel Deaconess Medical Center, Boston, MA, USA Chapter 9

Marcio Nucci, MD Associate Professor, Department of Internal Medicine, Chief, Mycology Laboratory, University Hospital, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil Chapter 88

Thomas B. Nutman, MD Head, Helminth Immunology Section and Head, Clinical Parasitology Unit, Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, Bethesda, MD, USA Chapters 104 and 105

Nigel O’Farrell, MD FRCP Consultant Physician, Ealing Hospital, London, UK Chapter 28

Juan P. Olano, MD Associate Professor, Department of Pathology, Director, Residency Training Program, Member, Center for Biodefense and Emerging Infectious Diseases, University of Texas Medical Branch, Galveston, TX, USA Chapters 1 and 125

Eng Eong Ooi, BMBS PhD Associate Professor, Program in Emerging Infectious Diseases, Duke-NUS Graduate Medical School, Singapore Chapter 75

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Luis S. Ortega, MD Chief, Asia Regional Program, Division of Global Migration and Quarantine, National Center for Emerging and Zoonotic Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 127

Mark A. Pallansch, PhD Chief, Polio and Picornavirus Laboratory Branch, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 60

Jean W. Pape, MD Professor of Medicine, Center for Global Health, Division of Infectious Diseases, Department of Medicine, Weill Medical College of Cornell University, New York, NY, USA, Director, Les Centres GHESKIO, Port-au-Prince, Haiti Chapter 95

Georgios Pappas, MD Physician, Institute of Continuing Medical Education of Ioannina, Ioannina, Greece Chapter 40

Julie Parsonnet, MD George DeForest Barnett Professor of Medicine and Professor of Health Research and Policy, Stanford University School of Medicine, Stanford, CA, USA Chapter 11

Geoffrey Pasvol, MA MB ChB DPhil FRCP FRCPE Professor, Wellcome Centre for Clinical Tropical Medicine, Imperial College London, Wright Fleming Institute, London, UK Chapters 6 and 137

Sharon J. Peacock, BM BA MSc FRCP FRCPath PhD Professor of Clinical Microbiology, Department of Medicine, University of Cambridge, Addenbrooke’s Hospital, Cambridge, UK Chapter 33

Richard D. Pearson, MD Professor of Medicine and Pathology, Division of Infectious Diseases and International Health, University of Virginia School of Medicine, Charlottesville, VA, USA Chapters 12, 13, and 100

Rosanna W. Peeling, PhD Professor and Chair of Diagnostic Research, London School of Hygiene and Tropical Medicine, London, UK Chapter 47

David A. Pegues, MD Professor of Medicine, Division of Infectious Diseases, David Geffen School of Medicine, Los Angeles, CA, USA Chapter 17

Jacques Pépin, MD FRCPC MSc Professor of Infectious Diseases, University of Sherbrooke, Sherbrooke, Quebec, Canada Chapter 98

C.J. Peters, MD Professor of Pathology and Microbiology and Immunology, Center for Biodefense and Emerging Infectious Diseases, WHO Collaborating Center for Tropical Diseases, University of Texas Medical Branch, Galveston, TX, USA Chapters 67, 68, 69, 71, 73, and 125

Chapter 92

William A. Petri, Jr, MD PhD Professor and Chief, Wade Hampton Frost Professor, Division of Infectious Diseases, University of Virginia Health Sciences Center, Charlottesville, VA, USA Chapter 92

Françoise Portaels, PhD Professor and Head, Unit of Mycobacteriology, Department of Microbiology, Institute of Tropical Medicine, Antwerp, Belgium Chapter 36

José Luiz Proença-Módena, PhD Research Fellow, Department of Cell Biology and Virology Research Center, University of São Paulo School of Medicine, Ribeirão Preto, SP, Brazil Chapter 58

Thomas C. Quinn, MD MSc Professor of Medicine, International Health, Epidemiology, and Molecular Microbiology and Immunology, Division of Infectious Diseases, The John Hopkins Medical Institutions, Baltimore, MD, USA Chapter 81

G. Raghurama Rao, MD Former Professor and Head of Department of Dermatology, Andhra Medical College, Andhra Pradesh, India Chapter 38

Didier Raoult, MD PhD Professor, Unité des Rickettsies, Faculté de Médecine, Marseille, France Chapter 50

Rino Rappuoli, PhD Global Head of Vaccine Research, Novartis Vaccines and Diagnostics, Siena, Italy Chapter 34

John H. Rex, MD Vice President Clinical Infection, AstraZeneca Pharmaceuticals; Adjunct Professor of Medicine, University of Texas Medical School, Houston, TX, USA Chapter 89

Steven J. Reynolds, MD MPH FRCPC DTM&H Senior Clinician, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Washington, DC, USA Chapter 81

José M.C. Ribeiro, MD PhD Chief, Vector Biology Section, Laboratory of Malaria and Vector Research, NIAID, National Institutes of Health, Rockville, MD, USA Chapter 8

Emmanuel Roilides, MD PhD Professor of Pediatrics, Third Department of Pediatrics, Aristotle University, Hippokration Hospital, Thessaloniki, Greece Chapter 89

Pierre E. Rollin, MD Deputy Branch Chief, Special Pathogens Branch, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 55

Allan R. Ronald, MD FRCPC MACP Distinguished Professor Emeritus, University of Manitoba, Winnipeg, Manitoba, Canada Chapters 26 and 138

Paul A. Rota, PhD Lead Scientist, Measles Team, Measles, Mumps, Rubella and Herpesviruses Laboratory Branch, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 55

Sharon L. Roy, MD MPH Medical Epidemiologist, Waterborne Disease Prevention Branch, Division of Foodborne, Waterborne, and Environmental Diseases, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 101

CONTRIBUTORS

Kristine M. Peterson, MD MPH Assistant Professor, Division of Infectious Diseases, University of Virginia Health System, Charlottesville, VA, USA

Ernesto Ruiz-Tiben, PhD Director, Dracunculiasis Eradication, The Carter Center, Atlanta, GA, USA Chapter 108

Edward T. Ryan, MD Associate Professor of Medicine, Harvard Medical School; Associate Professor of Immunology and Infectious Diseases, Harvard School of Public Health; Director, Tropical and Geographic Medicine, Massachusetts General Hospital, Boston, MA, USA Chapters 3 and 135

Debasish Saha, MBBS MS Clinical Epidemiologist, Bacterial Diseases Programme, Medical Research Council (UK) Laboratories, Banjul, The Gambia, West Africa Chapter 20

Mohammed A. Salam, MD Director, Clinical Sciences Division, International Centre for Diarrhoeal Disease Research, Bangladesh (ICDDR,B), Dhaka, Bangladesh Chapter 18

Amidou Samie, BSc MSC PhD Senior Lecturer, Department of Microbiology, University of Venda, Thohoyandou, South Africa Chapter 94

Julius Schachter, PhD Professor Emeritus of Laboratory Medicine, Chlamydia Laboratory, University of California, San Francisco, San Francisco, CA, USA Chapter 46

Peter M. Schantz, MD VMD PhD Epidemiologist and Senior Service Fellow, Division of Parasitic Diseases/NCZVED/CCID, National Center for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapter 120

W. Michael Scheld, MD Professor of Medicine, Division of Infectious Diseases and International Health, Department of Medicine, University of Virginia School of Medicine, Charlottesville, VA, USA Chapter 24

Elizabeth P. Schlaudecker, MD Pediatric Infectious Diseases Fellow, Division of Infectious Diseases, Cincinnati Children’s Hospital Medical Center, Cincinnati, OH, USA Chapter 27

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David A. Schwartz, MD MSHyg FCAP Associate Clinical Professor of Pathology, Department of Pathology, Vanderbilt University School of Medicine, Nashville, TN, USA Chapter 102

CONTRIBUTORS

Joseph D. Schwartzman, MD Professor of Pathology, Department of Pathology, Dartmouth Medical School, Lebanon, NH, USA Chapter 103

Chapter 91

Cynthia B. Snider, MD MPH Infectious Diseases Fellow, University of Virginia, Charlottesville, VA, USA Chapter 58

Arlene C. Seña, MD MPH Clinical Associate Professor, Division of Infectious Diseases, University of North Carolina at Chapel Hill, Chapel Hill, NC, Medical Director, Durham County Health Department, Durham, NC, USA Chapters 43 and 138

Daniel J. Sexton, MD FACP FIDSA Professor, Department of Medicine, Division of Infectious Diseases, Duke University Medical Center, Durham, NC, USA Chapter 49

Truman W. Sharp, MD MPH (Capt, MCUSN) Commanding Officer, US Naval Medical Research, Unit No. 3, Cairo, Egypt Chapter 128

Wun-Ju Shieh, MD MPH PhD Pathologist, Infectious Diseases Pathology Branch, Division of HighConsequence Pathogens & Pathology, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapters 45 and 68

Shmuel Shoham, MD Scientific Director, MedStar Clinical Research Center, Washington Hospital Center, Assistant Professor of Medicine, Georgetown University School of Medicine, Washington, DC, USA Chapter 88

Afzal A. Siddiqui, PhD Professor, Microbiology and Immunology, Internal Medicine, Pathology, School of Medicine, Texas Tech University Health Sciences Center, Lubbock, TX, USA Chapter 117

Upinder Singh, MD Associate Professor, Chief, Division of Infectious Diseases and Geographic Medicine, Stanford University School of Medicine, Stanford, CA, USA Chapter 92

David W. Smith, BMedSc MBBS FRCPA FACTM Clinical Professor, School of Biomedical, Biomolecular and Chemical Sciences and School of Pathology and Laboratory Medicine, University of Western Australia, Nedlands, Western Australia, Australia Chapter 78

Michael B. Smith, MD MS Terminology Manager, Clinical Informatics, SNOMED Terminology Solutions, College of American Pathologists, Northfield, IL, USA Chapter 82

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A. George Smulian, MD Chief, Infectious Disease Section, Cincinnati VA Medical Center, Associate Professor of Medicine, University of Cincinnati, Cincinnati, OH, USA

Bonnie L. Smoak, MD PhD MPH (Col, MCUSA) Associate Professor, Department of Preventive Medicine and Biometrics, Uniformed Services University of the Health Sciences, Bethesda, MD, Director, Division of Preventive Medicine, Walter Reed Army Institute of Research, Silver Spring, MD, USA Chapter 128

Tom Solomon, BA BM BCh FRCP DCH DTMH PhD Chair of Neurological Science, Head of Brain Infections Group and Director of Institute of Infection and Global Health, University of Liverpool, Liverpool, UK Chapters 76 and 136

Samba O. Sow, MD MS Associate Professor of Medicine, General Director, Center for Vaccine Development-Mali (CVD-MALI), ex-Institut Marchoux, Ministry of Health, Bamako, Mali Chapter 20

P. Frederick Sparling, MD Professor of Medicine and Microbiology and Immunology, University of North Carolina School of Medicine, Chapel Hill, NC, USA Chapter 43

Lisa A. Spencer, PhD Assistant Professor of Medicine, Harvard Medical School, Division of Allergy/Inflammation, Department of Medicine, Beth Israel Deaconess Medical Center, Boston, MA, USA Chapter 9

Lawrence R. Stanberry, MD PhD FAAP Reuben S. Carpentier Professor and Chairman, Department of Pediatrics, College of Physicians and Surgeons, Columbia University, New York, NY, USA Chapter 56

J. Erin Staples, MD PhD Medical Epidemiologist, Arboviral Diseases Branch, Division of Vector-borne Infectious Diseases, Centers for Disease Control and Prevention, Fort Collins, CO, USA Chapter 74

Robert Steffen, MD Hon FFTM/ACTM Emeritus Professor of Travel Medicine, University of Zurich Center of Travel Medicine, WHO Collaborating Centre for Travellers’ Health, Zurich, Switzerland Chapter 126

Theodore S. Steiner, MD Associate Professor of Medicine, University of British Columbia, Vancouver, Canada Chapter 15

Mark C. Steinhoff, MD Director, Global Health Center, Professor of Pediatrics, Division of Infectious Diseases, Cincinnati Children’s Hospital Medical Center, University of Cincinnati College of Medicine, Cincinnati, OH, USA Chapter 27

Dennis L. Stevens, PhD MD Chief, Infectious Disease Section, Veterans Affairs Medical Center, Boise, ID, Professor, Department of Medicine, University of Washington School of Medicine, Seattle, WA, USA Chapter 30

Kathryn N. Suh, MD FRCPC Associate Professor of Medicine, Division of Infectious Diseases, The Ottawa Hospital, Ottawa, ON, Canada Chapter 140

Diederik van de Beek, MD PhD Neurologist, Department of Neurology, Center of Infection and Immunity Amsterdam (CINIMA), Academic Medical Center University of Amsterdam, Amsterdam, The Netherlands

Chapter 87

Paul J. Szaniszlo, PhD Professor Emeritus, The University of Texas at Austin, Section of Molecular Genetics and Microbiology, Austin, TX, USA Chapter 84

Milagritos D. Tapia, MD Assistant Professor, University of Maryland School of Medicine, Center for Vaccine Development, Baltimore, MD, USA Chapter 16

Herbert B. Tanowitz, MD Professor of Pathology and Medicine, Director of the Diagnostic Parasitology Laboratory, Jacobi Medical Center, Albert Einstein College of Medicine, Bronx, NY, USA Chapters 118 and 121

Sam R. Telford III, ScD Associate Professor, Tufts University, Cummings School of Veterinary Medicine, North Grafton, MA, USA Chapter 97

Robert B. Tesh, MD Professor of Pathology and Microbiology and Immunology, Center for Biodefense and Emerging Infectious Diseases, University of Texas Medical Branch, Galveston, TX, USA Chapters 72 and 76

Nathan M. Thielman, MD MPH Associate Professor of Medicine, Director, Duke Global Health Residency Program, Division of Infectious Diseases and International Health, Duke Global Health Institute, Duke University, Durham, NC, USA Chapters 13, 15, and 129

Fernando J. Torres-Vélez, DVM PhD Veterinary Pathologist, Infectious Disease Pathogenesis Section, Comparative Medicine Branch (CMB), National Institute of Allergy and Infectious Diseases (NIAID), National Institutes of Health (NIH), Bethesda, MD, USA Chapter 2

Joseph D. Tucker, MD MA Instructor in Medicine, Harvard Medical School, Assistant in Medicine, Massachusetts General Hospital, Boston, MA, USA Chapter 43

Luis M. Valdez, MD Associate Professor of Medicine, Universidad Peruana Cayetano Heredia, Internal Medicine-Infectious Diseases Specialist, British American Hospital, Lima, Peru Chapter 129

Jesus G. Valenzuela, PhD Chief, Vector Molecular Biology Section, Laboratory of Malaria and Vector Research, NIAID, National Institutes of Health, Rockville, MD, USA Chapter 8

Pedro F.C. Vasconcelos, MD PhD Chief, Department of Arbovirology and Hemorrhagic Fevers, Director, WHO Collaborating Center for Research, Diagnostic Reference and Training on Arbovirus, Director, National Institute for Viral Hemorrhagic Fevers, Instituto Evandro Chagas/SVS/MS, Ananindeua, Brazil, Professor of Pathology, Pará State University, Belém, Brazil Chapter 72

Govinda S. Visvesvara, PhD Research Microbiologist, Division of Foodborne, Waterborne, and Environmental Diseases, Centers for Disease Control and Prevention, Atlanta, GA, USA

CONTRIBUTORS

Chapter 24

Khuanchai Supparatpinyo, MD Professor of Infectious Disease, Department of Medicine, Faculty of Medicine, Chiang Mai University, Chiang Mai, Thailand

Chapter 101

Victoria Wahl-Jensen, MS PhD Managing Consultant/Virologist, Integrated Research Facility, Division of Clinical Research/NIAID/NIH, Fort Detrick, Frederick, MD, USA Chapter 73

David H. Walker, MD Carmage and Martha Walls Distinguished University Chair in Tropical Diseases, Director, Center for Biodefense and Emerging Infectious Diseases, Professor and Chair, Department of Pathology, University of Texas Medical Branch, Galveston, TX, USA Chapters 1, 38, 39, 49, 50, 51, and 125

Douglas S. Walsh, MD FAAD Chief, Department of Immunology and Medicine, United States Army Medical Component, Armed Forces Research Institute of Medical Sciences (AFRIMS), Bangkok, Thailand Chapters 36 and 37

Thomas J. Walsh, MD FACP FAAM FIDSA Professor of Medicine, Director, Transplantation – Oncology, Infectious Diseases Program, Weill Cornell Medical College, New York, Presbyterian Hospital, New York City, New York, USA Chapters 88 and 89

David A. Walton, MD Instructor in Medicine, Department of Internal Medicine, Brigham and Women’s Hospital, Boston, MA, USA Chapter 4

Peter D. Walzer, MD MSc Associate Chief of Staff for Research, Cincinnati VA Medical Center, Professor of Medicine, University of Cincinnati, Cincinnati, OH, USA Chapter 91

Cirle A. Warren, MD Assistant Professor, Center for Global Health, Division of Infectious Disease and International Health, Charlottesville, VA, USA Chapter 22

Scott C. Weaver, PhD Professor, Department of Pathology, Scientific Director, Galveston National Laboratory, Director, Institute for Human Infections and Immunity, University of Texas Medical Branch at Galveston, Galveston, TX, USA Chapter 78

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Louis M. Weiss, MD MPH Professor of Pathology, Division of Parasitology and Tropical Medicine, Professor of Medicine, Division of Infectious Diseases, Albert Einstein College of Medicine, Bronx, NY, USA

Robert J. Wilkinson, MA PhD BM BCh DTM&H FRCP Wellcome Trust Senior Fellow, MRC Programme Leader, Imperial College London, MRC National Institute for Medical Research, University of Cape Town, Cape Town, South Africa

Peter F. Weller, MD FACP FIDSA Professor of Medicine, Harvard Medical School, Professor, Immunology and Infectious Diseases Department, Harvard School of Public Health, Chief, Infectious Disease Division, Vice Chair of Research, Department of Medicine, Beth Israel Deaconess Medical Center, Boston, MA, USA

Mary E. Wilson, MD FACP FIDSA Associate Professor of Global Health and Population, Harvard School of Public Health, Boston, MA, USA

CONTRIBUTORS

Chapter 102

Chapters 1, 12, 97, and 131

A. Clinton White, Jr, MD FACP FIDSA Paul R. Stalnaker MD Distinguished Professor, Director, Infectious Disease Division, Department of Internal Medicine, University of Texas Medical Branch, Galveston, TX, USA Chapters 118, 119, and 121

Nicholas J. White, OBE DSc MD FRCP FMedSci FRS Professor of Tropical Medicine, Mahidol University and Oxford University, Faculty of Tropical Medicine, Mahidol University, Bangkok, Thailand Chapter 96

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Chapter 6

Chapters 130 and 131

Murray Wittner, MD PhD Professor Emeritus, Director, International Health and Travel Medicine Clinic, Montefiore Medical Center, Bronx, NY, USA Chapters 118 and 121

Anita K.M. Zaidi, MBBS SM FAAP A. Sultan Jamal Professor of Pediatrics and Child Health, Department of Pediatrics and Child Health, Aga Khan University, Karachi, Pakistan Chapter 16

Sherif R. Zaki, MD Chief, Infectious Disease Pathology, Centers for Disease Control and Prevention, Atlanta, GA, USA Chapters 45 and 67

SECTION I: PRINCIPLES AND GENERAL CONSIDERATIONS

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CHAPTER 1 

Principles of Parasitism: Host–Parasite Interactions Juan P. Olano • Peter F. Weller • Richard L. Guerrant • David H. Walker

INTRODUCTION The relationship between two living organisms can be classified as parasitic, symbiotic, or commensal.1–3 This same classification scheme can be used to describe relationships between microorganisms and more complex living organisms that act as hosts. The term parasite is used here in its broad sense to mean a microorganism interacting with another organism (either vertebrate or invertebrate) in the same ecologic niche. The following definitions are used in this chapter: Parasitism: Association between two different organisms wherein one benefits at the expense of the other. All infectious agents causing illness belong to this category. Commensalism: Association between two organisms in which one derives benefit from the other without causing it any harm. This intermediate category is not uniformly accepted. Often, upon detailed analysis, the relationship turns out to be either parasitic or symbiotic.2 Symbiosis or mutualism: Both organisms benefit from the relationship. The type of relationship also depends on host factors. For example, bacteria normally inhabiting the bowel live in an apparent commensal or (by inhibiting potential pathogens) symbiotic relationship with humans. However, in cases of cirrhosis with consequent hepatic insufficiency, bacteria can become a dangerous source of ammonia that leads to hepatic encephalopathy. A commensal relationship can be transformed into a potentially harmful one.

MICROBIAL FACTORS Principles of Microbial Evolution and Classification The Earth is approximately 4.5 to 5 billion years old. There is good fossil evidence of microbial life approximately 3.5 billion years ago. Microbial life (stromatolites) was mostly photosynthetic, unicellular, and anaerobic.1,4 Eukaryotes, bacteria, and archaea evolved from a still hypothetical universal common ancestor.5–7 Eukaryotes then evolved into protozoans, metazoans, plants, and animals, as we know them today. Moreover, there is strong evidence that primitive eukaryotic cells established relationships with bacterial organisms that later evolved into cytoplasmic organelles, such as chloroplasts in plants and mitochondria in animals.8 To put things into perspective, approximately five-sixths of the history of life on Earth has been exclusively microbial. Human beings appeared on the planet only 2 million years ago as very late newcomers to the biosphere. Life was initially anaerobic, but with the appearance of photosynthetic organisms and chloroplasts, oxygen was released into the atmosphere for the first time.9 Radiation in the upper atmosphere created

the ozone layer from molecular oxygen, which then shielded the Earth’s surface from dangerous radiation. Nucleic acids were therefore protected from harmful mutations. Organisms had to evolve to survive in the pre­ sence of oxygen. A few of the ancient anaerobes were able to survive in the highly oxidant atmosphere, and they represent the anaerobes as we know them today. The phylogeny of living organisms is based on molecular approaches, particularly analysis of ribosomal RNA.5,6,10 Because of the antiquity of the protein synthesis machinery, these molecules are excellent evolutionary clocks. For prokaryotes, the 16S subunit of ribosomes is the most useful for classification purposes. Viruses deserve special comment because of their molecular simplicity and at the same time their importance as human pathogens and as possible agents of hereditary changes and cancer.11,12 A virus is a genetic element with either DNA or RNA coated by protein of viral origin and sometimes enveloped by lipid material of host origin. Some viruses have enzymes that are necessary for their replication. The only criterion that these organisms fulfill to be considered living organisms is that of reproduction. They are inert particles when outside of the host cell, and once they have access into a cell they become active and the cell is subverted to produce more viral particles. Sometimes the cell dies in the process, and sometimes the relationship is stable. Viral hosts include bacteria, protozoa, animals, and plants. The classification of viruses is based on different criteria than the ones used for other organisms. The major criteria are type of nucleic acid, presence or absence of an envelope, manner of replication, and morphologic characteristics.12,13 Simpler forms of self-replicating organisms include virusoids and viroids.14,15 The former are satellite RNAs that are found encapsidated in the proteins encoded by their helper virus (e.g., hepatitis caused by the hepatitis D virus delta agent in conjunction with hepatitis B virus). The viroids are mostly plant pathogens that consist of single-stranded circular RNA molecules. The concept of “infectious agent” was revolutionized by the dis­covery of proteinaceous infectious agents known as prions. These proteins are responsible for neurodegenerative diseases in animals and humans. The protein particles lack nucleic acids but still are able to reproduce and trigger conformational changes in host proteins, leading to cell death.16,17 In contrast, protozoa are nucleated, single-cell organisms that, depending on the species, replicate by means as simple as binary fission (e.g., Trichomonas) or as complex as involving multiple sexual and asexual stages in both animal and invertebrate hosts (e.g., plasmodia). Protozoa include amebae (e.g., Entamoeba histolytica), flagellates (e.g., Giardia lamblia), ciliates (e.g., Balantidium coli), and sporozoa (e.g., Cryptosporidium). Even more complex are helminths, which are multicellular metazoan organisms with highly developed internal organs, including alimen­ tary and reproductive tracts. The helminths include nematodes (roundworms), cestodes (tapeworms), and trematodes (flukes). Many helminths

1

SECTION I

have complex life cycles with multiple developmental stages both in the animal host and in intermediate invertebrate or vertebrate hosts. Because of their size, helminths, the macroparasites, are solely extracellular pathogens; because of their prolonged life cycles and generation times, their capacity for genetic alteration is diminished compared to smaller, simpler microbes (the microparasites).

PRINCIPLES AND GENERAL CONSIDERATIONS

Development of Microbial Virulence

  

 

Evolution of Virulence

2

The traditional view assumes that natural selection would favor evolution toward a benign coexistence between host and parasite.18,19 A modern view of evolution of virulence focuses on the tradeoff between the benefits that pathogens accrue through increased exploitation of hosts and the costs that result from any effects of disease that reduce transmission to susceptible hosts.19,20 From this point of view, virulence could be the evolved as well as the primitive stage of the association between host and parasite, depending on the development of enhanced rather than reduced transmission. According to Levin19 and Levin and Svanborg-Eden,20 there are three alternative models to explain evolution of a microparasite’s virulence: direct selection, coincidental evolution, and short-sighted within-host selection. The direct selection model states that there is a direct relationship between the parasite’s virulence and its rate of infectious transmission. The best documented and often cited example is that of the dramatic changes in virulence that the myxoma virus underwent after being released into the wild in Australia to “control” the population of wild rabbits. In the beginning, rabbit mortality and viral transmission rates were high. As the population of rabbits was decimated, the virulence of the virus decreased and its rate of transmission actually increased. This outcome is explained by the longer survival and duration of the period of shedding of the virus. At the same time, more resistant rabbits increased in number due to the selection process.21 According to the coincidental evolution model, the factors responsible for the virulence of a microparasite evolved for some purpose other than to provide the parasite with some advantage within a host or for its transmission to other hosts. Clostridial toxins are good examples in this category. There is no beneficial reason to kill a human host who became infected by Clostridium tetani spores from soil in order for the parasite to survive. They are mostly soil bacteria and do not need humans for their survival.19 Short-sighted within-host evolution posits that the parasites responsible for the morbidity and mortality of the host are selected for as a consequence of within-host evolution since that produces a local advantage for their survival within the host. The host dies and the rate of transmission would decrease. This is an example of evolutionary myopia in which the long-term consequences of killing a host would not matter to the parasite.22,23 Natural selection is a local phenomenon that happens at a given time and place and goes perfectly with this model. Bacteria such as Neisseria meningitidis that normally live attached to human pharyngeal epithelial cells sometimes invade the central nervous system (CNS) and kill the host. Their replication in the CNS is favored since competition is low and defenses are not as abundant as in the tonsillar areas.19 It follows from the previous paragraphs that evolutionary theories addressing virulence have predicted that virulence of microbes evolves in response to changes in conditions or tradeoffs. One of the most commonly considered “tradeoffs” is between the benefit of a high within-host microbial density (allowing efficient transmission to the next host) and the cost of a reduced longevity of the infection in the host due to high parasite density that tends to kill the host. Other approaches have been proposed such as the role of how pathogenic mechanisms affect virulence evolution. According to this model, pathogenic mechanisms that manipulate host immunity or escape from the host immune response (longer survival in the host) dominate as causes of virulence compared to mechanisms that incrementally alter transmission (higher microbial density and increased mortality).24,25

Furthermore, natural selection would act more strongly on mutations influencing clearance than on mutations influencing transmission.26 Another cited model to explain dynamics of virulence evolution is the so-called “source–sink” model in which evolution of bacterial pathogens is evaluated from ecological points of view that microbes switch from permanent (source) and transient (sink) habitats.27 The generation times of mammalian hosts are much longer than those of microorganisms. Therefore, genetic mutations in these hosts, on which natural selection acts, take longer to become part of a large population. Nevertheless, there is evidence that specific microorganisms can exert selective pressure on the gene pool of human hosts. The evidence is strongest for the potentially lethal infections caused by falciparum malaria. In regions of the world where falciparum malaria is endemic, including Africa, there is a high prevalence of genetic mutations that alter hemoglobin structure or synthesis, decreasing or abolishing falciparum malaria parasites’ survival.28 The selective pressure of malaria on human gene expression is not confined solely to affecting erythrocytes but also likely involves the immune system, cytokines, and other systems.29

Other Modes of Altering Virulence and Pathogenicity Although the selective pressures of evolution generally exert changes over a multitude of centuries, there are other mechanisms that may more rapidly alter microbial pathogenicity, virulence, and drug susceptibility. The expression of mutated genes in microorganisms is heightened when there are greater numbers of organisms and their generation times are brief. Hence, altered gene expression in helminths will be slow to be expressed, whereas in microparasites genetic alterations will be likely to develop. For mycobacterial infections, large numbers of bacilli that persist for a long time facilitate the genetic emergence of drug resistance to a single agent, and this likelihood underlies the principle of using more than a single drug to treat tuberculosis. Even more rapidly dividing microparasites can develop genetic alterations, and this is especially true when the fidelity of genetic replication is poor. This is prominent in human immunodeficiency virus type 1 (HIV-1), whose reverse transcriptase lacks a 3′ exonuclease proofreading activity.30 Alterations in cell tropism, pathogenicity, and drug sensitivity are frequent in HIV-1 infections. Again, several antiviral agents must be employed concomitantly to circumvent the highly frequent mutations that alter drug susceptibility in HIV-1 strains. In addition to their own genetic material, many classes of microparasites either contain or are capable of acquiring transferable genetic elements in the form of plasmids, transposons, or bacteriophages. These transferable genetic elements also provide a means for the spread of resistance to antibacterial drugs, an increasing problem in all regions of the world.31

Causes of Acute or Chronic Infections in Individuals One obvious impact of an infectious disease is on the individual infected. Hence, in any region of the world independent of other infectious diseases or malnutrition, the acute infection will cause morbidity and potential mortality in the infected human host. Among otherwise healthy people, the immediate impact of the infection is the symptomatic acute illness. For some infections that have prolonged courses, their impact may also continue over many years. Chronic infections include most of those caused by helminthic parasites, which characteristically live for years; persisting mycobacterial infections; and retroviral infections (HIV-1, HIV-2, and human T-cell lymphotropic virus type 1). Finally, the sequelae of some infections can include the development of neoplasms. Examples include hepatocellular carcinomas associated with chronic hepatitis B and C viral infections, bladder tumors with urinary schistosomiasis, cholangiocarcinomas with biliary fluke infections, and gastric adenocarcinomas and lymphomas associated with Helicobacter pylori infections.

Polyparasitism and Effects on Nutrition and Growth In an otherwise healthy and fully nourished person, a new infection is likely to be the only active infection in that person. In contrast, in regions where enteric and other infections are highly prevalent because of inadequate sanitation and poor socioeconomic conditions, adults and especially children may harbor several infections or be subject to repeated episodes of new enteric pathogens. Thus, the polyparasitism of multiple concurrent or recurrent infections adds a new dimension to the impact of acute infections, not often encountered in developed countries. Moreover, the subclinical impact of a number of tropical infectious diseases is beginning to become apparent. Increasing data suggest that even “asymptomatic” giardial,38 cryptosporidial,39 and enteroaggregative Escherichia coli40 infections may be very important in predisposing to malnutrition, thus reflecting a clinically important impact, even in the absence of overt clinical disease such as diarrhea. Likewise, chronic intestinal helminth infections also have a major impact on nutrition in those with already marginal nutrition. Anthelminthic therapy in these children, who lack symptomatic infections, has led to increases in growth, exercise tolerance, and scholastic performance.41,42

Chapter 1

Infectious diseases may affect not only individuals but also large groups of people or entire populations due to epidemic or highly endemic transmission. Throughout human history, a few microorganisms have been responsible for great epidemics and massive numbers of dead or crippled people as a result of infections spreading locally or throughout the world.32–35 Typhus has been associated almost always with situations that involve overcrowding, famine, war, natural disasters, and poverty. The outcomes of several European wars were affected by the morbidity and mortality inflicted by typhus or other diseases on the military. Typhus epidemics were common during the world wars of the twentieth century and in the concentration camps where the ecological conditions were ideal for such a disease to spread.30 Today, typhus and other rickettsioses are still public health problems in some countries, but overall the disease was brought under control after its life cycle was described and antibiotics, insecticides, and public health measures became available.34 Bubonic plague, caused by Yersinia pestis, is another disease that has shaped history, especially in Europe during the Middle Ages.35 Millions of people were affected by pandemics that spread throughout the con­ tinent. Tuberculosis, smallpox, and measles had a tremendous effect on the native populations of the Americas after Columbus’s voyages to the New World. It has been estimated that 90% of the population in Mexico was killed by these pathogens, which were novel to the native residents. Acquired immunodeficiency syndrome (AIDS) represents the modern pandemic that will continue to affect human history for at least decades. Other examples are cholera and influenza, which are capable of causing pandemics.36 In addition to widespread diseases caused by epidemic spread of infections, some infectious diseases, because of their highly endemic prevalence in populations, continue to affect large segments of the world’s population. These include enteric and respiratory infections, measles, malaria, tuberculosis and schistosomiasis. Furthermore, even the staggering mortality and morbidity of these tropical infectious diseases do not control populations but are associated with population overgrowth. This is true not only across the different countries of the world but also throughout the history of developed countries. Thus, the impact of these infections is not solely on the individual but, because of their highly endemic or epidemic occurrence, on populations. This has consequences on economic, political, and social functioning of entire societies.37

of infections with diverse microbes. The human immune system is composed of multiple elements, including those of innate immunity and those of adaptive immunity. Many of the elements of innate immunity are more primitive and found in invertebrate organisms, whereas the adaptive immune responses have evolved further in vertebrate hosts. Microorganisms that successfully infect human hosts must, at least in the short term, overcome elements of the host immune system, which then may react further to attempt to control these infections. The study of microbial pathogenesis has been revolutionized with the advent of comparative genomics and the multitude of genomic tools that have become available in the last two decades or so. The first bacterium whose genome was fully sequenced was that of a laboratory strain of Haemophilus influenzae in 1995, followed by Mycoplasma genitalium the same year. Since then over 250 bacterial genome sequences have become available. Discovery of virulence genes has therefore expanded rapidly and has benefited from other strategies such as powerful computational methods, genetic signatures, analysis of physical linkage to accessory genetic elements, and biochemical and genetic approaches that depend on comprehensive genome sequence information.43 Furthermore, proteomics-based methods have also been used in combination with genome-sequence analysis to define new virulence factors.43 For obligate intracellular pathogens, whose genetic manipulation is far more difficult than for other microbes, the use of genomic and post-genomic tools (genomic-microarray methods and proteomics) has been virtually the only path to discovery of virulence factors. In general, these tools have advanced the understanding of virulence factors in pathogenic micro­ organisms exponentially. Microorganisms that infect humans are exogenous to the host and must colonize or penetrate epithelial barriers to gain access to the host. Except for infections acquired during the intrauterine period, infectious agents must bridge host epithelial surfaces, the keratinized epithelium of the skin, or the mucosal epithelium of the respiratory, gastrointestinal, or genitourinary tracts. Ultimately, there are four types of microbial localization in the host (Fig. 1.1). Some microbes will enter intracellular sites either within the cytoplasm or within vesicular or vacuolar compartments in cells. Other microbes remain extracellular, either at epithelial surfaces or within the host in the blood, lymph, or tissues.

Principles of Parasitism: Host–Parasite Interactions

Causes of Widespread Infections in Populations

Interactions at Epithelial Barrier Surfaces The barrier functions occurring at epithelial surfaces are part of the innate host defenses and are important in determining the outcome of interactions of potential pathogens with the host. Interactions at epithelial barriers involved in defense against external microbes include not only the physical properties of the epithelial surfaces but also the overlying mucous phase, the ciliated or other propulsive activities facilitating microbe clearance, and the normal microbial flora.

Normal Flora Vertebrate warm-blooded organisms, such as humans, are an ideal site for the survival of many microbes and provide a rich source of organic material and a constant temperature and pH. Microbes coexist with us in and on our bodies, especially on epithelial surfaces where there is contact with the outside world, such as the bowel, upper respiratory tract,

Intracellular

MICROBIAL INTERACTIONS WITH HUMAN HOSTS

Locale: Cytoplasm

Just as microorganisms have evolved over centuries or longer, mammalian hosts have evolved to contain and limit the deleterious consequences

Figure 1.1  Microbial localization.

Vesicles, vacuoles

Extracellular

Blood, lymph, tissues

Epithelial surfaces

3

PRINCIPLES AND GENERAL CONSIDERATIONS

SECTION I

mouth, skin, and distal portions of the genitourinary tract.1,2,43 Most of these microorganisms are highly adapted to live with us and do not cause any harm. The presence of the same type of microorganisms at a particular site in the absence of disease is called colonization. Normal colonizing microbial flora help to limit access by potentially pathogenic micro­ organisms. One condition predisposing to infection is the alteration of the normal epithelial flora, as occurs with antibiotic therapy, since this may allow for the proliferation of pathogenic organisms normally held in balance by the endogenous normal microbial flora. Examples include Candida vaginitis or the development of pseudomembranous colitis due to toxigenic Clostridium difficile, which may complicate antibiotic therapy.

Adhesion to the Epithelium

  

 

Microorganisms maintain themselves in or on their host by adhesion to cells or the extracellular matrix. Adhesins are encoded by chromosomal genes, plasmids, or phages.44 They are usually divided into fimbrial and afimbrial adhesins.45 Fimbrial adhesins are present in organisms such as Neisseria gonorrhoeae and are in part responsible for the attachment to genitourinary tract epithelium, preventing the bacteria from being washed out by the urine stream.46,47 An example of an afimbrial adhesin is the filamentous hemagglutinin of Bordetella pertussis, which is responsible for the attachment of B. pertussis to epithelial cells in the respiratory tract.48 Adhesins attach to receptors in the host. These receptors include proteins, glycolipids, and carbohydrates exposed on the surface of cells or in the extracellular matrix.44 Integrins are one class of proteins present on eukaryotic cell surfaces that can serve as bacterial receptors.44 Helicobacter pylori binds to Lewis blood group antigen present in the gastric epithelium.49 Neisseria has a ligand that binds to CD66 molecules on epithelial cells. Some pathogens have even more evolved interactions with the host and activate signal transduction mechanisms in the host cell, which in turn upregulate other molecules that aid in the adhesion process.2,44 Certain strains of enteropathogenic E. coli possess type III secretion or contactmediated systems.50 In such cases, the secretion and synthesis of virulence factors is modulated by contact with host surfaces. The systems are complex (more than 20 genes are involved) and have not been elucidated completely at the molecular level.51,52

Penetration of the Epithelial Barriers

4

Some microbes do not have the means to penetrate skin barriers and are only able to gain access through bites produced by arthropods (e.g., rickettsiae, arboviruses, plasmodia, and filariae).53,54 In such cases, microbes may be introduced by direct inoculation (e.g., rickettsiae, arboviruses, and plasmodia) or may gain access by migrating through the puncture site (filariae). Other microbes (e.g., skin bacteria and fungi) depend on mechanical disruption of the skin (e.g., due to burns, trauma, or intravenous catheters) to invade deeper structures.55 Still others invade when defenses on mucosal surfaces are lowered due to combined local or generalized immunosuppression and altered mucosal integrity (mucositis) due to chemotherapy or malnutrition (e.g., Candida spp. and anaerobic and other enteric bacteria in the bowel). Some microbes do not invade tissues at all and affect the host locally and systemically by liberating toxins at the site of colonization (e.g., diphtheria exotoxin).44 For enteric pathogens, some, including poliovirus, Salmonella enterica serovar Typhimurium, Salmonella enterica serovar Typhi, Campylobacter jejuni, Yersinia enterocolitica, and Yersinia pseudotuberculosis, gain access to the host across the intestinal epithelium by utilizing uptake in specialized epithelial M cells.56 Internalization of some microorganisms is also achieved through other mechanisms, such as sequential “zipper-like” encircling of the organisms triggered by bacterial ligands and cellular receptors, as occurs in infections caused by Listeria monocytogenes.44 The trigger mechanism of the bacteria induces massive rearrangements of cytoskeletal proteins such as actin, which results in membrane ruffles, as occur with shigellosis and salmonellosis.44

In the genitourinary tract, invasion of some agents (e.g., HIV-1) is facilitated by mucosal erosions caused by other infectious agents.57

Spread from the Portal of Entry Once the organisms gain access to the body after overcoming the first lines of defense, they either spread to other sites of the body or reproduce locally and often invade surrounding tissues. Local spread is facilitated by a number of factors, including collagenases, hyaluronidases, fibrin­ olysis, and other enzymes. They are produced by a wide range of organisms, and the role of these enzymes in invasion is, in some cases, controversial.2 Lymphatic spread occurs in most cases once the organisms gain access to subepithelial tissues or serosal surfaces. Lymphatic vessels are distributed in most tissues of the body, with few exceptions such as the brain. Lymph is carried by lymphatic vessels to regional lymph nodes, where it circulates through the node and eventually returns to the systemic circulation through the thoracic duct and the great lymphatic vein. One to three liters of lymph is returned to the systemic circulation every day. Most pathogens are filtered in lymph nodes before reaching the systemic circulation, but some actually reproduce either in the endothelium of lymphatic vessels (e.g., Mycobacterium leprae)2,58 or in tissue macrophages present in the lymph nodes (e.g., Brucella spp.) or lymphocytes (HIV and herpesviruses, including Epstein–Barr virus).59 Some organisms reach the systemic circulation after overwhelming the defenses in the lymph nodes (e.g., Bacillus anthracis and Y. pestis). Microorganisms carried in the blood are transported either extracellularly (e.g., most of those causing bacteremia) or intracellularly. Intracellular pathogens are carried by red blood cells (e.g., Plasmodium, Babesia, Colorado tick fever virus, and Bartonella), monocytes (e.g., measles virus, cytomegalovirus, and Toxoplasma), or neutrophils (e.g., Anaplasma phagocytophilum, Ehrlichia ewingii, and some pyogenic bacteria).2,60 Once in the blood, by initial lymphatic or hematogenous spread, the microorganisms have access to virtually any site in the body. However, some pathogens exhibit tropism for certain tissues. This tropism depends on multiple factors, including the anatomy of the microcirculation in a given tissue (fenestrated capillaries versus continuous endothelial lining), receptors present on certain endothelial cells, and the presence of mononuclear phagocytic cells in organs such as bone marrow, liver, and spleen.2 Other less common routes of spread include peripheral nerves (e.g., rabies and varicella zoster virus), cerebrospinal fluid (after the organisms traverse the blood–brain barrier), and serosal cavities.

Localization in the Host Microbes that have gained access to the host at or through epithelial barriers then, depending on the properties and size of the pathogens, have the capacity either to seek intracellular sites or remain extracellular (see Fig. 1.1). Mechanisms of host immune responses to the microorganisms vary depending on their sites of localization.

Intracellular Localization Specific microorganisms use highly developed processes to gain access to and survive within host cells. The microorganisms may reside either in the cytoplasm or within vesicular or vacuolar compartments of targeted cells. Targeting and penetration of cells is governed by the interactions of microbial surface proteins that may engage host cell molecules that function as receptors for the microbial ligands. The entry of malarial parasites into erythrocytes is a good example, and the nature of the erythrocyte receptors used by different malarial parasite species governs which red blood cells are infected. Plasmodium vivax binds to the Duffy blood group antigens present on some people’s red blood cell membranes. The expression of the Duffy blood group antigen is genetically determined, and this antigen is present mostly in whites and Asians and largely absent

Chapter 1

Immune responses against microbes within macrophages rely heavily on class II MHC-mediated presentation of host antigenic peptides to T-helper 1 (Th1) types of CD4+ T cells, which then augment the microbicidal activities of the macrophages.

Extracellular Localization Some types of microbes that remain extracellular typically reside at epithelial surfaces, including bacteria such as N. gonorrhoeae, H. pylori, Vibrio cholerae, and E. coli, and helminths such as adult Ascaris lumbricoides, hookworms, and Trichuris trichiura. Mucosal immune responses, including IgA and leukocytes, participate in host immune reactions to these pathogens. Other microbes that survive extracellularly are present within the blood, lymph, or tissues of the host, and these organisms include fungi, viruses, bacteria, protozoa, and notably the helminths. Multicellular helminths, due to their large size, remain forever extracellular and may be found in the blood (e.g., microfilariae), lymph (adult lymphatic filarial worms), tissues (migrating larvae and adult stages of some helminths), and cerebrospinal fluid. Host defense against extracellular pathogens uses antibodies, complement, phagocytic cells, and, for helminths, IgE, eosinophils, and mast cells.85

Principles of Parasitism: Host–Parasite Interactions

in blacks of sub-Saharan African ancestry.61–64 This genetic absence of a receptor on red blood cells required for vivax malaria’s survival explains why vivax malaria is rare in regions of Africa. Plasmodium vivax also exhibits a characteristic restriction in the age of erythrocytes it infects. Only young red blood cells and reticulocytes are susceptible to infection, even though the Duffy blood group antigen is present on red blood cells of all ages. The basis for this restriction to younger red blood cells also rests with receptor-mediated limitations. Plasmodium vivax parasites contain reticulocyte-binding proteins, which recognize and bind to reticulocytespecific antigens on the red blood cell surface.65,66 Thus, host cell receptor–microbial ligand interactions have an impact on the geographic range of infections based on host genetic differences in requisite receptor expression and on the specific cells that a microbe may enter. Another example of the intricacies of microbe–receptor interactions has been recognized with HIV-1. Although CD4 is the primary cellular receptor for HIV entry, binding to CD4 alone is not sufficient for entry of HIV-1 into cells. Cellular coreceptors that are members of the chemokine receptor family of seven-transmembrane G protein-coupled molecules are also important. T-cell tropic strains use the CXCR-4 chemokine receptor, and macrophage tropic HIV-1 strains use the CCR-3 and CCR-5 chemokine receptors as coreceptors in concert with CD4. The differences among strains of HIV-1 in their capacities to bind to different chemokine receptor–coreceptors may help explain differences in cell tropism and pathogenicity, the lack of infectability of nonprimate cells, and, for those with genetically altered coreceptors, the apparent resistance to HIV-1 infection of some individuals.67–70 Typical of those etiologic agents that have an intracellular localization are viruses. The entry of these agents into cells is increasingly recognized to be dependent on their interactions with specific host cell proteins that act as their “receptors.” For instance, host cell molecules that function as viral receptors include multiple isoforms of membrane cofactor protein (CD46), a complement regulatory protein, for measles; the integrin, intracellular adhesion molecule-1 (ICAM-1), for rhinovirus; erythrocyte P antigen for parvovirus B19; and the C3d complement receptor (CR2) for Epstein–Barr virus.71–74 Microbes that exist principally within the cytoplasm are sequestered from many immune response mechanisms active on extracellular pathogens, including antibody and phagocytic cells. Viral intracellular proteins will be processed and displayed with class I major histocompatibility complex (MHC) proteins, which enable CD8 cytotoxic T cells to recognize and kill the virally infected cell. Other microbes are internalized within phagocytic cells, especially macrophages. Once internalized in host cells, organisms such as Salmonella, Mycobacterium, Chlamydia, and Legionella use an extraordinary assortment of mechanisms to prevent their phagocytic vacuole from fusing with the host cells’ acidifying lysosomes.75–77 For some parasites, the intracellular environment is an important determinant of parasitism. For example, Leishmania and Coxiella (unlike other pathogens) benefit from the acidic environment of the macrophage phagolysosome. Leishmania use the proton gradient across the lysosome to drive the energy-dependent uptake of two important substrates: glucose and proline.78 Thus, Leishmania amastigotes actually survive in the macrophage phagolysosome because they benefit from its proton gradient and because they avoid activating the processes that normally kill ingested microorganisms. Leishmanial lipophosphoglycan inhibits the action of β-galactosidase, chelates calcium, inhibits protein kinase C and the oxidative burst, and may scavenge toxic oxygen metabolites.79 Conversely, other intracellular pathogens such as Toxoplasma gondii survive within the macrophage by using an alternative pathway of entry that avoids fusion of the parasitophorous vacuole with lysosomes.76,80 In contrast, dead or antibody-coated T. gondii enter via the Fc receptor and are routed to a different intracellular compartment, which fuses with the lysosome, and are then killed in the phagolysosome.76,80,81 Other organisms, such as Shigella, Listeria, and Rickettsia, breach their vacuolar membrane to multiply freely in the cytoplasm and may also usurp host cellular actin to propel their further spread to neighboring cells, continuing to exploit their intracellular sanctuary.82–84

Tissue Damage There are multiple mechanisms by which microbes inflict damage on host tissues.

Direct Damage or Alteration of Host Cell Function Host cells can be killed directly by the infectious agent, as in some viral or bacterial infections that are highly cytopathic (e.g., yellow fever virus in hepatocytes and Salmonella in macrophages).86,87 Some microorganisms multiply intracellularly until the cell bursts and dies (e.g., Rickettsia prowazekii).34 Some bacteria, viruses, and other parasites, such as Shigella, HIV-1, and Listeria, can induce apoptosis of host cells.59,88,89 Apoptosis is triggered by different mechanisms, such as activation of the interleukinconverting enzyme pathway.90,91 This form of programmed cell death is probably more widespread as a mechanism of cell death in infectious diseases than previously thought. Damage is sometimes caused by toxins secreted by bacterial cells (exotoxins). In this case, bacteria can either invade host tissues or colonize mucosal sites and then release toxins at the mucosal site that are absorbed systemically and cause distant damage.92 Exotoxins can act through different pathways that damage the components of the cell membranes such as phospholipids88,93 or affect signaling pathways (e.g., V. cholerae).44,94 Other exotoxins, such as streptolysins and listeriolysins, alter membrane permeability. Still others, such as exfoliatin (e.g., Staphylococcus aureus) and elastase (e.g., Pseudomonas spp.), are capable of degrading extracellular elements.2 Some toxins are translocated to the intracellular environment, where they affect multiple enzymatic systems. These toxins are classified according to their enzymatic activity, such as adenosine diphosphate (ADP) ribosyl transferase (e.g., diphtheria toxin, P. aeruginosa exotoxin A, and pertussis toxin), depurinase (e.g., Shiga toxin), adenylate cyclase (e.g., pertussis hemolysin and anthrax edema factor), and zinc protease (e.g., tetanus).94 The end result ranges from blockade of protein synthesis and cell death or blockade of exocytosis (especially CNS neurotransmitters at the synaptic cleft)95,96 to increases of cyclic adenosine monophosphate (AMP) or cyclic guanosine monophosphate (GMP) and changes in cell permeability.94 Still other organisms, such as C. difficile, produce toxins that change basic cell signaling transducers such as Rho to alter cell function or affect their spread. Finally, organisms can interact with host cell or microbial transcriptional regulation of genes (such as iron-binding proteins for uropathogenic E. coli92,97) or cytokine release (such as H. pylori or enteroaggregative E. coli40,98–100) to enhance their survival or elicit pathogenic responses. The evolutionary advantages to a microbe of its remarkable array of traits that we call “virulence” hold many of the clues to their control, if we can but truly understand them.

5

SECTION I

Endotoxins are a subset of lipopolysaccharides present in the outer membrane of Gram-negative bacteria that can trigger a wide variety of responses in the host, including massive cytokine release leading to hypotension and shock.101,102 These deleterious effects occur with high-grade invasion of the blood by Gram-negative bacteria, including enteric Gramnegative bacteremias and meningococcemia.

Table 1.1  Mechanisms of Immune Evasion Used by Pathogenic Microbes Type of Immune Response

Immune Evasion Mechanisms

Innate immunity

Block natural killer cells Recognition avoidance Complement inactivation or blockage Avoidance of phagocytosis by macrophages and polymorphonuclear neutrophils Manipulation of cell surface Modulation of inflammatory response Interference with signaling pathways Interference with RNAi Antimicrobial peptide degradation Modulation of endosomal trafficking Modulation of cytoskeleton Induce apoptosis Interfere with receptors and signaling Modulation of antigen presentation and processing Modulation of cell maturation Superantigens Antigenic variation Phase shift Escape mutants

PRINCIPLES AND GENERAL CONSIDERATIONS

Indirect Damage

  

 

Damage to the host may also develop as a consequence of immune reactions to the infectious agents. One scheme for classifying immunopathologic responses divides the reactions into four types based on the elements of the immune response involved.103 Type I reactions involve elements of strong Th2 responses that lead to increased IgE, eosinophilia, and eosinophil and mast cell activation. Adverse reactions of this type include the development of urticaria (with several helminthic parasites), the occurrence of potentially life-threatening anaphylactic shock in IgE-mediated mast cell degranulation (e.g., triggered by systemic release of antigens from echinococcal cysts104), and exuberant eosinophilic infiltration of tissues due to migrating helminth larvae (e.g., Löffler’s pneumonia with the pulmonary migration of Ascaris larvae). Type II reactions are also dependent on elements of Th2 cell responses that lead to increased IgM and then IgG antibodies directed toward the infectious agents. These antibodies, if cross-reactive with host antigens, may lead to complement-mediated cytotoxicity or antibody-dependent cell-mediated cytotoxicity by natural killer cells, which have Fc receptors. An example of this type of immunopathologic response is the uncommon hemolytic anemia associated with Mycoplasma pneumoniae infection that is mediated by complement-induced hemolysis triggered by IgM (cold agglutinin) antibodies against erythrocyte I antigen. Type III reactions are caused by the deposition of immune complexes. When neither antibody nor antigen is present in excess of the other, the complexing of antibodies with soluble antigen results in the formation of immune complexes that may cause disease. This may develop acutely as antibody titers rise in the presence of microbial antigens, causing the syndrome of serum sickness. In addition, when soluble antigen is persistently abundant, sustained formation of immune complexes develops, leading to chronic immune complex-mediated tissue damage (especially glomerulonephritis), as found in subacute bacterial endocarditis, chronic hepatitis B antigenemia, and chronic Plasmodium malariae infections.105 Type IV reactions include adverse reactions mediated by macrophages and cytotoxic T cells. Examples are damage caused by granulomas in leprosy, tuberculosis, tertiary syphilis, and fungal infections. Likewise, granulomas developing around schistosomal eggs, depending on their location, may cause ureteral obstruction or hepatic presinusoidal lesions. Other deleterious inflammatory reactions in this category are mediated by parasite-elicited host cytokines, such as the hepatic fibrosis elicited by schistosomal eggs.

IMMUNE INTERACTIONS Immune Evasion

6

The human immune system has evolved in concert with microbes and is very sophisticated, especially with regard to host defenses against microbes, but the system is not perfect. Interactions of the immune system with microbes are an ongoing affair. Microbes have a high mutation rate compared to human beings. Microbes have evolved a diversity of mechanisms that can enable microorganisms to subvert immediate immunologically mediated elimination (Table 1.1). Persistence within the host is necessary for the propagation of some parasites. There are multiple mechanisms by which microbes can persist in the body and evade the immune system. Tolerance is defined as specific reduction in the response of the immune system to a given antigen.106,107 In the case of transplacental infection, the fetus develops a certain degree of tolerance to antigens to which it is exposed. The immune system of

Early induced response

Adaptive immunity

Protective immunity

fetuses is rather incompletely developed in utero, and microorganisms survive easily. Cytomegalovirus infects the fetus transplacentally and produces extensive damage to multiple tissues. After delivery, infants continue shedding virions for weeks to months because they are unable to destroy the virus. Other mechanisms include the production of superantigens that stimulate a large population of T cells, which then become deleted if the encounter occurs during early development. Exposure to massive amounts of antigen in the circulation can also lead to tolerance.2,103 Immunosuppression is a well-demonstrated phenomenon that occurs during certain infections caused by viruses, bacteria, protozoa, and helminths. These infections usually involve the lymphoid tissues and macrophages and hamper the immune response. Intracellular pathogens that are able to spread from cell to cell without exposure to the extracellular compartment can avoid exposure to some elements of the immune system. In other cases, pathogens reside in sites relatively inaccessible to the immune system, such as glandular luminal spaces or kidney tubules. In many infections, antibodies are produced but do not effect microbial killing. Sometimes, antibody avidity is low, the epitopes against which the antibody is directed are not critical to the microorganism’s survival, or the mechanism of immune elimination is not antibody dependent.2 Other microorganisms have developed means of counteracting specific elements of immune responses, such as production of an IgA-degrading enzyme, IgAase, by certain strains of N. gonorrhoeae.108 Some strains of amebae also produce proteases that destroy complement.2 Reactivation of infections in old age due to waning immunity has been well demonstrated in cases of tuberculosis and varicella zoster virus, allowing transmission to new hosts. One well-studied mechanism of immune evasion is the capability of changing the antigenic structure by genetic mutation or by pro­ grammed sequential expression of genes encoding different surface antigens.109Antigenic drift and recombination between influenzaviruses affecting humans and animals are well documented. Borrelia recurrentis and Trypanosoma gambiense are also capable of changing their surface antigens after antibodies control the initial bloodstream infection.110,111 The new antigens are not recognized by the antibodies, allowing relapse of the infection. Parasites in which sexual reproduction is possible benefit

EMERGING INFECTIOUS DISEASES The concept of emerging infectious diseases is not new but has been the focus of attention due to the resurgence of old infectious diseases that were thought to be controlled and the recognition of new pathogens as humans increase their interaction with the biosphere. By definition, an emerging infectious disease is one that has newly appeared in the population or has existed but is rapidly increasing in incidence or geographic range.114 Others define emerging infections as “new emerging or drug resistant infections whose incidence in humans has increased within the last two decades, or whose incidence threatens to increase in the near

Chapter 1

future.” Emerging infections are classified by some as newly emerging, re-emerging/resurging, and deliberately emerging. Since 1967, after a widely publicized statement from the US Surgeon General declaring victory in the war against infectious diseases, more than 85 new pathogens have been described. These include viruses, bacteria, protozoans, and helminths. These pathogens can cause a wide diversity of syndromes including acute respiratory infections (influenza A H1N5, H1N1, SARSCoV, Sin Nombre virus, human metapneumovirus), systemic diseases caused by viral hemorrhagic fever viruses (Lassa, Ebola, dengue), encephalitic syndromes (Nipah virus, West Nile virus), arthropod-borne agents (Borrelia burgdorferi, Rickettsia spp., Ehrlichia spp., Anaplasma), enteric pathogens (Cryptosporidium, microsporidia), chronic viral diseases (HIV-1 and 2, human T-cell lymphotropic virus types 1–3, human herpesviruses 6–8), hepatotropic viruses (hepatitis C and E), and other infectious agents. The factors involved in the emergence or reemergence of infectious diseases are complex and include ecological changes (deforestation, reforestation, flooding, and climatic changes), changes in human demographics and behavior (sexual, cultural, and war), increased international travel, technological advances (organ transplantation and antibiotics), microbial evolution with the appearance of antibiotic-resistant or antigenically distinct strains, and deficiencies in surveillance and public health policy.113,115–117 The classic triad of microbe, host, and environment is again exemplified.

Principles of Parasitism: Host–Parasite Interactions

enormously.112 Genetic variability introduced by crossing over during meiotic divisions is much greater than the variability introduced by asexual reproduction. As many as four crossovers on a single pair of chromosomes have been demonstrated in P. falciparum.113 Microparasites also have multiple mechanisms by which they can evade the initial line of defense provided by phagocytes. These strategies include killing of the phagocyte (e.g., Streptococcus pyogenes and Entamoeba histolytica), inhibition of chemotaxis (e.g., Clostridium perfringens), decreased internalization of microbes by phagocytic cells (e.g., T. gondii), inhibition of opsonins (e.g., Treponema pallidum), inhibition of phagolysosome fusion (e.g., M. leprae and Mycobacterium tuberculosis), and escape from the phagosome into the cytoplasm (e.g., Rickettsia spp., Trypanosoma cruzi, and Listeria).2,44,75,92 With cell-to-cell spread, microorganisms may be minimally exposed to complement, antibodies, or phagocytes in the extracellular or intravascular spaces.77,78 Rickettsial infections spread from cell to cell throughout the infected foci in the endothelial layer of the microvasculature.82,83,94 Macroparasites, the helminths, have evolved diverse mechanisms that enable them to survive in vivo.85 Characteristically, helminths live for months to years in infected hosts within the lumen of the bowel, within tissues, or in the blood or lymphatic vessels. Many helminths are in intimate and recurring contact with all elements of the immune system. As a consequence of their size, helminthic worms do not use intracellular mechanisms to evade immune responses but have evolved a number of capabilities that permit their survival. For instance, interference with antigen processing has been well documented in animal models and patients infected with the filarial nematodes Brugia malayi and Onchocerca volvulus. These helminths produce a family of proteins called the cystatins that are capable of inhibiting proteases responsible for antigen degradation and subsequent presentation through MHC class II pathways in antigenpresenting cells. These proteins are also capable of modulating T-cell proliferation and elicit upregulation of interleukin-10 (IL-10) expression. Other modulators include helminthic derivatives of arachidonic acid such as lipoxin A4, which is capable of blocking production of IL-12 in dendritic cells. Helminthic prostaglandins can also inhibit IL-12 production by dendritic cells. Since helminths have very complex genomes (~21 000 protein-encoding genes in some of them), they are capable of producing a large variety of proteins. Some of them are cytokines and related proteins also capable of modulating the host immune response to their advantage. For example, B. malayi has been shown to express transforming growth factor (TGF)-β-like proteins capable of binding TGF-β human receptors. Other cytokines include macrophage-migration inhibition factors produced by several nematodes including B. malayi. Blockade of effector mechanisms has also been demonstrated in some helminth infections, including proteases that target effector molecules such as eotaxin. Neutrophil proteases can also be inhibited by serpins.

TROPICAL INFECTIOUS DISEASES Globally, as assessed in terms of disability-adjusted life years (DALYs), which measure morbidity and mortality,118 infectious diseases in 1990 accounted for 36.4% of total DALYs. Infectious disease DALYs were considerably in excess of those attributable to cancer (5.9%), heart disease (3.1%), cerebrovascular disease (3.2%), or chronic lung disease (3.5%).119 However, these calculations admittedly miss the disproportionate impact of tropical infectious diseases on the still rapidly increasing populations living in impoverished, tropical areas, and they grossly underestimate the major developmental impact of common childhood enteric, helminthic, and other infections.38,120–122 For those caring for individual patients with infectious diseases, appropriate diagnosis and treatment are important considerations for the individual. Even more important is the consideration of approaches that will lead to diminished acquisition of infectious diseases. For some infectious agents, immunization holds promise, as witnessed by the successful global eradication of smallpox and the potential eradication of poliomyelitis. Greater progress in the control of infectious diseases, however, rests with improvements related to socioeconomic conditions of the population at risk. In developed countries, tuberculosis was diminished well before the introduction of the first antimicrobial agents active against M. tuberculosis, and the reduction was attributable to improved socioeconomic conditions. For the major infectious diseases of the tropics, improvements in sanitation, living conditions, and general public health will be critical in helping control the impact of the diverse infectious agents that currently contribute to human morbidity and mortality. The impact of these infections is related not only to their effect on the health of the infected individual but also to their contribution to the morbidity associated with malnutrition and to their larger societal impact as an impediment to the full development of the political, economic, and social potential of entire populations.

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7

SECTION I: PRINCIPLES AND GENERAL CONSIDERATIONS

CHAPTER 2 

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The Perfect Storm: Climate, Ecosystems and Infectious Diseases Fernando J. Torres-Vélez • Corrie Brown

8

The studies of emerging diseases and environmental changes are linked together in a manner reminiscent of the old adage about the forest and the trees. Each new disease that occurs is studied in significant depth, that is, with an intense focus, as on a single tree. However, it is often the overall environmental changes, or the proverbial forest, that is the greatest influence on the actual emergence of that disease. In recent decades the world has experienced a plethora of new and reemerging pathogens – human immunodeficiency virus, Hantavirus, Nipah virus (NiV), severe acute respiratory syndrome, Rift Valley fever, and bluetongue among others. A common denominator in the appearance of these diseases has been some modification of the environment, intensification or regional variations of normal atmospheric phenomena such as El Niño-Southern Oscillation (ENSO), as well as more specifically human-induced environmental changes.1 Some of these anthropogenic factors are habitat encroachment by the growing human population, putting humans in closer contact with animals harboring pathogens previously unknown to infect people. Other instances involve impacts that humans have been having on the environment for decades, brewing today’s global climate changes. Over the last decade, the scientific investigations into climate change have expanded logarithmically and now comprehensive and elegant multidisciplinary studies are beginning to illuminate potential disease problems due to climate change. Climate is usually defined in a specific area as the “average weather,” the mean and variation of atmospheric components such as temperature, precipitation, and wind, over time, traditionally 30 years. The overall global climate is powered by a fraction of the total solar radiation reaching the planet. Approximately one-third of all the incoming solar energy never makes it to the Earth’s surface but rather is reflected back to space by atmospheric components. Of the remaining two-thirds absorbed by the Earth, an equal amount must be radiated back to space, to prevent an ever-increasing surface temperature. Clouds and gases, such as water vapor and carbon dioxide (CO2), absorb much of the thermal radiation emitted by land and oceans, retransmitting it to Earth in a desirable greenhouse effect. This natural greenhouse effect allows for temperatures that are compatible with life processes as we know them. Water vapor and CO2 are the most important greenhouse gases and significant increases in either can cause an enhanced greenhouse effect which has an overall effect of climate warming. Oceans and plants serve as natural CO2 buffers by absorbing a great deal of what is generated. However, human activity is offsetting this natural system by burning fossil fuels and clearing forests.2 In addition to the anthropogenic contribution to global warming, external factors such as volcanic eruptions can have a major impact on the global climate. This was clearly illustrated in 1991 after Philippine’s Mount Pinatubo’s eruption when a global cooling and drying of the atmosphere was recorded.3 Also there are poorly understood, coupled air–sea climatic events, such as ENSO, which can alter the climate of vast geographical areas for extended periods of times. During the ENSO, there is a reversal of surface air pressures between the eastern and western

tropical Pacific, bringing a weakening and often reversal of trade winds. This allows for warmer water from the western Pacific to flow toward the east and build up off the coast of South America. The change in trade winds brings rain to the dry eastern Pacific, leaving extensive drought in the western Pacific. Volcanic eruptions or ENSO can be influenced by or combined with anthropogenic-induced global warming to augment further climate changes or contribute to adverse climatic events.4–7 A global authority on climate change was established by the United Nations Environment Programme and the World Meteorological Organization in 1989 in the form of the Intergovernmental Panel on Climate Change (IPCC). The IPCC is composed of scientists from around the world who review and assess scientific and technical data relevant to the understanding of climate change. They are charged with providing the world with a clear scientific view on the current state of climate change and its potential environmental and socioeconomic consequences. This panel has concluded that the observed climate changes cannot be explained by just natural factors. There is agreement that disease incidence and distribution are likely to change, although the manner in which this will happen is complex and difficult to specify.2 Below are three examples of infectious diseases that have emerged as a result of some of the climatic changes described above. The first, bluetongue, and its spread into northern Europe may be the only clear example of disease spread directly related to global warming. The second, Rift Valley fever, has reemerged several times in Eastern Africa in recent years, largely as a result of expanded ENSO impacts. The third, NiV, emerged in Malaysia due to a combination of direct anthropogenic effects and socioeconomic pressures, combined with ENSO effects.

BLUETONGUE Bluetongue virus extension into northern Europe is undoubtedly the best example of a livestock disease expanding its range based on climate change, although the intensive outbreaks experienced in the last 2 years demonstrate the multiplicity of factors that play a role. Bluetongue is an arthropod-borne disease, capable of infecting all ruminants, but with most severe clinical disease in sheep, in which there is severe depression, lameness, and often facial and tongue swelling, and abortion in pregnant animals. In cattle there is usually minimal clinical disease, but these animals can remain viremic for months, and so serve as the main reservoir of infection.8 Bluetongue virus (BTV) is a segmented dsRNA virus in the Orbivirus genome, family Reoviridae. The disease is endemic in much of Africa and over large segments of the Americas, with distribution coinciding with the range of Culicoides spp., the responsible insect vector. The main Asian-African vector is Culicoides imicola. The disease has occurred in North Africa and the Middle East, making sporadic incursions into southern Europe, as Culicoides are easily transported on wind. The “envelope” or range of usual C. imicola distribution includes the southernmost parts of Europe, which are the regions that have been

RIFT VALLEY FEVER Rift Valley fever virus is a segmented, enveloped, single-stranded RNA virus in the genus Phlebovirus, family Bunyaviridae. It has a wide variety of hosts but severe disease is seen most commonly in ruminants, especially sheep and goats, where it can present as an “abortion storm,” death of neonates, and hepatic disease. It is also zoonotic and in humans presents as a febrile, flu-like illness, with some experiencing severe liver pathology or vascular complications. Mortality in humans ranges from 1% to 5% but in some outbreaks has been much higher.13,14 The virus is transmitted transovarially in mosquitoes and, under dry climatic conditions, can survive in Aedes mosquito eggs for years and maybe decades. With excessive rainfall, the mosquito eggs will hatch, and carry virus to susceptible ruminants, which experience a high viremia. Then Culex mosquitoes serve to transport the infection from animal to animal and, occasionally, animal to human. The disease was first described around Lake Naivasha, in the Kenyan portion of Africa’s Rift Valley, in 1930, and remained confined to the larger geographic zone of the Rift Valley for more than 40 years, with sporadic outbreaks occurring, usually associated with heavy rainfall.15 However, since 1977, Rift Valley fever has been recorded outside this zone on multiple occasions, all of which are associated with increases in flooding allowing for extensive mosquito replication. In 1977, Rift Valley fever surfaced in Egypt, and there was an extensive outbreak involving thousands of human and animal cases. Building of the Aswan dam in the years prior to this outbreak was probably an important factor as this hydrological project permitted controlled flooding of agricultural lands and subsequent enhanced mosquito habitats.16 Then, in 1987, there was another outbreak, this time across the continent to West Africa, in Mauritania. In this case, Rift Valley fever occurred subsequent to

Chapter 2

completion of the Diama dam on the Senegal river, another project that created increased standing water for mosquito amplification.17 In both of these cases, Egypt and Mauritania, there were human-induced changes to the environment, in the form of dam-building which created more standing water and therefore greater habitat possibilities for vectors of disease. In 1997, there was a large outbreak in the Horn of Africa, beginning in northern Kenya and eventually moving to affect five different countries in the region, with large losses of domestic ruminants and extensive human infection.18 This outbreak was attributed to excessive rainfall, which was brought about by the ENSO effect.19 Then again in 2006–2007 there was another less extensive outbreak in the Horn of Africa, also attributed to heavy rainfall due to ENSO. However, by the time of this outbreak, climatic prediction models had been developed, and so sufficient warnings of a potential outbreak were distributed 2–6 weeks before the outbreak, dampening the overall impact because of preparedness and mitigation efforts, involving prompt suspicion and documentation of the diagnosis with aborted or ill animals and selected use of the available vaccine, isolation, and mosquito precautions and control.20

The Perfect Storm: Climate, Ecosystems and Infectious Diseases

sporadically affected by bluetongue over the last few decades. Farther north in Europe, there are other Culicoides species known as the Palearctic species, C. pulicaris and C. obsoletus. Although both had been proven to be capable of transmitting bluetongue experimentally, they were not functioning in the natural epidemiology of infection prior to 2008. Over the last several years, there have been documented increases in nighttime and winter temperatures and also changes in moisture conditions.9 Warming trends in Europe created an environment for enhanced replication and survival of all of these Culicoides species, and, in some cases, shortened cold periods allowed for the vectors to persist throughout the winter. The range of C. imicola has expanded in recent years with the expansion shown to be coincident with warming.10 The warming not only improved and expanded the populations of Culicoides, but also enabled enhanced viral replication within the vector. Several strains of BTV have made incursions into Europe since the late 1990s. One of these, BTV-8, came from Africa and moved through Italy and up into Germany and the United Kingdom, affecting areas that had never before experienced bluetongue disease. It is hypothesized that the enhanced C. imicola population in southern Europe helped to fuel an outbreak. With expanding populations of C. imicola and the Palearctic species, there was more extensive overlap of these different species. A “handover” event occurred probably multiple times, in which the new host species picked up the virus from the endemic host. Once the virus was established within the Palearctic species, it was able to move to more northerly climes.11 The increased temperatures in these more northerly areas, especially the shortened winter period, ensured that the Palearctic species would survive through the season during which BTV would have otherwise been expected to die back. As a result, there was sustained transmission of the disease in new areas. This extension of BTV into northern Europe resulted in disease in many countries that had never before experienced the disease. More than 3000 outbreaks of bluetongue were reported to the World Organization for Animal Health in 2008 from northern European countries, and hundreds of thousands of ruminants died or were euthanized to prevent spread.12

NIPAH NiV is a newly described virus belonging in the Henipavirus genus of the Paramyxoviridae family. It emerged in Malaysia in the late 1990s in both humans and swine. In pigs, the disease occurred primarily as a respiratory ailment, with severe coughing, and was initially called “barking pig syndrome.” The virus replicated in the respiratory tract of pigs and presumably was disseminated by aerosol to humans who developed a multisystemic disease, with predominant encephalitis and high case-fatality rate. During the course of the outbreak, there were 105 human deaths, almost all of which were closely associated with pig rearing or slaughter. To control the disease approximately one million pigs were slaughtered. The disease has reappeared, on a much smaller scale, in small clusters in humans in India and Bangladesh, but without any associated swine disease.21–24 Early molecular characterization of NiV revealed its relatedness to Hendra virus (HeV), another novel Paramyxovirus previously discovered in Australia only a few years before.25,26 As HeV was shown to have a reservoir in pteropid bats (large fruit bats called flying foxes), these animals were studied for a possible role in NiV epidemiology. Approximately 3 years after the initial outbreak in Malaysia, NiV was isolated from the urine of these bats as well as from partially eaten fruits consumed by the animals.27 Subsequent detection of NiV antibodies and RNA in bats’ saliva and urine in Thailand, Cambodia, and India further strengthened the hypothesis that flying foxes of the genus Pteropus were the natural reservoir.28–30 With subsequent outbreaks taking place without pigs as the intermediate host, the ecologic and epidemiologic picture of NiV became more intricate and complex than originally thought. What was the role of pigs during its first emergence? What factors promoted infection of pigs during the first emergence? During the 1990s, swine production in Malaysia moved to a more intensive system with high turnover of piglets. In addition, most farms were often combining fruit production with the use of pig waste as fertilizer, resulting in many trees overhanging pigpens. Extensive slash and burn practices in the region and an extended dry season brought up by the ENSO promoted changes in migration patterns of the Pteropus bats to areas of plentiful food availability, the pig/fruit farms.31 It is believed that the combination of these production techniques, land use, and climate changes provided the ideal conditions for NiV’s emergence in swine, from which it was aerosolized easily to humans. A unique set of factors brought two species with historically distinct ecological niches into close contact. Furthermore, the intense production practices allowed for a subtle introduction, circulation, and establishment of the virus into a new amplifying host, the pig. Retrospective serologic studies indicate that the first human cases in Malaysia predated the 1997 outbreak, indicating that the virus may have been circulating at low levels prior to the known emergence. The intensive management (high turnover of piglets) likely

9

Section I

allowed a small amount of virus in select pigs to be widely circulated through transport and extensive sales associated with the rapidly developing intensive swine industry. As more animals became infected, there was increased possibility for transmission to people, and an outbreak occurred in the human population.32 Recent outbreaks in India and Bangladesh have been attributed to direct contact with the bat’s body fluids.24,33

PRINCIPLES AND GENERAL CONSIDERATIONS

CONCLUSION As we continue contributing to global climate changes, modifying and encroaching into new habitats, there will undoubtedly be new instances

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10

of disease emergence and modifications of existing epidemiologic patterns. To pinpoint emerging problems at their source and help to prevent their spread, it will be critical that health professionals become aware of the potential for novel or unexpected infections and to work more closely with veterinary, agricultural, and climate experts to understand better and control these emerging and expanding global health threats.

SECTION I:

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PRINCIPLES AND GENERAL CONSIDERATIONS

CHAPTER 3  Epidemiology and Biostatistics Edward T. Ryan • Megan Murray

Epidemiology is the science of investigating the occurrence, causes, and prevention of disease in human populations. Epidemiologic tools may be used to estimate disease frequency, uncover or confirm associations between risk factors and disease occurrence, and define the impact of preventive and curative measures to combat disease. As one of the primary disciplines of the field of public health, epidemiology is of great importance to human health worldwide, especially in the developing world. A fundamental understanding of epidemiologic principles – including basic terminology; study design and hypothesis testing; and data collection, analysis, and interpretation – is, therefore, necessary to the understanding of biomedical sciences.

OVERVIEW AND TERMINOLOGY Measures of Disease Frequency Prevalence is a measure of the total number of existing cases of a disease or condition in a specific population at a particular time. Prevalence is usually expressed as a fraction or percentage of a population, but it can also be given as the number of cases per 1000, 10 000, or 100 000 people. In contrast to prevalence, which enumerates all cases of a disease, incidence is a measure of the number of new cases of disease occurring over a specified period. Incidence is, therefore, expressed as the number of cases that occurred in a population of a given size per a unit time. Thus, if a cohort of 1000 individuals is followed for 1 year and 100 of these individuals develop a specific disease, the incidence rate of that disease would be 0.1, or 100 per 1000 per year.

Measures of Effect In addition to measuring frequency of occurrence of a disease, epidemiologic studies measure the strength of an association between a specific risk factor and the incidence of a disease or medical condition, or between a specific intervention or treatment and the prevention or resolution of a disease. For instance, the effect of a risk factor on disease frequency can be estimated by comparing the incidence of disease in a group that has been exposed to a specific risk factor to the incidence of disease in a group that has not been exposed.

Infectious Disease Terms There are a number of definitions specific to the epidemiologic study of infectious diseases. A disease that occurs regularly in a population is said to be endemic. When a disease occurs at a frequency higher than is expected, it is said to be epidemic. A localized epidemic may be referred to as an outbreak. Diseases in animals are said to be enzootic or epizootic. After infection, there often follows a period of latency, defined as the duration of time from infection to onset of infectiousness. The period of

time immediately following infection may also be an incubation period, defined as the time from infection to development of symptomatic disease. If the incubation period is longer than the latent period for a specific disease, individuals may infect others prior to the onset of recognizable illness. An attack rate refers to the proportion of a population that develops an infectious disease over a given period. The term secondary attack rate refers to the proportion of exposed individuals who become ill. The secondary attack rate is often measured among the household members of a known index case, since it is relatively easy to count the number of exposed individuals and follow them over time. The basic reproductive number, often expressed as R0, is defined as the expected number of secondary infectious cases generated by an average infectious case in a population in which everyone is susceptible. This quantity determines the potential for an infectious agent to start an outbreak, the extent of transmission in the absence of control measures, and the ability of control measures to reduce spread. R0 can be expressed as a product of the number of contacts each infectious individual has per unit time (k), the probability of transmission per contact between an infectious case and a susceptible person (b), and the mean duration of infectiousness, D: R0 = bkD Although it is a highly simplified summary of a pathogen’s epidemic potential, R0 may be used to predict outcome following an introduction of an infection into a population. If R0 is greater than 1, the number of people infected will grow and an epidemic will take place; if R0 is less than 1, the disease will die out. In real epidemics, it is useful to replace the basic reproductive number with the effective reproductive number, denoted R, which is defined as the actual average number of secondary cases infected by a primary case. R is usually less than R0, since it reflects both the impact of control measures instituted over time and the depletion of a susceptible population as previously infected individuals acquire immunity. Herd immunity results when a vaccine not only protects a vaccinated individual from contracting an infection but also prevents that individual from spreading the infection to others.

STUDY DESIGN Understanding scientific–medical studies requires knowledge of a number of fundamental epidemiologic and statistical concepts, the first of which concerns study design. Studies may be designed to measure disease frequency (incidence and prevalence) or to measure an effect (e.g., how effective is drug A compared with drug B in the treatment of individuals with a certain medical condition). A classic type of study that measures disease frequency is a surveillance study, which tracks the frequency of a disease in a population over time. It is effectively a reporting system. It may be active, in which case people with disease are actively sought, or

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Section I PRINCIPLES AND GENERAL CONSIDERATIONS  

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passive where cases are reported that present to medical attention. It may be hospital- or clinic-based or community-based. It may report specific well-defined diseases or, if diagnostic capabilities are limited, it may report syndromes (e.g., the syndrome of ulcerative genital disease as opposed to genital herpes, chancroid, or syphilis, specifically). Surveillance systems are a fundamental tool in public health because they provide critical information on disease burden and changes in disease frequency over time that may alert public health authorities to epidemic disease. Studies that measure an effect may measure the effectiveness of a new drug or vaccine; alternatively, they measure the effect of a possible risk factor on disease frequency (e.g., the effect that smoking has on the risk of developing lung cancer). The optimal study for measuring effect is a clinical trial. Randomized clinical trials are prospective studies in which individuals are randomized to one of at least two study arms and followed for the outcome of interest over time. The major advantage of this type of study is that the random nature of group assignment ensures that people in one group will not differ systematically from people in another group in some way that would influence outcome. Another way to say this is that the purpose of randomization is to eliminate potential confounding factors (whether suspected or not suspected) that are associated with both exposure and outcome. If the random assignment is completely unknown to both the study participants and researchers, it is called a double-blind randomized trial. If one of the study arms receives a treatment or intervention and the other receives a placebo, the trial is called a placebo-controlled trial. Since it is unethical to offer one group a placebo if there is available a treatment of known benefit, many trials compare the efficacy of a new intervention to that of a standard therapy; these trials are called equivalence studies. An example of a randomized double-blind equivalence study would be one that compares the efficacy of two drugs, drug A and drug B, for the treatment of shigellosis. To ensure that both participants and researchers are blinded to the intervention, the drug preparations should look and taste the same, be administered on the same dosing schedule, and given by the same route. Follow-up of the two groups should be identical. It is often impossible to conduct a clinical trial, especially when the exposure of interest is not an intervention or a treatment but some kind of environmental or genetic factor. In this case, a cohort study can be conducted to estimate association between risk factor and outcome. In a cohort study, groups of individuals with different exposure histories are identified and followed over time. As an example, imagine that we want to study the relationship between smoking and lung cancer. We could identify individuals who smoke and those who do not and follow them over time to see if the incidence of lung cancer between the two cohorts is different. This type of study has several possible shortcomings, one of which is that it may take years or decades for an individual to develop a disease or outcome after initial exposure. Another problem inherent in this type of study is the fact that the two cohorts of exposed and unexposed people may differ in ways other than just the basis of the exposure of interest (a confounding influence). For example, a confounding influence in our study may be the effect of alcohol consumption on the development of lung cancer (for instance, if individuals who drink heavily were more likely to smoke than those who do not drink heavily). An alternative study design is a case-control study. In this type of study, exposures of people who experience an outcome of interest are compared to exposures of those who have not had such an outcome. This type of study is especially useful for a rare disease or when there is a prolonged period of time between exposure and outcome. Continuing our example of examining the relationship of smoking and lung cancer, in a casecontrol study, we would identify individuals who have developed lung cancer and a group of “controls” who have not developed lung cancer. We could then ascertain the smoking histories of individuals in the study and try to determine whether they were consistently different between cases and controls. Since case-control studies often involve retrospective collection of exposure data obtained after a subject knows his or her diagnosis, there is a potential for recall bias. Another potential problem in case-control studies involves the selection of controls. Ideally, cases should be chosen from the population that gave rise to cases and should

be selected without regard to exposure status. Case-control and cohort studies are also referred to as observational studies, since there is no intervention.

HYPOTHESIS TESTING When designing a study, researchers should first state the hypothesis that they want to test. For example, “we hypothesize that drug A is effective in treating salmonellosis.” The hypothesis should be stated before data are collected. A common pitfall of studies is to first collect data and then to analyze data for comparisons that reach statistical significance. Such a fishing expedition may uncover real differences, but may also uncover differences related to chance alone. A placebo-controlled double-blind randomized trial would be the best way to test our hypothesis that our new drug A is “effective” in treating individuals with salmonella gastroenteritis. When designing this study, researchers should first select relevant and measurable endpoints that will distinguish whether individuals who get the drug “do better.” Primary endpoints may be days of diarrhea; days of fever; duration of bacterial shedding of salmonella organisms in stool; and/or the presence or absence of infectious complications, bacteremia, or death. These outcomes should be as clinically relevant and precise as possible; that is, definitions of what constitutes diarrhea and fever should be established before the study is undertaken, the same amount of stool and blood should be collected and processed from all study enrollees to ensure equality of assessment, and data should be recorded and reported as accurately and completely as possible. After the hypothesis is stated, the researchers should next formulate the null hypothesis. In this step, the investigators should assume that no true difference exists between the two study groups (those who get drug A and those who get the placebo). A decision should then be made as to what constitutes a statistically significant result. Statistical significance is usually conveyed through a statistic known as the P value; results are often considered significant if the P value is less than a cutoff value (or “alpha level”) of 5%. The P value refers to the probability that one would observe a result equal to or more extreme than the study result under the null hypothesis. One way to interpret this is to say that if a difference is shown between the two groups, there is a 95% chance that the difference is true (or a less than 5% chance that the difference is due to chance alone). The alpha level is a cutoff value for a P value for a hypothesis test that is often set, somewhat arbitrarily, at 0.05. A type I error occurs when the null hypothesis is incorrectly rejected when it is in fact true, that is, when there is no difference between drug A and a placebo. A test with an alpha level of 0.05 should lead to type I errors no more than 5% of the time. Unlike the P value that varies with the data, alpha levels are chosen in advance and indicate the specific P value that will be considered significant. A type II (or beta) error occurs when the null hypothesis is not rejected even when there is truly a difference between the two arms of the study, that is, between drug A and the placebo. Type II errors may occur when a study is not large enough to detect a difference, or when individuals are not followed for an adequate amount of time for differences between groups to become apparent. Most well-designed studies aim for a type II error rate between 10% and 20%. The power of a study refers to the probability that the null hypothesis is rejected when it is false, and it is thus given the expression Power = 1 − Probability of a type II error Therefore, most studies aim for 80–90% power (i.e., an 80–90% chance that if the null hypothesis is not rejected it is correct). It is this power calculation that determines the number of individuals who need to be enrolled in a study. Only after the hypothesis has been stated and a study appropriately designed and adequately powered should data be collected and stored. Once this is completed, data analysis may begin. In this step, investigators determine the estimated effect of the intervention or exposure, and the

Data may be expressed in many ways. When an exposure or outcome is expressed in terms of a continuous variable such as age or weight, the differences between groups may be expressed by comparing mean or median values for the two groups. Both these statistics are measures of central tendency, meaning that they describe the middle, or average, value of the data. The mean is the arithmetic average, which is simply obtained by summing the observations and dividing the sum by the number of observations. For instance, if we measure the days of diarrhea following administration of drug A to patients with salmonellosis, we may find that one patient had diarrhea for 2 days, another for 3 days, another for 4 days, another for 5 days, and another for 20 days. The mean would be a summation divided by the number evaluated (2 + 3 + 4 + 5 + 20 (equals 34) divided by 5 = 6.8). The median is the value that divides the data in half; 50% of the observations have values lower than the median, and 50% have values greater than the median. The median is also referred to as the 50th centile. Using the median rather than the mean lessens the impact of outliers, since the actual values of extreme data points do not affect the median. Another way of reducing the effect of extreme outlier observations is to use the geometric mean, which is often used with data measured on a logarithmic scale. The geometric mean is calculated by multiplying the observed values and taking the nth root, where n is the number of observations. For the preceding example, this would be given by 5

(2)( 3 )( 4 )( 5 )(20 ) = 4.7

Ever smoked Never smoked

∑

X −X n −1

Number or frequency

where X represents the value of each individual observation, X represents the mean, and n represents the number of observations. The standard error of the mean indicates the degree of uncertainty in calculating estimate from a sample. A standard error may be calculated from the standard deviation by dividing the standard deviation by a square root of n (with n representing the number of values measured). Range refers to the interval from the minimum to the maximum value in a set of quantitative measurements. For instance, the arithmetic mean in our example would be 6.8, the geometric mean would be 4.7, the median would be 4, and the range would be 2 through 20. Data that are normal or normally distributed are symmetrically distributed around a mean. A classic example of normally distributed data is a bell-shaped curve (e.g., a population-based IQ evaluation; Fig. 3.1). Characteristics that we might want to study may be measured in a variety of ways. Observed data may be dichotomous, categorical, or continuous. If data can take only one of two values, they are defined as dichotomous. Returning to our smoking and lung cancer example, we could describe smoking in terms of the dichotomous variables “ever” or “never” smoked.

Variable

Figure 3.1  Normal distribution.

Lung Cancer

No Lung Cancer

Total

5 1 6

95 99 194

100 100 200

Under the null hypothesis, we assume that there is no difference in the incidence of lung cancer among smokers and nonsmokers. Given that 6 cases of lung cancer occurred among the 200 people we followed, we can come up with a table of “expected” frequencies under the null hypothesis.

The term standard deviation measures the spread of the individual observations around the mean. It is given by the formula s=

Chapter 3

Data Expression and Analysis

Categorical observations have values that fit into categories. For example, we might characterize race or ethnicity using a categorical variable. Some data categories describe ascending levels of intensity or severity. For example, we could describe smoking history as “none,” “light,” “moderate,” and “heavy.” When categorical data are ordered in this way, they are ordinal. Finally, data may be measured on a continuous scale. Again, referring to our smoking example, we could measure smoking in terms of the number of cigarettes consumed. At analysis, continuous data may be transformed into categorical data (but not vice versa). Once we have summarized our data into the groups we are comparing, we need to decide whether the data differ between groups. If data are dichotomous, we may compare proportions, for example, the proportion of “ever” smokers who develop lung cancer to “never” smokers who develop lung cancer. For instance, imagine in our study that 5 smokers develop lung cancer out of a cohort of 100, while only 1 of 100 non­ smokers develops lung cancer. These data could be presented in a table of observed frequencies as follows:

Epidemiology and Biostatistics

probability that the observed difference between the two study groups would occur if no true difference exists in the larger population.

Ever smoked Never smoked

Lung Cancer

No Lung Cancer

Total

3 3 6

97 97 194

100 100 200

We can use the chi-square test statistic to ask how likely it would be that we obtained the observed frequencies if the null hypothesis were true. The chi-square statistic is given by the formula

χ2 =

∑

ALL CELLS

(O − E )2 E

where O is observed frequency and E is expected frequency. If the chisquare test is small, this suggests that there is no difference between the groups; if it is large, we assume that a difference exists. Other statistical tests are available to analyze data. The choice of optimal test depends on a number of variables, including data type and number of groups. For instance, for continuous data that are normally distributed, we could compare the means of two groups using the t test for comparing means. When data are not normally distributed, other tests would be required; these usually are based on “order statistics” and include such nonparametric methods as the Mann–Whitney U test, the Kruskal–Wallis test and the Wilcoxon matched rank test, among others. The ANOVA (analysis of variance) test may be used to compare more than two groups that are normally distributed. The Mann–Whitney U test is used for evaluating two groups that are not normally distributed, and the Kruskal–Wallis test may be used for evaluating more than two groups that are not normally distributed. Analysis of data may disclose “associations.” For instance, returning to our example of smoking and lung cancer, we may find that smoking and lung cancer are statistically associated. Although variables that are found to be associated with an outcome are often called risk factors, a statistical association does not imply a cause and effect relationship between that variable and outcome. The relative risk is the probability of an outcome if a risk factor/association is present divided by the probability of the

13

No Lung Cancer

Total

A C A+C

B D B+D

A+B C+D A+B+C+D

RR = OR = =

A (A + B) C (C + D )

[ A ( A + C )] [C ( A + C )] [B ( B + D )] [D ( B + D )] A C AD = B D BC

Confidence intervals are a way of combining information about the strength of an association with information about the effects of chance in obtaining the observed results. A 95% confidence interval (CI) is most commonly used. An association is usually reported as an odds ratio (OR) or relative risk (RR) with a 95% CI. The final stage of analyzing a study is extrapolation. We may extrapolate to an individual or to a group. For instance, based on a relative risk or odds ratio of 5 for smoking and lung cancer, we could conclude that if an individual smoked, he or she may be five times more likely to develop lung cancer than if he or she did not smoke. We may also speak of an attributable risk percent. The advantage of this concept is that it allows us to think of a portion of the risk of developing a disease that may be eliminated among those who do not have the risk factor. Attributable risk percentage may be thought of as

(RR − 1 RR ) × 100%

14

In many instances, laboratory tests are part of a case definition (e.g., detection of serum antibodies against human immunodeficiency virus (HIV) in a study involving individuals infected with HIV). It should be recalled that a test is often a surrogate marker to distinguish a disease-free group from a diseased group. Assuming a normal distribution in both groups of whatever marker we are measuring (e.g., a serum antibody level), we can imagine that the disease-free and diseased groups do not overlap at all with regard to the specific blood test of interest (Fig. 3.2A). Often, however, the two groups do overlap, and some individuals in the diseased group will have tests with lower values than some of the individuals in the disease-free group (Fig. 3.2B). In establishing the utility of a test, therefore, we must first establish the reference interval for disease-free individuals. Sensitivity and specificity of a test are then measured compared with a “gold standard.” Sensitivity measures the probability that those with a disease will have a positive test when individuals with the disease are identified by the gold standard. Specificity measures the probability that those who do not have the disease will test negative by the test being evaluated. There usually is a tradeoff between the sensitivity and specificity of a specific test. For example, if we choose X as the cutoff value for a positive test on Figure 3.2B of overlapping curves, we will achieve 100% sensitivity – but at the cost of misclassifying many negative cases as positive ones, that is, reducing specificity. Conversely, we could maximize specificity by moving our cutoff value for a positive test rightward to the Y position, but in so doing, we would compromise our ability to identify a true case of disease. For example, imagine that we are evaluating a new test to diagnose schistosomiasis, and imagine that we will compare this test to a gold standard in a village with a population of 1000 individuals of whom 500 actually have schistosomiasis by our gold standard. Imagine that our new test correctly identifies 400 infected individuals but incorrectly identifies 100 truly infected individuals as not having schistosomiasis when in fact they are infected ( false negative). Also imagine that our new test incorrectly labels 50 individuals as having schistosomiasis who do not (false positive).

For instance, in our study, we found that smoking was associated with a relative risk of 5 of developing lung cancer. This may not seem like an overly large risk of developing lung cancer; however, the attributable risk percent is 5 − 1/5 = 80%. This suggests that 80% of lung cancer in our study population could have been prevented if our study participants had never smoked.

Number or frequency

  

 

Ever smoked Never smoked

Lung Cancer

UNDERSTANDING DIAGNOSTIC LABORATORY TESTS

Test negative

B

Test positive

Test results

A Number or frequency

PRINCIPLES AND GENERAL CONSIDERATIONS

Section I

outcome if the risk factor/association is absent. For instance, in our example of the cohort study of smokers and nonsmokers, we imagined a study in which we have followed 100 individuals who smoke and 100 individuals who never smoked. We saw that 5 of the 100 smokers developed cancer (probability 0.05), but that only 1 of 100 nonsmokers developed lung cancer (probability 0.01). The relative risk is, therefore, 0.05 divided by 0.01, or 5. A relative risk of 5 implies that individuals who smoke are five times more likely to develop lung cancer than individuals who do not smoke. In case-control studies, it is the researcher who determines how many study and control participants are evaluated, and so a true disease frequency in the population as a whole cannot be established. In this case, we cannot estimate the relative risk, since we do not actually know the risk of disease in the unexposed population. An approximation of the relative risk for case-control studies is the odds ratio. To understand the difference between a risk and an odds ratio, think of the probability (or risk) of throwing a six-sided die in a game of chance and having the die land with six black dots facing up (1 in 6 chance). The odds of throwing a six on the other hand will be the number of times the die will land with six black dots showing divided by the number of times six dots will not be uppermost (1 to 5). An odds ratio is, therefore, the odds of developing an outcome if an association is present, divided by the odds of an outcome if the association is absent. Both the relative risk and the odds ratio are easy to calculate from a 2 × 2 table.

Test negative

Test positive

X

Y

Test results

Figure 3.2  (A) Normal distribution in which two groups do not overlap. (B) Two groups that overlap.

Gold Standard Disease-Free

Positive Negative

400 (a) 100 (c) 500

50 (b) 450 (d) 500

Sensitivity and specificity are calculated as: a 400 = = 0.80 or 80% a + c 500 d 450 Specificity = = = 0.90 or 90% b + d 500 Sensitivity =

Our test will, therefore, have a sensitivity of 80% and a specificity of 90%. The actual utility of this test, however, will rest not only on its sensitivity and specificity but also on the prevalence of the disease in question in our population of interest. In our preceding example, there was a 50% prevalence of schistosomiasis (500 infected individuals who lived in a population of 1000). If we assume that the “true” pre­ valence of schistosomiasis in a different village is 20%, at a population level, the average individual in that village will have a 20% chance of having the disease before the test is performed (in reality, certain individuals will be at higher or lower risk of having schistosomiasis on the basis of age, sex, and other factors). Assuming that we are evaluating 1000 individuals (with 20% of them having the disease), and assuming that we are using our new test with a sensitivity of 80%, we would assume that 160 of the 200 individuals with the disease will be correctly identified by the test:

Test

Gold Standard Disease

Gold Standard Disease-Free

Positive Negative

160 40 false negatives 200

80 false positives 720 800

The remaining 20% of these individuals will be incorrectly labeled as negative (n = 40; false negatives). Specificity equals 90%; therefore, 90% of those who are disease-free will be correctly labeled as negative (90% of 800; 720 true negatives). The remaining 10% of individuals who are disease-free will be incorrectly labeled as positive (10% of 800; 80 false positives). Now let us imagine that we are applying our same test in a different village (Village Y) in which schistosomiasis is much more prevalent and 75% of the population has the disease:

Test

Gold Standard Disease

Gold Standard Disease-Free

Positive Negative

600 150 false negatives 750

25 false positives 225 250

Finally, let us imagine that we use our test in a third village (Village Z), in which schistosomiasis is much rarer and the true probability of disease is only 2% (only 2% of the population is infected). Then our table would look like this:

Test

Gold Standard Disease

Gold Standard Disease-Free

Positive Negative

16 4 false negatives 20

98 false positives 882 980

Now, let us analyze the predictive value of positive and negative test results in each of these villages. The positive predictive value refers to the probability that one who tests positive truly has the disease, while the negative predictive value refers to the probability that one who tests negative

actually does not have the disease. The crucial point to understand is that the predictive value depends not only on the sensitivity and specificity of the test itself but also on the disease prevalence in the population being evaluated.

Test Positive

Negative

Gold Standard Disease

Gold Standard Disease-Free

a = number of individuals diseased and positive c = number of individuals diseased and negative

b = number of individuals disease-free and positive d = number of individuals disease-free and negative

Chapter 3

Gold Standard Disease

a + b = total number of test positives

Epidemiology and Biostatistics

Test

c + d = total number of test negatives

The following formulae are used for calculating the predictive value of a positive test and the predictive value of a negative test: Proportionof a individuals with Predictive value of a positive test = apositive test a + b who actually have the disease Proportionof individuals with d anegative test Predictive value of a negative test = c + d who actually donothave thedisease Using the preceding calculated numbers, and assuming a 2% pretest probability of disease (as in Village Z): Predictive valueof apositive test = Predictive valueof anegative test =

a 16 = = 14% a + b 16 + 98

d 882 = = 99.5% c + d 4 + 882

Similarly, if we assume a 75% pretest probability as in Village Y: Predictive valueof apositive test = Predictive valueof anegative test =

a 600 = = 96% a + b 600 + 25 d 225 = = 60% c + d 150 + 225

Therefore, using exactly the same test, with exactly the same sensitivity and specificity, we can generate the following table of positive and negative predictive values of our test in villages with different prevalences of the disease in question:

Pretest Probability

Predictive value of a positive test Predictive value of a negative test

2% (Village Z)

20% (Village X)

75% (Village Y)

14%

66.7%

96%

99.5%

94.7%

60%

This means that there is an 86% chance that a positive test obtained in Village Z with a 2% pretest probability of disease is falsely positive. Similarly, there is a 40% chance that a negative result is in fact falsely

15

Section I

negative in Village Y with a pretest probability of disease of 75%. This is despite the fact that we are using the same test with the same sensitivity and specificity in each village. It is, therefore, crucial to understand that the interpretation of laboratory tests (whether in a study or in clinical practice) should be understood in context.

PRINCIPLES AND GENERAL CONSIDERATIONS

Access the complete reference list online at

  

 

http://www.expertconsult.com

16

ACKNOWLEDGMENTS We are grateful to Margaret Bikowski and Sam Riley for assistance with figures. This work was supported in part by a grant from the US Centers for Disease Control and Prevention, U19CI000514 (ETR).

SECTION I: PRINCIPLES AND GENERAL CONSIDERATIONS

CHAPTER 4 

Access the complete reference list online at

http://www.expertconsult.com

Social and Cultural Factors in Tropical Medicine: Reframing Our Understanding of Disease Rebecca Dillingham • David A. Walton • Paul E. Farmer

BARRIERS TO CARE AND THE PERSISTENT PATHOGENS: A BIOSOCIAL APPROACH By the close of the twentieth century, the world’s ranking infectious pathogens – from well-known malaria and filariasis to the more recently described human immunodeficiency virus (HIV) – could be controlled by the tools of modern medicine and public health. And yet, none of these diseases, or other important “tropical diseases,” has been brought to heel. None of them, even those without a nonhuman host, have been eradicated. Some epidemic diseases continue to expand. Still others mutate to become drug-resistant pathogens for which there are no clear therapies. Vaccines, in the event they exist, are not always delivered effectively in the settings in which they are most needed. The cumulative burden of these diseases remains unchanged, and in some cases is growing; this burden weighs most heavily on the world’s poorest billion inhabitants. This chapter examines this central irony of twenty-first century medicine: the persistence of morbidity and mortality due to infectious diseases years after effective vaccines, cures, suppressive treatments, and preventive strategies have been discovered. The forces inhibiting the successful application of existing scientific knowledge are not primarily biological or cultural; they are, rather, economic and structural barriers to change. Using what may be called a biosocial model, this chapter will describe these barriers to health and will highlight several approaches to overcome them. Because modern epidemics are almost invariably rooted in social conditions, some long-standing and others changing rapidly, inquiry that does not draw on the social sciences is unlikely to offer a comprehensive or accurate understanding of these plagues. The mechanisms by which social forces shape epidemics are not readily revealed by single methodologies, but rather by interdisciplinary research that links qualitative and quantitative methods. A biosocial model calls into question the very term tropical diseases, since in the past many of the pathogens discussed in this textbook caused significant mortality far from the tropics. The dozen or so diseases grouped under this rubric have little in common in terms of pathogenesis, chronicity of infection, modes of transmission (some are vector-borne, others are not), or efficacy of existing control and treatment strategies. But whether we consider what are often termed the “neglected diseases” (African trypanosomiasis, dengue, leishmaniasis, schistosomiasis, Chagas’ disease, lymphatic filariasis, and onchocerciasis) or the largest infectious killers (diarrheal and respiratory diseases, human immunodeficiency virus (HIV), tuberculosis (TB), and malaria), there are important similarities to consider. All of these diseases afflict the poor disproportionately and are linked to social conditions. For each disease, there exist important deliverables: new and improved diagnostics and therapeutics that could affect the distribution and outcomes of these epidemics. In each instance, new discoveries could substantially decrease the morbidity and mortality associated with these persistent pathogens, but social and economic barriers hamper effective control strategies.

The mechanisms by which social forces delay health progress are in large measure those leading to the propagation of new epidemics such as HIV. We begin this review with a case study of HIV and TB in rural Haiti, since many of the barriers to care are best seen by closely examining individual illness trajectories and since TB is often the leading cause of death among patients living with HIV disease in resource-poor settings. The case study is followed by a review of structural barriers to effective HIV prevention and care. Next, a brief description of the challenges of implementing insecticide-treated bednets (ITNs) and artemisinin-based combination therapies (ACTs) illustrates how poverty, the primary social barrier, limits effective implementation of proven prevention and treatment strategies. Finally, we will explore the implications of this review for effective control of these and other leading infectious killers.

AIDS AND TUBERCULOSIS IN HAITI: JOSEPH’S STORY On the afternoon of March 17, 2003, four men appeared at the public clinic in Lascahobas, Haiti, each carrying a leg of a makeshift stretcher. (The Lascahobas health clinic in central Haiti is a partnership between the nonprofit organization Partners In Health/Zanmi Lasante and the Haitian Ministry of Health. When it began providing voluntary counseling and testing and treatment for HIV in late 2002, Lascahobas became rural Haiti’s second full-service HIV clinic.) On the stretcher lay a young man, Joseph, eyes closed and seemingly unaware of the 5-mile journey he had just taken on the shoulders of his neighbors. When they reached the clinic after the 4-hour trip, they placed him on an exam table. The physician tried to interview him, but Joseph was already moribund. His brother recounted the dying man’s story. Joseph, 26, had been sick for months. His illness had started with intermittent fevers, followed by coughing, weight loss, weakness, and diarrhea. His family, too poor to take him to a hospital, brought Joseph to a traditional healer. Joseph would later explain: “My father sold nearly all that he had – our crops, our land, and our livestock – to pay the healer, but I kept getting worse. My family barely had enough to eat, but they sold everything to try to save me.” Joseph was bedridden 2 months later. He became increasingly emaciated and soon lost all interest in food. As he later recalled, “My mother, who was caring for me, was taking care of skin and bones.” Faced with what they saw as Joseph’s imminent death, his family purchased a coffin. Several days later, a community health worker employed by the Lascahobas clinic visited their house. The health worker was trained to recognize the signs and symptoms of TB and HIV and immediately suspected that the barely responsive Joseph might have one or both of these diseases. Hearing that their son might have one last chance for survival, Joseph’s parents pleaded with their neighbors to help carry Joseph to the clinic since he was too sick to travel on a donkey and too poor to afford a ride in a vehicle.

17

Section I PRINCIPLES AND GENERAL CONSIDERATIONS

A

B

C

Figure 4.1  The “Lazarus effect”: effective, accessible treatment for HIV and TB brought Joseph back from the brink of death. His experience, and that of hundreds of thousands of other impoverished patients in rural Haiti now receiving free medical care through a public–private partnership, is renewing his community’s faith in the health sector and restoring hope to the destitute sick everywhere.   (A) March 2003: Joseph, before treatment. (B) September 2003: Joseph after treatment for HIV and TB, with his healthy niece. (C) August 2004: Joseph speaks at a health and human rights conference. (A and B by David Walton; C by Joia Mukherjee. Copyright, Partners In Health. All rights reserved.)

  

 

At the clinic, Joseph was diagnosed, as per the community health worker’s suspicions, with advanced HIV and disseminated TB (Fig. 4.1A). He was hospitalized and given antiretrovirals and antituberculous medications. Like his family, however, Joseph had lost faith in the possibility of recovery. He remembers telling his physician early in the course of his treatment, “I’m dead already, and these medications can’t save me.” Despite his doubts, Joseph dutifully took his medications each day, and he slowly began to improve. Several weeks later, he was able to walk. His fevers subsided and his appetite returned. After discharge, he received directly observed therapy (DOT) for both HIV and TB from a neighbor serving as an accompagnateur. (For a more detailed discussion of community-based approaches to HIV treatment and care, see Farmer et al.1) After 4 months of therapy, Joseph had gained 30 pounds (Fig. 4.1B). Now, Joseph is employed as an HIV outreach worker, often speaking in front of large audiences about his experience (Fig. 4.1C). “When I was sick … I couldn’t farm the land, I couldn’t get up to use the latrine; I couldn’t even walk. Now I can do any sort of work. I can walk to the clinic just like anyone else. I care as much about my medications as I do about myself. There may be other illnesses that can break you, but AIDS isn’t one of them. If you take these pills, this disease doesn’t have to break you.”

STRUCTURAL BARRIERS TO EFFECTIVE HIV AND TUBERCULOSIS CONTROL

18

By the end of 2008, 4 million people in resource-limited settings had initiated antiretroviral therapy (ART), a 10-fold increase in 5 years. Despite this impressive scale-up of the provision of ART, Joseph’s experience is typical in many ways and instructive in most. In December 2008, only 44% of those who needed ART were receiving it. The rate of coverage among infected children was even lower.2 In addition, budgetary decisions by donor countries, including the United States, limit the expansion of ART programs, making rationing of the life-saving therapy increasingly common in some countries in Africa.3 Joseph’s story is similar to that of tens of millions worldwide suffering from HIV and TB. He lives in great poverty; he sought care unsuccessfully until he simply gave up. His family did what they could to save his life, selling off meager assets and receiving contradictory advice from neighbors and from traditional healers in a country in which trained medical personnel are rare and most care is fee-for-service. Joseph’s experience is exceptional in that he lived in an area of Haiti in which treatment for both HIV and TB had recently come to be considered a public good and made available to poor patients who could not afford user fees.4 Treatment

for these two chronic infectious diseases is supervised not by physicians or nurses – who do play a role in the diagnosis of illness – but by community health workers who are also one’s neighbors.5 In the following sections, we review these topics in greater detail by examining an emerging literature that explores socioeconomic barriers to effective care for HIV and tuberculosis.

The “Brain Drain” One significant and often-invoked barrier to effective care in resourcepoor settings is the lack of medical personnel. In what has been termed the “brain drain,” large numbers of physicians and nurses are leaving their home countries in order to pursue opportunities abroad, leaving behind health systems that are understaffed and ill-equipped to deal with the epidemic diseases that ravage local populations. A recent survey reports that one-fifth of African physicians and one-tenth of African nurses work abroad; some countries have lost as many as 70% of their health care professionals.6 The World Health Organization (WHO) estimates that there is a global deficit of 2.4 million doctors, nurses, and midwives, with the greatest number needed in sub-Saharan Africa.7 This deficit of health care workers, while appalling, conceals further inequalities in health care staffing within countries. Rural–urban disparities in health care personnel mirror disparities of both wealth and health. Fewer than 55% of all people in the world live in urban areas, but over 75% of physicians and more than 60% of nurses are concentrated there.7 In addition to inter- and intranational transfer of personnel, HIV itself is contributing to personnel shortages across Africa. Although data on the prevalence of HIV among health professionals are scarce, the available numbers suggest substantial and adverse impacts on an already overburdened health sector. The situation in Malawi is instructive, where a ranking cause of loss of health care workers is premature death, often from AIDS.8 The shortage of medical personnel in the areas hardest hit by HIV has profound implications for prevention and treatment efforts in these regions. The cycle of health sector impoverishment, brain drain, and consequent lack of personnel to fill positions, when they are available, conspires against ambitious programs to bring antiretroviral drugs to those living with both HIV and poverty. Furthermore, the education of medical trainees is jeopardized as the ranks of the health and academic communities continue to shrink due to migration or disease. A proper biosocial analysis of the brain drain reminds us that health personnel flight – almost always from poor to less poor regions – is not simply a question of desire for more equitable remuneration. Epidemiologic trends and access to the tools of the trade are also relevant, as are working conditions more generally. In many of the settings now losing skilled

Structural Adjustment and Access to Care The large-scale socioeconomic forces at work in the differential distribution of sickness and health often stem from decisions made far from hospital wards or even national capitals. Many African countries in fact registered improvements in health systems and indices in the years following independence. Kenya, for example, saw infant mortality decline by more than 58% between 1963 and the early 1990s.12 However, the economic crises of the 1970s and 1980s left many of these postcolonial nations mired in mounting debts to offshore creditors. African governments needed loans to pay interest on the outstanding debts and to meet basic social spending needs. The World Bank and the International Monetary Fund became the principal creditors in this arena, and these institutions soon found themselves the chief sources of funding for health and development projects in poor countries. Loans came with strict requirements such as the imposition of sweeping economic “reforms” in favor of a “free market” system designed to stimulate the economy and fix perceived imbalances in trade and government budgets. These economic austerity measures – termed “structural adjustment programs” – had profound and often deleterious effects on the health of many countries, as mandatory reductions of government budgets led to sharp declines in funding for the health sector. In Nigeria, for example, per capita expenditure on health care fell by more than 70% between 1980 and 1987.13 Some have argued that structural adjustment programs in fact increased risks for HIV infection in Africa.14 Cuts in health care spending led to the closing of public health posts, clinics, and hospitals in many parts of sub-Saharan Africa. Even basic services – from first-aid to prenatal care – were slashed from the

Chapter 4

operating budgets of many Ministries of Health. Between 1990 and 1992, 14 African countries – all mired in structural adjustment programs – saw at least a 10% decline in the level of polio vaccination. Botswana topped the list with a 24% reduction in polio vaccine coverage during this time.15 The collateral damage of economic austerity programs in sub-Saharan Africa also directly affected health care professionals. Thousands of doctors and nurses across the continent lost their jobs as a result of budget restructuring. Many emigrated overseas, and funds for health care worker education disappeared, threatening the future supply of trained professionals.16 For example, in late 2003, an estimated 4000 Kenyan nurses were unemployed secondary to economic policies that restricted recruitment of health workers into the public sector.17 The damage done by structural adjustment remains, but encouraging changes have occurred. The advent of substantial and sustained funding from large charitable foundations and donor countries to address the health problems of the poor has changed the discussion from whether to develop and support the systems needed to provide access to care to how to provide it.18

Social and Cultural Factors in Tropical Medicine: Reframing Our Understanding of Disease

health personnel, the advent of HIV has led to a sharp rise in tuberculosis incidence; other opportunistic infections have also become, in the eyes of providers, insuperable challenges. Together, these forces have conspired to render the provision of proper care impossible, as the comments of a Kenyan medical resident suggest: “Regarding HIV/AIDS, it is impossible to go home and forget about it. Even the simplest opportunistic infections we have no drugs for. Even if we do, there is only enough for a short course. It is impossible to forget about it … Just because of the numbers, I am afraid of going to the floors. It is a nightmare thinking of going to see the patients. You are afraid of the risk of infection, diarrhea, urine, vomit, blood … It is frightening to think about returning.”9 When providing care for the sick becomes a nightmare for those at the beginning of clinical training, physician “burn-out” soon follows among those who carry on in settings of impoverishment. In the publicsector institutions put in place to care for the poorest, the confluence of epidemic disease, lack of resources with which to respond, and user fees has led to widespread burn-out among health workers. Raviola and colleagues offer an important biosocial study linking epidemiology, the experience of providers, and an understanding of the political economy of the Kenyan health care system: “I don’t see a future for the medical profession here,” concluded a resident at Kenyatta National Hospital in Nairobi. “Why? It is expensive. You have to invest a lot. There is no government support. People can’t afford care here.”9 The WHO focused its 2006 report, “Working together for health,” on the issue of developing an equitable distribution of health care workers. The report highlights strategies to build stronger health care systems within which workers can actually provide care, to improve educational opportunities throughout health care workers’ careers, and to provide adequate, reliable compensation.7 While countries are still developing their national plans based on this report, an example of a successful program in Ghana, a country that lost 68% of its medical school graduates between 1993 and 2000, demonstrates how an emphasis on the development of in-country training and support of young physicians can persuade them to stay. In 2002, Ghana’s parliament allocated US$3 million to develop postgraduate training programs for physicians, the “Ghana College of Physicians and Surgeons.”10 The “Safe Motherhood Initiative” in obstetrics and gynecology, one of the new specialty training programs, had retained 37 of 38 of its trainees at the time of their report.11

User Fees Another legacy of economic austerity programs is the widespread institution of “user fees” for health services that were formerly free of charge, with the intent of generating revenue for the health sector. Instead of revitalizing the health sector, however, user fees often sharply reduce the ability of the poor to access medical care.19 For example, UNICEF and the WHO introduced the Bamako Initiative in 1987, with the approval of the Health Ministries of the WHO African Region, to try to improve access to primary care by creating a structure to generate revenue for clinics. The initiative includes the provision of generic essential drugs by donor agencies or national governments to district and village health management committees. These drugs are then sold to the public at a profit, which is then, theoretically, used to buy back the initial stock of drugs and to improve the quality of health centers. Despite widespread support from decision makers and a great deal of positive press, several studies have reported a decline in attendance rates for health services as a result of the user charges stipulated by the Bamako Initiative.20,21 A longitudinal study conducted in the Democratic Republic of the Congo revealed a 40% decrease in health service utilization between 1987 and 1991 after the adoption of the Bamako Initiative.22 Other studies also document deleterious effects of user fees. Despite exemptions for children and the indigent, Mbugua and colleagues found that attendance at rural Kenyan government health facilities plummeted by 41% after user fees were introduced, and rebounded after fees were abolished.23 In a cross-sectional study of 37 countries in sub-Saharan Africa, even the money-minded World Bank concluded that user fees decreased utilization of health services.24 Another study revealed that the number of men reporting to sexually transmitted disease (STD) public clinics in Kenya fell by 40% after user charges were introduced in 1989; the authors concluded that the introduction of user fees in STD clinics likely increased the number of untreated STDs in the population.25 This association has clear implications for HIV treatment and prevention efforts, as HIV is a sexually transmitted disease in much of the world. While user fees remain a topic of debate among some donors, many countries have developed strategies to eliminate them.26 Uganda eliminated user fees in 2001 with a subsequent increase in utilization of services and without any decline in measured or perceived quality of service.27 Other countries, like Rwanda and Cambodia, have implemented programs to provide government-subsidized health care to the poor through health insurance and a health equity fund respectively.26,28 In Haiti, a careful analysis by Médecins du Monde demonstrated that the elimination of user fees in clinics that the organization co-manages with the Haitian Ministry of Health resulted in a dramatic increase in clinic usage with a coincident reduction in cost per unit of care delivered. The authors of the report make a strong argument and provide important empiric evidence of the benefits of both augmenting the “supply-side” of the health

19

Section I

care equation with new clinics and personnel as well as the “demand-side” by eliminating the often insurmountable barrier of user fees.29

Economic Costs and Adherence to HIV Therapy

  

 

PRINCIPLES AND GENERAL CONSIDERATIONS

As HIV treatment programs in resource-poor settings undergo expansion, advocates and pundits alike agree that medication adherence is paramount to the success of these initiatives. Requiring patients to pay for ART is an enormous barrier to adherence. Data from individual programs have shown that adherence is significantly reduced at sites in which patients are forced to pay even nominal fees for their medications.30,31 In addition, dramatic clinical consequences, beyond poor adherence, result from charging the poor for ART. Ivers et al report in a metaanalysis of clinical cohorts from resource-limited settings that providing free medications contributes most to increasing the proportion of patients with a suppressed viral load, a critical parameter for HIV-infected patients.32 Similarly, the Antiretroviral Therapy in Lower-Income Countries (ART-LINC) collaboration documents results from 23 clinical cohorts in Africa, Asia, and South America. In analyses from this group, the risk of death for patients in fee-for-service programs was nearly fivefold higher than in programs that provided medications at no cost (HR: 4.64; 95% CI: 1.11–19.41).33,34 These studies suggest that charging for ART excludes the poor from access to care, increases the likelihood that acquired resistance will develop through inadequate therapy, and leads to unnecessary loss of life. As Mukherjee has noted, “a human rights-based – rather than marketbased – approach is the only realistic strategy for an epidemic that is concentrated in the poor and marginalized communities who have neither access to health care nor the ability to pay for treatment.”35 Unfortunately, the direct costs of ART are not the only financial burdens faced by HIV-infected individuals and their families. The economic toll of HIV before the patient is even diagnosed leads to further impoverishment and delays in seeking and receiving effective care. Lessons from the long-entrenched TB epidemic are instructive. In Bangladesh, where all TB services and medications are provided free of charge, the average total loss of income before proper diagnosis and treatment was estimated to be US$245, or nearly 4 months of a family’s yearly income.36 In Thailand, even after diagnosis at a government hospital, 80% of patients incurred significant costs for travel and food during hospital visits. In the same study, up to 15% of patients were forced to sell household assets and use bank loans to cope with illness-related expenses.37 These data on the deleterious effects of direct and indirect costs of therapy have obvious implications for the successful provision of lifelong HIV treatment. Fortunately, some programs have addressed the problem of hidden costs. One such example, the HIV Equity Initiative, the program in which Joseph was treated, covers indirect costs with a package of benefits that includes a monthly transportation stipend and emergency transportation coverage at a cost of US$60 per patient per year. In partnership with the World Food Programme, the HIV Equity Initiative also provides nutritional support to those in need, at a cost of approximately US$450 per family per year.38 Ensuring access to food combats another critical threat to adherence to ART: hunger.39,40

Civil Conflict and Natural Disaster

20

The noxious synergy of civil strife and poverty has a well-documented negative impact on the health of the poor. Conflict contributes to the deterioration of existing public health infrastructure in addition to creating conditions that decrease access to clean water and shelter, which can in turn lead to increased incidence of infectious pathogens. In an editorial on the violence that engulfed Haiti in 2004, Farmer laments the near-total collapse of the health sector.41 As easy targets for violence, as well as often being overwhelmed by the consequences of violence, hospitals are increasingly prone to acute staff shortages, stockouts of medications and

supplies, and even closure.42 More lasting effects also follow conflict due to exacerbation of the “brain drain.” A study of the migration of African health professionals documented the loss of more than 40% of the physicians between 1990 and 2000 from the war-ravaged countries of Angola, Congo-Brazzaville, Guinea-Bissau, Liberia, Mozambique, Rwanda, and Sierra Leone.6 The direct impact of conflict on HIV and TB epidemics has also been documented. In Sierra Leone, 23% of tuberculosis clinics closed between 1990 and 1994 secondary to violence and war.43 The Rwandan genocide in 1994 is believed to have contributed to the expansion of the HIV epidemic to rural areas of that country.44 The civil war in Guinea-Bissau in 1998–1999 is linked to a doubling of the HIV-1 seroprevalence in that country.45 At the time of this writing, just weeks after a 7.0 earthquake devastated Haiti’s capital city, it is also impossible to ignore the effects of natural disasters on the health sector. Like human-generated violence, events such as earthquakes, tsunamis, and hurricanes tend to reinforce preexisting social inequalities and strain health systems. In addition to leaving over 200 000 people dead and nearly 300 000 more wounded, the January 12, 2010 earthquake in Haiti collapsed the country’s largest nursing school, killing 100–150 students and faculty, including the entire second-year nursing class. Clearly, this heartbreaking loss will have significant long-term repercussions for Haiti’s already-strained health sector human resources.

Stigma The role of stigma as a major barrier to care for HIV patients has been extensively reviewed elsewhere.46–49 However, recent experiences in resource-poor settings suggest that the introduction of effective HIV treatment and care programs can help destigmatize what was once considered a fatal disease.49 In communities in rural Haiti, dramatic, visible recoveries after initiation of antiretroviral treatment, dubbed the “Lazarus effect,” have led to a sharp decline in HIV-related stigma.50 While the contours of discrimination vary from country to country, AIDS-related stigma will likely decrease as access to treatment improves and as people come to realize that HIV is a manageable disease.

THE COST OF MALARIA Malaria’s human toll is enormous. Nearly 250 million people suffer from malarial disease each year, and the disease annually kills almost 1 million people, mostly pregnant women and children under 5. The poor disproportionately suffer the consequences of malaria. Fifty-eight percent of malaria mortality occurs in the poorest 20% of the world’s population; 90% of malaria mortality is registered in sub-Saharan Africa.51 The differential magnitude of this burden of mortality is greater than that associated with any other disease.52 Despite suffering the greatest consequences of malaria, the poor are precisely those least able to access effective prevention and treatment tools.53

Rolling Back Malaria Due in part to differences in vector distribution and climate, resourcerich countries offer few blueprints for malaria control and treatment that are applicable in tropical settings. In 2001, African heads of state endorsed the WHO Roll Back Malaria (RBM) campaign, which prescribes strategies appropriate for sub-Saharan African countries. RBM recommends a three-pronged strategy to reduce malaria-related morbidity and mortality: the use of insecticide-treated bednets (ITNs), combination antimalarial therapy with an artemisinin-containing medication (ACT), and indoor residual spraying. In the first years of this program, uptake of these strategies was disappointing, but beginning in 2004, a growing number of malaria-endemic countries have adopted these principles and implemented programs.53 This may in part reflect a fivefold increase in international funding for malaria programs between 2003 and 2009 from

Chapter 4

Emulating the strategy pioneered to increase access to medications for drug-resistant TB may be helpful for ACTs as well, and a global subsidy fund has already been created to purchase therapeutics for countries that need them.64,65 In addition, careful attention to the features of programs that successfully ensure appropriate diagnosis and treatment of malaria – such as the employment of community health workers and the use of blister packs – will provide important guidance in the struggle to bring proven prevention and treatment techniques to the poorest, especially in rural settings.65

Social and Cultural Factors in Tropical Medicine: Reframing Our Understanding of Disease

US$0.3 billion to US$1.7 billion.51 A closer look at the differential success in increasing access to ITNs and to ACTs is useful when considering the importance of careful attention to the economic and social complexities revealed through a biosocial lens. The initially limited success of the scale-up of ITN coverage was indicative of the campaign’s inadequate acknowledgment of the economic barriers that preclude the destitute sick from accessing critical preventive technologies. Despite proven efficacy and what are considered “reasonable costs,” the 2003 RBM report revealed disappointing levels of ITN coverage. In the 28 African countries surveyed, only 1.3% (range, 0.2– 4.9%) of households owned at least one ITN, and less than 2% of children slept under an ITN.54 The RBM strategy initially emphasized the importance of commercial markets as sources of ITNs for African populations.55 A precedent supporting this emphasis was the prior existence in countries such as Madagascar and Mali of local markets for untreated bednets. Presumably, therefore, a demand for bednets existed prior to the RBM campaign, as did a distribution system with points of sale.56 However, this market approach, even with the application of subsidized social marketing strategies, did not result in large increases in coverage in the first years of the RBM campaign. Several studies attempted to define willingness to pay (WTP) and actual payment for ITNs in African countries in order to understand why market-based strategies were unsuccessful. Policy-makers often utilize WTP to determine appropriate pricing for social marketing projects and to project revenue and demand.57,58 Guyatt and colleagues provide an important critique of WTP studies from their work in highland Kenya. The 2002 study compared the attitudes of people living in homesteads that had been provided with heavily subsidized ITNs (n = 190) to households that had no bednets and had not been targeted by other health care initiatives (n = 200). Ninety-seven percent of all households expressed willingness to pay for nets; however, only 4% of those willing to pay offered spontaneously to meet the suggested price of 350 KSh (Kenyan shillings). After being prompted that “nets are expensive,” 26% of respondents expressed willingness to pay the full price. This study did not actually offer nets for sale; therefore, the number of nets actually purchased cannot be compared. However, this study did contextualize the hypothetical WTP for ITNs by comparing it to other household costs: the price of an ITN equaled the cost of sending three children to primary school for a year. By contextualizing the nets’ relative cost, the authors called into question the likelihood that families in this district, over half of whom fall below the Kenyan poverty line, would actually be able to purchase ITNs despite their “willingness to pay.”59 Given the barriers to purchasing ITNs, especially among the poorest of the poor, many researchers and development professionals involved in malaria programs called for the free distribution of nets, stressing their importance as a public health measure: “the priority for Africa should be to adopt ITNs as a public good – like childhood vaccines.”60 A great deal of progress has been made over the past 5 years, and observational studies of large, community-based programs to distribute no-cost ITNs confirm their ability to dramatically increase coverage by ITNs as well as the feasibility of combining distribution with other existing public health services such as vaccination campaigns and antenatal care clinics.61–63 The 2009 RBM report reflects the success of the change in policy: the WHO estimates that at the end of 2008, 31% of households owned ITNs, compared to 17% in 2006 and 40 kg: 1 adult tablet daily

Chloroquine phosphate (Aralen and generic)

Prophylaxis only in areas with chloroquinesensitive malaria

300 mg base (500 mg salt) orally, once/ week

5 mg/kg base (8.3 mg/kg salt) orally, once/week, up to maximum adult dose of 300 mg base

Doxycycline (many brand names and generic)

Prophylaxis in all areas

100 mg orally, daily

≥8 years of age: 2 mg/kg up to adult dose of 100 mg/day

Hydroxychloroquine sulfate (Plaquenil)

An alternative to chloroquine for prophylaxis only in areas with chloroquinesensitive malaria Prophylaxis in areas with mefloquinesensitive malaria

310 mg base (400 mg salt) orally, once/ week

5 mg/kg base (6.5 mg/ kg salt) orally, once/week, up to maximum adult dose of 310 mg base

Begin 1–2 days before travel to malarious areas. Take daily at the same time each day while in the malarious area and for   7 days after leaving such areas. Contraindicated in persons with severe renal impairment (creatinine clearance 40 mg salt/kg in the past 48 hours). For quinine there is considerable variability in apparent volume of distribution, so the full loading dose is necessary to ensure that the majority of patients do have adequate blood concentrations. In the past there was too much concern over the dangers of potential quinine toxicity and insufficient concern over the dangers of undertreatment in severe malaria. The riskto-benefit ratio changes as treatment progresses. The majority of deaths from malaria occur within the first 48 hours following admission to hospital; undertreatment may have fatal consequences. In practice, quinine toxicity in the first 24 hours of treatment is rare. The maintenance dose is 10 mg/kg every 8 hours. After 48 hours of treatment, if there is no clinical improvement in severe malaria, or if the patient is in ARF, the dose should be reduced by one-third to avoid continued accumulation to potentially toxic concentrations. The therapeutic range has not been well defined, but total plasma concentrations of between 8 and 15 mg/L are

Where Parenteral Treatment Cannot be Given In most areas of the rural tropics, severe malaria occurs far from medical attention. Since delay in starting treatment can have fatal consequences, rectal formulations of artemisinin have been developed and shown to reduce mortality. These should be given by village health workers pending transfer to a facility where parenteral treatment and supportive care can be given.127–129

Adjuvant Treatments Many adjuvant treatments have been evaluated in severe malaria, but none has proved beneficial and several were harmful. Exchange blood transfusion in severe malaria has been recommended by some on the basis of uncontrolled studies which showed benefit. There is insufficient evidence to provide a clear recommendation.

Supportive Care Patients with severe malaria require intensive nursing in an intensive care unit if possible. Following initial assessment and commencement of antimalarial treatment, clinical observations should be made as frequently as possible, including recording of vital signs, with an accurate assessment of respiratory rate and pattern, assessment of the coma score, and urine output. The blood glucose should be checked, with rapid stick tests every 4 hours if possible, particularly in unconscious patients. Convulsions should be treated promptly with anticonvulsants such as IV or rectal benzodiazepines or IM paraldehyde. Each patient’s fluid requirements should be assessed individually. Although many patients with severe malaria are dehydrated on admission and need immediate rehydration, adults with severe malaria are vulnerable to fluid overload and the physician may tread a narrow path between underhydration, and thus worsening renal impairment, and overhydration, with the risk of precipitating pulmonary edema. If the patient becomes oliguric (1 year) prophylaxis with currently recommended prophylactic drugs has not been clearly established.

Malaria Drug Prophylaxis for Residents of Malaria-Endemic Areas The routine use of malaria drugs for suppressing malaria disease lost favor during the 1970s–80s due principally to the presumption that the use of drugs on a mass basis at subtherapeutic doses would encourage drug resistance to valuable therapeutic drugs, notably chloroquine. In recent years selective use of malaria drugs in a preventive mode but at therapeutic doses (intermittent preventive treatment, or IPT) has been documented to be effective in reducing the adverse effects of malaria in pregnant women, infants, and children.

Malaria Control in Malaria-Endemic Areas During the past 5 years there has been a renewal in the global commitment to control the enormous health and economic burden that malaria extracts. Under the banner of the Roll Back Malaria (RBM) partnership, national governments and financing and technical agencies such as WHO and UNICEF have mobilized, with a focus on the enormous malaria burden in the Africa region. The malaria control landscape in mid-2010 reflects the remarkable mobilization of resources, players, commitment, and national programming action that has occurred around malaria over the past 3–5 years. Underlying this accelerated effort is a highly cost-effective set of malaria control interventions that, despite technical limitations such as drug resistance, have been demonstrated to control both malaria transmission and disease. While the treatment of malaria illness remains a constant component of malaria control, prevention has become the predominant cost-effective approach. The major challenge to the control of malaria has been the development of effective program delivery systems

Insecticide-Treated Nets Based on evidence from five randomized controlled trials and populationbased program effectiveness studies in different malaria transmission settings, ITNs can reduce the number of under-5 deaths from all causes by about 20% and clinical episodes of malaria by about half135 (Fig. 96.11). Since malaria causes approximately 20% of under-5 mortality in Africa, the protective efficacy of nets on malaria-specific mortality in children less than 5 years of age is estimated at 80%, in the range of the protective efficacy of most childhood vaccines.136,137 Recent evidence shows that ITNs protect not only those who sleep under them but also those in the same dwelling and those living nearby. Current ITNs, termed long-lasting or wash-durable, are effective in community usage for 3–5 years. ITNs are less effective in parts of Asia, where the main malaria vectors bite outdoors early in the evening.

Indoor Residual Spraying IRS has been a highly effective intervention in many parts of the world, and in particular in the Americas, in Asia, and in Southern Africa. IRS, through its house-to-house, publicly funded and managed approach, can achieve very high coverage and thus have a major impact on malaria disease burden. IRS programs are most feasible in urban areas in countries with stable malaria endemicity, especially those where there has been no tradition of managing programs of such logistic complexity.138

Intermittent Preventive Treatment IPT is the preferred approach to reduce the adverse consequences of malaria during pregnancy in high-transmission areas where P. falciparum is SP-sensitive; it involves the administration of full curative treatment doses of SP at predefined intervals during pregnancy, beginning in the second trimester after quickening. Where effective, IPT with SP reduces

Prompt and Effective Case Management of Malaria Illness The prompt recognition that a febrile illness could be Plasmodium infection is the key to limiting the risk of progression to severe or fatal disease. In most of the malaria-endemic world, malaria is self-diagnosed and treated; many episodes of fever not caused by malaria are therefore self-treated with antimalarials. Standard microscopy – and increasingly, rapid diag­ nostic tests (see section on diagnosis) in certain settings – allow for an accurate diagnosis of malaria and increase the potential that antimalarial drugs are used for the correct purpose and not simply in response to an unspecified fever episode. While drug efficacy should be the principal determinant of which drugs are used to treat malaria infections, in many settings there are many substandard and counterfeit drugs in the market place and the majority of malaria drugs come from commercial outlets without the advice of a health care provider. A major effort has been to extend the availability of rapid diagnostics and certified drugs to the community and household level, through community health workers. While this strategy should increase the promptness and effectiveness of malaria and fever management, it also raises concerns about quality control and the widescale deployment of malaria drugs and the risks of drug resistance.

Chapter 96

the incidence of severe anemia in the mother and lowers the proportion of LBW babies.139,140 There is uncertainty how to deploy IPT in the increasing areas of SP resistance and whether IPT should be deployed in low-transmission settings.

Malaria

to attain and sustain high population coverage with these robust interventions. Based on malaria epidemiology and the capacity of local health system infrastructures to deliver malaria control interventions, there are currently several distinct settings for malaria prevention and control efforts. In areas where malaria transmission is either highly seasonal or of low intensity, principally in the Indian subcontinent and Southeast Asia and most of the Americas, malaria control focuses around the prompt and effective treatment of malaria illness and selective use of antimosquito measures. In many of these areas there is a continuation of control methods developed during the malaria eradication programs of 1950–75, with the use of the indoor spraying of residual insecticides (currently limited use of DDT and more typically use of less environmentally threatening chemicals) to interrupt malaria transmission. Malaria control programming and funding have in recent years focused on developing national-level programs to stem the health and economic burden in Africa. Based on WHO and RBM guidance, the following are the key malaria control and prevention interventions advocated for the Africa region.

Program Goals for Malaria Control In the Africa region currently at least a dozen African nations are at some stage of scaling up at national level malaria control programming. There is an increasing evidence base on these national efforts. A consistent profile emerging is that in the hands of national program partnership the delivery of program to cover 60% with ITNs can happen in 2–3 years, and the national programs are achieving or exceeding the impact on under-5 mortality (20–26% reduction) documented in earlier trials of ITNs.141 The Global Fund against AIDS, Tuberculosis, and Malaria (GFATM) has in a decade become the predominant financing agency for malaria control programming. Through GFATM alone, over US$1 billion have been allocated for malaria programming since 2000.

Development of New and More Effective Malaria Control and Prevention Tools The 2007 call for malaria eradication by the Bill and Melinda Gates Foundation has energized ambitions, constructively. The global community has rallied around the call for a long-term goal of eradicating malaria. In 2008 the RBM partnership launched the Global Malaria Action Plan (GMAP), which mapped a programming sequence from scale-up to elimination.142 The GMAP will serve as a consensus platform for partners in coordinating investments, and assuring investments in research and development logically linked to programming priorities. Although there are highly efficacious interventions for malaria control and intervention, continued emphasis on the development of new and more efficacious methods is required since the evolution of drug and insecticide resistance is assumed. In the context of the GMAP, a consensus strategic plan for investments in research, termed MalERA, is being developed.143

Response to Drug Resistance and the Development of New Malaria Drugs Figure 96.11  Children sleeping under an insecticide-impregnated bednet. (Courtesy of C. Campbell.)

Since the early 1960s, there has been increasing resistance of malaria parasites to antimalarial drugs (Table 96.2). Resistance has had a profound

673

Section II PATHOGENS  

PART H: Protozoan Infections 

impact on malaria control and prevention efforts globally. The development of novel malaria drugs has received renewed attention in recent years, notably under the guidance of several multinational initiatives such as the Medicines for Malaria Venture (www.mmv.org), and the Drugs for Neglected Diseases Initiative, public–private partnerships devoted to malaria drug development. There are promising leads, but cost and the long lead time required to move efficacious compounds to licensure remain serious impediments to the control and prevention of malaria globally.

Development of Malaria Vaccines Malaria vaccines could benefit those living wherever there is malaria and travelers to those areas. The primary goal, however, must be to prevent the enormous numbers of deaths and cases of severe malaria in infants and young children in Africa caused by P. falciparum. Thus, most malaria vaccine development efforts are focused on P. falciparum. There are no commercially available vaccines for human parasites. Parasites present a greater challenge than do the viruses and bacteria for which we have vaccines because they are more complex. They have much larger genomes coding for more proteins. They have multistage life cycles in which they express many different proteins at different times (Fig. 96.12). As a result, protective immune responses against the extracellular sporozoites that enter during the bite of a mosquito may have no direct effect on the parasites that later emerge from the liver and infect red blood cells (the asexual erythrocytic-stage merozoites). The P. falciparum parasite in particular has enormous variability in its proteins. These characteristics are critical to the parasite’s survival because they enable it to evade host immune defenses. They also mean that a vaccine containing just a single sequence of a single protein, or a few proteins, may fail to have a large, sustainable impact on the disease. A truly effective malaria vaccine may need to induce both antibody and T-cell responses.

Antibodies

Antibodies

T cells Cytokines Antibodies Antibodies RBC

Antibodies Cytokines Free radicals

Figure 96.12  Schematic of the life cycle of Plasmodium falciparum with indication of which immune responses can affect which stage of the life cycle. RBC, red blood cells. (Reproduced from Hoffman SL (ed.) Malaria Vaccine

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Development: a Multi-Immune Response Approach. Washington, DC, ASM Press, 1996. Courtesy of the American Society of Microbiology.)

Antibodies have the potential to block sporozoites as they enter the body (Fig. 96.12) but have to act within minutes to block entry into the liver. They can also prevent infection of red blood cells, help destroy those that are already infected, and prevent infection of mosquitoes. T cells have the potential to kill infected liver cells, thereby controlling and even eliminating infection and potentially, through release of soluble mediators, have an impact on infected erythrocytes. Both types of response may have to be directed against multiple different proteins, at different stages of the complex parasite life cycle, and at the same time.144–147 If so, malaria vaccine developers face a technical problem that has never been solved. Scientists attempting to develop a malaria vaccine have generally kept two observations in mind. The first is that most malaria-associated deaths and severe disease in sub-Saharan Africa occur in infants, young children, and pregnant women. Nonpregnant adolescents and adults who have had multiple previous infections rarely develop severe disease or die when infected with P. falciparum. They have presumably developed natural immunity that limits parasite replication and severe forms of malaria but does not prevent infection, resulting in milder symptoms.42 Pregnant women, especially with their first child, apparently lose this immunity. This observation has led to the idea that a vaccine would be worthwhile even if it only limited the severity of disease for those most at risk without preventing infection or moderate disease. Such a vaccine would probably not be useful for travelers, and as malaria control efforts become increasingly effective in reducing severe disease and death, may be less useful for other populations in endemic areas. The second observation is that when volunteers are exposed to more than 1000 bites from P. falciparuminfected Anopheles mosquitoes that have been irradiated to weaken the sporozoites they carry, they develop protective immunity against multiple strains of P. falciparum. If these volunteers are exposed to normal sporozoites, more than 90% are completely protected against developing erythrocytic-stage infection.148 This is the strongest evidence that development of a highly effective malaria vaccine is possible, and accordingly there are efforts to develop vaccines that prevent all infections with P. falciparum in a majority (>85%) of recipients.149 During the last few years control efforts have had a profound effect on malaria in a number of areas of sub-Saharan Africa and the rest of the world. This has led to the call for elimination of malaria from defined geographic areas and eventual eradication of malaria. However, until elimination/eradication is accomplished, as the transmission intensity of P. falciparum decreases, it is likely that immunity in the population will decrease, and the susceptibility of a population to P. falciparum will increase. Thus, to prevent severe disease and death, paradoxically, more of the population may need to be immunized than has always been anticipated. While transmission will undoubtedly be interrupted in a number of malaria-endemic areas in the next decade, new tools will be needed to eliminate malaria caused by P. falciparum in the areas of the world where transmission is most intense. A pre-erythrocytic-stage vaccine that prevents transmission by preventing asexual erythrocyticstage infection will be the ideal vaccine.150 There are three main strategies by which it may be possible to achieve the preceding goals. The first is to create vaccines that counter sporozoites as they enter the body and invade and reproduce in the liver (preerythrocytic-stage vaccines). These have the potential to limit or prevent infection altogether, and thereby prevent transmission entirely. The second is to limit parasite invasion of erythrocytes and subsequent multiplication and pathologic effects (asexual erythrocytic-stage and antitoxin vaccines). Such vaccines would limit only severe disease – they would not prevent infection or mild disease, and be unlikely to have a major impact on transmission. The third strategy is to prevent the spread of viable parasites to other people with sexual and mosquito-stage vaccines. These stimulate the production of antibodies that are ingested with the parasite and destroy the parasite within the vector’s gut (Fig. 96.12), thereby reducing or preventing transmission without having any effect on disease manifestations in the immunized individual. It may be necessary to combine all three strategies to have maximum success. Current vaccine candidates in clinical trials, however, contain just one or a few proteins.

Chapter 96

reduces the rate at which malaria is acquired by 30–50%, have been initiated in multiple countries in sub-Saharan Africa. Initial trial results should be available in the next 1–2 years. In addition to the modern subunit vaccine approaches, there is a major effort to develop a nonreplicating, metabolically active whole sporozoite P. falciparum vaccine.149,157 This pre-erythrocytic-stage vaccine is intended to achieve the high level of protection demonstrated when volunteers were immunized by the bite of irradiated, P. falciparum sporozoiteinfected mosquitoes. The first such vaccine, the PfSPZ Vaccine, has been manufactured, and is now being assessed in clinical trials. It currently costs US$0.5–1 billion to bring a vaccine to market. There are large numbers of vaccine candidates in preclinical and clinical development, and many more candidates are likely to enter efficacy field trials in the next 5 years. Furthermore, emerging genomic and proteomic studies of P. falciparum will lead to the development of even more candidate vaccines.158 It will unquestionably take creative and committed public–private partnerships to bring a malaria vaccine to the populations who need it most. We look forward to the time when a malaria vaccine is widely used to successfully prevent the morbidity and mortality of P. falciparum in the 20–25 million infants born annually in sub-Saharan Africa, and to eliminate and eventually eradicate malaria.

Malaria

In contrast, the protective immune responses elicited by natural exposure to malaria or by immunization with radiation-attenuated sporozoites could be directed at many – perhaps hundreds or even thousands – of the proteins encoded by the 5300 genes in the P. falciparum genome. According to WHO (http://www.who.int/vaccine_research/links/ MaVa/en/index.html), in 2008 more than 50 subunit malaria vaccine candidates were in different stages of development. Despite these efforts only one P. falciparum protein, the P. falciparum circumsporozoite protein (PfCSP),151 has been repeatedly evaluated in clinical trials and shown to provide complete protection in a portion of volunteers. The protein was discovered in 1979 by the Nussezweig group, shown to be protective in mice in 1980, and first shown to protect humans as a vaccine in 1987. The lead candidate152 based on this protein has been RTS,S/AS02A, and it protects approximately 30–45% of nonimmune volunteers against experimental challenge with P. falciparum for 2–3 weeks.151,153 In 1–4-year-old children154 in Mozambique during 6 months, it reduced the incidence of new clinical attacks of malaria by 22.6%, new P. falciparum infection by 10.4%, and the incidence of severe disease by 58%.155 Subsequently, the adjuvant has been changed from AS02A to AS01E and it appears that this improves the protective efficacy of the vaccine.154,156 Phase III studies of this vaccine, which does not prevent malaria but

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SECTION II: PATHOGENS

PART H: Protozoan Infections

CHAPTER 97 

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Babesiosis Sam R. Telford III • Peter F. Weller • James H. Maguire

INTRODUCTION Babesiosis, a tick-borne malaria-like zoonosis, appears to have plagued humans since antiquity in its veterinary form. Human babesiosis has been reported mainly from Europe and the eastern or western United States, but cases are increasingly being reported from Asia. The prevalence of babesiosis afflicting domestic animals in tropical regions suggests that human infection is not rare in the tropics and perhaps is mistaken for falciparum malaria.1

THE AGENT The genus Babesia was named after the Romanian bacteriologist Victor Babes, who in 1888 attributed “hemoglobinuric fever” of cattle to inclusions he detected within erythrocytes. Babesia are obligately intraerythrocytic in vertebrates but, unlike plasmodia, replicate by budding rather than schizogony, and do not produce hemozoin. Currently Babesia are classified as apicomplexans of the order Piroplasmidora and family Babesiidae.2,3 Over 100 species of Babesia have been described from domestic and wild mammals on the basis of morphology and life cycle.3 Recent molecular taxonomic analyses suggest that there is even greater diversity than what has previously been recognized.4–7 The powerful new methods of molecular phylogenetics will eventually stimulate the development of a new classification for the piroplasms.

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In 1893, Theobald Smith8 described the life cycle of Babesia bigemina, the agent of bovine red-water fever, and demonstrated for the first time transmission of an infectious agent by arthropods. All piroplasms for which life cycles have been described require a tick as the definitive host (Fig. 97.1). Upon ingestion of infectious blood from a vertebrate host, babesiae undergo syngamy and replicate in the intestinal epithelium of the tick vector and develop further in the salivary glands, ovaries, and other tissues. Sporozoites in salivary glands are deposited in the skin of vertebrate hosts during the tick’s blood meal. Because of transstadial transmission, ticks infected as larvae generally remain infected as nymphs and adults. Transovarial transmission in the tick has been documented for some species such as B. bigemina but not for others such as B. microti; such a major life history difference has long been considered to represent a significant phylogenetic divergence. The common belief that sporozoites enter erythrocytes directly (no pre-erythrocytic phase) has not been critically examined. Exoerythrocytic forms, demonstrated in lymphocytes with B. equi and the closely related Theileria species,9,10 may occur in infections with B. microti.10 The process by which extracellular merozoites invade erythrocytes (induced endocytosis) is similar to that of the plasmodia. In the rat babesia, B. rodhaini, complement facilitates invasion by modification of either the erythrocyte

surface or that of the merozoite; with B. divergens, sialic acid appears to be an important ligand for erythrocyte invasion.11 Following entry into erythrocytes, pear-shaped trophozoites (piroplasms) replicate by asynchronous budding rather than by schizogony as occurs in malarial parasites. During replication, double-membraned segments develop and pinch off from the parental piroplasm, resulting in both asexually reproducing merozoites and nonreplicating sexual parasites (gametocytes).12 Asexual forms appear as simple rings, pairs, or tetrads and are difficult to distinguish from sexual stages by light microscopy.

EPIDEMIOLOGY AND DISTRIBUTION Two major epidemiologic patterns of human babesiosis are apparent. The first involves splenectomized or otherwise compromised persons and diverse babesiae, some of which have been distinguished only by molecular phylogenetic methods. A bovine-infecting species then identified as B. bovis (perhaps conspecific with B. divergens)13 was the cause of the first well-documented case of human babesiosis in 1957. A 33-year-old splenectomized farmer living in Yugoslavia and who had been exposed to cattle died as a result of hemolytic anemia, hemoglobinuria, and renal failure.14 Since then, more than two dozen sporadic cases have been reported from Ireland, Yugoslavia, France, the British Isles, Spain, the Canary Islands, Portugal, Germany, Sweden, and Russia.15 All cases were severe and often fatal despite treatment. The vector, Ixodes ricinus, also transmits Lyme disease, granulocytic ehrlichiosis, and tickborne encephalitis. Reservoirs of B. divergens, other than ticks, include cattle and perhaps deer. In 1992, a splenectomized man living in rural Missouri with an unspecified exposure history died of infection with a parasite closely related to B. divergens, designated MO-1.16 A splenectomized Kentucky resident, potentially exposed while hunting rabbits, recovered from a similar infection17 with a species whose sequenced 18S rDNA demonstrated 99.8% identity with sequences of bovine-derived B. divergens and 100% identity with an agent maintained among cottontail rabbits.18 A third case was reported in a splenectomized resident of Washington.19 Although the degree of DNA sequence similarity between the American divergens-like parasites and the European cattle and deer parasites is within that which might be expected due to geographic variation of a single species, the former fails to propagate by subinoculation into gerbils or cattle whereas the latter readily does so.20 Although European “divergens babesiosis” has heretofore been solely attributed to infection by B. divergens, molecular analysis suggests that another closely related babesia, designated EU-121 and perhaps representing B. capreoli (an agent maintained by I. ricinus among red or roe deer),22 also parasitizes humans and could be mistaken by inexperienced microscopists for B. divergens infection. Thus, it appears that “divergens babesiosis” may be due to diverse parasites that are geographically widespread.

Transfusion Transplacental 0.5–3 mm Ixodes usually nymphs

Transstadial transmission (for Ixodes spp.)*

0.5 mm larvae Late summer through winter

1– 2 mm nymphs May– July

Transovarial not documented with B. microti

vs

2– 3 mm adults Fall

B. divergens which is maintained transovarially

White-footed mice Deer maintain adult Ixodes ticks (which are competent for B. divergens, but not B. microti) *Ixodes species include I. scapularis (for B. microti), I. ricinus (for B. divergens), and probably I. granulatus and ovatus (for B. microti in Asia)

Figure 97.1  All piroplasms for which life cycles have been described require a tick as the definitive host. CMI, cell-mediated immunity.

A variation on this epidemiologic pattern has emerged along the Pacific Coast of the United States. B. duncani (previously known as WA-1) is closely related by DNA sequencing to the canine pathogen B. gibsoni.23,24 CA-type (DNA sequences designated CA-1, CA-2, etc.) agents25 are most closely related to parasites of mule deer and bighorn sheep.6 Both agents cause a similar disease and appear morphologically indistinguishable on blood smears, with abundant Maltese-cross forms within parasitized erythrocytes. Of 9 reported cases of B. duncani or CA-type babesiosis, 4 occurred among splenectomized persons, and 1 involved an apparently healthy 41-year-old man. The others were transfusion-induced cases involving a spleen-intact elderly man with multiple medical problems26 and a premature infant.27 It appears that most infections are subclinical because surveys have shown seroprevalences of 3.5–20% among persons with exposure in rural and semirural areas of California.28 As with B. divergens, splenectomized or otherwise compromised individuals appear at risk for more severe infections and cases are sporadic. The second major epidemiologic pattern is that of endemic infection due to the widely distributed rodent B. microti.29 The risk of human infection with this organism is not increased in the absence of a spleen, although infected persons without spleens are at high risk of becoming severely ill. The global distribution of B. microti parallels that of Borrelia burgdorferi and other agents transmitted by ticks in the Ixodes persulcatus species complex.30 Human babesiosis due to B. microti has been reported from Taiwan, Japan, and Germany.31–33 Additional sporadic cases of

Chapter 97

Replicate by asynchronous budding

babesiosis due to B. microti likely will be detected from Eurasia, given the wide enzootic distribution of this babesial species complex,7 greater physician awareness, and increasing availability of molecular diagnostic tools.34 In the United States, B. microti babesiosis largely remains limited to the terminal moraine islands of New York, Massachusetts, and Rhode Island, and focal areas in Massachusetts, Connecticut, New Jersey, Wisconsin, and Minnesota.35 Because most cases are asymptomatic or mildly symptomatic, the incidence is difficult to estimate and differs from year to year due to fluctuations in the densities of the rodent reservoir and tick vectors. While there have been only several hundred reported cases of symptomatic infection with B. microti since the first reported case in 1969, serologic surveys indicate that in highly endemic areas as many as 9–21% of persons have been infected.36 The incidence of babesiosis has been rising in parts of southern New England since the 1990s, and on Block Island, Rhode Island, there may be as many as 900 cases per 100 000 residents per year.37 There have been at least 70 cases of babesiosis acquired from blood products from asymptomatically infected donors,38 most commonly packed erythrocytes, but also platelets and frozen-deglycerolized erythrocytes and 1 case of vertically transmitted infection.39 Few cases of babesiosis have been reported outside the United States and Europe, but this may be changing due to the wider diagnostic use of polymerase chain reaction (PCR). There have been case reports demonstrating active infection (a photomicrograph of a blood smear) in China,40 India,41 and South Africa,42 and serosurveys suggesting human babesial infection in Latin America and West Africa. An unusual case of babesiosis due to a Babesia sp. most closely related to those that infect sheep was reported from a 75-year-old, splenectomized female Korean resident who was treated and survived.43 Because of the ubiquity of ticks and the diversity of Babesia species and animal hosts, it is likely that transmission to human beings occurs wherever humans are greatly exposed to ticks.

Babesiosis

Anemia, hemoglobinemia/uria, jaundice; severe if splenectomy, elderly or depressed CMI

Seasonality and Ecology Human B. divergens cases occur mainly in cattle-raising regions during summer months when the presumed vector, I. ricinus, is most active and the incidence of red-water fever in cattle is greatest.15 The only known reservoir hosts for B. divergens are cattle and reindeer, although deer are also suspected as potential hosts. The American B. divergens-like agent (MO-1) is most likely transmitted by I. dentatus.18 Although long thought to be specific for rabbits and birds, this tick appears to feed on humans more frequently than appreciated. B. capreoli (EU-1) is maintained by I. ricinus, probably with roe and red deer as amplifying hosts.22 The few reported cases of B. duncani and CA-type babesiosis had their onset between June and August. Vectors and reservoirs remain to be described although canids, ungulates, and their associated ticks are suspected given the genetic relatedness of these agents to B. gibsoni and the mule deer or bighorn sheep babesias, respectively.6 Vectors for KO-1 or other agents reported from India or South Africa remain undescribed. In contrast, the ecology of B. microti has been well studied.44 Its enzootic cycle in the northern United States depends on the interaction of subadult deer ticks, I. dammini (thought by some to be an aggressively human-biting variant of I. scapularis) and their main source of blood meals, the white-footed mouse. Deer are the hosts on which adult ticks primarily feed, but are incompetent reservoirs. Adult ticks feed during the fall, and lay eggs during the spring. Eggs hatch in late July, and emergent larvae feed mainly during August and September, at which time they may acquire babesial infection from an infected mouse. Fed larvae overwinter and molt to the nymphal stage during the spring. The life cycle of the tick is complete when nymphs that have fed on a mouse or other host during the summer molt to the adult stage in the fall. Humans become infected primarily by the nymphal ticks (Fig. 97.2) rather than adults, because survival of B. microti during the nymphal to adult tick molt is poor.45 In addition, people are likely to discover the larger adult deer tick (albeit smaller than the dog tick) and remove it before the tick has fed long enough (>60 hours for B. microti) to deliver

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PART H: Protozoan Infections 

A

B

Figure 97.2  Ixodes dammini (deer tick) vector of Babesia microti. Ixodes ricinus, the European vector of B. divergens, is morphologically very similar. (A) Berlese fluid-cleared nymphal tick demonstrating heavily sclerotized legs and articulations as well as capitulum (“head”). The backward-facing spikes (denticles) of the mouth parts (hypostome) can be seen; this structure helps to anchor the tick and is the basis for the difficulty with which ticks can be removed from the skin. (B) Two nymphal deer ticks with a millimeter scale. Most babesiosis cases do not remember the infecting tick bite. Note that in living specimens of unfed or partially fed nymphs, prominent dark gut diverticula are seen through the body wall, giving the impression of an external color pattern.

an infectious inoculum of sporozoites. Accordingly, in >80% of reported cases, onset of illness occurred between May and August, when nymphs are most abundant.46 In Taiwan and Japan, zoonotic vectors of B. microti remain to be described, although I. granulatus and I. ovatus are probably enzootic vectors. Various small rodents (Apodemus spp., Rattus spp.) are frequently parasitized.47,48 In the German case of B. microti babesiosis, acquired from a transfusion, the likely vector is I. ricinus. B. microti appears to be part of a characteristic microbial assemblage with the agents of Lyme disease, granulocytic ehrlichiosis, and tick-borne encephalitis virus throughout the Holarctic.30,49

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Lysis of parasitized erythrocytes leads to anemia, hemoglobinemia, hyperbilirubinemia, and, in severe cases, intense jaundice, massive hemoglobinuria, and renal shutdown. Electron microscopy shows parasite-mediated damage to erythrocyte membranes with perforations, protrusions, and inclusions.50,51 There is reduced deformability of infected erythrocytes that enhances their removal by the spleen. In experimental B. bovis infections, increased peroxidation of membrane lipids appears to be responsible for a further decrease in erythrocyte survival and adherence of infected erythrocytes to the microvascular endothelium. However, endothelial adherence was not observed in tissues of a B. microti fatality.52 In several European cases, autopsies showed parasites in erythrocytes in congested capillaries of various organs, especially in hepatic sinusoids.53,54 Other findings in these cases and in fatal cases due to B. microti suggest a cytokine-mediated shock syndrome. Indeed, laboratory studies of rodents infected by B. microti or B. duncani demonstrate overproduction of the proinflammatory mediators tumor necrosis factor-α (TNF-α) and interferon-γ.55 Mice with a genetic disruption in the TNF-α pathway were less likely to die of fulminating B. duncani infection, as were CD4 and CD8 gene knockout mice, whereas in γδ T-cell knockout mice and control mice B. duncani infection terminated fatally. Thus, CD8+ T cells may contribute to B. duncani-induced pathology.55 In addition, depletion of macrophages and natural killer cells seems to increase susceptibility.56 In humans, acute tubular necrosis, ischemic necrosis of liver, spleen, pancreas, and heart, noncardiac pulmonary edema, and swelling and congestion of the brain and other organs have been demonstrated, even in cases in which parasites had been cleared by treatment.57 Other findings at autopsy have included hemophagocytosis, hypercellularity of the bone marrow, extramedullary hematopoiesis, and hemosiderin deposits

in the Kupffer cells and the kidneys, which correspond to findings in histopathologic studies of rodent infections.58,59 Asplenia, advanced age, and depressed cellular immunity are associated with severe clinical illness.60,61 The spleen phagocytizes parasitized erythrocytes, thus limiting parasitemia, particularly early in infection. Nevertheless, persons with intact spleens have died of overwhelming infections, and persons without spleens have recovered from babesiosis, even without specific therapy.61 Severity of illness is greater in persons older than 50 years than in younger adults, and overt illness in children seems unusual.36 As a rule, younger persons who become ill are asplenic or immunocompromised or have other underlying medical conditions.62 B. microti infections may be severe in persons who are infected with human immunodeficiency virus-1 (HIV) or are receiving corticosteroids or other immunosuppressive therapy, particularly rituximab (anti-CD20 B-cell monoclonal antibody).63 Similarly, hamsters that receive antilymphocyte serum and athymic mice experience high parasitemias and mortality when exposed to B. microti.64,65 CD4 gene knockout mice sustained B. microti parasitemias for longer periods of time than did congeneic controls.55 Humoral immunity appears to be less important than cellular immunity in controlling infection. Passive transfer of immune serum to immunodeficient severe combined immunodeficient (SCID) and nude mice fails to protect them from B. microti infection.66 B-cell-deficient mice remain less susceptible to B. microti infection, whereas T-cell receptor-deficient mice are readily infected.67 Whether sterilizing immunity develops is questionable. Human infection may last over a year, even in the absence of underlying illness.68 Reinfection has not been reported and would be difficult to distinguish from recrudescence of an earlier infection.

THE DISEASE Depending on the species of Babesia and host factors, infection can be subclinical, cause a self-limited febrile illness, produce a moderate to severe illness resembling malaria, or progress rapidly to death. Infection with B. divergens occurs almost always in splenectomized persons and runs a fulminant, usually fatal, course without treatment.15 After an incubation period of 1–4 weeks, B. divergens-infected patients become acutely ill with high fevers, prostration, rigors, diaphoresis, headache, myalgia, jaundice, and hemogloblinuria. Nausea, vomiting, and diarrhea are prominent, and the liver may be enlarged and painful. Most patients develop acute respiratory distress. Renal failure induced by intravascular hemolysis and hypotension ensues and is followed by coma and death, usually within a week of onset of symptoms. The 3 patients with the American B. divergenslike infection, 1 of whom died, presented with fever, headache, and rigors; thrombocytopenia, hemoglobinuria, and proteinuria developed. Most infections with B. microti and likely also with B. duncani or CA-1 are subclinical. Subclinical infections have been detected during sero­ surveys and investigation of blood donors following transfusion-induced cases.26,27,69 Animal inoculation has confirmed viable parasites in smearnegative asymptomatic persons with positive serologic tests for antibodies to Babesia.27,36 Subpatent parasitemia in the absence of symptoms may last over a year.70 Because of the small number of reported cases, the spectrum of illness among persons with B. duncani or CA-type infections remains to be defined. Four of 7 persons with symptomatic infections had been splenectomized, including 2 patients who had severe disease with high parasitemias and multisystem organ failure, 1 of whom died.25 The index case, a healthy, spleen-intact man, had a moderately severe illness with a 3% parasitemia, high fevers, and a slow recovery.23 A similar illness occurred in an elderly man with multiple medical problems who acquired infection from a blood transfusion.26 Persons with symptomatic B. microti infections typically experience an influenza-like illness 1–4 weeks after a tick bite or 4–9 weeks following transfusion. There is a gradual onset of malaise, anorexia, and fatigue followed within a week by sustained or intermittent fever as high as 40°C, drenching sweats, and myalgia.69 Nausea, vomiting, headache, shaking

DIAGNOSIS The nonspecific symptoms and absence of a history of a tick bite in most cases preclude the ability to make a diagnosis of babesiosis on clinical grounds alone. The definitive diagnosis usually follows demonstration of organisms parasitizing erythrocytes on conventional Giemsa-stained thin films. The small nucleus of Babesia can be difficult to identify on thick films, particularly when parasitemia is sparse. As with malaria, sequential blood smears may be required to detect organisms when the level of parasitemia is low. Babesia is distinguished on blood smear from Plasmodium falciparum by a combination of criteria, including the demonstration of basket-shaped and frequently extracellular merozoites (Fig. 97.3) with white or lightcolored cytoplasm, erythrocytes containing two or more parasites, and the presence of tetrad forms (Maltese crosses). Tetrad forms, however, are rarely encountered with B. microti infection, in contrast to acute B. divergens (Fig. 97.4), B. duncani, and MO-1 (Fig. 97.5) infections, in which

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Babesiosis

chills, dark urine, emotional lability and depression are not uncommon.71 Physical examination may show mild splenomegaly and, less often, hepatomegaly. Most patients have mild to moderate anemia, thrombocytopenia, a normal or depressed white blood cell count, mildly elevated hepatic enzymes, and evidence of hemolysis, including hyper­bilirubinemia, elevated serum lactate dehydrogenase, and decreased haptoglobin levels. Parasitemias may range from 1% to 20% in spleen-intact persons. After several days to weeks, fever and more intense symptoms may resolve spontaneously, but weakness, malaise, and fatigue persist for months.71 In other persons, largely the elderly, those without spleens, and those who are immunosuppressed or have other underlying medical problems, illness can be severe with intense hemolysis leading to jaundice, severe anemia, and renal failure. Parasitemias can reach 85% in asplenic patients.50 Severe pancytopenia in some cases is due to hemophagocytosis.72 Disseminated intravascular coagulation, hypotension, and adult respiratory distress syndrome have been seen in fatal cases. Noncardiac pulmonary edema, at times requiring mechanical ventilation, may occur in persons with less severe illness several days after beginning therapy. No relationships are apparent with parasitemia, splenectomy antibody or immune status; onset may be either early or late in the infection; and pulmonary edema resolves with supportive care.73 In an early series of 136 patients with babesiosis due to B. microti, mortality was 5% despite treatment.60 Babesiosis in persons with acquired immunodeficiency syndrome (AIDS) is characterized by a prolonged duration and frequent relapses. There have been five reports of B. microti infection in persons with advanced HIV infection.63,74–77 Two splenectomized persons had parasitemias in excess of 40% and more severe illness than the 3 who were spleen-intact, who had up to 20% parasitemia. Recrudescences occurred in 4 patients and lasted as long as 400 days after the initial episode; further recurrences were prevented by continual administration of antimicrobial agents. Borrelia burgdorferi (see Chapter 44) or the agent of human granulocytic ehrlichiosis (Anaplasma phagocytophilum) (see Chapter 52) may coinfect ticks with Babesia microti and may be concurrently transmitted.78,79 Persons with B. microti infection who are coinfected with Borrelia burgdorferi experience more symptoms and longer duration of illness than persons infected with either organism alone.80 In a fatal case of a man with coexistent babesiosis and Lyme disease, the cause of death appeared to be related to pancarditis caused by the Lyme disease spirochete.81 Another man with coinfection developed a more severe case of transverse myelitis than that which is seen with Lyme disease alone.82 Coinfection with B. microti and Ehrlichia chaffeensis led to multiorgan failure and death in an 85-year-old man.83 The differential diagnosis for babesiosis includes other tick-borne diseases, including Lyme disease, ehrlichiosis, typhoidal tularemia, and Rocky Mountain spotted fever, all of which may be endemic in the same region, as occurs in New England. Other illnesses that may be confused with babesiosis include viral hepatitis, bacterial sepsis, infectious mononucleosis, leptospirosis, malaria, and relapsing fever.

B

Figure 97.3  Babesia microti, human infection, Nantucket Island. Pleomorphic parasites with high parasitemia, multiple parasite infection of single cells, and range of developmental forms from newly invaded merozoites to mature trophozoites. A, mature trophozoite with prominent dark chromatin that extends nearly around the circumference of the cell; whitish cytoplasm; and threadlike band of chromatin across the middle of the parasite cytoplasm. B, mature trophozoite with prominent chromatin dot; white cytoplasm, and threadlike chromatin band across the cytoplasm. C, five small merozoites, perhaps comprising four daughter cells from one tetrad, with an additional newly invaded merozoite. D, extracellular merozoite. In heavy infections clumps of extracellular parasites may be observed; the presence of a discrete nucleus for each such parasite distinguishes them from platelets.

they are frequent. The absence of hemozoin (malarial pigment) is considered diagnostic for the piroplasms, but early ring stages of the plasmodia also lack pigment. The presence of schizonts, gametocytes, or erythrocyte stippling may be characteristic of malaria but not of Babesia. B. divergens and related infections may be identified by the presence of accole forms and paired divergent pyriforms occupying no more than 20–25% of the erythrocyte area, along with small single oval or round merozoites; high parasitemias are often apparent. Although it is difficult to identify Babesia spp. specifically by morphology inasmuch as host cell factors influence this, there are discriminating features for mature infections. Note that for all species, simple ring forms may be all that is observed on a thin blood smear. Serologic testing is useful, particularly in diagnosing chronic B. microti infections in which the parasitemia is subpatent. The indirect fluorescent antibody test using antigen derived from infected hamster erythrocytes84 is sensitive and specific, and is currently the serologic method of choice. In cases in which parasitemia is difficult to detect, detection of specific IgM confirms a clinical diagnosis of babesiosis.85 Absence or low titers of specific antibodies against B. microti when blood smears contain parasites suggests an infection by another Babesia sp. or an immunocompromised patient (splenectomy, HIV, recent infusion of anti-B-cell antibody). The sensitivity and specificity of serology for B. duncani are not known. Because specific antibodies do not become detectable until at least 1 week after onset of illness, serology is not reliable for diagnosis of the rapidly fulminating B. divergens babesiosis.15 Serology for divergens-like babesiosis (EU-1, MO-1) may be accomplished by the use of B. divergens antigen because these parasites are cross-reactive. Inoculation of a sample of patient blood into hamsters or SCID mice facilitates diagnosis of B. microti or B. duncani infection when smears are

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TREATMENT AND PROGNOSIS B

 

PART H: Protozoan Infections  680

negative. Approximately 300 parasites are sufficient to induce persistent parasitemia, which becomes detectable 1–6 weeks following inoculation. B. divergens can be isolated by inoculation of gerbils, but this procedure is only retrospectively useful for confirming the diagnosis of this rapidly progressive infection. PCR-based assays, which can detect Babesia DNA corresponding to three parasites in a 100-µL sample of blood, have the advantage of yielding a diagnosis in less than a day.86 In addition, sequencing of the amplification products may provide more rapid specific identification than immunological analysis or animal inoculation studies when the identity of the infecting parasite is questioned.

C

Figure 97.4  Babesia divergens, gerbil. The chromatin of this parasite usually appears thick and well stained. A, B, classic paired pyriforms. C, mature trophozoite with prominently staining chromatin that extends around the circumference of the parasite. Compared with B. microti and B. duncani, B. divergens-like parasites tend to produce two pairs of daughter cells as opposed   to a tetrad.

Figure 97.5  Babesia duncani, hamster. The classic Maltese-cross form is more frequently seen in B. duncani, with daughter cells typically a third of the diameter of the erythrocyte. In contrast,   B. microti tetrads have “short leaves” with daughter cells shorter and more compact. B. divergens-like infections in the United States demonstrate a mixture of paired pyriforms with classic Maltese crosses.

The combination of quinine (650 mg orally tid and clindamycin, either 1.2 g intravenously bid or 600 mg orally tid) can be used to treat babesiosis due to all species.87,88 Pediatric doses are quinine 30 mg/kg/day and clindamycin 20–40 mg/kg/day, both in three divided doses.87 Treatment should be continued for at least 7 days or until parasitemia remits. Evidence of excellent in vitro efficacy and the successful resolution of the index case for KO-1 babesiosis suggests that clindamycin alone warrants further study as an effective treatment for babesiosis.43,89,90 Because B. divergens parasitemias increase very rapidly, any case of babesiosis acquired in Europe should be regarded as an emergency requiring prompt treatment and frequent monitoring of blood smears.91 Exchange transfusion should be considered in severely ill patients with parasitemias in excess of 10% and evidence of severe hemolysis or organ failure.92 In particularly severe cases of B. divergens and B. microti babesiosis, complete blood exchange transfusion (2–3 blood volumes) should be undertaken, followed by treatment with clindamycin and quinine.91 Given that patients may be disproportionately ill even with virtually undetectable parasitemia, a one-log reduction of preprocedure parasitemia, rather than a targeted universal recommendation for posttransfusion parasitemia, has been suggested.93 Babesiosis due to B. duncani appears to respond to the combination of quinine and clindamycin, but the small number of cases that have been treated to date does not permit strong conclusions about the efficacy of this regimen. Because illness due to B. microti is often mild and short-lived, not all cases require treatment, with the caveat that such patients be deferred from donating blood. For persons with B. microti infections that are not life-threatening, the combination of atovaquone (750 mg PO bid × 7–10 days) and azithromycin (600 mg PO daily × 7–10 days) may be used.87 A prospective randomized trial demonstrated that patients treated with atovaquone and azithromycin cleared parasitemia as effectively as did those receiving clindamycin and quinine, and with fewer side effects.94 Combination therapy including all four of these drugs has been used as an alternative for treating human babesiosis when other regimens have failed or patients have developed side effects while receiving standard therapy.63 Quinine and clindamycin should be given if the symptoms are sufficiently severe or if the patient is elderly, splenectomized, or immunocompromised. Treatment may occasionally fail, especially in such highrisk patients, or in those who must discontinue quinine due to side effects such as severe tinnitus. In most patients who complete the full regimen, parasite DNA seems to become undetectable by PCR within a month.68 In a minority of cases, particularly in immunocom­promised patients, symptoms may resolve, but parasitemia persists.63 Following treatment, patients with AIDS should remain on suppressive therapy to prevent relapses.63,95 The ideal regimen and duration of suppression have not been determined, but combinations of quinine, clindamycin, and other drugs have been used for periods of 6 months or maintained for life. A variety of other agents, including chloroquine, co-trimoxazole, and pentamidine, have been tried without success for treating infected humans. Doxycycline in combination with other agents appears to have efficacy in refractory cases in persons with AIDS,63 but patients who are treated with a tetracycline for Lyme disease appear to require specific antibabesial treatment if they are coinfected (T.J. Lepore, personal communication).

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Chapter 97

Repellents or arthropod toxicants are useful for personal protection. Permethrin-based formulations applied to shoes, socks, and pants are particularly active against ticks although ticks may attach and feed briefly before succumbing to permethrin intoxication. Common diethyltoluamide (DEET)-based repellents will also repel most ticks, but may require frequent reapplication. Preventing ticks from crawling underneath clothing by simply tucking pants cuffs into socks reduces the risk. Because

delivery of an infectious inoculum usually requires more than a day of tick attachment (>60 hours for B. microti), the body surface of persons who have visited a site where transmission is intense should be examined daily and all ticks should be removed promptly with forceps. Various public health strategies may be followed to protect human populations against zoonotic babesial infection. Acaricidal spraying of vegetation, host-targeted acaricides, and reduction of the reproductive hosts for the vector ticks (mainly deer) are all effective community-level interventions.96–98

Babesiosis

PREVENTION AND CONTROL

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PART H: Protozoan Infections

CHAPTER 98 

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African Trypanosomiasis (Sleeping Sickness) Jacques Pépin • John E. Donelson

INTRODUCTION Human African trypanosomiasis (HAT), or sleeping sickness, is a purely African disease caused by two morphologically identical subspecies of trypanosomes – Trypanosoma brucei gambiense and Trypanosoma brucei rhodesiense – transmitted to humans by tsetse flies. Clinically, T. b. gambiense HAT is characterized by an early stage during which trypanosomes are found in the blood or lymph node aspirates of mostly asymptomatic persons and by a late stage during which there is involvement of the central nervous system (CNS) with somnolence, other neurologic symptoms, and trypanosomes in the cerebrospinal fluid (CSF). T. b. rhodesiense HAT, which is sometimes seen in short-term visitors to eastern and southern Africa, is a much more acute febrile illness occurring within days of the infective bite and which, if untreated, can be fatal in a matter of weeks.

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T. b. gambiense and T. b. rhodesiense are morphologically identical to a third salivarian trypanosome, Trypanosoma brucei brucei, which infects domestic and wild animals but does not survive in humans because it is lysed by specific components in the high-density lipoprotein fraction of human serum.1,2 This group of three subspecies is often referred to as the T. brucei complex. Several other African trypanosome species are important pathogens of domestic livestock and wildlife in Africa but do not infect humans and are not discussed here. The genus Trypanosoma occurs within the order Kinetoplastida. The single large mitochondrion of these organisms contains an appendage called the kinetoplast that lies in close proximity to the basal body at the base of the flagellum (Fig. 98.1). In the T. brucei complex, the kinetoplast contains as much as 10% of the total DNA of the organism. This DNA is organized into homogeneous maxicircle DNA molecules (about 20 000 base pairs each), which are equivalent to mitochondrial DNAs of other organisms, and heterogeneous minicircle DNA molecules (about 1000 base pairs each). A unique feature of organisms in the order Kinetoplastida is that some of their kinetoplast DNA genes are transcribed into RNAs that must be extensively edited by insertions or deletions of uridine nucleotides at many locations along the messenger RNA (mRNA) molecule before the correct mitochondrial proteins can be synthesized.3 In the T. brucei complex, as many as 50% of nucleotides in some kinetoplast mRNAs are added or deleted after the initial synthesis of the RNA molecule. The reasons for the existence of this kinetoplast RNA editing and, indeed, the need for the kinetoplast itself are unknown, but this distinctive property of trypanosomes and related organisms offers an attractive target for future drug development. The sequence of the 35 million basepair DNA genome in the nucleus of T. b. brucei has been determined4 and also provides the foundation for identification of unique trypanosome enzymes and metabolic pathways that are potential new drug targets.

During their life cycle members of the T. brucei complex cycle between the mammalian bloodstream and several species of tsetse flies of the genus Glossina. When a tsetse fly bites an infected person, or an animal in the case of T. b. rhodesiense, trypanosomes in the bloodstream can be ingested with the blood meal (Fig. 98.2). In the fly, the infected blood moves to the lumen of the midgut, where the bloodstream trypanosomes transform during a 2- to 3-day period into the procyclic stage. This transformation is accompanied by many developmental changes including an increase in body length, a switch from anaerobic to aerobic metabolism, and a change in the main nutritional carbon source from glucose to proline, which is also used as the predominant energy source by the tsetse fly itself during flight.5,6 In addition, the major protein on the surface of bloodstream trypanosomes, the variant surface glycoprotein (VSG), is replaced by an invariant surface protein called procyclin. After 2–3 weeks of multiplication in the midgut, the procyclic trypanosomes migrate to the salivary gland, where they undergo several additional developmental changes, culminating in the formation of the mature metacyclic stage. Metacyclic trypanosomes reacquire a VSG coat and stop multiplying. An infected tsetse fly can harbor as many as 10 000 to 20 000 metacyclic trypanosomes, of which one is potentially sufficient to initiate the mammalian infection if transmitted during the fly bite. Each trypanosome of the mature metacyclic population is completely covered with about 107 copies of a single VSG, but the population as a whole expresses 10–15 different metacyclic VSGs.7 Moreover, one study showed that these metacyclic VSGs gradually diverged over a 20-year period,8 indicating that a vaccine directed against the metacyclic VSGs would not be successful. The tsetse fly injects the metacyclic trypanosomes into the connective tissue of the skin, where a temporary local inflammation, called a chancre, often develops. From this initial portal of entry, parasites enter the draining lymphatics, pass into the bloodstream through the thoracic duct, begin to multiply by binary fission and transform back into the bloodstream form, returning to anaerobic glycolysis as the main source of adenosine triphosphate (ATP). They continue, however, to express the metacyclic VSGs on their surface for about 5 days after the initiation of infection. They then switch from the expression of metacyclic VSGs to the bloodstream VSGs, one of which is sometimes the same VSG as on the surface of trypanosomes ingested by the fly.9 The slender bloodstream form of the parasite actively divides every 5–10 hours, whereas the shorter, stumpy form does not divide but has a more developed mitochondrion and is thought to be more infective to the insect. These bloodstream forms can traverse the walls of the blood and lymph capillaries to the connective tissues and eventually enter the CSF and the brain. The parasite’s life cycle is completed when a tsetse fly takes up the bloodstream forms while feeding on an infected mammal (Fig. 98.2). Extracellular trypanosomes, in constant contact with their host’s immune system, have evolved sophisticated mechanisms to evade immune attack. The VSG is a crucial component of one of these evasion

VSG coat Glycosome

Kinetoplast Flagellar pocket A

EPIDEMIOLOGY

B

Figure 98.1  (A) Schematic diagram of a bloodstream trypanosome showing some of its subcellular organelles and the location of the variant surface glycoprotein (VSG). (B) Scanning electron micrograph of a bloodstream Trypanosoma brucei rhodesiense adjacent to a rat red blood cell (×5500).

mechanisms, a phenomenon called antigenic variation. As early as 1910, HAT was reported to be characterized by successive waves of parasites in the blood.10 It is now well known that these peaks of parasitemia are due to trypanosomes expressing antigenically different VSGs.11 The nuclear DNA of the T. brucei complex contains as many as 1000 different VSG genes.4 Usually each trypanosome expresses one, and only one, of these VSGs at a given instant. The 107 copies of a VSG on a trypanosome represent about 5% of the total protein of the organism. The major function of all this protein on the surface of the living trypanosome is to serve as a barrier to protect other invariant constituents of the organism’s outer membrane from attack by the immune system. The three-dimensional structures of two different VSGs have been shown by X-ray crystallography to be cylindrical, allowing them to pack very closely together on the parasite’s surface. During infection, antibodies are continually raised against the VSG, but trypanosome populations manage to escape total destruction because individual parasites occasionally undergo antigenic variation by switching spontaneously from the expression of one VSG to another. New antibodies must be directed against the VSG of the switched parasite and its descendants, enabling the population as a whole to stay one step ahead of this humoral (B-cell) immune response. The switch rate of the bloodstream trypanosome’s antigenic variation ranges between 10−2 and 10−6 per parasite per doubling time.12 The switch itself is often associated with spontaneous VSG gene rearrangements that maneuver duplicated copies of silent VSG genes into special gene expression sites located near the ends of some chromosomes, where they are transcribed into mRNA. Sometimes these rearrangements result in the formation of new VSG genes that are mosaic or mutated versions of preexisting genes.13,14 This ability to create new VSG gene versions during the DNA rearrangements likely means that bloodstream trypanosomes have the potential to express sequentially a much larger repertoire of VSG proteins than they have VSG genes. Hence, the

HAT is endemic only in sub-Saharan Africa between latitudes 15° N and 15° S, corresponding to the distribution of its vector.15,16 Its incidence increased considerably at the beginning of the twentieth century as a consequence of population migrations brought on by colonization. In some areas HAT caused half of the overall mortality. Trypanosomiasis control was organized around mobile case-finding teams that regularly examined the population of endemic foci through cervical lymph node palpation and aspiration to detect and then treat infected but asymptomatic persons. The goal was to reduce transmission by shortening the duration of infectiousness. In the late 1940s and 1950s, millions of people were given pentamidine “chemoprophylaxis” twice a year, but it probably corresponded more to early treatment than to prevention. Such initiatives were highly successful, and the incidence of HAT decreased dramatically. After the end of colonial rule, HAT remained at a low level or disappeared in countries able to maintain some case-finding activities, but increased considerably when such programs were interrupted for just a few years at the end of the twentieth century. As a result, the distribution of sleeping sickness ten years ago paralleled that of wars or civil conflicts that devastated parts of Africa, the highest incidences occurring in the Democratic Republic of Congo (DRC), Angola, and Sudan. Once more, substantial investment into its control proved successful: in 2006, a total of 11 382 cases of T. b. gambiense trypanosomiasis were reported to the World Health Organization (WHO), down from 37 385 cases in 1999.17 T. b. rhodesiense trypanosomiasis is much less common, with about 500 cases reported annually.17 Even though there is some degree of underdiagnosis and underreporting,18 the burden of HAT is now currently less than what was estimated in 2003 (48 000 deaths per year and 1.5 million disability-adjusted life-years lost).19 Gambian HAT is endemic in western and central Africa and Rhodesian HAT in eastern and southern Africa (Fig. 98.3). Uganda is the only country in which both subspecies are present. In West Africa, the disease has disappeared from Senegal, The Gambia, Guinea-Bissau, Sierra Leone, and Ghana for ecologic reasons that are poorly understood, since significant tsetse fly populations remain. A few dozen cases are diagnosed each year in Ivory Coast, Guinea, and Nigeria. The country with by far the highest incidence is the DRC, where 8023 cases were reported in 2006, followed by Angola (1105 cases), Sudan (809 cases) and the Central African Republic (460 cases). Within the DRC, the epidemic reached Kinshasa, where almost 1000 cases were diagnosed in 1999, mostly acquired in its peri-urban areas.20 In southeast Uganda, an epidemic of Rhodesian HAT has been successfully controlled by tsetse fly trapping and case-finding, but Gambian HAT persists in northwest Uganda. In 2006, 245 cases of Rhodesian HAT were reported in Uganda, 125 in Tanzania, 58 in Malawi, 57 in Zambia, and only a handful in Kenya and other countries of the region.17 Each year, ~30 cases of Rhodesian HAT are diagnosed in tourists, especially visitors to the Serengeti,21 where this risk has now been reduced through

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Mitochondrion

African Trypanosomiasis (Sleeping Sickness)

Flagellum

development of a HAT vaccine directed against bloodstream VSGs is also unlikely. The tsetse fly vector belongs to the Glossina genus, found only in Africa, which contains 22 species, only a few of them being involved in the transmission of HAT: the riverine palpalis group (G. palpalis, G. tachinoides, and G. fuscipes) for T. b. gambiense, and the morsitans group (G. morsitans, G. pallidipes, and G. swynnertoni) for T. b. rhodesiense. Tsetse flies live in hot, humid, and dark ecologic niches, and their distinct requirements relate to epidemiologic differences between Gambian and Rhodesian HAT. The palpalis group is found mostly in forests on the banks of a river and humans are a variable source (10–40%) of their blood meals, while the more zoophilic morsitans group is seen in the woodland and thickets of East African savannas. Once infected with trypanosomes, tsetse flies remain infected for the rest of their lives (a few months). Less than 5% are infected in epidemic areas.

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Trypanosomes mature, divide in blood and lymph and eventually invade CNS causing: Intermittent fever and rash Posterior cervical adenopathy (Winterbottom’s sign) Wasting Meningoencephalitis (including obtundation) (early with T. b. rhodesiense, after months to years with T. b. gambiense) Trypanosomes at bite site may cause a:

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African sleeping sickness Trypanosoma brucei gambiense/rhodesiense

Painless trypanosomal chancre

Metacyclic trypanosomes from tsetse fly (Glossina) salivary gland enter bite site

Bloodstream trypanosomes taken with blood meal by tsetse fly

Procyclic trypanosomes develop into epimastigotes and metacyclic trypanosomes in tsetse salivary gland

Figure 98.2  Life cycle of Trypanosoma brucei gambiense/rhodesiense infections (African Sleeping Sickness).

0°

T.b. gambiense

T.b. rhodesiense

Figure 98.3  Distribution of African trypanosomiasis. (Redrawn from World Health Organization. Control and surveillance of African trypanosomiasis. Report of a WHO expert committee. World Health Organization Technical Report Series No. 881, 1998.)

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vector control. About 20 cases of Gambian HAT are diagnosed annually in Africans who have migrated outside the endemic areas.22 The major determinants of the epidemiology of Gambian HAT are (1) the long duration (up to several years) of infection in human hosts, with cycles of intermittent parasitemia; (2) human–fly contact and infection rates among tsetse flies; and (3) the efficacy of passive and active casefinding.15 Human–fly contact can be enhanced by changes in tsetse fly

density, distribution, and feeding habits or in human behavior that results in people spending more time in areas of high fly density. Breakdowns in active case-finding and, to a lesser extent, in passive detection by multipurpose health facilities allow the human reservoir to expand and increase the percentage of infectious tsetse flies. Significant immunity appears after an adequately treated first episode of Gambian HAT,23 which might explain the resurgence of HAT in the same foci as 50 years earlier. After years of high incidence and successful treatment of most cases, a large proportion of the population becomes immune and decades are necessary before the pool of immunologically naïve persons is large enough to sustain an epidemic. Familial clustering is also observed, the risk of HAT in a child being higher if the mother (but not the father) had had HAT before, presumably reflecting shared exposure or congenital infections rather than a genetically determined susceptibility.24 There seems to be little or no interaction between HAT and human immunodeficiency virus (HIV) infection.25 Because of the long incubation period, there is no seasonal variation in Gambian HAT incidence. Domestic animals (pigs, goats, dogs, sheep, cattle, and even chickens) can be infected with T. b. gambiense. The importance of these animal reservoirs is probably marginal in high-incidence countries of central Africa, but they might play a role in the persistence of the disease in lowincidence countries, where the size of the human reservoir is much smaller.15 In contrast, Rhodesian HAT is a zoonosis and to some extent an occupational disease, with many species of game animals and cattle harboring the parasite and sustaining sporadic transmission to humans.15 Person-to-person spread occurs only during epidemics, but this subspecies offers less potential for large-scale epidemics because of its acute nature. Changes in agricultural practices and movements of cattle have been incriminated as favoring the spread of the parasite.26

PATHOGENESIS AND IMMUNITY The immune response and pathogenesis that develop during HAT are complex and poorly understood. From an immunologic perspective, a trypanosome may be regarded as a package of thousands of nonvariable antigens surrounded by 10 million copies of a variable antigen, the VSG.5 Since bloodstream trypanosomes are destroyed during an infection, the

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In Gambian HAT, the asymptomatic phase can last many months, if not years. Intermittent fever then appears with nonspecific symptoms such as headache, myalgia, and malaise. These symptoms last a few days before subsiding and reappearing weeks later, and correspond to successive waves of antibodies produced in response to antigenic variation of the VSGs. Eventually weight loss and asthenia become significant, and pruritus can be troublesome. Transient edema, mostly of the face, is seen in fewer than 10% of patients. After months of nonspecific symptoms, evidence of CNS involvement makes the diagnosis obvious. Somnolence is seen in 80% of late-stage patients, and only the occasional patient experiences both diurnal somnolence and nocturnal insomnia. Most patients then complain of constant and severe headaches unresponsive to analgesics. Behavior change or psychosis is seen in 5–10% of cases. If untreated, the somnolence progresses to stupor and coma, and the ultimate cause of death is often a bacterial superinfection such as aspiration pneumonia. Convulsions are rare in adults but more common in children (up to 30%), among whom retardation in psychomotor development, language, or walking, and irritability can be the presenting symptoms. In children, lymphadenopathy is less prominent and the disease progresses more rapidly to overt CNS involvement than in adults. Various endocrine disorders have been reported: hypogonadism, amenorrhea, infertility, and low triiodothyronine (T3) or low thyroxine (T4) syndromes but rarely frank hypothyroidism. There are exceptional cases of healthy carriers who remained parasitemic and asymptomatic for years without treatment, but untreated HAT is ultimately fatal to almost all infected persons. An inoculation chancre is rarely recognized in persons repeatedly bitten all year long by various insects but is more frequent in the rare white individuals who acquire Gambian HAT.22,27 Fewer than half of asymptomatic patients actively detected during surveys, but up to 85% of those passively detected in a hospital, have the classic Winterbottom cervical lymphadenopathies: soft, painless, 1–2 cm in diameter, numerous, and rather mobile. Lymphadenopathy can be less typical, and in persons from high-incidence areas, any lymph node large enough to be aspirated should be aspirated. As the disease progresses, lymphadenopathy regresses. Cutaneous lesions, the trypanids, are seen only in white people. Modest splenomegaly occurs in 10–20%; hepatomegaly is rare (1%). On neurologic examination, patients do not have neck stiffness apart from the occasional patient with a very high CSF white blood cell (WBC) count. Focal signs (hemiparesis, hemiplegia) are unusual, but in a patient from a high-incidence community, any neurologic sign should be considered as potentially caused by HAT until proved otherwise. The hand–chin reflex can be elicited in half the patients. Tremors are not unusual; choreoathetosis is seen mostly in patients with multiple relapses. Despite in vitro evidence of immunosuppression, superinfections seen in HAT patients relate to the altered level of consciousness rather than to opportunistic pathogens. In Rhodesian HAT, the clinical presentation is similar but much more acute. Most patients have been sick for less than a month when the diagnosis is made.28 Inoculation chancres are more common than in Gambian HAT, and trypanosomes can sometimes be seen microscopically in fluid expressed from them. Lymphadenopathy is less frequent than in Gambian HAT and is often submandibular, axillary, or inguinal rather than cervical.28 Myocarditis is an uncommon complication. In tourists from nonendemic countries, the inoculation chancre is often present when the patient develops an acute disease that, if untreated, can rapidly progress to multiorgan involvement and disseminated intravascular coagulation.29

immune system is continually exposed to massive amounts of invariant antigens and VSGs, but the immune responses to these foreign antigens are not protective because the invariant antigens inside the living trypanosome are inaccessible and the readily accessible VSGs periodically switch via antigenic variation. Hence, many of the immune events observed in HAT are likely the result of the perpetual presence of ever-changing VSGs combined with many other invariant antigens, a scenario that mimics successive infections by related but nonidentical organisms. In experimental laboratory animals, the immune responses to a trypanosome infection are dominated by three overwhelming phenomena: nonspecific polyclonal B-cell activation, macrophage activation and generalized suppression of some humoral (B-cell) and cellular (T-cell) immune functions.30,31 The massive polyclonal B-cell activation results in a large production of immunoglobulin M (IgM), the first class of antibody to be generated by the appearance of new foreign antigens. This activation is not triggered entirely by the continually changing epitopes of the different VSGs, however, since the newly synthesized antibodies do not react solely with VSGs and other trypanosome antigens. They frequently are heterospecific in their reactivity and can be autoantibodies directed against the proteins and nucleic acids of the host. It has been proposed that VSGs or unknown non-VSG molecules of trypanosomes cause this massive nonspecific expansion of B cells and the subsequent increase in immunoglobulin concentration.30 The greatly elevated levels of antibody and resultant antigen–antibody complexes in turn cause hyperplasia of the reticuloendothelial system, especially the spleen and lymph nodes, and are likely responsible for many of the pathogenic characteristics of the disease. The other striking immune features of HAT are activation of macrophages and subsequent suppression of immune responses other than the initial B-cell activation, which affects a large variety of both B-cell and T-cell functions and seems to inhibit many secondary immune events.30,31 The concentrations of the cytokines interferon-gamma (INF-γ) and tumor necrosis factor are greatly increased in experimental animals infected with T. brucei. Macrophage activation may be triggered by increases in INF-γ, and the elimination of trypanosomes by antibodies is thought to be mediated by opsonization and destruction by liver macrophages, rather than by complement-mediated lysis. Levels of some other cytokines (e.g., interleukin-2) decrease during trypanosome infections, which may contribute to the lack of T-cell proliferation. Although the coexistence of polyclonal B-cell expansion, macrophage activation, and significant immunosuppression appears at first glance to be counterproductive, these phenomena obviously generate an environment conducive to perpe­ tuation of the infection. Much remains to be learned about the myriad of complex immune events occurring during HAT. HAT is often accompanied by anemia, which may be caused by hemolysis induced by the immune complexes, although other mechanisms are involved.32 Platelet destruction and increased vascular permeability occur, and the parasites readily infiltrate the interstitial spaces and lymphatic system, where they continue to multiply. There is widespread lymphadenopathy due to increased lymphocyte proliferation, which can be followed by fibrosis. The spleen is sometimes enlarged, with generalized cellular proliferation, congestion, and focal necrosis. The spleen and lymph nodes can both develop endarteritis with perivascular infiltration by trypanosomes and lymphocytes. The heart can also be affected, particularly in Rhodesian HAT, in which a pancarditis can develop that involves all cardiac structures, including valves and the conduction system, resulting in cardiac failure and electrocardiographic changes.33 The late-stage of HAT is defined by the detection of trypanosomes in the CSF via lumbar puncture (LP) or CSF pleocytosis (which appears earlier in CSF obtained by cisternal puncture than by LP). The pressure and total protein content of the CSF are increased, and there is infiltration by immune complexes, white blood cells, and sometimes small numbers of eosinophils and morular (Mott) cells, which are thought to be IgMcontaining plasma cells that fail to secrete their antibodies. In the brain, the parasites are found mainly in the frontal lobes, the pons, and the medulla, where they are associated with diffuse meningoencephalitis, parenchymal edema, and dura–arachnoid adhesions. Lymphoid cells often

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infiltrate the brain through the space surrounding the blood vessels, and hemorrhages can occur. Widespread multifocal white matter degeneration develops, especially during the final stages of the disease, but there are no structural nerve cell alterations.34 The actual cause of the cerebral damage in late-stage HAT remains uncertain but the lesions may be mediated by immunologic reactions that occur around the blood vessels and affect the brain parenchyma.

DIAGNOSIS Anemia, thrombocytopenia, an increased erythrocyte sedimentation rate, hypergammaglobulinemia, and hypoalbuminemia are frequent but nonspecific. Eosinophilia is not seen, but an elevated serum IgM (up to 16 times the normal concentration) is suggestive of HAT. The diagnosis requires, however, the detection of trypanosomes. Examination of lymph node aspirates is the classic detection method in Gambian HAT. A 25-gauge needle is inserted in the node, which is massaged for 1 minute while the needle is rotated, the needle is withdrawn and with a syringe the lymph node “juice” is pushed out onto a slide, which must be examined immediately at ×400. Trypanosomes can be seen moving for 15–20 minutes afterwards and are more numerous on the edges of the coverslip. In the absence of lymphadenopathy or if the aspirate is negative, the diagnosis can be made on a wet smear (a drop of unstained blood between slide and coverslip) or a Giemsa-stained thick smear. The latter is more sensitive, but a microscopist who must examine 50 smears per day usually prefers the wet smear in which trypanosomes can be readily detected by their high mobility. Because of the low level of parasitemia and its fluctuation over time, repeated examinations on consecutive days may be necessary before a trypanosome is seen. Patients in whom HAT is suspected but for whom those assays are negative should be tested with other techniques. The most sensitive is the mAECT (miniature anion-exchange centrifugation technique). Blood is filtered through a resin that retains blood cells but not trypanosomes. The eluate is centrifuged, followed by a direct microscopic examination of the pipette in a viewing chamber. The hematocrit centrifugation technique for an examination of the buffy coat is an alternative. The lower limits to the number of parasites detectable by the different methods are 104/mL for the wet smear, 5 × 103/ mL for the thick smear, 5 × 102/mL for the hematocrit centrifugation, and 102/mL for the mAECT. The quantitative buffy coat technique using acridine orange stain is also sensitive.35 The polymerase chain reaction remains a research tool unavailable in endemic countries.36 As the disease progresses, it becomes more difficult to find parasites in the blood and lymph nodes, and more likely that only CSF will reveal trypanosomes. An LP should be performed on all patients for whom parasites have been documented in any of the preceding tests, and on patients with symptoms of sleeping sickness but negative assays. The most sensitive techniques for detecting CSF trypanosomes are the double centrifugation (6–8 mL of CSF is obtained and, after a first centrifugation, the sediment drawn into a capillary tube, centrifuged again, and examined immediately) or simple centrifugation in a sealed Pasteur pipette.35,37 The CSF WBC is measured; patients with a WBC higher than 5/mm3 are arbitrarily considered to be in late-stage. Many patients without neurologic symptoms have a slightly elevated CSF WBC; those with overt somnolence usually have a WBC between 100 and 500/mm3. The CSF is clear, and its white blood cells are mononuclear. Measuring CSF proteins adds little information because they are elevated along with the WBC. Large eosinophilic plasma cells (Mott cells) “typical” of HAT are rarely seen. In endemic areas, clinicians sometimes treat patients with a presumptive diagnosis: no trypanosomes have been found but the typical symptoms, residence in a known focus, positive serologic reactions, and elevated CSF WBC make the diagnosis virtually certain. Many serologic techniques have been developed for Gambian HAT to improve the efficacy of case-finding surveys by identifying serologically positive persons on whom parasitologic assays are selectively performed. Because of drug toxicity, treatment is generally given only to patients in whom trypanosomes are subsequently documented. The CATT (card

agglutination test for trypanosomes, Institute of Tropical Medicine, Antwerp, Belgium), can be performed without electricity, and results are available within 10 minutes. It has good (90–98%) sensitivity and a specificity of ~95% (owing to cross-reactivity with animal trypanosomes).35 Depending on the prevalence of HAT in the population to be tested, its positive predictive value is 66–89% in passive case-finding in a hospital but only 20–30% in active case-finding surveys. Using the CATT on diluted (1 : 10) serum rather than whole blood increases specificity but decreases sensitivity.38 The CATT can be reliably performed using a micromethod on filter paper or diluted blood, reducing the cost of screening.39 In Rhodesian HAT, parasitemia is higher and trypanosomes easier to find in the blood using the above methods. Lymph node aspirates are rarely feasible or necessary. Since the disease progresses rapidly, an abnormal CSF WBC or CSF trypanosomes are frequent. There are no antibody-detection assays. Rare patients diagnosed with late-stage African trypanosomiasis in industrialized countries and investigated with modern tools have been found by magnetic resonance imaging to have hyperintense signal changes in the basal ganglia and deep within gray and white matter as well as meningeal thickening.40,41 Mediastinal, hilar, and para-aortic lymphadenopathy can be seen on CT scan.40

Differential Diagnosis The differential diagnosis of early-stage HAT is that of prolonged fever, associated with a long list of other diseases in the tropics: malaria, HIV infection, typhoid, tuberculosis, etc. The cervical lymphadenopathy of HAT is softer and smaller than in tuberculous lymphadenitis and cancer but similar to HIV-associated lymphadenopathy. In late-stage HAT, other causes of chronic lymphocytic meningitis must be considered, especially tuberculous meningitis and HIV-associated cryptococcosis. In tourists with T. b. rhodesiense trypanosomiasis, the disease needs to be distinguished from severe malaria and African tick bite fever.

TREATMENT AND PROGNOSIS There are few diseases for which most recommended drugs have been used for so long: suramin since 1925, melarsoprol and pentamidine since the 1940s.42 Eflornithine, the only new drug, is now widely available through the generosity of Sanofi-Aventis. To select the best treatment for a patient in whom trypanosomes have been found, two questions must be answered: (1) Is it T. b. gambiense or T. b. rhodesiense? (2) Is the patient in early-stage or late-stage? The first is easily addressed by geographic considerations. For the rare traveler potentially exposed to both subspecies, detection of the serum resistance-associated gene can confirm that the infection is caused by T. b. rhodesiense.43,44 To answer the second, an LP must always be performed. Patients with a CSF WBC >5/mm3 or with CSF trypanosomes are in late-stage, all others are in early-stage. Intrathecal IgM synthesis and interleukin-10 have been proposed to diagnose late-stage disease45 but are generally unavailable. Pentamidine is the standard treatment for early-stage Gambian HAT; suramin is less effective and melarsoprol, though very effective, is avoided because of toxicity. Pentamidine isethionate, given intramuscularly or intravenously (IV), cures 93% of early-stage patients. Its efficacy has remained remarkably uniform over space and time. Pentamidine injections are exquisitely painful and may cause sterile abscesses. During pentamidine treatment, 1% of patients die. In rural African hospitals, the causes of such deaths are difficult to determine, but potentially severe adverse effects include hypotension, hypoglycemia (during treatment), diabetes (after treatment), hypocalcemia, hyperkalemia, renal failure, neutropenia, and ventricular arrhythmias. As soon as the CSF WBC count increases, the efficacy of pentamidine decreases: it should not be used in patients with >5 WBC/mm3.46 For late-stage Gambian HAT, eflornithine, a selective and irreversible inhibitor of ornithine decarboxylase, remains the drug of choice for

Chapter 98

repeated after 1–2 months. The trend in the CSF WBC is more important than the absolute value, since many genuinely cured patients have a slightly elevated CSF WBC 6 months after treatment. Patients who relapse after eflornithine should be given melarsoprol (and vice versa). Patients with a CSF WBC greater than 10/mm3 after pentamidine treatment of early-stage Gambian HAT are considered to be relapsing and should be treated with eflornithine or melarsoprol. Those with borderline results (6/mm3 to 9/mm3) should be retested earlier than the routine interval. Most late-stage adults cured by melarsoprol or eflornithine, even if comatose before treatment, develop no obvious sequelae, but children do so more frequently (poor school performance). In early-stage Rhodesian HAT, suramin is the treatment of choice and is superior to pentamidine (melarsoprol would be effective but is avoided because of the risk of encephalopathy). A test dose is traditionally given although anaphylaxis is rare. Several regimens are recommended,16 but this probably does not matter given the drug’s half-life of ~50 days. Suramin is 99.7% protein-bound and has no CSF penetration. The failure rate with suramin varies from 0% to 31%.42 Adverse effects include fever, proteinuria, and urticaria. T. b. rhodesiense is intrinsically resistant to eflornithine. Melarsoprol is the only effective treatment for late-stage Rhodesian HAT patients, having a 95% cure rate. Several regimens are used; Table 98.1 lists one of those recommended by WHO16 which favors starting with a small dose of melarsoprol and increasing it progressively. However, a similar regimen was more toxic in patients with Gambian HAT54 and a clinical trial is under way evaluating a regimen of 10 consecutive daily injections. Pretreatment with suramin is generally advocated on the theoretical grounds that it might prevent seeding of the CSF when the LP is conducted. Melarsoprol-induced encephalopathy is more common in Rhodesian (5–18%) than Gambian HAT, and mortality during treatment is higher (3–12%). It seems reasonable to administer prednisolone at the same dosage as in Gambian HAT. Encephalopathy should be treated along the same lines. Follow-up LPs should be carried out every 3 months during the first year, and every 6 months during the second year. Most patients who relapse after melarsoprol treatment of Rhodesian HAT are cured by a second course of melarsoprol. For those who relapse a second time, a combination of melarsoprol and nifurtimox might be considered. Given the lack of financial incentives, it appears unlikely that novel drugs will be developed in the near future. Orally administered pafuramidine (DB289) unfortunately failed in clinical trials.

African Trypanosomiasis (Sleeping Sickness)

clinicians working in developed countries. It is as effective as melarsoprol but less toxic. The 14-day regimen cures ~95% of new cases and 98% of post-melarsoprol relapses, presumably because of higher CSF eflornithine levels in the latter.47,48 For new cases, a 7-day course is clearly inferior to the 14-day regimen.47 Eflornithine is less effective in HIV-seropositive patients, who should be treated with melarsoprol,42 and in children, who should be given higher doses (125–150 mg/kg every 6 hours for 14 days). Eflornithine is less effective in patients from Uganda, for unclear reasons.47 For relapsing cases, a 7-day course (same dosage) seems adequate and could be considered in conditions of drug shortage.47 Anemia, leukopenia, and thrombocytopenia are common adverse effects but not clinically significant. Convulsions (6–8%) are related to high CSF drug levels, usually subside when eflornithine is withheld, and do not recur when resumed after 24–48 hours. Fatalities (~2%) during eflornithine treatment are related to advanced HAT rather than toxicity. Oral eflornithine is less effective than IV eflornithine owing to its 55% bioavailability and the osmotic diarrhea it induces. Nifurtimox cures at most two-thirds of patients49 and cannot be recommended as monotherapy. However, there has been much interest in using it as a component of combination therapy. In small case series, encouraging results were documented when nifurtimox was combined with low-dose melarsoprol or a 7-day course of eflornithine.49–51 A recent multicenter randomized trial documented that the combination of oral nifurtimox (5 mg/kg every 8 hours for 10 days) and intravenous eflornithine (200 mg/kg every 12 hours for 7 days) was as effective as the standard eflornithine 14-day regimen described above.52 The combination shortens the duration of treatment and decreases the number of intravenous injections to be administered, which makes it an attractive option for hospitals in endemic areas, where it should be considered now as the treatment of choice. Nifurtimox carries substantial toxicity (mostly anorexia and nausea, sometimes tremors) and it is recommenced that each dose should be administered under direct supervision. The trivalent arsenical derivative melarsoprol should be used in cases of late-stage Gambian HAT if eflornithine is not available. It is remarkably effective (94–97% cure rate) but very toxic (4–6% death rate). It should be given as 10 daily IV injections on successive days, rather than the traditional scheme of three series of three daily injections separated by drug-free intervals. A randomized trial has shown that these two regimens had comparable efficacy and toxicity.53 A graded regimen of melarsoprol was less effective and more toxic.54 Reactive encephalopathy (4–8%, 15% if CSF WBC is >100/mm3) with grand mal seizures, coma, or behavior changes can be a severe, unpredictable complication of melarsoprol treatment, with a case-fatality ratio >50%. Prednisolone, which reduces by two-thirds the risk of encephalopathy without increasing the risk of treatment failure,55 should be given to all melarsoprol-treated patients, as well as ivermectin or albendazole to avoid disseminated strongyloidiasis. Pretreatment with pentamidine or suramin has been given for decades in the hope of reducing the risk of encephalopathy, but this remains unproved. Encephalopathy should be treated with anticonvulsants and dexamethasone to reduce cerebral edema. Dimercaprol, a heavy metal chelator, is useless in the treatment of encephalopathy whose pathogenesis is an immunologic reaction. Polyneuropathy (up to 10%) is a toxic effect of arsenic, which if neglected can progress to paraplegia or quadriplegia. When a patient complains of paresthesias, melarsoprol should be withheld for a while and thiamine (100 mg three times a day) administered. Tremors respond well to β-blockers. Fever can be caused by lysis of trypanosomes, but superinfections should be sought, especially pneumonia. Cutaneous reactions (1%) can be troublesome. Phlebitis and cellulitis at the injection sites are caused by the propylene glycol solvent. After treatment, all patients should be followed with an LP every 6 months for 2 years, or sooner if symptoms recur. A relapse is certain if trypanosomes are found in the CSF (rarely in blood or lymph nodes), but most relapses are characterized only by an elevated CSF WBC. Patients should be considered as having post-eflornithine or post-melarsoprol relapse if their CSF WBC is >50/mm3 and higher than the previous determination, or 20–49/mm3 and higher than the previous measurement with recurrence of symptoms. When in doubt, the LP should be

PREVENTION AND CONTROL For Gambian HAT, case-finding remains the cornerstone of disease control. Infected individuals can remain asymptomatic and contagious for months or years before developing overt sleeping sickness, becoming less contagious when CNS involvement progresses since trypanosomes are then found more in the CSF than in the bloodstream. The only way to break the chain of transmission is to identify and treat asymptomatic persons. Populations of endemic foci should be examined twice a year by mobile teams. Previous case-finding surveys based only on detection and aspiration of cervical lymphadenopathy gave excellent results when 95% of the inhabitants were present. If only 50% of the population participates, as is often the case nowadays,56 and if only this traditional method is used, not enough carriers will be identified to modify the epidemic dynamics. However, using the CATT to identify suspects on whom parasitologic assays are concentrated doubles the number of parasitemic persons detected (half of whom have no lymphadenopathy) and compensates for the lower participation. The best approach is to perform the CATT on all inhabitants. CATT-positive subjects should have blood examinations, and lymph node aspiration if feasible. Most control programs treat only persons in whom trypanosomes have been seen.Whether the systematic treatment of individuals with a positive CATT and negative parasitologic assays would lead to more rapid disease control is unknown, but many such individuals have merely transiently positive CATT assays as a result of infection with animal trypanosomes.57 Pentamidine

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PART H: Protozoan Infections 

Table 98.1  Treatment Regimens for Gambian and Rhodesian Trypanosomiasis

Trypanosoma brucei gambiense

Trypanosoma brucei rhodesiense

Early stage

Pentamidine IM 4 mg/kg up to 300 mg/day for 7 days

Late stage

First choice in endemic countries: eflornithine IV 200 mg/kg q12h for 7 days plus nifurtimox 5 mg/kg q8h for 10 days First choice in non-endemic countries: eflornithine IV 100 mg/kg q6h for 14 days Alternative: melarsoprol IV 2.2 mg/kg daily for 10 days None

Suramin IV 5 mg/kg on day 1, then 20 mg/kg (up to 1.0 g) on days 3, 5, 12, 19, 26 Melarsoprol IV (dose in mg/kg): 1.80 mg/kg (day 5), 2.16 (day 6), 2.52 (days 7, 14), 2.88 (day 15), 3.24 (day 16), 3.6 mg/kg up to 180 mg (days 23, 24, 25)

Pretreatment before melarsoprol Other drugs Melarsoprol-induced encephalopathy   Prevention

  Treatment Relapses

Ivermectin or albendazole

Prednisolone 1 mg/kg up to 40 mg/day, started 1–2 days before first dose of melarsoprol, continued until last dose, tapered over 3 days (30 mg, 20 mg, 10 mg) Anticonvulsants, IV dexamethasone Post-pentamidine: eflornithine or melarsoprol as above Post-eflornithine: melarsoprol as above Post-melarsoprol: eflornithine as above

As in T. b. gambiense

As in T. b. gambiense Post-suramin: melarsoprol as above Post-melarsoprol: second course of melarsoprol IV, 3 × 4 daily injections, all doses at 3.6 mg/kg (up to 180 mg)

IM, intramuscular; IV, intravenous; q12h, every 12 hours; q8h, every 8 hours; q6h, every 6 hours.

“chemoprophylaxis” of the whole population of endemic areas would not be acceptable today because of its adverse effects and the risk of transmission of bloodborne pathogens. Certainly no traveler is at sufficiently high risk of HAT to warrant consideration of individual chemoprophylaxis. To what extent HAT case-finding can be integrated within the activities of multipurpose health centers has been long debated.58 Most cases passively detected by health centers already have an abnormal CSF, and their treatment must have only a modest impact on transmission. Vector control by tsetse fly trapping is a useful but costly adjuvant, to be considered only for high-incidence villages. Given the lack of commercial potential and the capacity of the parasite to undergo continuous antigenic variation, it is unlikely that a vaccine will ever be developed. For Rhodesian HAT, the experience of Uganda showed that an epidemic can

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Suramin IV 5 mg/kg on day 1, 20 mg/kg on   day 3 Ivermectin or albendazole

be controlled through surveillance and case detection by sleeping sickness orderlies, combined with intensive tsetse fly trapping. The international public health community will need to address whether T. b. gambiense trypanosomiasis can be eliminated. Such a strategy would imply that for many years donors must be willing to pay a very high cost for each case detected and treated, with the understanding that these expenditures would be reimbursed over time should elimination be accomplished. The contribution of the animal reservoir of T. b. gambiense will need to be understood before elimination can be contemplated, and modern molecular biology diagnostics will need to be deployed in endemic countries so as to identify and treat the residual human reservoir. T. b. rhodesiense infection is a zoonosis and its elimination may not be feasible, unless at least the cattle reservoir can be controlled.

SECTION II: PATHOGENS

PART H: Protozoan Infections

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CHAPTER 99  American Trypanosomiasis (Chagas Disease)* Louis V. Kirchhoff

INTRODUCTION

EPIDEMIOLOGY

American trypanosomiasis (Chagas disease) is a zoonosis caused by Trypanosoma cruzi, a protozoan parasite found only in the Americas. The geographic range of T. cruzi infection in humans and other mammalian hosts is primarily determined by the distribution of the various species of blood-sucking triatomine insects that act as its vectors. This range extends from the southern United States to Central Argentina (Fig. 99.1).

Distribution of Trypanosoma cruzi

THE AGENT The genus Trypanosoma contains several dozen species, but only T. cruzi and the two African trypanosome subspecies (see Chapter 98), Trypanosoma brucei gambiense and T. b. rhodesiense, cause disease in humans.1 T. rangeli, found only in the Americas, can be transmitted to humans, but does not cause a persistent infection and is not pathogenic.2 T. cruzi was first described in 1909 by the Brazilian physician Carlos Chagas, who saw the motile parasites while doing microscopic examinations of dissected intestines of triatomine insects.3 The complex life cycle of T. cruzi involves insect vectors as well as mammalian hosts (Figs 99.2 and 99.3). Vectors become infected when they ingest blood from mammals that have circulating trypomastigotes, which are nondividing but infective forms of the parasite (Fig. 99.4). Once inside the midgut of an insect host, the parasites undergo transformation to epimastigotes, which are flagellates having a distinct morphology, and these organisms then multiply extracellularly. After migration to the hindgut, epimastigotes transform into nondividing metacyclic trypomastigotes, which are then discharged with the feces around the time of a subsequent blood meal. Transmission to a second mammalian host occurs when breaks in the skin, mucous membranes, or conjunctivae are contaminated with insect feces containing the infective metacyclic forms. Inside the new host, these parasites enter a variety of host cell types and, after transformation into amastigotes, multiply intracellularly. When proliferating amastigotes fill the host cell, they differentiate into trypomastigotes and the cell ruptures. The parasites released invade adjacent tissues and spread hematogenously to distant sites where they initiate further asynchronous cycles of intracellular multiplication, thus maintaining a parasitemia infective for vectors. T. cruzi can also be transmitted by transfusion of blood donated by infected persons,4–6 from mother to fetus,7–10 and in laboratory accidents.11,12

*All the material in this chapter is in the public domain, with the exception of the borrowed figures.

Epizootiology Infection with T. cruzi is a zoonosis, and involvement of humans in the cycle of transmission is not necessary for perpetuation of the parasite in nature. T. cruzi is found only in the western hemisphere, where it primarily infects wild and domestic mammals and insects.13 Triatomine vectors that transmit T. cruzi are found in spotty distributions, from central Argentina to the southern half of the United States. Hollow trees, burrows, palm trees, and other animal shelters are places where transmission of T. cruzi occurs among infected vectors and nonhuman mammalian hosts. Piles of wood, old vegetation, and stacks of roof tiles near houses have also been found to harbor large numbers of insects.14,15 T. cruzi has been found in more than 100 species of domestic and wild mammals,16 from the southern United States to Central Argentina.17–22 Opossums, wood rats, armadillos, raccoons, dogs, and cats are typical hosts, but T. cruzi is not a problem in livestock. Nontypical hosts can become infected when held in zoos in areas in which T. cruzi is enzootic.23,24 This lack of species-specificity, combined with the fact that infected mammals have lifelong parasitemias, results in an enormous domestic and sylvatic reservoir in enzootic areas.

Epidemiology of Chagas Disease in Latin America Historically, humans have become part of the cycle of T. cruzi transmission as farmers and ranchers open up land in enzootic regions. When this development takes place, vectors such as Triatoma infestans, Rhodnius prolixus, and Panstrongylus megistus invade the nooks and crannies of the primitive wood, mud-walled, and stone houses that are typical of rural Latin America. In this manner the vectors become domiciliary, establishing a cycle of transmission which involves humans and peridomestic mammals and is largely independent of the sylvatic cycle.25–27 For the most part, Chagas disease has been a problem of poor people living in rural areas. In recent decades, however, large numbers of infected people have migrated to cities, thus urbanizing the disease and resulting in frequent transmission by blood transfusion prior to the implementation of effective serologic screening.4,28 Early reports indicated that most cases of acute Chagas disease that came to medical attention occurred in children.29 Prevalence data support this view, but few age-specific and geographic incidence data have been available because most cases of the acute illness go undetected due to its mild nature and the lack of access to medical care among those at highest risk. The endemic range of human Chagas disease includes Mexico,5,30,31 as well as all the countries in Central and South America. There is no Chagas disease in the Caribbean islands. The Pan American Health Organization (PAHO) has estimated that 8 million people are infected

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Section II

Chagas disease* Trypanosoma cruzi Bloodstream trypomastigotes circulate to smooth muscle and autonomic ganglia in heart, esophagus, colon (converting to amastigotes) causing chronic disease:

PATHOGENS

Dysrhythmias Cardiomyopathy Megaesophagus Megacolon

 

PART H: Protozoan Infections  690

Bloodstream trypomastigotes

Trypomastigotes enter local cells become intracellular amastigotes, multiply and spread systematically causing acute disease: Trypanosoma cruzi – Human infection Prevalence 1%

Figure 99.1  Geographic distribution of human infection with Trypanosoma cruzi (Chagas disease). Endemic countries in which the overall prevalence rate is ≥1% are shown in pink and those in which the prevalence rate is less than 1% are shown in yellow.

with T. cruzi and that roughly 20 000 deaths each year are attributable to Chagas disease.32 In 2000 the aggregate cost of the morbidity and death associated with Chagas disease was estimated to be more than US$8 billion.33 Despite this enormous public health burden, however, in recent years the epidemiology of T. cruzi infection has improved markedly in much of the endemic range, as vector and blood bank control programs have achieved striking successes. As a consequence, prevalence rates in younger age groups have been decreasing in many areas.34–37 A major international eradication program in the “southern cone” countries of South America (Brazil, Paraguay, Uruguay, Argentina, Chile, and Bolivia) has provided the framework for much of this progress. Uruguay, Chile, and Brazil were certified as transmission-free in 1997, 1999, and 2006 respectively, and marked reduction of transmission has been achieved in Argentina.38 Similar control programs have been established in the Andean countries39 and in Central America.40 In Mexico, expansion of vector control and blood donor screening for Chagas disease are being implemented.30,41,42 In the context of these successes, nonetheless, 41 200 instances of vector-borne transmission were estimated to have occurred in 2005 in the endemic range.32 Moreover, several outbreaks of oral transmission of T. cruzi through contaminated food or drink have been reported, and increased transmission in the Amazon basin has been documented.43–49 The obstacles hindering the elimination of T. cruzi transmission to humans throughout the endemic range are economic and political, and no major technological advances are necessary for its completion. The only caveat to this broad statement is that there is no clearly effective approach for blocking congenital transmission, which occurs in 2–10% of pregnancies in women with chronic T. cruzi infection.50,51 An estimated 14 385 instances of congenital transmission occurred in the endemic countries in 2005.8,32 The epidemiology of symptomatic chronic Chagas disease is note­ worthy. Roughly 70–90% of persons who harbor T. cruzi chronically never develop associated cardiac or gastrointestinal (GI) symptoms. In the past, however, the relatively high frequency of sudden death among young adults in some areas was attributed to arrhythmias due to chronic Chagas disease, and several decades ago in one highly endemic area of Brazil chagasic cardiac disease was found to be the most frequent cause of death in adults.52 Taking a broader perspective, however, the annual diseasespecific death rate among all T. cruzi-infected persons may be about 1 in 400, or 0.25%.

Fever Local swelling (chagoma) Periorbital edema (Romaña’s sign) Myocarditis

Trypomastigotes taken up by triatomine vectors (Triatoma, Rhodnius, Panstrongylus) with blood meals transform into epimastigotes

Metacyclic trypomastigotes, shed in triatomine bug feces, infect the bite site, mucosal surfaces or conjunctivas; enter local cells, multiply and spread systematically

Epimastigotes migrate to hindgut and differentiate into metacyclic trypomastigotes

Epimastigotes multiply in midgut

*Reservoir hosts include armadillos, opossums, dogs, cats, rats and many other mammals

Figure 99.2  Life cycle of Trypanosoma cruzi.

Figure 99.3  Eggs, first and second instar nymphs, and adult of Rhodnius prolixus, a triatomine vector of Trypanosoma cruzi.

Interestingly, there is considerable geographic variation in the relative prevalence of cardiac and GI megadisease in patients with chronic Chagas disease. In most South American countries, megadisease is nearly as common as chagasic cardiac disease, but in Colombia, Venezuela, Central America, and Mexico megadisease is virtually unknown. Parasite strain differences (e.g., genotypes TcI and TcII) may play a major role in this variability of clinical penetrance, and host factors likely are important as well.53,54

Chapter 99 Epidemiology of Chagas Disease in the United States Despite the presence of the sylvatic cycle of T. cruzi in many parts of the southern and western United States, only six instances of autochthonous transmission have been reported there: three in Texas and one each in Louisiana, Tennessee, and California.55–57 Moreover, the blood donor screening program that started in January 2007, in which more than 35 million donations have been tested, has uncovered only a small number of T. cruzi-infected donors who appear to have acquired the infection here. Our relatively high housing standards and low overall vector density probably underlie the rarity of transmission of T. cruzi to humans in the United States. In the past three decades only about 15 imported and laboratory-acquired cases of acute Chagas disease have been reported to the Centers for Disease Control and Prevention (CDC), but none in the latter group occurred in returning tourists. Three instances of tourists returning to Europe from Latin America with acute T. cruzi infection have been described.58,59 In contrast, the number of persons in the United States with chronic T. cruzi infections has grown enormously in recent years. It is currently estimated that 23 million persons born in countries in which Chagas disease is endemic currently live in the United States. Approximately 17 million of these individuals are from Mexico,60 where the overall prevalence of T. cruzi infection may be 0.5–1.0%.5 Moreover, a large proportion of these immigrants have come from Central America, a region in which T. cruzi prevalence is relatively high (e.g., 3.4%, 3.1%, and 2.0% in El Salvador, Honduras, and Guatemala).32 T. cruzi infection rates of 1 in 5995 and 1 in 3285 were found in blood donors in Tucson and Los Angeles,61 and a T. cruzi prevalence of 1.0% was found in 782 Hispanic immigrants attending health fairs in Los Angeles (personal communication, Dr Sheba Meymandi). Recently it was estimated that 300 000 T. cruzi-infected persons currently live in the United States.62 Before blood donor screening began in 2007, the presence here of infected immigrants posed a risk of transfusionassociated transmission of T. cruzi,63 and seven such instances in the United States and Canada had been reported.64,65 All these cases occurred in immunosuppressed patients in whom the diagnosis of T. cruzi infection was made because of the fulminant course of the illness. Since most transfusions are given to immunocompetent persons in whom acute T. cruzi infection would be a mild illness, it is reasonable to infer that other cases have occurred here but have not been noticed. The question as to how best to avoid transmission of the parasite via transfusion in the United States has been debated since the first known instance occurred in 1988, although prior to December 2006 this was done in the context of a lack of a Food and Drug Administration (FDA)-approved screening test. Common sense suggested that if screening is warranted in the endemic countries from which the 23 million immigrants living here have come, then they should be tested when they present for donation here. At the end of the day governmental and blood industry authorities adopted this perspective; and shortly after the Ortho T. cruzi ELISA Test System (Ortho-Clinical Diagnostics, Raritan, NJ) was approved for donor

Figure 99.5  Romaña’s sign (unilateral painless periorbital edema) in a Brazilian patient with acute Chagas disease. (Courtesy of Dr Mário Shiroma, São Paulo, Brazil.)

American Trypanosomiasis (Chagas Disease)

Figure 99.4  Trypanosoma cruzi trypomastigote in human blood (Giemsa stain, ×625). (Courtesy of Dr. Maria Aparecida Shikanai Yasuda, São Paulo, Brazil.)

screening by the FDA, near-universal screening began. To date more than 1200 T. cruzi-infected donors have been identified and deferred permanently from donation, and the confirmed rate of T. cruzi infection among donors has been about 1 in 29 000.6 No cases of transfusion transmission of T. cruzi have been detected since screening started. In an effort to reduce the enormous cost of universal screening ($100–200 million per year) a selective screening protocol based on negative test results on a prior donation is being implemented in some areas. Sizable numbers of persons have also emigrated from the countries in which Chagas disease is endemic to Europe, and T. cruzi infection in these people has been studied.66,67 Finally, the transplantation here of organs from three persons with chronic T. cruzi infections resulted in acute infection in five recipients, one of whom died of the acute illness.68,69

THE DISEASE The clinical manifestations of acute and chronic Chagas disease are different and are described separately. Acute Chagas disease results from a host’s first encounter with the parasite whereas chronic symp­ tomatic Chagas disease involves late sequelae resulting from persistent infection.

Acute Chagas Disease Acute Chagas disease usually occurs in children but can occur at any age. The first sign can be a chagoma, which is an erythematous and indurated lesion at the site of parasite entry that appears a week or two after transmission has occurred.70 When the conjunctiva is the portal of entry, the patient may develop Romaña’s sign, which is unilateral and painless periorbital edema (Fig. 99.5). Romaña’s sign is found in only a small proportion of patients with acute T. cruzi infection and similar findings can be the result of several other processes. Systemic spread of the parasites from the site of initial multiplication may be accompanied by malaise, fever, and edema of the face and lower extremities, as well as hepatosplenomegaly and generalized lymphadenopathy. A peripheral lymphocytosis may accompany the high parasitemias of acute Chagas disease, and mild elevation of transaminases may also be present. Muscles, including the heart, can be heavily parasitized, and

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Figure 99.6  Trypanosoma cruzi in the heart muscle of a child who died of acute Chagas disease. The infected cardiomyocyte shown contains several dozen amastigotes.

severe myocarditis with functional impairment occasionally causing death develops in a small proportion of patients29,55,71 (Fig. 99.6). Nonspecific electrocardiographic (ECG) changes can occur, but the life-threatening conduction defects common in chronic cardiac Chagas disease are usually not seen. T. cruzi also can invade the central nervous system (CNS),72 but neurologic findings are not common. Rarely, meningoencephalitis develops, and it is associated with a poor prognosis.73 In the vast majority of patients, the acute illness resolves spontaneously in 4–8 weeks and they then enter the indeterminate phase of the infection, which is characterized by a lack of symptoms, subpatent parasitemias, and generally detectable antibodies to a variety of T. cruzi antigens.

Figure 99.7  Barium esophagogram in a Brazilian patient with dolichomegaesophagus caused by chronic Chagas disease. Contrast material is pooled in the distal esophagus, which is markedly enlarged. (Reproduced from Neva FA, Brown HW. Basic Clinical Parasitology, 6th ed. Norwalk, CN: Appleton & Lange; 1994.)

Chronic Chagas Disease Chronic Chagas Cardiopathy Although most persons chronically infected with T. cruzi remain in the indeterminate phase for life, approximately 10–30% develop symptomatic chronic Chagas disease, usually years or even decades after the infection was initially acquired. Cardiac problems are the most frequent complications of chronic Chagas disease.74–79 Hearts obtained at autopsy from patients who died of chagasic heart disease usually have a global appearance due to chamber enlargement, and mural thrombi are frequently present, often in the right atrium and the apex of the left ventricle. Left ventricular apical aneurysm is typical in patients with advanced cardiac Chagas disease. At a cellular level, the process that underlies these gross pathologic lesions is a chronic inflammation with mononuclear cell infiltration and diffuse fibrosis that affects the conduction system as well as the cardiac muscle.80–82 Parasites are rarely seen on histological examination, but they can often be detected by polymerase chain reaction (PCR) assays.83–85 The process results in a variety of dysrhythmias, including atrial bradyarrhythmias and fibrillation; premature ventricular contractions; bundle branch blocks, often of the right bundle (RBBB); and complete atrioventricular block. Most instances of sudden death in persons with chronic T. cruzi infection probably result from complete atrioventricular block, ventricular tachycardia, or ventricular fibrillation. The symptoms associated with chronic cardiac Chagas disease reflect the congestive failure, rhythm disturbances, and thromboembolism that result from the fibrosing cardiopathy.86–88 The arrhythmias can cause dizziness and syncope, and sudden death is a common occurrence. The cardiomyopathy frequently affects the right side of the heart more than the left, and thus symptoms of right-sided failure are often present.

Chronic Gastrointestinal Chagas Disease (Megadisease)

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Dysfunction of the GI tract is the second most common consequence of chronic T. cruzi infection.89,90 As in the case of chagasic cardiopathy, GI Chagas disease usually occurs years or even decades after infection with T. cruzi is acquired. Dysfunction related to megaesophagus (Fig. 99.7) is the most typical clinical manifestation, but symptoms due to megacolon

Figure 99.8  Barium enema examination of a patient with megacolon caused by Chagas disease. Markedly increased diameters of the ascending, transverse, and sigmoid segments of the large bowel are marked with opposing arrows.

(Fig. 99.8) are also frequent. The process underlying megadisease is a loss of neurons in the gut.91 Quantitative assessments of this degenerative process have shown that in severely affected patients as many as 85% of the neurons in the esophagus and 50% of those in the colon may be lost. The factors that determine the rate and pattern of the neuronal destruction are not known. Pathologic examination of esophageal specimens obtained surgically or at autopsy from patients with megaesophagus have shown dilatation and varying degrees of thickening of the muscular wall. As in the case of cardiac tissue, microscopic examination shows mononuclear cell infiltration and fibrosis, but finding parasites is unusual. The most common symptom associated with chagasic megaesophagus is dysphagia. Many patients experiencing this sense the accumulation of swallowed food in the esophagus and take in water or more food, or even eat in a standing position, to facilitate its passage into the stomach. Pain, typically starting in the lower substernal area and spreading upward, is also a frequent symptom in patients with megaesophagus. In patients with severe degrees

Trypanosoma cruzi Infection in Immunosuppressed Patients Immunosuppression of persons who chronically harbor T. cruzi can lead to a recrudescence of the infection, frequently with an intensity that is atypical of acute Chagas disease in immunocompetent patients.96–99 The incidence of reactivation of T. cruzi in patients who become immunosuppressed is not known. Persons immunosuppressed by the human immunodeficiency virus (HIV) and infected with T. cruzi are also at risk for reactivation of the latter. To date, dozens of such cases have been described,100–103 and it is noteworthy that many of these patients developed T. cruzi brain abscesses, which do not occur in immunocompetent persons with acute or chronic Chagas disease, and may be difficult to distinguish radiographically from those of cerebral toxoplasmosis.104

DIAGNOSIS The approaches used to diagnose acute and chronic Chagas disease are quite different and will be considered separately.

Acute Chagas Disease The first step in diagnosing acute Chagas disease is determining that the patient has a history consistent with exposure to T. cruzi. Risk factors include residence or a blood transfusion in an endemic country, birth to a mother known or suspected of being infected with T. cruzi, or a laboratory accident involving the parasite. Patients with acute Chagas disease may develop a variety of local and systemic signs, but these are usually mild. Occasionally, severe myocarditis develops, leading to nonspecific ECG changes, radiographic signs of cardiomegaly, or pericardial effusion. Invasion of the CNS by the parasites can lead to abnormal cerebrospinal fluid (CSF) values.72 The differential diagnoses of all of these clinical findings are quite broad, however, and in the absence of a parasitologic diagnosis they can only be viewed as suggestive of acute Chagas disease. A definitive diagnosis of acute Chagas disease is made by detecting parasites. Serologic assays for T. cruzi-specific IgM are not accurate enough to warrant their use.105 Circulating trypomastigotes are highly motile and frequently can be seen in wet preparations of anticoagulated blood or buffy coat. The parasites often can be seen in Giemsa-stained

Chronic Chagas Disease As mentioned previously, patients with chronic cardiac Chagas disease can develop a variety of dysrhythmias, including RBBB, which is the most representative conduction abnormality of Chagas cardiopathy. Nonetheless, it is a nonspecific finding since many persons with RBBB do not have Chagas disease. Echocardiographic and radiographic signs of chronic cardiac Chagas disease are similar to those found in patients with cardiomyopathies caused by other processes. Megacolon and megaesophagus are best diagnosed by barium contrast radiographic tests. It is important to keep in mind, however, that all of these diagnostic studies are useful primarily for defining the degree of associated abnormalities in patients known to be infected with the parasite. Chronic Chagas disease is usually diagnosed by detecting IgG antibodies that bind specifically to parasite antigens, and in the vast majority of instances isolating the organism is not important. At the present time, more than 30 assays for the serologic diagnosis of T. cruzi infection are available commercially. Most of these assays are based on ELISA, indirect immunofluorescence, and indirect hemagglutination formats,110,111 and they are used widely in the endemic countries for screening donated blood and clinical specimens. Many of the assays have sensitivities and specificities that are less than ideal, and false-positive reactions typically occur with samples from patients having illnesses such as leishmaniasis, paracoccidioidomycosis, syphilis, malaria, and other parasitic and nonparasitic diseases.112–115 Because of these shortcomings, the PAHO recommends that samples be tested in two assays based on different formats before decisions regarding infection status are made. A number of assays in which recombinant proteins serve as target antigens are available commercially or in development and hold the promise of improved diagnostic accuracy.116–119 Two FDA-approved assays are available in the United States for clinical testing, but not for screening donated blood (Chagas kit, Hemagen Diagnostics, Inc., Columbia, MD; Chagatest ELISA Recombinante v3.0, Wiener Laboratories, Rosario, Argentina).120 The Ortho T. cruzi ELISA Test System and the Abbott PRISM Chagas (Abbott Laboratories, Abbott Park, IL), which is based on recombinant antigens, have been approved by the FDA for blood donor screening.117,121 These assays currently are used to screen most of the 15.2 million units of blood donated in the United States each year. A Clinical Laboratory Improve­ ment Amendments-approved radioimmune precipitation assay (Chagas RIPA)115,122 is used for confirmatory testing of donated units that are repeat positive in the Ortho and Abbott tests and also is available for testing clinical and research samples in my laboratory. The possibility of using PCR assays for detecting T. cruzi infection has been the focus of considerable study. Although the number of parasites in the blood of chronically infected persons is extremely low, PCR assays have the potential for detecting their presence because they have large

Chapter 99

smears as well. In immunocompetent patients with acute Chagas disease, examination of blood preparations is the cornerstone of detecting T. cruzi. In immunocompromised patients suspected of having acute Chagas disease, however, other specimens such as bone marrow, lymph node aspirates, endomyocardial tissue, skin lesion biopsies, CSF, and pericardial fluid should be examined microscopically. When these direct methods fail to detect organisms in any patient with clinical and epidemiologic histories suggesting a diagnosis of acute Chagas disease, as well as in infants born to serologically positive mothers, samples should be tested in a PCR assay (see below). Considerable data have accumulated supporting the concept that PCR assays are more sensitive for detecting the presence of T. cruzi than are the direct methods described above. Another option is to culture blood or other samples in specialized liquid medium,106–108 but the usefulness of this approach is limited by low sensitivity (50–70% for hemoculture in the best of hands) and by the fact that a minimum of 2–3 weeks is required for cultures to turn positive. In infants who are negative by direct examinations and a PCR assay right after birth, serology for T. cruzi-specific IgG should be done 6–9 months later, by which time maternal antibodies will have cleared.109

American Trypanosomiasis (Chagas Disease)

of megaesophagus, regurgitation can become a problem, and if the underlying problem is not treated, it can lead to intermittent aspiration with associated chronic cough, bronchitis, and pneumonia. As in the case of chagasic megaesophagus, colonic disease is manifested by dilatation and typically the sigmoid colon is the most affected segment. As the disease progresses, the colon can become markedly enlarged in both length and diameter, and the thickening of the wall can become less pronounced. The pathologic changes evident on microscopic examination of affected colonic tissue are similar to those found in the esophagus. The cardinal symptom associated with Chagas disease of the colon is constipation. Pain is also a common symptom, resulting from accumulation of feces and flatus, as well as ineffective and recurrent colonic contractions. Other GI and urinary viscera can be affected in persons with chronic Chagas disease, but this is much less common.92 The most frequent occurrence is hypertrophy of the parotid glands, which is present in as many as 25% of patients with chagasic megaesophagus. The stomach may also be affected, and hypoperistalsis, hypotonia, decreased acid secretion, and delayed emptying of the stomach have been documented in patients with megaesophagus, but dilatation of the stomach is not found frequently.93 The pathogenesis of the cardiac and GI lesions of chronic Chagas disease has been debated for decades. Recently, convincing evidence has accumulated supporting the concept that the low-level presence of parasites in chronically affected cardiac tissue, detectable by molecular methods, stimulates a chronic inflammatory response that over time leads to the pathologic changes observed microscopically.82–85,94,95

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numbers of highly conserved nuclear and kinetoplast DNA (kDNA) sequences. Two decades ago Moser and coworkers123 described a PCR assay in which a 188-basepair nuclear repetitive DNA sequence is amplified using primers called TCZ1 and TCZ2. Each parasite contains approximately 100 000 copies of this sequence, and in contrived experiments as little as 0.5% of the genome of a single organism gave a positive result. A study in mice with acute and chronic T. cruzi infections indicated clearly that TCZ1–TCZ2 assay is much more sensitive than microscopic examination of blood.124 In another PCR assay, described by Sturm et al.,125 a 330-basepair segment of the T. cruzi kinetoplast minicircle is amplified using primers designated S35–S36. It is estimated that each parasite has 120 000 copies of this sequence, and in mixing experiments the authors detected 0.1% of one parasite genome. To date three studies have been published in which the sensitivities of these two assays were compared head to head, and in all three the TCZ1–TCZ2 assay appeared to have an edge in terms of sensitivity.120,124,126 PCR assays for T. cruzi detection based on several other primer pairs have been described, but none appears to have any advantage over tests based on TCZ1–TCZ2 and S35–S36, probably because the parasite has fewer copies of the DNA sequences they amplify. Since these two original reports were published in 1989, more than 100 articles have appeared that deal with the detection of T. cruzi by PCR tests. In nine key human studies published in the 1990s, the sensitivities of the PCR assays ranged from 44.7% to 100%, with most results falling slightly over 90%.107,108,127 These disappointing results likely are the result of a sampling phenomenon in that the large numbers of amplifiable sequences are not dispersed, but rather are contained in the rare parasites that may or may not be swept up with blood drawn for testing. Clearly the level of sensitivity achieved with PCR assays for T. cruzi infection is not high enough to allow their use for confirmatory testing of donated blood.128 Nonetheless, PCR assays are potentially useful for detecting T. cruzi in persons with possible T. cruzi infection who have borderline serologic results, in persons suspected of having acute or congenital Chagas disease in whom parasites are not detected microscopically, and in infected patients who have received specific treatment. In all such individuals, only positive PCR results can be taken as truly indicative of infection status. Recently a group of Latin American experts met to standardize a PCR protocol for T. cruzi testing with the purpose of increasing accuracy and facilitating comparability of results. Parasitologic diagnosis of chronic T. cruzi infection also can be attempted by hemoculture or xenodiagnosis. Due to technical issues the latter largely has become a thing of the past. As noted, hemoculture may have a sensitivity as high as 50–70% in persons with chronic T. cruzi infection.106 Its use should be limited to the groups of patients listed above in whom PCR assays also may have a role.

TREATMENT AND PROGNOSIS Antiparasitic Drugs

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Treatment for T. cruzi infection is unsatisfactory and the need for a parasitologically curative drug regimen is the most important current challenge in Chagas disease research. Many dozens of drugs have been tested for activity against T. cruzi, including those active against other parasitic protozoans, and only two have been found to be clinically useful.129,130 The first of these is nifurtimox (Lampit, Bayer 2502, Leverkusen, Germany), a nitrofuran derivative with which extensive clinical experience has accumulated during the three decades it has been available.131 In patients with acute Chagas disease, nifurtimox reduces the severity and duration of the illness and lessens mortality. Parasitologic cures are achieved with nifurtimox only in approximately 70% of these patients. Importantly, the cure rate is greater than 90% in babies with congenital Chagas disease who are treated during the first year of life.7,132 Those not cured enter the indeterminate phase of T. cruzi infection and over time are at risk of symptomatic chronic Chagas disease. In general, cure rates may be 600 mmH2O), and CSF may show a neutrophil-predominant pleocytosis, normal or decreased glucose concentration, and elevated protein content (100–1000 mg/ 100 mL). A wet mount of the CSF examined immediately may reveal actively moving trophozoites. Smears of CSF should be stained with Giemsa or Wright stain to identify the trophozoite by delineating the nucleus with a centrally placed, large nucleolus. Gram stain is not useful. CT scans of the head without contrast are unremarkable or show only cerebral edema with PAM. Contrast may show meningeal enhancement of the basilar cisterns and sulci, but these are not specific for amebic infection.1,20 Serologic tests usually are of no value in the diagnosis of N. fowleri infections, since most patients die too soon in the disease process (within 5–7 days) to mount a detectable immune response. Molecular techniques such as PCR, nested PCR, and real-time PCR assays have been developed for the specific identification of N. fowleri in cultured amebae from patients and the environment as well as N. fowleri DNA in the environment.1,69,70 Sequencing of the 5.8S rRNA gene and the internal transcribed spacers 1 and 2 (ITS1 and ITS2) of N. fowleri has shown that the genomic region can be used to identify specific genotypes. Epidemiologic typing of N. fowleri has also been done by analyzing the 5.8S rRNA gene and the ITS of clinical isolates.69,70 For example, N. fowleri amebae isolated from two cases who had visited the same hot spring in California at different times belonged to the same genotype (II), and these in turn differed from genotypes of other N. fowleri strains examined.70 A real-time PCR assay provides results in about 5 hours and is currently being used at CDC to identify N. fowleri, Acanthamoeba spp., B. mandrillaris, and Sappinia spp. in CSF or brain tissue.

TREATMENT AND PROGNOSIS Granulomatous Amebic Encephalitis Because most cases of GAE have been diagnosed postmortem, or just before, experience with specific treatment is limited, and current recommendations are based on in vitro studies of the susceptibility of isolates to different antimicrobial agents and a few cases of successful treatment. Treatment is more likely to succeed with early diagnosis and treatment before the infection disseminates, particularly to the CNS. Several patients with Acanthamoeba GAE and Acanthamoeba cutaneous infection without CNS involvement have been successfully treated with multidrug regimens consisting of various combinations of pentamidine isethionate, sulfadiazine, 5-flucytosine, fluconazole or itraconazole, trimethoprim– sulfamethoxazole, and topical application of chlorhexidine gluconate for skin ulcers.1,3,5–7 In vitro experiments suggest that azithromycin and miltefosine, a phosphocholine analog, may also be of value in the treatment of Acanthamoeba GAE.71,72 An immunocompromised HIV-negative patient with Acanthamoeba GAE and disseminated disease was successfully treated with miltefosine.7 Cure in a few cases with involvement of the nasal mucosa and paranasal sinuses has included surgical debridement of diseased tissue. Three patients with GAE caused by B. mandrillaris also have recovered from the disease following treatment with pentamidine isethionate, sulfadiazine, azithromycin or clarithromycin, fluconazole, and, in two cases, flucytosine as well.16,17

Unlike GAE, AK often can be cured by prompt aggressive application of topical antimicrobial agents that achieve high tissue levels, and surgery when necessary. In early infections, debridement may remove infectious organisms or improve the penetration of antimicrobial drugs. Medical cure has been achieved with the application of polyhexamethylene biguanide or chlorhexidine gluconate, either alone or in combination, with or without propamidine isethionate (Brolene) or hexamidine. When medical treatment has failed, a combination of debridement and penetrating keratoplasty has been used with good results in some cases.1,23–25

Primary Amebic Meningoencephalitis PAM is almost always fatal. Only a few patients have survived this disease. A well-studied patient, a California girl, who survived the infection was aggressively treated with intravenous and intrathecal amphotericin B, intravenous and intrathecal miconazole, and oral rifampin. At 4 years’ follow-up, she was found to be completely healthy and free of neurologic deficits.35 Miconazole is no longer available in the United States. Another well-documented patient from Mexico also survived PAM.33

PREVENTION AND CONTROL Granulomatous Amebic Encephalitis Diseases produced by A. castellanii spp. and by B. mandrillaris have occurred most often in hosts with weakened immune systems, and presently no clearly defined methods are available for the prevention of infection with these amebae.

Acanthamoeba Keratitis Contact lenses, which are being used not only for vision correction but also for cosmetic purposes, are the major risk factor for AK. During

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2004–2007, an outbreak of AK occurred in the United States associated with a particular brand of multipurpose contact lens solution found to have deficient disinfection activity against Acanthamoeba.24 A previous AK outbreak during 1985–86 involving contact lens users implicated contaminated homemade saline solutions.22,74 Most of the contact lens solutions marketed currently in the United States do not have sufficient disinfection capacity to kill Acanthamoeba cysts.75 Education of patients regarding the proper care of contact lenses is important in the prevention of AK. Contact lenses should not be used during swimming or while performing water sport activities.

Primary Amebic Meningoencephalitis As a thermophilic ameba N. fowleri proliferates when the ambient temperature exceeds 30°C. Hence global warming might expand the range of N. fowleri in the environment.37 Since N. fowleri is susceptible to chlorine at 1 part per million, it can be controlled readily by adequate chlorination of swimming pools. However, unchlorinated lakes and ponds remain a risk. Therefore, people should assume that there is always a slight risk when entering warm fresh water for recreation, such as swimming and waterskiing. Fortunately the risk of infection is very low, with only 33 reported infections in the United States from 1998 to 2007.75 Only avoidance of water-related activities can prevent Naegleria infection. However, one might reduce the risk by limiting exposure to contaminated water through known routes of entry, such as getting water up the nose. These measures include: • avoiding water-related activities in warm fresh water, hot springs, and thermally polluted water (e.g., water around power plants) • avoiding water-related activities in fresh water during periods of high water temperature and low water levels • holding the nose shut or using nose clips during water-related activities in warm fresh water • avoiding stirring up the sediment during water-related activities in warm, shallow, fresh water.

Pathogenic and Opportunistic Free-Living Amebae: Acanthamoeba spp., Balamuthia mandrillaris, Naegleria fowleri, and Sappinia pedata Chapter 101

Acanthamoeba Keratitis

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PART H: Protozoan Infections

CHAPTER 102 

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Microsporidiosis Louis M. Weiss • David A. Schwartz

INTRODUCTION Microsporidia were identified in 1857 as the cause of pebrine, a disease of the economically important silkworm.1 Microsporidia are worldwide in distribution, with infections reported from all continents except Antarctica (Fig. 102.1).2–10 They are obligate intracellular eukaryotic parasites. Serologic studies of travelers and residents of the tropics suggest that the frequency of infection is higher in tropical countries.11,12 These organisms are most likely zoonotic and/or waterborne infections. Microsporidia infect nearly every animal phyla including other protists. They are important veterinary and agricultural parasites in insects, fish, birds, laboratory rodents, rabbits, dogs, primates, and many other mammals.13,14 Some microsporidia have been used as biologic pesticides for the control of destructive insects.15 Microsporidia were first recognized in mammalian tissue more than 75 years ago16 and suspected as causing human disease in 1959.17

THE AGENTS

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The class or order Microsporidia was elevated to the phylum Microspora in 1977,18 and the phylum was suggested to be renamed Microsporidia in 1998.19 Although traditionally considered “primitive” protozoa, recent molecular phylogenetic analysis suggests that the Microsporidia are related to the fungi.20–23 Microsporidia are currently classified on the basis of ultrastructural features, including size and morphology of the spores, number of coils of the polar tube, developmental life cycle, and host– parasite relationship. Excellent overviews describe the history, ultrastructural and structural characteristics, and life cycle differences among taxa of Microsporidia.18,24–28 Sequence data of rRNA from the Microsporidia have been used to develop diagnostic molecular tests and in the study of phylogenetic relationships.29 The genome size of the microsporidia varies from 2.3 to 19.5 Mb,30 making many Microsporidia among the smallest eukaryotic nuclear genomes so far identified.31,32 Chromosomal analysis suggest they are diploid.33 There are almost no introns in the compact 2.9 Mb genome of Encephalitozoon cuniculi, the gene density is high, and proteins are shorter than the corresponding genes in Saccharomyces cervisiae. There appears to be a high degree of gene conservation among the Microsporidia.34 A genome sequence survey has been completed for Enterocytozoon bieneusi.35 Genome data on many of the Microsporidia are available at the BioHealthBase Bioinformatics Resource Center (http://www. biohealthbase.org) and on EuPathDB (http://eupathdb.org/eupathdb; and http://microsporidiadb.org). The phylum Microsporidia contains more than 1100 species distributed in over 150 genera, of which the following have been demonstrated in human disease (Table 102.1)13,28: Nosema (N. corneum renamed Vittaforma corneae36 and N. algerae initially renamed Brachiola algerae37 and then renamed Anncaliia algerae38), Pleistophora, Encephalitozoon,

Enterocytozoon,39 Septata40 (reclassified as Encephalitozoon41), Trachipleistophora,42,43 Brachiola37 (reclassified as Anncaliia38) and Microsporidium.13 In the immunosuppressed host (e.g., those treated with immunosuppressive drugs or those infected with human immunodeficiency virus (HIV)), microsporidian infection can produce a wide range of clinical diseases. Reports of diarrheal syndromes in patients with acquired immunodeficiency syndrome (AIDS) due to microsporidiosis were first published in 1985.39 Microsporidia can infect virtually any organ system, and cases of encephalitis, ocular infection, sinusitis, myositis, and disseminated infection are well described.13,44 These organisms have also been reported in immune-competent individuals. Other intestinal pathogens may occur simultaneously or sequentially with microsporidiosis.45 The Microsporidia are eukaryotes containing a nucleus with a nuclear envelope, an intracytoplasmic membrane system, and chromosome separation on mitotic spindles, as well as vesicular Golgi46 and a mitochondrial “remnant.”47 Microsporidia form characteristic unicellular spores (Figs 102.2 and 102.3) that, for the human pathogenic microsporidia, range from 1.0 to 3.0  µm by 1.5 to 4.0  µm in size.13,48 The spore coat consists of an electron-dense, proteinaceous exospore, an electron-lucent endospore composed of chitin and protein, and an inner membrane or plasmalemma.49 A defining characteristic of all Microsporidia is an extrusion apparatus consisting of a polar tube that is attached to the inside of the anterior end of the spore by an anchoring disc and coils around the sporoplasm in the spore (Fig. 102.4). During germination, the polar tube rapidly everts, forming a hollow tube that brings the sporoplasm into intimate contact with the host cell. The polar tube provides a bridge to deliver the sporoplasm to the host cell. The mechanism by which the polar tube interacts with the host cell membrane is not known, but this may require the participation of the host cell.50 If a spore is phagocytosed by a host cell, germination will occur and the polar tube can pierce the phagocytic vacuole, delivering the sporoplasm into the host cell cytoplasm. The overall process of germination and formation of the polar tube inoculates the sporoplasm directly into a host cell, functioning essentially like a hypodermic needle.51,52 The general features of the microsporidian life cycle are as follows (Fig. 102.5): 1. Spores are ingested or inhaled and then germinate, resulting in the extrusion of the polar tube, which injects the sporoplasm into the host cell. 2. Germination is followed by merogony, during which the injected sporoplasm develops into meronts (the proliferative stage) that multiply, depending on the species, by either binary fission or multiple fission with the formation of multinucleate plasmodial forms. 3. Sporogony follows, during which meront cell membranes thicken to form sporonts that, after subsequent division, give rise to sporoblasts that go on to form mature spores without additional

Microsporidiosis Chapter 102 Microsporidia

Figure 102.1  The Microsporidia are worldwide in distribution, with infections reported from all continents except Antarctica.

Table 102.1  Microsporidia Identified as Pathogenic to Humans Genus (Species) Encephalitozoon   Enc. cuniculi

  Enc. hellema   Enc. intestinalisa

Enterocytozoon bieneusi Trachipleistophora   T. hominisa   T. anthropoptheraa Pleistophora sp.   P. ronneafiei   Pleistophora sp. Anncaliia (Brachiola)   A. vesicularam   A. algeraea   A. connori Nosema   N. ocularum Vittaforma   Vittaforma corneaea

Microsporidium   M. africanus   M. ceylonesis

Reported Infections Hepatitis, peritonitis, nephritis, encephalitis,b urethritis, cellulitis, prostatitis, sinusitis, keratoconjunctivitis, cystitis, diarrhea,b disseminated infection Keratoconjunctivitis, sinusitis, prostatitis, pneumonitis, nephritis, urethritis, cystitis, diarrhea, disseminated infection Diarrhea,b intestinal perforation, keratoconjunctivitis, cholangitis, nephritis Diarrhea,b wasting syndrome, cholangitis, rhinitis, bronchitis Myositis, keratoconjunctivitis, sinusitis Encephalitis, keratoconjunctivitis, disseminated infection Myositis Myositisb Myositis Keratoconjunctivitis, myositis, skin infection Disseminated infection Keratoconjunctivitisb Keratoconjunctivitis,b urinary, tract infection b

Corneal ulcer Corneal ulcerb

Ex En AD M PI VPI PT Sp Nu Pm

PV

Figure 102.2  Structure of a microsporidian spore. Depending on the species, the size of the spore can vary from 1 to 10  µm, and the number of polar tubule coils can vary from a few to 30 or more. The extrusion apparatus consists of the polar tube (PT), vesiculotubular polaroplast (VPl), lamellar polaroplast (Pl), the anchoring disk (AD), and manubrium (M). This organelle is characteristic of the microsporidia. A cross-section of the coiled polar tube is illustrated. The nucleus (Nu) may be single (such as   in Encephalitozoonae and Enterocytozoonae) or a pair of abutted nuclei termed a diplokaryon (such as in Nosema). The endospore (En) is an inner, thicker, electron-lucent region, and the exospore (Ex) is an outer electron-dense region. The plasma membrane (Pm) separates the spore coat from the sporoplasm (Sp), which contains ribosomes in a coiled helical array. The posterior vacuole (PV) is a membrane-bound structure. (From Chapter 6 “The Structure, Function and Composition of the Microsporidian Polar Tube”. Page 199, Figure 4, in Wittner M, Weiss LM, ed. The Microsporidia and Microsporidiosis. Washington, DC: ASM Press; 1999.)

a

Organism can be grown in tissue culture. Cases reported in immunocompetent hosts.

b

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Section II

multiplication. Once a host cell becomes distended with mature spores, the cell ruptures, releasing mature spores into the environment and completing its life cycle. The combination of multiplication during both merogony and sporogony results in a large number of spores being produced from a single infection and illustrates the enormous reproductive potential of these organisms.

PATHOGENS

EPIDEMIOLOGY

PART H: Protozoan Infections 

 

Microsporidia appear to be common self-limited or asymptomatic enteric pathogens in immunocompetent hosts.10,53 Multiple reports describe Enterocytozoon bieneusi in travelers and residents of tropical countries,4,53–62 as well as Encephalitozoon intestinalis.63 Serosurveys in humans demonstrate

Figure 102.3  Transmission electron microscopy of mature spores of Encephalitozoon hellem in a parasitophorous vacuole. The spore wall consists of an electron-dense exospore (open arrows) and an electron-lucent endospore (large solid arrows). The characteristic coiled polar tube, also known as the polar filament, is seen in cross-section (arrowheads). One can also see the polaroplasts (p) within the sporoplasm (s) as well as ribosomes and rough endoplasmic reticulum. Some extruded polar tubules are seen in cross-section (small arrows), and empty spores (e) are also seen that have discharged their contents. (Original magnification ×7000.)

Figure 102.4  Scanning electron microscopy of a single mature spore of Encephalitozoon hellem from tissue culture demonstrating an extruded polar tube. (Original magnification ×8000.) (From Schwartz DA, Sokottka I, Leitich GJ, et al: Pathology of microsporidiosis. Emerging parasitic infections in patients with the acquired immunodeficiency syndrome. Arch Pathol Lab Med 120:173–188, 1996.)

Microsporidiosis

Enc. intestinalis in epithelial cells, endothelial cells, or macrophages

Intracellular multiplication via merogony and sporogony

Polar tubule pierces host epithelial cell, injects sporoplasm

Ent. bieneusi in epithelial cell

Chronic diarrhea Cholangitis Sinusitis Bronchitis Nephritis Cystitis/prostatitis Keratoconjunctivitis

Presumed ingestion or respiratory acquisition of spores

Person-to-person, zoonotic, waterborne or foodborne transmission?

716

While Ent. bieneusi is primarily in the GI tract other species may invade the lung or eye or disseminate to cause:

Diagnostic spores present in stool, urine, respiratory fluids, CSF or various tissue specimens

Spore-laden host epithelial cells sloughed into lumina of GI, respiratory or GU tracts

Sloughed cells degenerate; spores shed in bodily fluids

Figure 102.5  The general features of the microsporidian life cycle. CSF, cerebrospinal fluid; GI, gastrointestinal; GU, genitourinary.

chronic diarrhea with wasting has been reported to be in excess of 50%.116,117 In patients undergoing liver and bone marrow transplantation,68–76,118–121 clinical manifestations have included watery, nonbloody diarrhea, nausea, and diffuse abdominal pain. Ent. bieneusi can also invade cholangioepithelium,117 leading to sclerosing cholangitis, AIDS cholangiopathy, and cholecystitis,122 with associated abdominal pain, nausea, vomiting, and fever. Imaging studies (abdominal ultrasound, computed tomography, endoscopic ultrasonography, and endoscopic retrograde cholangiopancreatography) usually demonstrate dilated biliary ducts, irregularities of the bile duct wall, and gallbladder abnormalities such as thickening, distension, or the presence of sludge. Systemic dissemination is rare. There is a case report describing this organism in nasal mucosa123 and reports of respiratory tract involvement with Ent. bieneusi associated with chronic diarrhea, persistent cough, dyspnea, wheezing, and chest radiographs with interstitial infiltrates.114,124 Enc. cuniculi, Enc. hellem, and Enc. intestinalis have been associated with gastroenteritis, keratitis, sinusitis, bronchiolitis, nephritis, cystitisureteritis, urethritis, prostatitis, hepatitis, fulminant hepatic failure, peritonitis, and cerebritis, as well as disseminated infection.13,48,113,125–131 An Encephalitozoon sp. has also been reported in a case of nodular skin lesions.132,133 Enc. intestinalis most commonly causes diarrhea134 but can also cause cholangitis,40,135 keratoconjuctivitis, ostemyelitis of the mandible,136 upper respiratory infections, renal failure, keratoconjunctivitis, and disseminated infection in patients with AIDS.135–139 Elimination of this parasite from patients with diarrhea following treatment with albendazole correlates with the resolution of symptoms.137,140 Enc. cuniculi has been associated with hepatitis,141 peritonitis,128 hepatic failure,129 disseminated disease with fever,130 renal insufficiency, and intractable cough.142 Cases of encephalitis and seizures due to Enc. cuniculi have been reported in AIDS patients.130,131 These infections have been reported to respond to albendazole.125,130,131,142 Enc. hellem has been reported to cause disseminated disease associated with renal failure, nephritis, pneumonia, bronchitis, sinusitis, and keratoconjunctivitis.11,143–145 Most reports of ocular infection due to Encephalitozoonidae, have implicated Enc. hellem, including three cases originally classified as Enc. cuniculi.146,147 Patients present with bilateral coarse punctate epithelial keratopathy and conjunctival inflammation resulting in redness, foreign body sensation, photophobia, excessive tearing, blurred vision, and changes in visual acuity. Ocular microsporidial infection in HIV-1infected patients has been restricted to the superficial epithelium of the cornea and conjunctiva (i.e., superficial keratoconjunctivitis) and rarely progresses to corneal ulceration. Physical examination reveals conjunctival hyperemia with superficial punctate keratopathy, and slit-lamp examination usually demonstrates punctate epithelial opacities, granular epithelial cells with irregular fluorescein uptake, conjunctival injection, superficial corneal infiltrates, and a noninflamed anterior chamber (Fig. 102.6). Ocular infection is often associated with disseminated

Microsporidiosis Chapter 102

a high prevalence of antibodies to Enc. cuniculi and Enc. hellem, suggesting asymptomatic infection is common.5,64 In HIV-positive Czech patients, 5.3% were seropositive to Enc. cuniculi and 1.3% to Enc. hellem.65 In Slovakia, 5.1% of slaughterhouse workers were seropositive to Encephalitozoon spp.66 Singh and colleagues found positive antibody titers in 8.6% of healthy adults in England, 43% of Nigerians with tuberculosis, 19% of Malaysians with filariasis, and 36% of Ghanaians with malaria.11 In another study, 12% of travelers returning from the tropics were seropositive and no control nontravelers were positive.12 Antibodies to Enc. intestinalis were found among 5% of pregnant French women and 8% of Dutch blood donors.67 Enterocytozoon bieneusi causes the majority of infections in patients with AIDS and presents as diarrhea with wasting syndrome. Infections with Ent. bieneusi have been reported in liver and in heart–lung transplantation recipients, and Encephalitozoon spp. infections have been reported in patients with kidney, pancreas, liver, or bone marrow transplantation.68–76 Reported prevalence rates in the 25 studies conducted on patients with HIV infection before the widespread use of highly active antiretroviral therapy (1989–1998) varied between 2% and 70%.2,8–10,44,77–82 When combined, these studies identify 375 cases of Ent. bieneusi among 2400 patients with chronic diarrhea for a prevalence rate of 15% in this population. With immune reconstitution due to highly active antiretroviral therapy, the incidence of microsporidiosis has decreased. Microsporidian spores are commonly found in surface water, and human pathogenic species have been found in municipal water supplies, tertiary sewage effluent, recreational bathing water, and ground­ water.56,83–87 Water contact has been found to be an independent risk factor for microsporidiosis in some studies88,89 but not in others.90,91 Enc. cuniculi spores are viable for at least 6 days in water.92 Most microsporidian infections are transmitted by oral ingestion of spores, with the site of initial infection being the gastrointestinal tract. Microsporidia of the genus Encephalitozoon are widely distributed parasites of mammals and birds,88 and the onset of microsporidiosis has been associated with exposure to livestock, fowl, and pets,10,14,93–97 suggesting that microsporidiosis may be a zoonosis. Ent. bieneusi has been reported in pigs,98,99 dogs,100 dairy cattle,101 chickens,102 pigeons,103 simian immunodeficiency virus-infected rhesus monkeys,104,105 and recently, falcons.106 A classification system has been published for the description of isolates of Ent. bieneusi from different reservoir hosts.107 Viable infective spores of Microsporidia are present in multiple human body fluids (e.g., stool, urine, and respiratory secretions) during infection.108 Person-to-person transmission is supported by concurrent infections in cohabiting homosexual men; however, there are no confirmed person-to-person outbreaks of microsporidiosis.9 It has been possible to transmit Encephalitozoon spp. via rectal infection in rabbits, suggesting the possibility of sexual transmission.109 Enc. hellem has been demonstrated in the respiratory mucosa, prostate, and urogenital tract of patients, raising the possibility of respiratory and sexual transmission in humans.110,111 Although congenital transmission of Enc. cuniculi has been demonstrated in rabbits, mice, dogs, horses, foxes, and squirrel monkeys, it has not been demonstrated in humans.112

THE DISEASES Microsporidian Infection in Immunosuppressed Hosts Although the majority of reported cases of Microsporidia involve diarrhea, the spectrum of diseases caused by these organisms has expanded to include keratoconjunctivitis, disseminated disease, hepatitis, myositis, sinusitis, kidney and urogenital infection, ascites, cholangitis, and asymptomatic carriage.13,44,113 Ent. bieneusi infection usually involves chronic diarrhea of 3–10 bowel movements per day,44,48,114–116 anorexia, weight loss, and bloating without associated fever. It occurs more commonly in patients with AIDS having CD4+ counts less than 50 cells/mm3. Diarrhea is often associated with malabsorption, weight loss, and wasting syndrome.114 The mortality of patients with advanced HIV disease and

Figure 102.6  Slit-lamp photomicrograph demonstrates punctate epithelial keratopathy in a patient with AIDS and microsporidian keratoconjunctivitis due to Encephalitozoon hellem. (From Schwartz DA, Visvesvara GS, Diesenhouse MC, et al. Pathologic features and immunofluorescent antibody demonstration of ocular microsporidiosis (Encephalitozoon hellem) in seven patients with acquired immunodeficiency syndrome. Am J Ophthalmol. 1993;115:285–292.)

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disease,143,148–151 and thus urine examination often demonstrates microsporidian spores.143,148–151 Trachipleistophora hominis has been described in several patients with AIDS as a cause of disseminated disease,42 with associated myositis, sinusitis, and keratoconjunctivitis. T. anthropophthera was described in several patients with AIDS having encephalitis associated with myositis and keratoconjunctivitis.43,152 Anncaliia (Brachiola) vesicularum was reported as a cause of myositis,37 as was Pleistophora sp. (P. ronneafiei).153–156 The presentation of microsporidian myositis includes myalgias, weakness, elevated serum creatinine phosphokinase and aldolase levels, and abnormal electromyography consistent with inflammatory myopathy.37,42,154,155 A fatal infection in a 4-month-old athymic male infant with severe diarrhea and malabsorption was due to Anncaliia (Brachiola/Nosema) connori at autopsy, where microsporidia were seen in the lungs, stomach, small and large bowel, kidneys, adrenal glands, myocardium, liver, and diaphragm.157 A case of urinary tract infection and prostatitis due to Vittaforma corneae has also been reported in a patient with AIDS.158 In a child with leukemia, skin infection was due to Anncaliia (Brachiola) algerae and spores were seen infecting the cellular elements of the dermis.159 A. (Brachiola) algerae infection also occurred in a patient with rheumatoid arthritis treated with steroids and monoclonal antibody to tumor necrosis factor (TNF)-α.160

Microsporidian Infection in Immunocompetent Hosts In patients with or without HIV infection, the most common symptom of microsporidian infection is diarrhea.3,4,9,54,58,63,68,69,88,161 Ent. bieneusi has been identified as a cause of self-limited diarrhea in immunocompetent patients and travelers9,53,54,60–63,162,163 and has been found in up to 10% of African children with diarrhea.2,164,165 Enc. intestinalis was found in 8% of the stools of patients in a survey for the etiology of diarrhea in Mexico88 and has been seen in travelers with chronic diarrhea.63 Cerebral infections due to Enc. cuniculi are described in several mammals but have been reported only rarely in immunocompetent humans. In a 3-year-old boy with seizures and hepatomegaly, Encephalitozoon infection was suggested by positive immunoglobulin G (IgG) and IgM indirect immunofluorescence assays (using Enc. cuniculi as the antigen).64 Similarly, Encephalitozoon spp. infection was reported in a 9-year-old Japanese boy with headache, vomiting, spastic convulsions, and recurrent fever.17 An unidentified microsporidium responsive to albendazole was also described as causing multiple cerebral lesions in a 33-year-old man in Japan.166 Pleistophora spp. have been identified in the skeletal muscle of an HIV-negative patient with myositis.153–155,167 Ocular infections with ulcer or deep cornea stroma infection associated with eye pain have been reported in immunocompetent patients. In 1981, corneal microsporidiosis due to Microsporidium africanus168 was described, and in 1973 infection due to M. ceylonesis was described.169 Other cases of microsporidian keratitis have since been identified in immunocompetent hosts.13 One of these organisms was classified as N. ocularum170 and the other, which was successfully propagated in vitro, was named N. corneum173 (now V. corneae36). A. (Brachiola) algerae infection of the cornea has also been reported.159 In these immunocompetent patients with corneal infections, one patient required enucleation,168 one underwent unsuccessful penetrating keratoplasty,169 one was successfully treated with a corneal transplant,170 and the last was maintained on a variety of topical agents without effect until keratoplasty.172 Encephalitozoon spp. and T. anthropophthera corneal infections have been described in contact lens wearers.173,174 In a report from India on 40 immunocompetent patients, epidemic keratoconjunctivitis has been associated with microsporidia.175

PATHOGENESIS AND IMMUNITY 718

Infection of the epithelium of the gastrointestinal tract (small intestine and biliary epithelium) is the most frequent presentation of

microsporidiosis. Ent. bieneusi infection does not produce active enteritis or ulceration, but infection results in variable degrees of villus blunting and crypt hyperplasia. The organism is located on the apical surface of the enterocytes of the small intestine and epithelial cells of the biliary tract and pancreas. Spores are rarely found on the basal surface or in the lamina propria.127,176 Infection may be associated with increased intraepithelial lymphocytes and epithelial disarray. Enc. intestinalis and other Encephalitozoon spp. are invasive; spores are found in the apical and basal sides of infected intestinal enterocytes and in the lamina propria.177 Histopathology can demonstrate areas of necrosis and mucosal erosion. Encephalitozoon spp. infect the genitourinary system in most mammals, including humans,53,127,137,178 in which infection discovered in any organ (eye, gastrointestinal tract, liver, central nervous system, etc.) is often associated with the shedding of spores in the urine. Granulomatous interstitial nephritis composed of plasma cells and lymphocytes is the most frequent pathologic finding. This is associated with tubular necrosis, with the lumen of the tubules containing amorphous granular material. Spores are located in the necrotic tubes and sloughing tubular epithelial cells.70,74,119,121,148 As spores and infected tubular cells are shed into the bladder, they can infect other epithelial cells of the urogenital tract, causing ureteritis, prostatitis, and cystitis127 and infection in macrophages, muscle, and supporting fibroblasts of the associated mucosa. Lower respiratory tract infection due to Encephalitozoonidae has demonstrated erosive tracheitis, bronchitis, and bronchiolitis.126,148 In most cases, organisms are found in intact or sloughed epithelial cells. Sinus biopsies in AIDS patients with chronic sinusitis and microsporidiosis have demonstrated spores in epithelium as well as in supporting structures.145,179–185 Infection with Enc. cuniculi, Enc. hellem, or Enc. intestinalis can result in punctate keratopathy and conjunctivitis characterized by multiple punctate corneal ulcers (e.g., a superficial epithelial keratitis). Microsporidian spores are present in corneal and conjunctival epithelium that can be obtained by scraping or biopsy of the lesions. The organisms do not invade the corneal stroma but remain limited to the epithelium. Inflammatory cells are rarely present.93,108,141,186,187 Infections in immunocompetent hosts with other species of Microsporidia have usually involved deeper levels of the corneal stroma with associated necrosis and acute inflammatory cells, with some giant cells in several cases. Clinically, these patients have a corneal stromal keratitis and occasionally a uveitis. There are scant data on the immune response to microsporidia in humans. It is clear that a strong humoral response occurs during infection and that it includes antibodies that react with the spore wall and polar tube. The immunosuppressive states associated with microsporidiosis (e.g., AIDS and transplantation) are those that inhibit cellmediated immunity. Microsporidiosis is usually seen in HIV-infected patients when there is a profound defect in cell-mediated immunity (e.g., a CD4+ cell count less than 100/mm3); spontaneous cure of microsporidiosis can be induced by immune reconstitution with antiretroviral treatment.188–190 Overall, these data are consistent with observations on the immunology of the mouse model of microsporidiosis in which interferon (IFN)-γ, IL-12, and CD8+ cells have been implicated as critical in the immune response to infection.191,192 It is possible that, in humans, administration of IFN-γ or IL-12 could be useful adjuncts for treating microsporidiosis.

DIAGNOSIS Coprodiagnosis Examination by light microscopy of stool specimens using special stains is the practical method for the diagnosis of gastrointestinal microsporidiosis. Experience is greatest with Weber’s chromotrope-based stain.193 Some laboratories prefer the Ryan modification,194 which uses aniline blue in place of fast green. Using these chromotrope 2R stains, spores appear as 1–3  µm ovoid light pink structures with a beltlike stripe girding them diagonally and equatorially. Enterocytozoon spores are smaller (1.5  µm)

Cytologic Techniques In body fluids other than stool (e.g., urine, cerebrospinal fluid, bile, duodenal aspirates, bronchoalveolar lavage fluid, and sputum), Microsporidia have been easily visualized using a variety of stains (e.g., chromotrope 2R, chemofluorescent optical brightening agents, Giemsa, Brown–Hopps Gram stain, acid fast staining, and/or Warthin–Starry silver staining).48,199,200 Since Microsporidia often infect mucosa or epithelium, cytologic preparations are useful for diagnosis.48,201 Microsporidia have been easily demonstrated in intestinal and biliary epithelium, epithelium of the cornea and conjunctivae, epithelium of the sinonasal and tracheobronchial regions, renal tubular epithelium, and urothelium. Microsporidian keratitis cell samples obtained by gentle rubbing over the conjunctiva and cornea with a tissue swab usually reveal multiple, Grampositive, oval organisms within epithelial cells (Fig. 102.7).

Histologic Techniques Using routine procedures, microsporidian spores are discernible with a modified tissue chromotrope 2R or tissue Gram stain (Brown–Hopp or Brown–Brenn) in biopsy and autopsy tissue specimens (Figs 102.8 and 102.9). In transbronchial biopsies, spores are best identified in bronchial or bronchiolar epithelium, but they can also be found within alveolar spaces. Microsporidia are usually Gram positive and some are also acidfast positive. Due to their thick wall, unstained spores are refractile and as such can be birefringent in unstained tissue sections. With experience, microsporidia can also be seen on hematoxylin and eosin stain. Other stains that may be useful include periodic acid–Schiff, Giemsa, and Steiner silver stains. Biopsy or autopsy material should, if possible, also be placed in electron microscopic fixative when microsporidiosis is suspected because definitive diagnosis of species requires ultrastructural information. Although molecular methods such as PCR can be performed using formalin-fixed tissue, better results are obtained with unfixed tissue or tissue fixed in ethanol. Similar to its role in the understanding of other emerging infections, the autopsy has proven to be a valuable method for understanding the pathogenesis, mechanism(s) of acquisition, and potential transmission, and organ-based pathology of the emerging microsporidial pathogens.148,202,203

Figure 102.7  Conjunctival smear from a patient with AIDS and Encephalitozoon hellem keratoconjunctivitis stained with Gram stain. Numerous Gram-positive microsporidian spores can be seen within the cytoplasm of the epithelial cells as well as outside of these cells.

Microsporidiosis Chapter 102

than Encephalitozoon spores (2.5–3.0  µm). The Gram-chromotrope stain combines chromotrope 2R staining with a Gram-staining step and results in violet-staining spores.195 Spores can also be visualized by ultraviolet (UV) microscopy using chemofluorescent optical brightening agents such as Calcofluor White M2R (fluorescent brightener 28, Fungi-Fluor)196 or Uvitex 2B (Fungiqual A),197 which stain chitin in the spore wall. Chemofluorescent stains also stain fungi, but microsporidian spores can be distinguished from yeast because they have a uniform oval shape and are nonbudding. The limit of detecting microsporidia by these light microscopy techniques appears to be 50 000 organisms/mL.198 Overall, the sensitivity of the chemofluorescent brightener-based stains is slightly higher than that of chromotrope-based stains (especially when low numbers of spores are present in a sample); however, the specificity of the chemofluorescent stains is lower (90% versus 100% in one study).198 Microsporidian spores in food can give a false-positive result, but this does not appear to be a common problem in fecal diagnoses. Because renal involvement with shedding of spores in the urine is common in all of the species of Microsporidia that disseminate, urine specimens should be obtained whenever the diagnosis of microsporidiosis is considered. This has therapeutic implications because the Microsporidia that disseminate (e.g., Encephalitozoon spp.) are usually sensitive to albendazole, but those that do not disseminate (e.g., Ent. bieneusi) are resistant. Definitive identification of the Microsporidia requires either ultrastructural examination (e.g., electron microscopy) or molecular techniques (e.g., species-specific polymerase chain reaction (PCR)). If stool examination is negative in the setting of chronic diarrhea (more than 2 months’ duration), then endoscopy should be performed.

Figure 102.8  Encephalitozoon intestinalis in a duodenal biopsy plastic-embedded section stained with toluidine blue. Unlike Enterocytozoon, the spores of Encephalitozoon intestinalis develop in a vacuole, termed the parasitophorous vacuole, as shown in both superficial enterocytes and in a lamina propria macrophage. (Magnification ×1000).

Figure 102.9  Enterocytozoon bieneusi in a small intestinal biopsy from a patient with AIDS and chronic diarrhea. Microsporidian spores (arrow) are visible in the apical cytoplasm of an enterocyte in this tissue section stained with Gram stain (Brown and Hopps). (Magnification ×1000.)

Electron Microscopy, Immunofluorescence, Molecular Diagnostic Methods, Serology, and Tissue Culture Transmission electron microscopy (TEM) is a useful technique for confirmation of microsporidian infection in patient tissues and fluids, as well as for detailed ultrastructural studies of microsporidian life cycles and host–parasite relationships that are required for the description of new species of Microsporidia (see Figs 102.2 and 102.3). TEM should be performed on any new or atypical infections that are diagnosed as being due to Microsporidia. Scanning electron microscopy (SEM), while not as widely available as is TEM, is highly useful in understanding the threedimensional structure of microsporidia, and their relationship with their host cell.126,127

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Fluorescent antibody reagents can be used for the species-level identification of various Microsporidia in both cytologic and tissue specimens.204–208 Monoclonal antibodies to Enc. hellem, Enc. intestinalis, and Ent. bieneusi209,210 have been described and several of these antibodies have been demonstrated to be useful for the examination of stool specimens and have demonstrated good sensitivity and specificity. Detection kits for microsporidia in stool and environmental samples using antibodies to Encephalitozoonidae and Ent. bieneusi are now commercially available (Waterborne, Inc., New Orleans, LA). Serologic tests for the diagnosis of microsporidiosis have been developed and utilized for epidemiologic studies, but they have not been useful for the diagnosis of microsporidiosis in AIDS patients.211 Molecular diagnostic techniques such as PCR can dentify Microsporidia at the species level in intestinal biopsies, stool specimens, and other tissues (for a review see Weiss and Vossbrinck30).212–215 Currently, these tests are available in reference laboratories such as the Centers for Disease Control and Prevention. The in vitro cultivation of several humaninfecting Microsporidia has enhanced our understanding of these pathogens (for a review see Visvesvara216). Vittaforma corneae,171 Enc. cuniculi, Enc. hellem,186 T. hominis,42 and Enc. intestinalis217 have been cultivated in vitro, whereas Ent. bieneusi has not. The isolation of Microsporidia from clinical specimens is not a routine procedure and is available only in a few specialized research laboratories. The Microsporidia are generally considered after the more common pathogens of the gastrointestinal, hepatobiliary, respiratory, genitourinary, ocular, musculoskeletal, and central nervous systems have been considered and ruled out. The index of suspicion for microsporidiosis should be highest in patients with severe immunosuppression, especially, but not exclusively, those infected with HIV or having undergone organ transplantation. Intestinal microsporidiosis should be considered in any patient with chronic diarrhea or hepatobiliary disease of uncertain cause. The differential diagnosis includes other infectious agents characterized by chronic diarrhea. Intestinal microsporidiosis should be considered in cases of presumptive traveler’s diarrhea in which other routine pathogens have been excluded. A diagnosis of microsporidiosis should also be

considered in cases of unexplained keratoconjunctivitis or corneal ulcers, persistent sinusitis or diffuse lower respiratory disease, unexplained renal insufficiency or abnormalities in urinary sediment, and myositis. Since dissemination can occur, microsporidiosis may affect virtually any organ system, including bone and the central nervous system. Therefore, the identification of Microsporidia in any specimen should prompt a thorough search in all other readily available sources, including stool, urine, sputum, nasal and conjunctival swabs, and possibly cerebrospinal fluid, with consideration of more invasive approaches for those patients requiring a tissue-based diagnosis (e.g., myositis).

TREATMENT AND PROGNOSIS For a review of drugs used in microsporidiosis in humans and animals, see Costa and Weiss218 (Table 102.2). Although albendazole has significant activity against many Microsporidia, such as the Encephalitozoonidae, it has limited efficacy for Ent. bieneusi infection,219–221 with relapse of symptoms rapidly occurring with the discontinuation of therapy in patients who reported improvement of symptoms with treatment. Other studies have found that albendazole has no efficacy in Ent. bieneusi infection.222 Analysis of the tubulin sequences of Ent. bieneusi and V. corneae demonstrates that both of these microsporidia have amino acid substitutions associated with resistance to albendazole.223,224 Fumagillin is used to treat honeybees infected with the microsporidian Nosema apis and has been used to treat microsporidiosis in aquaculture.225,226 Fumagillin and its semisynthetic analog, TNP-470, were found to have activity in vitro and in vivo against Enc. cuniculi, Enc. hellem, Enc. intestinalis, and V. corneae.227–232 Fumagillin has also been demonstrated in both a dose-escalation trial and a randomized clinical trial to be effective for the treatment of human infection with Ent. bieneusi at a dose of 60 mg/ day (20 mg three times daily).227,228 Treatment was associated with resolution of diarrhea, clearance of spores, improvement of Karnofsky scores, and improvement in D-xylose absorption tests. It has also been demonstrated in case reports to be effective in the treatment of Ent. bieneusi in

Table 102.2  Treatment Options for Microsporidiosis Organism

Drug

All microsporidian infections

Restoration of immune function can be critical in control of infection. Patients with AIDS should have highly active antiretroviral therapy optimized No effective commercial treatment Oral fumagillin 20 mg tid (e.g., 60 mg/day has been effective in a clinical trial Albendazolea resulted in clinical improvement in up to 50% of patients in some studies; however, it was not effective in other studies

Ent. bieneusi

Encephalitozoonidae infection (systemic, sinusitis, encephalitis, hepatitis, etc.)   Enc. cuniculi   Enc. hellem   Enc. intestinalis Encephalitozoonidae (keratoconjunctivitis)

Trachipleistophora hominis Anncaliia (Brachiola) vesicularum a

Albendazole Albendazole ± Itraconazole

400 mg bidb 400 mg bid 400 mg bid 2 drops every 2 hours for 4 days, then 2 drops 4 times a dayd 400 mg bid 400 mg bid 400 mg qd

Albendazole 400 mg bid. The duration of treatment for microsporidiosis has not been established. Relapse of infection has occurred upon stopping treatment. Patients should be maintained on treatment for at least 4 weeks and may require prolonged treatment. c Fumagillin bicylohexylammonium. d Eyedrops should be continued indefinitely; relapse is common on stopping treatment. bid, twice daily; tid, three times daily; qd, every day. (Adapted from Costa SF, Weiss LM. Drug treatment of microsporidiosis. Drug Resist Update. 2000;3:384–399.) b

720

Albendazole Albendazole Albendazole Fumagillin solutionc (Fumidil B 3 mg/mL, fumagillin 70  µg/mL) Patients may also need albendazolea if systemic infection is present

Dosage and Duration

treatment. Polymyxin B, propamidine isethionate 0.1% (Brolene), gramicidin, neomycin sulfate, and tetracycline have limited efficacy and should not be used except as treatment for secondary bacterial infections. Keratoplasty provides temporary improvement in some cases, and debulking by corneal scraping may be useful in cases not responding to medical treatment. Steroids may be useful in decreasing the associated inflammatory response.

PREVENTION AND CONTROL The presence of infective spores in various bodily fluids suggests that body substance precautions in health care settings and general attention to hand washing and other personal hygiene measures should be useful in preventing primary infections. Hand washing may be particularly important in the prevention of ocular infections, which may occur as a result of inoculation of conjunctival surfaces by fingers contaminated with respiratory fluids or urine. Whether respiratory precautions are necessary for people with spores in sputum or other respiratory secretions is unknown. It is reasonable to screen close contacts of index cases of microsporidiosis. Spores survive and remain infective in the environment for prolonged periods of time.92 In a typical hospital environment, Enc. cuniculi spores can survive and remain infectious for at least 1 month but can be rendered noninfectious by a 30-minute exposure to most common disinfectants and by the methods employed for sterilization. Therefore, the procedures used to clean most hospital rooms should be sufficient to limit infection. It is likely that these organisms are food- or waterborne pathogens and the usual sanitary measures that prevent contamination of food and water with the urine and feces of animals should decrease the chance for infection. Existing guidelines for the prevention of opportunistic infections that address food, water, and animal contact may be useful in preventing microsporidiosis. Severely immunocompromised patients may wish to consider using bottled or filtered water in some settings. No prophylactic antiparasitic agents have been identified for these organisms. Microsporidiosis has developed in patients on trimethoprimsulfamethoxazole prophylaxis249 and those receiving dapsone, pyrimethamine, itraconazole, azithromycin, and/or atovaquone.250 No studies have evaluated albendazole for prophylaxis, but given its relative lack of efficacy for Ent. bieneusi infections, it is unlikely to be effective for prevention. The most effective prophylaxis is a restoration of immune function in immunocompromised hosts. In AIDS patients, several studies have demonstrated that highly active antiretroviral therapy can produce remission of intestinal microsporidiosis.234–237

Microsporidiosis Chapter 102

the setting of organ transplantation.76 The main limiting toxicity of treatment was thrombocytopenia, which was reversible on stopping fumagillin treatment. Despite a few case reports that indicated that metronidazole was effective for Ent. bieneusi infection, the majority of studies have demonstrated that this drug is not effective.178,229,233 Other medications used without success in the treatment of gastrointestinal microsporidiosis are azithromycin, paromomycin, and quinacrine. Clinical studies have demonstrated that improved immune function can result in the clinical response of patients with gastrointestinal microsporidiosis, with elimination of the organism and normalization of the intestinal architecture.234–238 Relapse has been reported in patients who developed failure of their antiretroviral therapy associated with a decline in immune function and falling CD4+ counts. Overall, these observations suggest that part of the primary treatment of microsporidiosis in the setting of AIDS is the institution of effective antiretroviral therapy. There have been no reports of immune reconstitution syndromes in patients with microsporidiosis treated with antiretroviral therapy. There are numerous case reports that demonstrate the efficacy of 2 to 4 weeks of 400 mg of albendazole twice daily for the treatment of microsporidian infections due to Encephalitozoon spp. A double-blind, placebocontrolled trial of eight patients with AIDS and diarrhea due to Enc. intestinalis demonstrated that albendazole (400 mg twice daily for 3 weeks) treatment led to a resolution of the diarrhea and elimination of the organism, similar to observations in case reports.76,137,138,140,178,182,239,240 In case reports of chronic sinusitis, respiratory infection, and disseminated infection due to Enc. hellem, treatment with 400 mg of albendazole twice daily resulted in resolution of symptoms and clearance of the organism.241,242 In a patient with disseminated Enc. cuniculi infection involving the central nervous system, conjunctiva, sinuses, kidney, and lungs, clinical improvement was demonstrated with albendazole treatment.131 The successful use of albendazole has also been reported in gastrointestinal infection in the setting of organ transplantation76 as well as in cases of urethritis,243 renal failure,244 and disseminated infection142 due to Encephalitozoon spp. In addition, albendazole has been reported to have activity in the treatment of disseminated microsporidian infection with T. hominis and in myositis due to a Anncaliia (Brachiola) vesicularum.37,42 Ocular microsporidiosis can be treated with a solution of 3 mg/mL of Fumidil B, fumagillin bicylohexylammonium, in saline (fumagillin 70  µg/mL);230,245–248 however, recurrence is known to occur when topical therapy is discontinued. Although clearance of microsporidia from the eye can be demonstrated, the organism is still present systemically and can often be demonstrated in urine or nasal smears. The use of albendazole as a systemic agent is thus also reasonable for the treatment of ocular infection and should probably be used in addition to topical

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721

SECTION II: PATHOGENS

PART H: Protozoan Infections

CHAPTER 103 

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Toxoplasmosis Joseph D. Schwartzman • James H. Maguire

INTRODUCTION Toxoplasma gondii is a versatile intracellular parasite that has adapted to infect many animal species and is capable of causing a wide spectrum of disease, the preponderance of which is asymptomatic. Invasion of host cells leads to their eventual death; a complex interplay of host cellmediated immune responses arrests acute infection and maintains continued suppression of the persistent encysted zoites, usually for the life of the host. The wide host range of T. gondii is an exception in contrast to other members of the phylum Apicomplexa. Toxoplasma also has a wide geographic range: its single species is found worldwide. The ability to infect many species by at least two routes and its broad distribution are responsible for the high prevalence of infections in humans; perhaps a third of the world’s people are chronically infected by T. gondii. Serious illness is unusual except among persons with deficient cell-mediated immunity and infants infected in utero. Toxoplasmosis is an important cause of abortion in sheep, swine, and goats.1,2

THE AGENT History and Taxonomy

722

In 1908, Nicolle and Manceaux3 identified T. gondii in a laboratory rodent, the North African gondi; and Splendore4 noted identical forms in a laboratory rabbit. Recognition of the importance of this parasite for humans came in 1937 when it was identified as a cause of “granulomatous encephalomyelitis.”5 The role of chronic infection was appreciated in the 1950s by demonstrating Toxoplasma in eyes previously thought to have been involved with tuberculosis or syphilis.6 The high prevalence of toxoplasmosis was only realized after the serologic dye test was developed by Sabin and Feldman in 1948.7 Congenital toxoplasmosis in infants was recognized before either generalized disease in adults or the lymphaden­ itis of primary Toxoplasma infections in adults was appreciated.5 Roles of reactivated latent infections in producing disease in immunosuppressed adults were recognized at the outset of solid organ transplantation.8 The onset of acquired immunodeficiency syndrome (AIDS) brought recognition of central nervous system (CNS) reactivation causing multifocal encephalitis.9 T. gondii is a genetically homogeneous single species.10 Three genetic types make up 95% of isolates from North America and Europe.11,12 South American strains appear to harbor more genetic diversity. Type II strains are most commonly recovered from humans with congenital and acquired toxoplasmosis.11 Type I strains are more virulent in outbred mice, more frequent in congenital infections in some geographic areas,13 found more frequently in immunosupresssed humans,14 and, along with “atypical” strains (recombinant genotypes of type I and type II strains), are overrepresented in serious ocular infections.15 Evidence suggests that the genetic lineages were separated between northern continents and South

America about 1 million years ago and were introduced into South America approximately 10 000 years ago associated with highly clonal spread, perhaps because of concurrent acquisition of efficient oral infectivity.16–19

Life Cycle Asexual stages of T. gondii are pathogenic for humans and animals.20 Two forms are produced: rapidly dividing tachyzoites invasive in all tissues, and slowly dividing bradyzoites in cysts predominantly found in brain and muscle21 (Figs 103.1 and 103.2). Tachyzoite replication causes acute disease, while bradyzoite cysts are long-lived, with slow turnover, and are responsible for latency and reactivation. Reservoirs of human infection are birds and rodents ingested by cats, as well as human food animals, especially pigs, goats, and sheep, that can carry infectious cysts in meat.1,2,22 The sexual stage, found in the small intestinal epithelium of both wild and domestic Felidae, yields oocysts that are resistant to environmental conditions, tend to float in watered soils, and thereby may be ingested by contamination of vegetables or hands.23,24 Oocysts sporulate within 12–24 hours to several days after passage from the cat and are thereafter infectious (Fig. 103.3).

ECOLOGY, EPIDEMIOLOGY, AND DISTRIBUTION Humans may be infected both by eating cysts in meat and by ingestion of oocysts from contaminated soil. The relative risk of infection in industri­ alized countries is higher from the ingestion of undercooked meat, especially lamb and beef, but in societies with little meat in the diet, oocysts are more important.25 Birds and rodents are important in picking up oocysts from soil and scavenging bradyzoite cysts from infected animals.26 Grazing food animals (e.g., sheep) are probably infected by soil oocysts, but swine are omnivores and may also ingest infected rodents.1,2 The prevalence of Toxoplasma in swine is quite variable. Bovine and fowl Toxoplasma levels are low.27,28 Although sexual recombination can take place in cats, it appears to be rare in nature.15,23,24,29 The distribution of T. gondii is worldwide; all genotypes are found on all continents except in Antarctica and on some islands in the Pacific and along the coast of Central America.25,30,31 Hot, dry climates have a lower incidence of toxoplasmosis than temperate, moist climates, and rates are low at high altitudes.25,32,33 Geographic foci of transmission by oocysts (cat cycle) have been described in societies that do not consume meat.34 The role of the cat in the transmission of toxoplasmosis is established, but the epidemio­ logy of transmission includes the possible role of dogs. Dogs may be vectors of oocysts based on associations in epidemiologic surveys and their habit of rolling in or eating cat feces.26 River water contaminated with oocysts was believed to be the source of an outbreak in a Panamanian jungle.35 Public water supplies were implicated in an outbreak in British Columbia, and drinking unfiltered municipal and surface water was

Figure 103.1  T gondii tachyzoites from culture, with no other cells (×1000). (Courtesy of Joseph D. Schwartzman.)

Toxoplasmosis Chapter 103

Figure 103.2  Congenital toxoplasmosis. Tissue cysts (large arrows) in uvea of human eye with numerous intracellular bradyzoites (small arrows). (Hematoxylin and eosin, ×1000.) (Courtesy of the Department of Tropical Public Health, Harvard School of Public Health, Boston, MA.)

Toxoplasma gondii Transmission via blood transfusion or organ donation

with development of immunity

Asexual cycle Tachyzoites multiply in all nucleated cell types and disseminate through blood and lymphatics causing:

may reactivate with loss of immunity causing:

Acute Lymphadenopathy Hepatosplenomegaly ± fever

Congenital CNS defects if early Chorioretinitis if late

Tissue cysts (10–200 µm) with many bradyzoites form in muscle, myocardium, CNS, et al

Immunocompromised patient CNS lesions Chorioretinitis Pneumonitis Myocarditis

Bradyzoites (from tissue cysts) or sporozoites (from oocysts) are released in intestine and invade surrounding cells becoming fragile 3 × 7 µm tachyzoites

Infection acquired via ingestion of either oocysts from cat feces or tissue cysts in meat

Buoyant oocysts sporulate over 2–3 days to become infectious (>>1 year)

Sheep, pigs, cats, rodents, cows, deer, et al, like humans, are infected from tissue cysts or fecal oocysts and develop throughout their bodies tissue cysts with many bradyzoites

Hardy 10–12 µm oocysts shed over 1–3 weeks in cat feces 3–34 days after cat becomes infected

Figure 103.3  Toxoplasma gondii life cycle.

Enteroepithelial sexual cycle in cats

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Section II PATHOGENS  

PART H: Protozoan Infections  724

associated with a high level of endemicity in Rio de Janeiro State, Brazil.30 Increased risk of infection has also been associated with ingestion of raw shellfish and raw goat’s milk.36 Toxoplasma infection also is acquired by transplacental transmission and, less commonly, through organ trans­ plantation8 and laboratory accident.37 Although Toxoplasma DNA can be detected by polymerase chain reaction (PCR) in blood from chronically infected persons,38 transmission of toxoplasmosis by transfusion of banked erythrocytes has not posed a public health problem.39 Serologic surveys demonstrate prevalences of infection from 90% in various geographic locations.40–50 In parts of France, where rates of infection may exceed 90% by the fourth decade,40,41 transmission is related to ingestion of rare meat. In contrast, rates in England and Finland are approximately 20%.42,43 In tropical areas of Latin America and subSaharan Africa, where cats are abundant and the climate favors survival of oocysts, prevalences may approach 90%.25,26,44–48 In comparison, rates in hot, dry regions such as North Africa usually do not exceed 20%.49 Rates in the United States vary, ranging from 3% in Denver, Colorado, to 17% in Massachusetts, and to 30% in Birmingham, Alabama.50 The seroprevalence of toxoplasmosis among persons aged 12–49 years in the United States remained stable at around 16% throughout the 1990s.51

PATHOGENESIS, PATHOLOGY, AND CLINICAL MANIFESTATIONS Primary Disease The primary route of infection is oral, with progression of infection through the gastrointestinal tract to local lymphatics and spread to other organs documented in the mouse, but this has not been documented in humans.21,50,52 In mice fed bradyzoites, the first step is local invasion of the small intestinal epithelium. The bradyzoite and tachyzoite are both capable of active invasion of many cell types and replicate within a parasite-modified vacuole.52–58 Bradyzoites rapidly convert to tachyzoites in vivo. In vitro, the formation of bradyzoite cysts can be stimulated by various maneuvers that stress the infected cells.59,60 The key step in spreading infection from the localized initial site is likely infection of circulating monocytes in the lamina propria; this cell subset is permissive for T. gondii replication in both mice and humans and may therefore be responsible for transport of the parasite widely throughout tissues.21,52,61 Tachyzoites are found in all organs in acute infection, most prominently in muscle, including heart, and in the liver, spleen, lymph nodes, and the CNS.50,52 The initial pathologic lesion is necrosis caused by death of parasitized cells, with a vigorous acute inflammatory reaction. As the disease progresses, more lymphocytic infiltration develops, but true granulomas are not formed. If the host controls the replication of tachyzoites effectively, tissues are restored to anatomical integrity without scarring, and cysts containing the long-lived bradyzoites remain without sign of host reaction.62 The humoral immune response is rapid and may be capable of killing extracellular tachyzoites (and is of use diagnostically), but it is not protective in the mouse model.61 Control of disease appears to depend on the elaboration of cytokines including interleukin (IL)-2, IL-12, and interferon-γ (INF-γ)63–65 followed by a specific cell-mediated immunity, with CD8+ helper T cells apparently the most important subgroup.66 Virulence of the parasite includes its ability to modulate the host response, a proinflammatory response that may result in increased tissue damage and a higher mortality in experimental infections. Parasite factors responsible for enhanced inflammation have been identified.67,68 Subclinical or unrecognized infections are the usual outcome of primary infection in immunocompetent persons.35,69–74 When symptoms occur, the most common manifestation is painless lymphadenopathy, with or without fever.50,73 Usually a single cervical node is enlarged, but there may be multiple enlarged nodes at one site or generalized lymphadenopathy. Toxoplasmic lymphadenopathy may persist for 4–6 weeks, raising the suspicion of lymphoma.73,75 Occasionally, a syndrome of fever, headache, malaise, myalgia, lymphadenopathy, hepatosplenomegaly and atypical lymphocytosis develops after an incubation period of 5–20

days.35,70–72 The course of illness may last weeks or months, suggesting a diagnosis of infectious mononucleosis. Pneumonitis, myocarditis, meningoencephalitis, polymyositis, and death are rare complications in otherwise healthy persons.76,77 In Brazil and British Columbia, a high incidence of acute acquired retinochoroiditis has been described.78,79 Retinochor­ oiditis in primary acute infection is more common than previously recognized and may be associated with particular genotypes.80

Congenital Disease Primary infection of the mother with consequent infection of the placenta is the mechanism by which almost all congenital disease is transmitted.33,81–83 The placenta allows transmission to the fetus in 30–50% of infections acquired during pregnancy.81 Congenital infection is exceptionally rare when the mother acquires infection before gestation,84,85 and most of the few reported cases have occurred when maternal infection occurred within 3 months of conception or in the setting of maternal immunosuppression due to human immunodeficiency virus (HIV) infection or immunosuppressive therapy.86,87 Reinfection may rarely be caused by exposure to a new genetic type of Toxoplasma.88 Acute infection is apparent in fewer than 20% of mothers, but both symptomatic and asymptomatic infections place the fetus at risk. The rate of transplacental transmission and the severity of disease vary with time of gestation.86 If maternal infection occurs during the first trimester, the risk of fetal infection is only around 10%, but disease is usually severe. Rates of congenital infection rise to about 65% for maternal infection during the third trimester and approach 100% at term, but neonatal infection in these instances usually is asymptomatic.33 Overall, fewer than 15% of infants with congenital toxoplasmosis have severe impairment of the brain or eyes, and about 80% are asymptomatic at birth or have mild disease that is not detected by routine physical examination.81,89–93 However, more than 85% of those with asymp­ tomatic infection will develop adverse sequelae of the CNS or eyes in subsequent years.91,92 Early fetal infections lead to spontaneous abortion, stillbirth, or severe neonatal disease.52,83 In the brain, there are foci of necrosis, microglial nodules, and perivascular mononuclear inflammation in association with free and intracellular tachyzoites.52 Vascular thrombosis may lead to large areas of coagulation necrosis, and necrotic brain may become calcified. Necrosis around ventricles or the aqueduct of Sylvius may lead to hydrocephalus. Neurologic sequelae include seizures, psychomotor retardation, deafness, hydrocephalus, microcephalus, and intracerebral calcifications visible on computed tomography (CT) scan (Fig. 103.4). A common feature of severe congenital toxoplasmosis is bilateral retino­ choroiditis manifested by necrosis of the retina and granulomatous inflammation of the choroid.94 Lesions are near to or in the macula, and vitritis and uveitis are frequently present. There may be micro-ophthalmia, strabismus, cataracts, glaucoma, and optic atrophy. Systemic manifestations such as fever, hypothermia, jaundice, vomiting, diarrhea, lymphadeno­ pathy, hepatosplenomegaly, pneumonitis, myocarditis, and rash may be present.90 Laboratory studies show anemia, thrombocytopenia, high cerebrospinal fluid (CSF) protein, and CSF pleocytosis. While the majority of infants who acquire infection late in pregnancy appear normal at birth, meticulous examination often shows abnormalities such as retinal scars (Fig. 103.5) or abnormal CSF.89,90 Healthyappearing infants occasionally develop severe CNS or ocular disease during the first months of life. More commonly, persons with asymptomatic infections at birth experience recurrent episodes of retinochoroi­ ditis or impaired psychomotor development during the first 10–20 years of life.91,92

Ocular Disease Toxoplasmosis may account for one-third of cases of retinochoroiditis.94–96 Most cases occur in teenagers and young adults and previously were ascribed to reactivation of congenitally acquired infection. Populationbased studies in southern Brazil and recent experience with clusters of

Figure 103.4  Intracerebral calcifications and mild hydrocephalus in an infant with congenital toxoplasmosis. CT scan of the head without contrast.

Scars likely due to previous active chorioretinitis

Macula

Optic Nerve Head

Active lesion

Figure 103.5  Active and healed retinochoroiditis in the periphery of the fundus of the eye of an adult who had documented congenital toxoplasmosis. (Photography by Thomas Monego, Dartmouth Hitchcock Medical Center.)

acute disease in adults have shown that acquired infection may account for more cases of retinochoroiditis than congenital infection.78–80,97,98 Active disease causes pain, photophobia, and blurred vision in the absence of constitutional symptoms. On funduscopic examination, the vitreous is hazy, and elevated, pale-yellow, or white cotton-like patches are seen in the retina. Healed scars are pale with distinct margins and prominent black spots of choroidal pigment. Recurrent retinochoroiditis usually occurs at the borders of scars and may lead to blindness.

Disease in Persons with AIDS and Other Causes of Immunodeficiency Toxoplasmosis is life-threatening for persons with impaired cellular immunity, in particular those with HIV infections and CD4 + T-cell counts 695 million persons in 51 countries have thus far participated.125 Not only has there been success in eliminating lymphatic filariasis in some defined areas, but also collateral benefit in averting disability (estimated 32 million disability-adjusted life-years averted)122 and treating intestinal helminths and other conditions (e.g., scabies, lice). The strategy of the global program is being refined and guided by ongoing research aimed at understanding the ecology and transmission efficiency of the various mosquito vector species and integration with other mass treatment strategies (e.g., deworming programs, malaria control, trachoma control).126–132

SECTION II: PATHOGENS

PART I: Nematode Infections

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CHAPTER 105  Loiasis and Mansonella Infections Amy D. Klion • Thomas B. Nutman

LOIASIS

EPIDEMIOLOGY

INTRODUCTION

Loa loa causes an estimated 3–13 million chronic infections in residents of endemic areas in western and central Africa,22 including the coastal plains of northern Angola, southeastern Benin, Cameroon, Central African Republic, Chad, Republic of the Congo, Equatorial Guinea, Gabon, Nigeria, Sudan, and the Democratic Republic of Congo.23,24 Rare cases have been reported in the region from Ghana to Guinea25,26and in Uganda,27,28 Mali,29 Zambia,23,30 and Ethiopia.31 The occurrence of Loa loa-related encephalopathy following mass treatment of onchocerciasis has led to renewed interest in mapping the distribution of loiasis in Africa, and subsequently to the validated implementation of new epidemiologic techniques, including remote sensing for suitable Chrysops spp. habitats32 and RAPLOA, a rapid assessment method to determine the proportion of community members with a history of eye worm.33,34 Most infected people have histories of prolonged exposure (>4 months’ residence in an endemic area),2,35 although infections can occur after repeated short stays,36 and anecdotally after 1–2 weeks of intense exposure. In hyperendemic areas, exposure, defined by the presence of filaria-specific antibodies, may approach 100%,37 and up to 40% of residents may be clinically infected (i.e., have clinical symptoms or microfilaremia).1 Although nonhuman primates can be experimentally infected with Loa loa, natural infection is restricted to humans.38 Alterations in the ecology of rainforests, primarily due to rubber plantations, which have a lower canopy and scant undergrowth, have led to an increased prevalence of loiasis in some regions.39 Conversely, village development with replacement of forests with farmland may decrease transmission by reducing vector populations.40 The need for both

Loiasis, infection with the filarial nematode Loa loa, is limited to and highly endemic in western and central Africa. Although generally associated with low morbidity, loiasis is the third most common reason for a medical visit in some hyperendemic regions1 and the most common filarial infection acquired by travelers,2 affecting as many as 30% of longterm visitors to endemic areas.3 Characteristic clinical features include Calabar swellings (transient localized angioedema) and subconjunctival migration of the adult parasite (eye worm). Severe complications of infection, including cardiomyopathy, nephropathy, and fatal encephalitis, are rare, but do occur.

THE AGENT Although extraction of an adult worm from the eye of an African slave was reported in 1770,4 clinical manifestations of loiasis were not described until 1781.5 The association between eye worm (synonyms: Filaria loa, F. oculi humani, F. lacrimalis, and F. subconjunctivalis), Calabar swellings, and Microfilaria diurna (Manson 1891) – a novel species of microfilaria in day blood samples of persons from the Congo – remained debatable until the parasite’s life cycle was elucidated in the early 1900s.6,7 Infective larvae are transmitted to humans by bites of infected female Chrysops species flies.8 Over 6–12 months, these larvae develop into white, threadlike adult worms that migrate through subcutaneous tissues, including the subconjunctiva (hence eye worm), at a rate of up to 1 cm/ minute. Adult worms may live for 17 years.9 In bisexual infections, offspring microfilariae (Fig. 105.1) are released into the bloodstream. Ingestion of microfilariae in a blood meal by the Chrysops vector completes the cycle (Fig. 105.2). Blood microfilarial levels range from undetectable to >100 000 parasites/mL and are remarkably constant in infected individuals over time.10 The diurnal periodicity of bloodborne microfilariae coincides with the temporal feeding patterns of the principal vectors, Chrysops silacea and C. dimidiata.11 These day-biting flies living primarily in the canopy of the rainforest,12 are attracted by movement,13 dark skin and clothing, and wood smoke,14,15 and bite most frequently in shaded areas or indoors. With simian strains of Loa loa, morphologically similar to human Loa loa,16,17 microfilariae demonstrate nocturnal periodicity and infections are transmitted by night-biting Chrysops species that do not bite humans.18 Unlike most filarial parasites that infect humans, Loa loa does not harbor the bacterial endosymbiont Wolbachia.19–21

Figure 105.1  Microfilaria of Loa loa in a hematoxylin-stained thick smear of peripheral blood (magnification, ×180). (From Armed Forces Institute of Pathology.)

735

Microfilariae released (during daytime) into peripheral blood after 6–12 months (diagnostic stage)

Adults develop Transient subcutaneous Calabar swellings over 3 months and live 4–17 years Urticaria causing: May migrate through conjunctiva

PART I: Nematode Infections 

 

PATHOGENS

Section II

Loa loa

1st–3rd stage larvae develop over 10–12 days in fat body and migrate to proboscis

Red (tabanid) fly (Chrysops) bites, injecting infective 3rd stage filariform larvae

Red (tabanid) fly (Chrysops) day-biting takes up microfilariae with blood meal

Chrysops breed in wet mud beneath high canopy of rainforests

Figure 105.2  Life cycle of Loa loa.

Figure 105.3  Calabar swelling of the right hand.

reservoirs of infected persons and conditions that support the insect vector is illustrated by the lack of establishment of endemic foci of loiasis in Cameroon following immigration of numbers of microfilaremic people to a nonendemic region,41 or in Louisiana (US), where Chrysops vectors capable of transmission of loiasis to humans exist,42 but microfilaremic individuals are exceedingly rare.

THE DISEASE

736

Whereas the majority of infected persons from endemic areas are asymptomatic despite high levels of microfilaremia, visitors to endemic areas are often symptomatic, with “allergic symptoms,” including pruritus, urticaria, and transient, migratory angioedema (Calabar swellings).35,43 Clinical complications, with the exception of renal abnormalities, are also more common in nonendemic patients, in whom bloodborne microfilariae are rarely detectable. Characteristic laboratory abnormalities in loiasis include eosinophilia (often >3000/mL) and elevated serum IgE, both of which are more pronounced in symptomatic amicrofilaremic patients.

Figure 105.4  Subconjunctival migration of an adult Loa loa.

(From Armed

Forces Institute of Pathology.)

Calabar Swellings Calabar swellings (Fig. 105.3) may occur anywhere on the body, but are most common on the face and extremities, and may be brought on following local trauma. These are evanescent, migratory angioedematous swellings in which edema is often preceded by local pain or itching lasting 1–2 hours after which a 10–20 cm non-erythematous, non-pitting swelling develops. The swelling generally resolves in 2–4 days, but may last as long as several weeks. Recurrences are common in the same site, but may develop anywhere on the body. The precise cause of Calabar swellings remains unproven, although they are thought to represent hyper­sensitivity responses to antigens or microfilariae released by a migrating adult parasite.

Eye Worm Migration of the adult worm across the conjunctiva occurs with equal frequency in natives of and visitors to endemic areas43 (Fig. 105.4). Conjunctival migration, often associated with transient intense edematous conjunctivitis, resolves without ocular sequelae.

Renal involvement, manifest by hematuria or proteinuria, occurs in up to 30% of infected persons and may be transiently exacerbated by treatment.35,43,44 Proposed mechanisms include immune complex glomerulonephritis or mechanical trauma due to filtration of large numbers of bloodborne microfilariae,44–47 although microfilariae are rarely seen in the urine.46,48 Azotemia or progression to renal failure is uncommon. Encephalitis that may develop after antifilarial chemotherapy is the most serious complication of Loa loa infection.49 It is most common in persons with high levels of microfilaremia (>5000 microfilariae/mL blood) and is associated with the presence of microfilariae in the cerebrospinal fluid (CSF).50,51 Symptoms range from headache, irritability, and insomnia, to coma and death. In fatal cases, autopsies demonstrated generalized acute cerebral edema or encephalitis with necrotizing granulomas around degenerating microfilariae.49 More recently, mass distribution of ivermectin in areas where Loa loa and Onchocerca volvulus are co-endemic has revealed similar posttreatment central nervous system effects.47,52 Although mechanisms underlying these neurologic sequelae of ivermectin treatment remain unclear,53 hemorrhages in the palpebral conjunctiva and retina may be an early marker and are more frequent with high levels of Loa loa microfilaremia.54 Perivascular inflammation and vascular wall thickening in the brain parenchyma have been described in a fatal case of post-ivermectin Loa loa-related encephalopathy, although no microfilariae were seen, likely due to the delay (54 days) between administration of ivermectin and the fatal outcome.55 Other less common complications of loiasis include entrapment neuropathy,56,57 psychiatric disturbances,58 arthritis,59,60 lymphadenitis,61 hydrocele,62 pleural effusion,63,64 retinal artery occlusion,65 posterior uveitis,66,67 macular retinopathy,68 blindness,69 and endomyocardial fibrosis (EMF).35,70,71 Circumstantial evidence linking EMF to loiasis includes their similar geographic distribution and the detection of antifilarial antibodies in some persons with EMF.72 Clinical resolution of biopsyproven EMF in a nonendemic patient with loiasis following antifilarial treatment provides additional support for this association,35 most likely secondary to massive reactive eosinophilia and eosinophil infiltration of the endomyocardium.

Other Microfilariae and adult worms have been detected in pathologic and cytology specimens from various unusual anatomical locations,73,74 and calcified adult worms may be detected by routine radiography in some infected persons.75,76

PATHOGENESIS AND IMMUNITY Many clinical manifestations of loiasis, including Calabar swellings, are likely immunologically mediated and are more severe in visitors to endemic areas than in the endemic population.35,43 This clinical hyperresponsiveness is accompanied by hypergammaglobulinemia, marked eosinophilia, increased serum IgE levels, and vigorous humoral and cellular immune responses to filarial antigens.77 In contrast, asymptomatic persons tend to have high levels of microfilaremia and relatively suppressed immune responses to filarial antigens.43 Mechanisms underlying these differences, while ill-understood, may involve genetic factors,78 prenatal sensitization to filarial antigens, or differences in duration or degree of exposure to parasite antigens.79–81 As with most helminth infections, Loa loa infection elicits eosinophilia (see Chapter 131) and elevations of serum IgE, which are most pronounced in persons with symptomatic infections.43 Eosinophils in these patients express surface markers associated with cellular activation, suggesting that activated eosinophils may contribute to anti-parasite immunity82 and/or may mediate pathologic changes. The presence of blood eosinophilia has been associated with secretion of interleukin (IL)-5 by mitogen-stimulated peripheral blood mononuclear cells (PBMCs) from

patients with loiasis and other filarial infections.83 Further, filarial antigens can induce IL-4 and IL-5 secretion from PBMCs or CD4+ T cells in vitro.84,85 Similarly, IgE can also be induced by parasite antigens in vitro, and persons with loiasis have markedly elevated numbers of B cells and T cells capable of responding to parasite antigen in vitro.85–88 Complement activation with deposition of C1q on the surface of microfilariae has been described, although a recent study demonstrating that complement regulators, including complement factor H and C4 binding protein, bind to the surface of Loa loa microfilariae in vivo suggests that inactivation of C3b and C4b may help the parasite down-modulate inflammatory responses.89 The presence of long-term residents of hyperendemic areas who have neither detectable microfilaremia nor clinical symptoms of loiasis suggests that protective immunity to loiasis may develop in some people.90 Definitive proof of naturally occurring immunity will require diagnostic tests sensitive and specific enough to exclude occult infections.

DIAGNOSIS Loiasis should be suspected in any person returning from an endemic area who presents with urticaria, localized swellings, visualization of an adult worm beneath the conjunctiva and/or eosinophilia. Definitive diagnosis can be made by extracting an adult worm from subcutaneous or subconjunctival spaces or identifying Loa loa microfilariae (or their DNA) in peripheral blood. Adult male worms are approximately 3.5 cm × 0.5 mm and female worms are 5–7 cm × 0.3 mm.91 The cuticle is thick, unstriated, and covered with irregularly spaced bosses, which are useful in identifying fragments of the worm. Loa loa microfilariae, approximately 290 × 7.5  µm in size, are distinguished from microfilariae of other species (most notably Wuchereria bancrofti, Brugia malayi, and Mansonella perstans) by their diurnal periodicity, sheath, and the presence of three or more terminal nuclei.23 The periodicity of Loa loa microfilariae is affected by differing time zones and variations in body temperature,92 supporting the hypothesis that blood microfilarial levels are linked to the host’s circadian rhythm. In those evaluated soon after leaving endemic regions, the optimal time of blood sampling for diurnal periodicity may still reflect timing in the endemic countries. Serology may be useful for confirming the diagnosis in visitors to endemic areas who have suggestive clinical symptoms or unexplained eosinophilia; however, currently available methods using crude antigen extracts from Brugia or Dirofilaria species do not differentiate between Loa loa and other filarial pathogens.93,94 The utility of serologic testing in endemic areas is limited by the often high prevalence of species nonspecific antifilarial antibodies.37 In a small study of patients with parasitologically proven loiasis and/or M. perstans infection living in an endemic area, a detectable IgG4 response to Loa loa adult antigen was 92% sensitive and 94% specific for Loa loa infection.95 Cross-reactivities in subjects with onchocerciasis and lymphatic filariasis were not assessed, and difficulties in obtaining adult Loa loa worms will limit the use of serologic tests based on extracts of Loa antigens. More recently, serologic assays using recombinant Loa loa antigens have been developed, including a sensitive and specific rapid assay that uses a luciferase immunoprecipitation system to detect antibodies to the recombinant Loa loa antigen LLSXP-1.96 Diagnostic testing for Loa loa antigen has yet to be developed. The identification of Loa loa-specific DNA sequences97,98 enabled the development of polymerase chain reaction (PCR)-based strategies both for speciation of Loa loa from pathologic specimens difficult to identify on morphologic grounds and for sensitive diagnostic strategies. Using different Loa-specific targeted sequences, a PCR-based multiplex technique has been developed that is 10 times more sensitive than traditional blood filtration. Further, a colorimetric detection system that can be performed at ambient temperature has simplified the PCR readout, facilitating its use in endemic areas.99 Differential diagnosis of Calabar swellings includes angioedema associated with C1 inhibitor deficiency, infection with other filariae (particularly M. perstans and Onchocerca volvulus), specific nematode and trematode infections (e.g., trichinellosis, gnathostomiasis), and hypereosinophilic

Loiasis and Mansonella Infections Chapter 105

Complications

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Section II PATHOGENS

TREATMENT AND PROGNOSIS

PART I: Nematode Infections 

 

syndromes. Although “eye worm” in the setting of a compatible exposure history is extremely suggestive of loiasis, subconjunctival migrations of a few nematodes, including Dirofilaria repens, a dog and cat filarial parasite (see Chapter 107), and Thelazia californiensis, deer eye worm (see Chapter 112), have been reported.100,101 Finally, the symptoms of Loa loa infection may sometimes be difficult to distinguish from those of often co-endemic onchocerciasis or bancroftian filariasis.

Diethylcarbamazine (DEC) is effective against both microfilariae and adult worms and, at a dose of 8–10 mg/kg daily for 21 days, remains the drug of choice for the treatment of Loa loa infection in amicrofilaremic patients, including most long-term visitors to endemic areas.102 Although curative after a single course in 45–50% of such patients, multiple courses of DEC are often necessary and recrudescence may occur up to 8 years post treatment.103 Mild side effects of treatment are common and include Calabar swellings, pruritus, arthralgias, fever, nausea, diarrhea, right upper quadrant discomfort, and a sensation of creeping under the skin.35,104 Antihistamines or steroids may reduce the occurrence and severity of these symptoms. Occasionally, adult worms become motile under the skin following DEC treatment and may be removed with forceps through a small skin incision104 or by excisional biopsy.105 More serious treatment-related complications, including renal failure, shock, coma, and fatal encephalitis, are related to microfilarial burden and may be triggered by the massive microfilarial clearance that occurs with DEC treatment.106,107 Historically, gradual increases in doses of DEC and pretreatment with antihistamines and steroids were advocated to prevent these complications; however, numerous reports demonstrated that these strategies do not prevent encephalitis.50 The level of microfilaremia that presents a significant risk for serious encephalitic complications is unknown, but 2500/mL blood has been suggested. Alternatives for persons with high levels of microfilaremia include no treatment, removal of circulating microfilariae by cytapheresis prior to DEC treatment,108,109 and newer drugs, including ivermectin110–114 and albendazole.115 Ivermectin reduces microfilarial levels in patients with Loa loa infection, but is ineffective against adult worms.110,113 Furthermore, side effects secondary to microfilarial clearance occur in 30–70% of patients with high levels of microfilaremia, and, in rare cases, may be lifethreatening.110,111 This has become a major problem for mass drug admini­ stration programs using ivermectin as unexpected deaths have occurred in subjects with concomitant loiaisis.116 Although reduction in microfilarial levels has been described after prolonged administration of high dose mebendazole in some patients with loiasis,117,118 clearance of microfilaremia is rare and at least one study has failed to demonstrate any effect of mebendazole on Loa loa microfilarial levels.119 Recently, albendazole, a benzimidazole with better oral bio­ availability, has been shown to significantly decrease Loa microfilarial levels in a double-blind, placebo-controlled study when used at a dose of 200 mg twice daily for 3 weeks.115 Adverse effects were not observed, even in persons with greater than 50 000 microfilariae/mL blood. Unfortunately, shorter higher dose regimens do not appear effective.120,121 The gradual decrease in blood microfilarial levels over the course of several months suggests that albendazole may have preferential effects on adult parasites, explaining the lack of adverse effects associated with acute microfilarial clearance. Sequential therapy with albendazole and a microfilaricidal agent (DEC or ivermectin) may provide an alternative to pretreatment cytapheresis in patients with high level microfilaremias.

PREVENTION AND CONTROL 738

Weekly chemoprophylaxis with DEC (300 mg) is effective for prevention of loiasis in long-term travelers to endemic areas.3 Vector control programs have achieved only limited success, primarily because of the

dense vegetation in endemic areas and difficulties accessing Chrysops breeding sites.40

MANSONELLA STREPTOCERCA INTRODUCTION Streptocerciasis, infection with the filarial nematode Mansonella strep­ tocerca, is limited to the tropical rainforests of central Africa, although a small focus has been found in western Uganda.122 Whereas infection can be completely asymptomatic, characteristic clinical features include a chronic, pruritic dermatitis and lymphadenopathy, often indistinguishable from onchocerciasis. Because adult male and female worms have been found in the skin of chimpanzees and because microfilariae have been found in gorillas, streptocerciasis may be a zoonosis.

THE AGENT Microfilariae of M. streptocerca (synonyms: Dipetalonema streptocerca, Tetrapetalonema streptocerca) were first identified in the skin in 1922.123 Adult female124 and male worms were recognized in 1972.125 Infective larvae are transmitted to human hosts by bites of infected Culicoides grahamii midges.126,127 Over months to years, these larvae develop into white, threadlike adult worms (males 17–18 mm × 40– 50  µm; females 27 mm × 65–85  µm) that live in the dermal layer of the skin. In bisexual infections, microfilariae (sheathless, 2.5–5.0  µm × 180– 240  µm) are produced and reside in the upper dermal and collagen layers of the skin.128 M. streptocerca has also been found in nonhuman primates,129 which appear to be additional definitive hosts for this parasite.

EPIDEMIOLOGY M. streptocerca is present in tropical rainforests of northern Angola, Cameroon, Central African Republic, Republic of the Congo, Equatorial Guinea, Nigeria, Uganda, and the Democratic Republic of Congo. In endemic areas, 40% of people may be infected (i.e., have clinical symptoms or detectable skin-dwelling microfilariae).

THE DISEASE The major clinical manifestations of streptocerciasis are dermatologic125,128 and include pruritus, papular rashes, and pigmentation changes. A chronic, pruritic dermatitis, characterized by nonanesthetic, hypopigmented macules, is most common.125 These macules may be discrete or confluent, and are located predominantly over the shoulder girdle and thorax. Dermal thickening is often present. Papular eruptions occur relatively less frequently. Inguinal adenopathy is extremely common in streptocerciasis; in several studies, often as frequent as 100%.125 Although massive lymphedema (elephantiasis) has been reported as a result of M. streptocerca infection, this causality remains unproven. Infection is associated with blood eosinophilia without leukocytosis.125,128

PATHOGENESIS AND IMMUNITY Most pathologic changes seen in streptocerciasis are dermal and consist of sclerosis of the papillae, edema, fibrosis, and perivascular infiltration with lymphocytes and eosinophils.125,128 Dermal lymphatics are dilated. Similar to other filariae, M. streptocerca is well adapted to human hosts and provokes little inflammatory reaction in the absence of treatment. With filaricidal treatment, however, inflammatory reactions appear around adult worms and microfilariae, potentially due to antigens or constituents released from treated parasites.125,128

microfilariae (sheathless, 3.5–4.5  µm × 100–200  µm) are produced and are found in the blood without any significant periodicity.145 Like most filarial parasites that infect humans, M. perstans appears to harbor the bacterial endosymbiont Wolbachia,146 although this may not be true for M. perstans from all geographic areas.20,21

Streptocerciasis should be suspected in persons returning from endemic areas who present with pruritus, a rash, or bilateral inguinal adenopathy. Definitive diagnosis is made by identifying M. streptocerca microfilariae in skin snips (see Chapter 106). Microfilariae are distinguished from microfilariae of other species (most notably skin-dwelling O. volvulus) by a characteristic tapered hook-shaped tail (shepherd’s crook) and nuclei extending to the very end of the tail.125,130,131 Adult worms may be found in skin biopsies, but this is rare. M. streptocerca DNA can be identified in skin biopsies using a nested PCR-based assay,132 which is specific and more sensitive than skin snips for diagnosis of streptocerciasis. Serology is not of diagnostic value. Streptocerciasis may be confused with onchocerciasis and other chronic dermatitides. Hypopigmented macules can resemble those of lepromatous leprosy. Although the nonanesthetic nature of nodules and their location on the upper body may suggest streptocerciasis, histopathologic evaluation is often necessary to distinguish leprosy from M. strep­ tocerca infection.

At least 30 million residents of endemic areas are infected with M. per­ stans.22 Distributed mainly in sub-Saharan Africa, from Senegal east to Uganda and south to Zimbabwe, and in South America along the northern coast of the entire continent, minor foci have been identified in Tunisia and Algeria. In highly endemic areas, close to 100% of people may have microfilaremia. Although humans are principal reservoirs of infection, nonhuman primates (gorilla147 and chimpanzee129) can also be hosts for the parasite. The principal vectors of M. perstans are Culicoides milnei and C. grahamii, but C. austeni, C. fulvithorax, C. kingi, and C. furens can also transmit infection.139–144

TREATMENT AND PROGNOSIS

THE DISEASE

DEC (6 mg/kg daily in divided doses for 14–21 days) is effective in killing both microfilariae and adult worms.125,128,133,134 As in onchocerciasis, increased pruritus, urticaria, and papular eruptions and systemic findings (arthralgias, myalgias, headaches, fever, nausea, and vomiting) may accompany treatment. Generally these symptoms occur within 24–48 hours of treatment and can be treated symptomatically with antihistamines and anti-inflammatory agents. Nevertheless, because DEC is contraindicated in most of Africa because of concerns with post-treatment reactions in onchocerciasis, its use in this infection is limited. Ivermectin 150  µg/kg is an effective microfilaricide both early (6–12 days)135 and later (1 year) following treatment.136

The clinical and pathologic features of M. perstans infection are poorly defined. Although most patients are asymptomatic,148–151 a wide range of clinical manifestations have been described,148 including transient angioedema and pruritus of the arms, face, or other parts of the body (analogous to the Calabar swellings of loiasis) and recurrent urticaria.148,150,152 Less commonly, fever, headache, arthralgias, and right upper quadrant pain can occur.148,150,152–155 Pericarditis,72,156 hepatitis157–159 meningoencephalitis, neuropsychiatric disturbances148,151,159 and rarely, conjunctival granulomas160,161 and intraocular (retinal) lesions162 have been reported.

PREVENTION AND CONTROL

EPIDEMIOLOGY

PATHOGENESIS AND IMMUNITY

MANSONELLA PERSTANS

Little is known about immune responses or the pathogenesis of symptomatic infections. Whereas blood eosinophilia and IgE elevations are common,157,160,163–165 inflammatory reactions in this infection have been difficult to document.163 There is some evidence that, when inflammation occurs, it is granulomatous.160,161 Live adults induce little host response,128 and pathologic findings are rare.166Although uncommon, secondary complications of hypereosinophilia due to M. perstans (e.g., valvular heart disease) have been reported.165

INTRODUCTION

DIAGNOSIS

Perstans filariasis, an infection caused by Mansonella perstans, is distributed across the center of Africa, parts of North Africa, the Caribbean basin, and in northeastern South America.23 Although generally associated with little morbidity, clinical manifestations may include transient angioedema and pruritus of the arms, face, or other parts of the body (analogous to the Calabar swellings of loiasis), fever, headache, arthralgias, and right upper quadrant pain. Occasionally pericarditis and hepatitis occur.

The diagnosis is made by finding microfilariae in blood or serosal effusions. Concentration techniques may be necessary in light infections.130,167–169 More recently, PCR has been developed for specific identification of M. perstans.146 Rarely, adult worms may be recovered. Occasionally, CSF and urine contain microfilariae. M. perstans filariasis is often associated with blood eosinophilia and antifilarial antibody elevations.153,164,170,171

THE AGENT

TREATMENT AND PROGNOSIS

Microfilariae of M. perstans (synonyms: Dipetalonema perstans, Tetrapeta­ lonema perstans, Acanthocheilonema perstans) were first identified in 1890,137 and the adult form of M. perstans was first collected from the mesentery in 1898.138 Infective larvae are transmitted to humans by bites of several species of infected midges (Culicoides spp.).139–144 Over 9–12 months, these larvae develop into creamy-white, threadlike adult worms (males, 35–45 mm × 50–70  µm; females, 60–80 mm × 100–150  µm) that live in serous cavities – pericardial, pleural, and peritoneal – as well as in the mesentery and in the perirenal and retroperitoneal tissues. In bisexual infections,

Treatment protocols have been attempted, including (1) DEC 8–10 mg/ kg daily for 21 days,150,151,154,172,173 (2) mebendazole 100 mg twice daily for 30 days,114,115,165,169,170,174,175 (3) mebendazole combined with levamisole,171,176 (4) ivermectin,114,177–180 (5) albendazole (up to 400 mg twice daily for 10 days),181 and (6) combination ivermectin/albendazole.180–182 None has been effective in M. perstans filariasis, although some regimens reduce microfilaremia to a small degree. Consistent with the presence of Wolbachia endosymbiouts within M. perstans, a randomized trial in Mali demonstrated the utility of doxycycline (200 mg daily for 6 weeks) treatment for this infection.183

There have been no studies of chemoprophylaxis for prevention of streptocerciasis. Although specific control programs have not been attempted, mass drug administration campaigns for W. bancrofti and O. volvulus control in Africa may help to dampen transmission of M. streptocerca.136

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DIAGNOSIS

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Section II PATHOGENS  

PART I: Nematode Infections 

PREVENTION AND CONTROL There are no studies of chemoprophylaxis for prevention of M. perstans infection, and control programs have not been attempted, although repeated doses of ivermectin (used for onchocerciasis control) have reduced the prevalence of M. perstans microfilaremia.184,185

MANSONELLA OZZARDI INTRODUCTION Infection with the filarial nematode Mansonella ozzardi is restricted to Central and South America and certain Caribbean islands. Although infected persons are usually asymptomatic, varied nonspecific symptoms have been ascribed to this infection.

THE AGENT M. ozzardi microfilariae were first described in blood films in 1897,186 and the adult worm was described in 1898.192 Infective larvae are transmitted to humans by bites of infected midges (Culicoides furens188–190 and other species191,192) or black flies (Simulium amazonicum192–194). There appears to be no difference between the parasites that are transmitted by the different vectors.195,196 Over months to years, these larvae develop into slender, threadlike adult worms (females, 32–51 mm × 130–160  µm; males, 24–28 mm × 150  µm), which probably inhabit the thoracic and peritoneal cavities.187,197 Adult worms have also been found in the lymphatics.198 In bisexual infections, microfilariae (sheathless, 3–5  µm × 170–240  µm) are produced and are found in the skin and blood, generally without periodicity.199–203

EPIDEMIOLOGY The number of people infected with M. ozzardi is unknown.23 The distribution of M. ozzardi is restricted to Central America, South America (Colombia, Venezuela, Guyana, Suriname, Brazil, Argentina, Bolivia) and certain Caribbean islands (Puerto Rico, Antigua, Guadeloupe, Nevis, Dominican Republic, Haiti, Martinique, St Kitts, St Lucia, St Vincent, and Trinidad). In highly endemic areas, 65–70% of people may have

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740

microfilaremia.204 Although nonhuman primates, other mammals, and certain birds and amphibians can be infected with M. ozzardi, humans are the only significant reservoir of infection. Like many other pathogenic filariae, M. ozzardi contains an endosymbiont of the Wolbachia genus.205

THE DISEASE Although M. ozzardi is generally thought to cause little or no disease in humans, several reports have clearly associated this infection with urticaria, lymphadenopathy, articular pains, pruritic skin eruptions, edema, headache, and pulmonary symptoms.171,204,206–213 Blood eosinophilia is commonly found.171,204,206,211,214–216

PATHOGENESIS AND IMMUNITY The pathogenesis of M. ozzardi infection is poorly characterized. Evidence from small numbers of expatriates suggests that immediate hypersensitivity may be responsible for some of the pathologic changes, eosinophilia, and IgE elevations seen in this infection.

DIAGNOSIS Definitive diagnosis of M. ozzardi infection is made by identifying microfilariae in blood or skin biopsies or by PCR in skin biopsies.217

TREATMENT AND PROGNOSIS Treatment of M. ozzardi infection has been problematic, since DEC207,218–222 and the benzimidazoles are ineffective against this parasite. In a single case report, ivermectin was effective in reducing symptoms and microfilaremia.215 In another study, four annual single doses of ivermectin (6 mg) reduced M. ozzardi microfilaremia levels by 82%.223 The presence of Wolbachia in M. ozzardi suggests that doxycycline may be an effective treatment.

PREVENTION AND CONTROL There have been no studies of chemoprophylaxis for prevention of M. ozzardi infections, and control programs have not been attempted.

SECTION II: PATHOGENS

PART I: Nematode Infections

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CHAPTER 106  Onchocerciasis Achim M. Hoerauf

INTRODUCTION Onchocerciasis, caused by the filarial nematode Onchocerca volvulus, is transmitted by black flies within tropical Africa, limited geographic areas of Latin America, and in Yemen. Pathology and disease symptoms such as keratitis, chorioretinitis, and forms of dermatitis are caused by offspring larvae, microfilariae, which migrate through the skin and the eye after release from adult worms that reside in subcutaneous nodules. Apart from this typical pattern of dermal and ocular disease in conjunction with rather high infection loads seen in people from endemic areas with a lifetime of exposure to the worms, a different syndrome develops in short-term visitors, with often stronger immunologic responses and usually light infection.1,2 In endemic areas, onchocercal disease is an important factor in higher mortality and economic loss.

THE AGENT O. volvulus is one of eight filarial species infecting humans. Several veterinary Onchocerca species not infecting humans but transmitted by the same black fly vector coexist in some endemic areas. When dissected from captured black flies, larvae of O. ochengi, O. gibsoni, O. lienalis, and O. cervicalis are morphologically indistinguishable from those of O. volvulus, but can be differentiated by polymerase chain reaction (PCR).3 O. volvulus uniformly harbors Wolbachia endosymbiotic intracellular bacteria,4 which are essential for fertility and reproduction5–8 as well as survival of the parasites.9 Substances released from dying Wolbachia play roles in treatment-induced reactions7 and in ongoing pathogenesis of human onchocerciasis.

History O’Neill in 1875 associated microfilariae present in skin with a papular dermatitis in Ghana.10 Leuckart in 1893 described adult worms in subcutaneous nodules from persons in West Africa. In 1917, Robles, a Guatemalan physician, demonstrated the association between nodules, skin lesions, anterior ocular lesions, and the presence of microfilariae.11 Several descriptions of ocular pathology including histologic proof of microfilarial presence in ocular tissues were made by Hisette in what was then the Belgian Congo.12

Life Cycle (Fig. 106.1) Humans are the only definitive host for O. volvulus, although chimpanzees have served as definitive hosts in experimental onchocerciasis. Black flies of the genus Simulium (Fig. 106.2A) are obligatory intermediate hosts. O. volvulus, like all nematodes, has a five-stage life cycle involving four molts. Infection begins with the bite of an infected Simulium vector.13 Infective larvae (L3 stage) are deposited into the skin of a new host, where

6–12 months is required for two molts and development of mature adult females (L5 stage) capable of producing new L1-stage larvae, called microfilariae. The white, hairlike adult worms live coiled in subcutaneous or deeper intramuscular tissues surrounded by a fibrous capsule (Fig. 106.2B) containing blood and lymph vessels. Adult males, which are 3–8 cm long, appear to migrate from nodule to nodule to inseminate the much larger females, which range from 30 to 80 cm in length. Microfilariae (220–360  µm long) are released from nodules to migrate through subcutaneous, conjunctival, and intraocular tissues. Adult females can live for up to 14 (average 9–10) years, each producing on average more than 700 microfilariae per day. Considerable numbers of adult females may lie in deep impalpable nodules.14 Microfilariae, which live for 6–30 months, are ingested by black flies. The two molts in the vector and maturation of the infective larvae (L3 stage) take 1–3 weeks. Any increase in adult worm burden necessarily implies reexposure to infective L3 from a vector, because O. volvulus adult worms do not reproduce within humans.

Vectors Six species of the Simulium damnosum sensu lato complex, identified based on fly morphology and on chromosome banding patterns, are vectors of O. volvulus in West Africa. Based on habitats and mitochondrial gene sequences,15,16 further grouping into a savanna clade (S. damnosum sensu stricto, S. sirbanum), a rain forest clade (S. yahense, S. squamosum), and a transition zone clade (S. leonense, S. sanctipauli) is possible. S. neavei is the principal vector in eastern and central Africa;17,18 S. ochraceum is the vector in Central America;19 and several other species transmit onchocerciasis in South America. Most Simulium species lay their eggs attached to rocks and vegetation submerged in highly oxygenated stretches of rivers and streams, where larval and pupal stages develop. In savanna areas in Africa, the vector breeds in large rivers such as the Volta, while in forest areas of Africa and the Americas the vector breeds in streams and rivulets that are less accessible to larviciding by aircraft. Larvae of S. neavei attach to crabs living in small creeks for further development. Flight range of Simulium is 12 km, and therefore transmission areas are limited to these distances away from breeding sites.

EPIDEMIOLOGY Onchocerciasis is endemic in 34 countries: 27 in Africa, 6 in Latin America, and 1 (Yemen) in the Arabian peninsula (Fig. 106.3). From data derived from a new mapping technology based on palpation of nodules in randomly selected communities,20 approximately 90 million people are at risk, with 37 million people infected. About 270 000 individuals developed blindness from onchocerciasis and another 500 000 have severe visual disability.7,21 Global disease burden in 2002 was 0.95

741

Larvae develop into adults in subcutaneous nodules where they may live up to 15 years

PART I: Nematode Infections 

Microfilariae released and migrate throughout subcutaneous tissue (diagnostic) and eye for 1–2 years causing: Pruritus, dermatitis (including sowda) Lymphadenopathy Punctate and sclerosing keratitis, blindness

Human Extrahuman

 

PATHOGENS

Section II

Onchocerca volvulus

Infective 3rd stage larvae enter skin with black fly (Simulium) bite

1st–3rd stage larvae develop in thorax muscles and proboscis of black fly over 7–9 days

Microfilariae taken up by black fly bite (with salivary anticoagulant)

Black flies breed in oxygen-rich flowing streams and rivers

Figure 106.1  Life cycle of Onchocerca volvulus.

have a higher risk of infection and become infected at a younger age and with increased microfilaria intensity, suggesting intrauterine O. volvulusspecific immunosuppression.33 In regions outside the savanna, skin manifestations are the main complications of disease. The profound psychosocial implications of unremitting pruritus and disfiguring skin lesions make onchocercal skin disease a major public health problem and more than 50% of DALYs lost due to onchocerciasis are due not to blindness but to skin disease.34,35 On a community basis, there is a strong correlation between prevalence of pruritus and O. volvulus endemicity in the community. In communities hyperendemic for O. volvulus (defined as >60% of people with microfilardermia and >30% with palpable nodules (onchocercomas)), 30–40% have symptomatic skin disease and 50% of children aged 5–9 are already infected. Adult persons may harbor up to 50 adult worms and >100 microfilariae/ mg skin. In mesoendemic areas, 30–60% of people and in hypoendemic areas 8000/mL blood (see Chapter 105). Rapid epidemiologic assessment tools have been developed to allow identification of a community as high risk for such occurrence and providing a tool for decision making in co-endemic areas.138,139 An alternative may be the administration of doxycycline in such communities, which does not affect Loa (since Loa does not contain Wolbachia) and reduces O. volvulus microfilariae only slowly; a pilot study is under way in Cameroon.140 OEPA (Onchocerciasis Elimination Programme in the Americas) was launched in 1991 and aims, by 2012, to eliminate transmission, which is defined as 40% of white blood cells), lymphadenopathy, and elevated liver enzymes (ALT) as compared with patients with other causes for fever. Among 41 patients with acute illness associated with HIV seroconversion,130 the most common symptoms were fever, sore throat, fatigue, weight loss, and myalgia. The median duration of symptoms was 14 days though symptoms persisted 4 weeks or longer in 32%.130 Physical findings in patients with acute HIV infection often include rash (70%) and lymphadenopathy (77%).246 In the evaluation of a person with prolonged fever, it is important to determine the HIV status, since the range of infections and the probabilities of each will be altered if the person is HIV infected. Causes of chronic or persistent fevers include protozoa, e.g., malaria, amebic liver abscess, trypanosomiasis (American and African), and visceral leishmaniasis; chronic bacterial infections such as brucellosis,247 tuberculosis, bartonellosis, Q fever, and melioidosis; fungal infections, such as histoplasmosis, coccidioidomycosis, and paracoccidioidomycosis;248 and a few helminthic infections, such as visceral larva migrans, fascioliasis, and schistosomiasis. Visceral leishmaniasis is an important cause of chronic fevers that can be acquired in some temperate as well as tropical areas of the world.249–251 For example, of 89 patients with visceral leishmaniasis in France in 1986– 1987, 70 (79%) had acquired the infection in France. Imported cases came primarily from other Mediterranean countries.249 In recent years visceral leishmaniasis has emerged as an important opportunistic infection in persons with acquired immunodeficiency syndrome (AIDS) living in southern France, Spain, and Italy.252 Melioidosis can reactivate years after the initial exposure. Manifestations are protean. Infection can involve lung, skin, and soft tissues and any part of the body via bacteremic spread. Pulmonary changes, including cavitation, can mimic tuberculosis.253,254 In a series of 602 patients seen in one Thai hospital, in-hospital mortality was 42%. Among 118 adult patients with long-term follow-up, 23% had culture-proven relapses, occurring 1–290 weeks after discharge (median, 21 weeks).255 Recrudescent infection has been reported as long as 26 years after initial infection.256 Ehrlichiosis is another potentially treatable cause of prolonged fever. In a series of 41 cases of human ehrlichiosis, 6 patients manifested protracted fever (range 15–51 days) as the principal finding.257 Chronic or recurrent fevers can be the result of uncommon complications of an infection; for example, Q fever endocarditis, brucella osteomyelitis or endocarditis, and splenic or liver abscesses secondary to melioidosis.258 In a prospective study of 530 patients with brucellosis in Spain followed for at least 1 year after treatment, 86 patients relapsed (97 relapse episodes). Among recognized relapses, 95% occurred within 6 months of the end of therapy.259 Relapsing fevers are commonly seen due to Borrelia260–263 and with malaria. They can also occur with cholangitis (which can be associated with parasites as well as with stones and other biliary disease), chronic meningococcemia,264 rat-bite fever (Streptobacillus moniliformis/Spirilum minus), brucellosis, filariasis, infective endocarditis, visceral leishmaniasis, trypanosomiasis, Whipple’s disease,198 Hodgkin’s disease, familial Mediterranean fever, and many other diseases.265,266 Pyomyositis can cause undifferentiated fever in its early stages. It is usually seen in persons who have lived in tropical areas. The fever that accompanies acute schistosomiasis (Katayama syndrome) and fascioliasis235 can persist for weeks. In an outbreak of

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syndrome.54 In an outbreak of acute schistosomiasis after exposures on safari in Tanzania, 37% developed a rash, usually urticarial (typically beginning 3 weeks or longer after exposure), often with onset prior to development of fever.233 Among 75 patients with acute schistosomiasis seen in Israel, 71% had fever and 45% had skin lesions, but both were present at the same time in only 26%.234 Erythema nodosum and other skin lesions may be present in acute coccidioidomycosis, histoplasmosis,187 and mycobacterial infections including tuberculosis. Urticarial skin lesions in the person with fever in addition to hypersensitivity reactions related to drugs should bring to mind early hepatitis B and several helminthic infections (e.g., acute schistosomal infection or Katayama syndrome, fascioliasis, trichenellosis, loiasis, gnathostomiasis, and others). Fever is not a prominent finding in many helminthic infections, but of 20 patients with acute fascioliasis diagnosed at a university hospital in Spain, 12 (60%) had fever and 4 (20%) had urticaria. Although patients with fascioliasis can have undifferentiated fever, the presence of abdominal or right upper quadrant pain and eosinophilia are findings that can suggest this infection.235 Vasculitic skin manifestations are associated with a variety of infections, collagen vascular diseases, and antimicrobial drugs.236

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Box 130.4  Persistent and Relapsing Feversa Bacterial Bartonellosis Brucellosis Ehrlichiosis Endocarditis (multiple causes) Leptospirosis Lyme disease Melioidosis Q fever Relapsing fever Rickettsial infections (several) Syphilis Tuberculosis and nontuberculous mycobacteriosis Tularemia Typhoid fever Fungal Blastomycosis Coccidioidomycosis Cryptococcosis Histoplasmosis Paracoccidioidomycosis Penicilliosis (Penicillium marneffei)

Protozoan Amebic liver abscess Babesiosis Leishmaniasis (visceral) Malaria Toxoplasmosis Trypanosomiasis Viral Cytomegalovirus infection Human immunodeficiency virus infection Helminthic Angiostrongyliasis due to Angiostrongylus costaricensis Clonorchiasis Fascioliasis Filariasis Gnathostomiasis Loiasis Opisthorchiasis Paragonimiasis Schistosomiasis Toxocariasis Trichinellosis

Reactivation of Latent Infection or Recrudescence of Inapparent Infection Brill–Zinsser disease (Rickettsia prowazekii) CNS Chagas’ disease Coccidioidomycosis Histoplasmosis Leishmaniasis Malaria (Plasmodium malariae) Melioidosis Paracoccidioidomycosis Strongyloidiasis Toxoplasmosis Tuberculosis Persistent Infection and Recurrent Symptoms or Progressive Disease Chagas’ disease Filariasis Leprosy Loiasis Onchocerciasis Syphilis

a

List includes infections that may cause fevers with duration exceeding 3 weeks. Infections and other illnesses unrelated to tropical exposures (e.g., endocarditis, cholangitis, Hodgkin’s disease, others) also can be associated with chronic and relapsing fevers.

schistosomiasis among travelers, 15 of 29 who were infected developed Katayama syndrome. The median duration of symptoms was 12 days (range 4–46 days).54 Visceral larva migrans267 can cause prolonged or intermittent fevers. The associated findings of leukocytosis, eosinophilia, and enlarged liver suggest the diagnosis. Box 130.4 lists some of the more common causes of persisting and intermittent or relapsing fevers.

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Box 130.5  Sequelae of Infections Acquired more than 10 years earlier: Mechanisms and Examples

Some infections can become clinically manifest years or decades after a person has left the place of acquisition. Late manifestations also may result from chronic sequelae of an earlier infection, even if active infection no longer is present. Persons with long residence in tropical areas are more likely to have acquired infections with late sequelae (Box 130.5). The mechanisms by which remotely acquired infections can cause disease are several. Latent infection may reactivate, causing fevers and focal or systemic symptoms. Examples include tuberculosis, histoplasmosis, paracoccidioidomycosis, melioidosis, and leishmaniasis. Low-grade, persistent infections may expand (recrudesce), causing symptoms leading to diagnosis (e.g., malaria, Brill–Zinsser disease). The balance between host and microbe may tip in favor of the pathogen in persons whose immunity has been altered by disease, drugs, or age. Strongyloides larvae may disseminate widely outside their usual gastrointestinal habitat; for example, in persons with cell-mediated immunosuppression, particularly those taking corticosteroids or coinfected with human T-cell lymphotropic virus type 1. Salmonella typhi may be carried in the biliary tree and remain inapparent unless mechanical (e.g., biliary obstruction) or other factors alter the local milieu. Worms with long life spans may breach a tissue barrier or cause chronic inflammatory changes leading to acute symptoms.

Mechanical Effects; Loss of Normal Function; Scarring, Fibrosis, Tissue Destruction Cysticercosis Echinococcal cysts Liver flukes Onchocerciasis (blindness) Schistosomiasis Tuberculosis (e.g., cavities and bronchiectasis in lungs; scarring and obstruction in urinary tract; adrenal insufficiency) Malignancya Schistosoma haematobium (bladder cancer) Epstein–Barr virus Hepatitis B and hepatitis C viruses (hepatocellular carcinoma) Human immunodeficiency virus Human T-cell lymphotropic virus type 1 Liver flukes Allergic, hypersensitivity reactions Rupture of echinococcal cyst

a

Papillomaviruses are also associated with cancers, but fever is not a feature of these infections.

Migration of ascaris can provoke acute pancreatitis or cause biliary obstruction. Remote infection may cause scarring, obstruction, or alteration in structure that may predispose to superimposed infections or affect function. Scarring associated with renal tuberculosis can increase the risk of urinary tract infection with Escherichia coli and other uropathogens, even if tuberculosis is inactive or has been treated and cured. Scarring of the urinary tract secondary to Schistosoma haematobium can be associated with bacterial infections and an increased risk of squamous cell carcinoma of the bladder. Late sequelae of schistosomiasis may include portal and pulmonary hypertension, liver cirrhosis, and polyps and fistulas of the bowel. Echinococcal cysts can impinge on the biliary tree, causing acute obstructive symptoms, or erode into the biliary tree and cause symptoms of acute cholangitis. Rupture of an echinococcal cyst can cause acute allergic symptoms, even anaphylaxis. Seizures can occur in patients with cysticercosis, even if parasites are no longer viable. Late complications of Chagas’ disease typically do not include fever unless progression of infection occurs (e.g., in persons with AIDS or immunocompromised transplant patients).268–270 Clinical findings of hepatocellular malignancy related to remotely acquired, persistent infection with HBV or HCV may include fever secondary to tissue necrosis and the effects of the tumor.

Processes Other Than Infection Causing Fever After Travel Travel itself may predispose to problems that cause fever. In their series of 24 patients with drug-induced hypersensitivity syndrome requiring hospitalization between 2004 and 2008, Ben m’rad and colleagues demonstrated that 50% of culprit drugs were antibiotics (n = 8) or antimalarials (n = 4), all of which could be used by the traveling population.271 One hundred percent of cases had fever and skin involvement,

Special Patients (HIV-infected, Immunocompromised, Pregnant) Many infections are more common, more severe, or have altered clinical expression in persons who are immunocompromised. Opportunistic infections, such as tuberculosis, visceral leishmaniasis,278 American trypanosomiasis, and histoplasmosis and other fungal infections may reactivate and become clinically apparent long after the infection was acquired. An evaluation of 50 HIV-infected patients in Spain with unexplained fever lasting at least 4 weeks found the most frequent diagnoses were tuberculosis (42%), visceral leishmaniasis (14%), and disseminated Mycobacterium avium complex (14%).279 Disseminated infection with the fungus Penicillium marneffei is occasionally seen in travelers, typically persons immunocompromised by HIV infection or other disease.280 In its endemic area in Thailand and Southeast Asia it is an important opportunistic pathogen.281 It is important to know during the evaluation whether a female patient is pregnant. The cause of fever may threaten the pregnancy, diagnostic tests and therapies may be harmful to the fetus, some infections can be transmitted transplacentally, and other infections, such as malaria, influenza, and hepatitis E, are more severe in pregnant than in nonpregnant women. Although Q fever is likely to be asymptomatic in pregnant women, infection is significantly associated with fetal morbidity and mortality.282

EVALUATION OF THE PATIENT WITH FEVER Patients with fever and recent tropical exposures always deserve careful evaluation even though similar symptoms might be treated casually in a person without a history of travel. It is helpful to construct a differential diagnosis encompassing potential bacterial, viral, parasitic, and fungal pathogens based on initial history, geographic exposures, and clinical findings. Patients with confusion, hypotension, hypoxemia, or hemorrhagic skin rash need immediate attention and care. A more leisurely pace may be appropriate in patients with subacute or chronic fevers. Any patient with a potential exposure to malaria who is febrile or gives a history of fevers or chills must be evaluated promptly for malaria.283–286 In general, most infections with the potential for rapid progression and fulminant course (e.g., falciparum malaria, hemorrhagic fevers, meningococcemia, plague, rickettsial infections) become manifest within a month of exposure. In thinking about the causes of fever in a person with diverse geographic exposures, one should always think first about possible diagnoses had the person not traveled, and then expand the differential diagnosis to include infections that may be related to exposures during travel. Omitting the first step can lead to pursuit of obscure or exotic diseases when the patient has a common readily treatable infection, such as acute pyelonephritis or streptococcal pharyngitis. Initial studies will generally include a complete blood count (CBC) with a differential leukocyte and platelet estimate, thick and thin malaria smears, blood cultures, urinalysis, and liver function tests. Serum should be saved early in the course in the event serologic studies are indicated. Chest radiographs should be obtained in patients with pulmonary symptoms or signs and in persons with persistent, unexplained fever. Serologic studies for HIV infection should be requested in persons with persistent fevers and those with possible exposures during travel. Studies that may yield useful information if initial studies are unrevealing include repeated malaria smears, blood cultures, serial CBCs, differentials, liver function

Box 130.6  Approach to Patient with Fever and History of Tropical Exposure Initial Evaluation of Acute Fever Complete blood count with differential Liver enzymes and function tests Blood culture: bacterial, viral, fungal (rarely) Urinalysis (culture if abnormal) Blood smears for malaria Other tests • Serologic tests (save acute serum) • Urinary antigens (e.g., Legionella species, Histoplasma) • Blood smears for Babesia, Borrelia causing relapsing fever • Bone marrow aspiration/ biopsy: pathology and culture Tests to Consider for Focal Symptoms and Findings Cough: chest film; sputum Gram stain, acid-fast stain, stains for fungi; cultures for bacteria, mycobacterial species, fungi; wet preparation for ova (Paragonimus species) or larvae (hyperinfection with Strongyloides stercoralis); bronchoscopy Diarrhea: examination for fecal leukocytes (lactoferrin where available); blood (stool guaiac); toxin and culture for Clostridium difficile; ova and parasites, stool cultures; fecal antigens; endoscopy Sore throat: rapid streptococcal antigen test, culture, infectious mononucleosis absorption test (Monospot) (in appropriate clinical setting) Skin lesions: aspirate, scrapings, biopsy, Gram stain, acid-fast stain, fungal stain, Wright– Giemsa stain (Leishmania); culture for bacteria, fungi, mycobacteria, Leishmania

Lymphadenopathy: aspirate or biopsy, acid-fast stain, fungal stain, Wright–Giemsa stain (trypanosomes); culture Genital lesions or symptoms: pelvic examination, dark-field examination of ulcers, cultures and urine sample for Neisseria gonorrhoeae and Chlamydia trachomatis, wet preparation for white blood cells, Trichomonas vaginalis, Candida albicans, and clue cells (bacterial vaginosis), vesicle exudate for electron microscopy CNS: CT scan, MRI, lumbar puncture with opening pressure and examination of CSF for cells, protein, glucose, bacterial antigens; cultures for bacteria, Mycobacterium tuberculosis, fungi, viruses; India ink for Cryptococcus neoformans and cryptococcal antigen; VDRL, PCR for specific antigens/ microorganisms; rarely, brain biopsy Abdominal pain: liver enzymes, amylase, lipase, lactate, stool examination (see above), abdominal radiographic series and/or ultrasound, CT scan, MRI Arthritis: aspirate – examine fluid for cells, protein, crystals; stains and cultures for bacteria, fungi, Mycobacterium species Cardiac findings: echocardiogram (transesophageal or transthoracic), blood cultures (at least three sets; hold for 21 days; consider lysis centrifugation), serology for Q fever, and brucellosis, PCR for Tropheryma whipplei, scan chest and abdomen for other possible foci of infection

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while 58% had peripheral edema, 54% had evidence of hepatic dysfunction, and 42% had hypotension.271 Pulmonary emboli after prolonged air flights272,273 may be associated with fever.274 Noninfectious diseases also cause fevers and should be considered if studies do not document infection.239,266,275,276 Biopsy may be necessary to confirm a diagnosis in processes such as inflammatory bowel disease, malignancies, and lymphadenopathy of Kikuchi.277 Hyperthyroidism may present with fever and diarrhea. In the Belgian study, 2.2% of the fevers after tropical travel were not caused by infections.13

CNS, central nervous system; CT, computed tomography; MRI, magnetic resonance imaging; CSF, cerebrospinal fluid; VDRL, Venereal Diseases Research Laboratory; PCR, polymerase chain reaction.

testing, and abdominal ultrasound. Repeated physical examinations may yield new information with the development of new skin findings, lymphadenopathy, or a tender liver or spleen. Although many of these findings may not give the diagnosis, they may help refine the list of considerations, allowing a more focused and efficient path to the diagnosis. Occasionally a patient will have two unrelated infections, both requiring specific interventions.287 Box 130.6 outlines an approach to patients with fever and a history of tropical exposures.

EMERGING DISEASES AND DIFFICULTIES Diseases are not fixed in distribution, clinical expression, or response to antimicrobials.28 As the hepatitis A and hepatitis B vaccines are now more

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widely used, these infections have become uncommon in travelers to developing countries. Conversely, increasing antimicrobial resistance of bacteria and protozoa may increasingly limit choices for prophylaxis, empirical therapy, and treatment. Persons who are older and those with chronic diseases are traveling more regularly and to more remote destinations. Long-term travel is also becoming increasingly popular, and carries its own set of unique risks.288 Multiple diverse geographic exposures are

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becoming a common part of the carefully taken medical history. New technologies, such as polymerase chain reaction, may reduce some of the frustrations currently encountered in making rapid specific diagnoses. Even with these advances in diagnosis and communication, evaluation and management of the patient with fever will continue to be a challenge, requiring a thoughtful and systematic approach and broad knowledge or access to it.

Section III: PRACTICE: APPROACH TO THE PATIENT IN THE TROPICS

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CHAPTER 131  Eosinophilia Mary E. Wilson • Peter F. Weller

INTRODUCTION Eosinophilia develops as an immunologically mediated response in association with diverse processes, including allergic, neoplastic, and infectious diseases (Box 131.1). Eosinophilia is a hematologic marker that warrants attention and serves as a clue to help direct a diagnostic workup. Although many diseases are associated with eosinophilia, the presence of eosinophilia in a person with tropical exposures suggests the possibility of specific parasitic infections. Eosinophilia is notably common in association with helminthic parasites, especially those in tissues.1 Before embarking on an exhaustive search for parasitic infections in a patient with eosinophilia, however, it is prudent to consider noninfectious causes of eosinophilia, such as allergic diseases and drug-induced hypersensitivity reactions.2 Eosinophilia in parasitic infections is the result of dynamic interactions that are influenced by host factors, the stage of parasite development, the location of the parasite within the human host, and the parasite burden, among other factors. Eosinophilia may wax and wane. Parasites often associated with eosinophilia may not provoke an identified eosinophilic response in all patients or at all times. Parasites may elicit a localized eosinophilic tissue response that is not reflected by an increase in peripheral blood eosinophil numbers. This chapter considers the immunobiology of eosinophils, the causes of eosinophilia with special attention to parasitic infections, and the evaluation of patients with eosinophilia.

Overview Most unicellular pathogens do not provoke an eosinophilic response.1,2 In contrast, multicellular, helminthic parasites are common, potent stimuli for eosinophilia (Box 131.2). Relative to most bacterial and viral pathogens, helminths are large and have a long life span, often measured in years or even decades. Unlike viral and bacterial pathogens that multiply in human hosts, most helminths cannot undergo reproductive cycles in humans; hence the numbers of adult helminths is limited by the number of eggs or larval forms acquired by the human. Prolonged or repeated exposures may be required for a human to acquire sufficient parasites to cause symptomatic infection. Many helminthic infections cause few or no symptoms or only intermittent findings. In addition, because of the lengthy developmental period and longevity of helminthic parasites, symptoms may begin months or years after exposures in endemic regions. Thus, the time period between possibly relevant exposures and the onset of clinical or other evidence of infection may be longer than usually considered in obtaining a medical history for most clinical evaluations. Most protozoan infections are not associated with eosinophilia. Exceptions are Isospora belli3 and possibly Dientamoeba fragilis,4 which have been reported to be associated with eosinophilia. In some reports of these infections, other causes of eosinophilia had not been excluded. Eosinophilia and eosinophilic myositis have been reported in association with the coccidian parasite Sarcocystis.5 In general, however, the finding of eosinophilia

in a patient with a protozoan infection should prompt a search for a concomitant helminthic infection or an alternative noninfectious process to explain the eosinophilia. Only a few fungal and, rarely, specific viral infections are associated with eosinophilia (Box 131.3). Acute bacterial and viral infections characteristically produce eosinopenia.2 Likewise, acute protozoan infections, including malaria,6,7 will suppress eosinophilia, including that caused by helminthic infections, during the intercurrent infection. Bacterial superinfections that complicate hyperinfection strongyloidiasis contribute to suppression of eosinophilia that might otherwise be a salient clue to the presence of strongyloidiasis. When eosinophilia is observed in bacterial infections, it may be consequent to hypersensitivity reactions to antibiotics used for treatment. The fungal diseases associated with eosinophilia are: aspergillosis, in the form of allergic bronchopulmonary aspergillosis,8 coccidioidomycosis,9 and basidiobolomycosis.10,11 Blood eosinophilia, peaking during the second or third week of illness, occurs with primary coccidioidomycosis. Eosinophilia also may develop with disseminated coccidioidomycosis.12,13 Eosinophilia has been reported in children with paracoccidioidomycosis, an infection whose involvement of adrenal glands14 perhaps is causing eosinophilia due to hypoadrenalism. A few reports note eosinophilia in patients with ectoparasitic infestations caused by myiasis15 and scabies.16

PATHOGENESIS Eosinophils are bone marrow-derived leukocytes. Eosinophilia develops when three specific cytokines, granulocyte–macrophage colonystimulating factor (GM-CSF), interleukin (IL)-3, and IL-5 stimulate enhanced eosinophilopoiesis. Of these, IL-5 is principally responsible for increases in eosinophilopoiesis and eosinophilia in helminthic, allergic, and other diseases. In humans, enhanced eosinophilopoiesis within the marrow requires over a week to increase blood eosinophilia. IL-5 also acts more rapidly to increase blood eosinophilia by mobilizing a marginated pool of preformed eosinophils resident within the marrow. IL-5 is produced by Th2-like CD4+ T lymphocytes (as well as other cells, including eosinophils); and eosinophilia is frequent with immune responses characterized by Th2-like T-cell activation, including those elicited by helminthic parasite infections and associated with allergic diseases.17 In both of these, enhanced immunoglobulin E (IgE) production is usually present.17 GM-CSF, IL-3, and IL-5 also act on mature eosinophils to prolong their survival by antagonizing apoptosis and to enhance their effector functions. In other diverse diseases associated with eosinophilia (Box 131.1), mechanisms leading to eosinophilia are not yet delineated. Although Th2-like T-cell responses lead to eosinophilia, it remains uncertain how specific parasitic or allergic diseases characteristically stimulate Th2-type responses. Specific features of helminthic antigens, as well as their particulate presentation, may influence the initial antigen presentation process that leads to Th2-like lymphocyte responses and consequent eosinophilia. As noted below, eosinophilia is most prominent

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Box 131.2  Key Concepts: Eosinophils in Parasitic Infections

“Allergic” Diseases Atopic and related diseases Medication-related eosinophilias

PRACTICE: APPROACH TO THE PATIENT IN THE TROPICS

Hematologic and Neoplastic Disorders Hypereosinophilic syndromes, including forms of chronic eosinophilic leukemia, lymphocytic variants, and idiopathic forms Lymphomas, especially nodular sclerosing Hodgkin’s; some Tand B-cell lymphomas Tumor-associated: on occasion with large-cell nonkeratinizing cervical tumors, large-cell undifferentiated lung carcinomas, squamous carcinomas (vagina, penis, skin, nasopharynx), adenocarcinomas (stomach, large bowel, uterine body), and transitional bladder cell carcinoma Mastocytosis

Elevation is caused primarily by helminthic parasites that reside in tissues. Most protozoan parasitic infections are not associated with eosinophilia Eosinophils may be prominent during only one stage of parasite development. Migration of parasites within tissues is often associated with high-grade eosinophilia. Chronic parasitic infections in antigenically sequestered sites (e.g., echinococcal cysts) or solely within the gut lumen (e.g., adult Ascaris or tapeworms) may not provoke an eosinophilic response. Release of antigenic material when parasites die or when walls protecting parasites are breached may lead to increased eosinophilia Eosinophil levels may wax and wane; many factors unrelated to the helminthic infection influence the level of blood eosinophilia. Acute bacterial, viral, and protozoal (e.g., malaria) infections suppress eosinophilia High-grade eosinophilia and symptoms may be prominent during the prepatent period and before the diagnosis can be confirmed by finding eggs or diagnostic forms of the parasite in tissues. Serologic tests may help in early diagnosis

  

 

Box 131.1  Eosinophil-associated Diseases and Disorders

Infectious Diseases Parasitic infections, mostly with helminths Specific fungal infections Other infections – infrequent

Pulmonary diseases (e.g., acute or chronic eosinophilic pneumonia, allergic bronchopulmonary aspergillosis) Gastrointestinal diseases (e.g., eosinophilic gastroenteritis) Neurologic diseases (e.g., eosinophilic meningitis) Rheumatologic diseases (e.g., Churg–Strauss syndrome vasculitis) Cardiac diseases (e.g., endomyocardial fibrosis) Renal diseases (e.g., druginduced interstitial nephritis, eosinophilic cystitis, dialysis) Immunologic Reactions Specific immunodeficiency diseases (hyper-IgE syndrome, Omenn’s syndrome) Transplant rejection Endocrine Hypoadrenalism Other Atheroembolic disease Irritation of serosal surfaces Inherited

Diseases with Specific Organ Involvement Skin and subcutaneous diseases (e.g., blistering diseases, bullous pemphigoid, pemphigus vulgaris, dermatitis herpetiformis, herpes gestationis, and drug-induced lesions)

with helminthic infections when parasites are migrating through or beginning to localize within tissues.

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Eosinophils are primarily tissue-dwelling leukocytes, normally several hundredfold more abundant in tissues than in blood. Eosinophil numbers are greatest in tissues with a mucosal epithelial interface with the environment, including the gastrointestinal (GI) and lower genitourinary (GU) tracts. The life span of eosinophils is longer than that of neutrophils. Eosinophils probably survive for weeks within tissues. The immunologic functions of eosinophils remain the subject of intense investigation. In addition to functioning as endstage effector leukocytes, eosinophils likely have additional roles in interacting with lymphocytes and other cells.18 As effector cells, eosinophils are capable of releasing specific lipid mediators, such as leukotriene C4, and are a source of a range of preformed, granule-stored cytokines. In addition, eosinophils uniquely contain specific, highly positively charged (“cationic”) proteins within their cytoplasmic granules. These eosinophil granule cationic proteins include major basic protein and eosinophil cationic protein. The release of these granule proteins can damage host tissues and more beneficially can contribute to the killing of helminthic parasites. The extent to which eosinophils are involved in the immune responses to helminthic parasites, and especially their early larval forms, to help kill these parasites in vivo is still not certain.17,19

The degree of eosinophilia and other clinical manifestations caused by helminthic infections may differ between long-term residents of endemic regions and short-term visitors, being more prominent in the latter Many infections associated with eosinophilia cause no or intermittent symptoms; first symptoms may develop months or years after exposure. Many helminths are long-lived with life spans that can exceed a decade Multiple infections may be present, especially in long-term residents of tropical areas. Finding intestinal eggs or certain parasites does not confirm that they are the cause of the eosinophilia. In a patient with Ascaris eggs and moderate or high-grade eosinophilia, the search for the cause of eosinophilia should continue Absence of eosinophilia does not exclude a parasitic infection typically associated with eosinophilia

Box 131.3  Infections Associated with Eosinophilia Viral Human immunodeficiency virus-1 – on occasion, may be associated with skin eruption

Protozoan Isospora belli Dientamoeba fragilis

Fungal Aspergillus – only with allergic bronchopulmonary aspergillosis Coccidioides immitis – acute and sometimes later disseminated infections Basidiobolomycosis Paracoccidioidomycosis

Infestations Scabies Myiasis

Helminthic Many (see Table 131.1)

In patients with marked eosinophilia, eosinophils may cause organ damage, most notably to the heart. The damage to the heart, ranging from early necrosis to subsequent endomyocardial thrombosis and fibrosis, is the same with varied eosinophilic conditions. Thus, endomyocardial thrombosis and fibrosis develop with eosinophilia associated with the hypereosinophilic syndromes,20 carcinomas and lymphomas, and parasitic infections, including at times trichinellosis, visceral larva migrans, loiasis, or other filarial infections.21–25 Pathologically, eosinophilic endomyocardial fibrosis26 is identical to tropical endomyocardial fibrosis.27,28 While diverse eosinophilic diseases can cause identical forms of cardiac disease, many patients with sustained eosinophilia never develop cardiac disease. Thus, the pathogenesis of eosinophil-mediated cardiac damage involves both the presence of increased eosinophils and other, as yet ill-defined, stimuli for recruitment or activation of these leukocytes.

Acute bacterial and viral infections; acute malaria Corticosteroids – endogenous and exogenous Acute stress Pregnancy (counts especially low during stress of delivery)

Immunosuppressive states (also less elevation of immunoglobulin E; sensitivity of serologic tests may be lower) Epinephrine (sharp fall after initial transient rise; can be inhibited by β-blockers)

Host Factors that Influence Eosinophil Responses Blood eosinophilia is clearly present when eosinophils are in excess of 450 cells/mL blood. Published studies use cutoffs that range from 350 to 500 cells/mL blood. Increased percentages, but not absolute numbers, of blood eosinophils (pseudoeosinophilia) due to leukopenia in other white blood cell lines can be misleading. Many host factors and other stimuli influence peripheral blood eosinophil levels (Box 131.4). Blood eosinophil numbers vary diurnally, with levels being higher in the early morning and lower at about noon. Levels are higher in the neonatal period and decline with age. Eosinophil levels fall during pregnancy; during the stress of delivery they almost disappear from the peripheral circulation.29 Epinephrine causes a sharp fall in circulating eosinophils, after a transient increase. Injection of 100 mg hydrocortisone is followed by a decrease in peripheral eosinophils to 35% of control levels within 1 hour, and blood eosinophils are nearly absent 4 hours after the steroid administration.30 Corticosteroids inhibit tissue accumulation of eosinophils, probably by several mechanisms, including promoting eosinophil apoptosis.31 Acute bacterial and viral infections and other processes causing acute inflammation are associated with a decline in and transient suppression of blood eosinophilia.30,32 Eosinophils drop during acute malaria, even if previously elevated.6,7 Depression of eosinophils in malaria persists beyond the clearance of fever and parasitemia, and is followed by an increase in eosinophils in convalescence.7 When it is important to know whether a patient has eosinophilia, a blood differential count should be repeated after a patient has recovered from an acute intercurrent infection.

Patterns Characteristics of the eosinophil response that may be useful in assessing more likely causes of eosinophilia include level, duration, pattern (constant versus intermittent), and associated symptoms. If a person has had tropical exposures, the timing of onset of eosinophilia in relation to those exposures may help in the evaluation. Eosinophilia should be characterized by level, not simply as present or absent. Absolute eosinophil counts of greater than 3000/mL are categorized as marked or high-grade eosinophilia. Several different patterns of eosinophilia may be seen in relation to parasitic infections. Factors that influence these patterns include the type of parasite, the stage of parasite development, the location within the human host, the integrity of barriers between parasite and host, and the viability of the parasite, as well as many attributes of the host. In some helminth infections, eosinophilia is prominent only during one stage of parasite development. An example is the intestinal parasite Ascaris lumbricoides, which provokes eosinophilia principally during the initial stage of larval migration through the lungs (see Chapter 115). Adult worms residing in the lumen of the gut typically do not cause an eosinophilic response. Some parasites, such as the tapeworms Diphyllobothrium latum and Taenia saginata, whose entire life in the human host takes place in the lumen of the gut, cause little or no eosinophilic response even though they can grow to impressive lengths and survive for decades. Table 131.1 delineates the many helminthic infections associated with eosinophilia, the main body site affected by each and candidate diagnostic tests for each; fuller details are found in pathogen-specific chapters.

Several helminthic parasites may induce high-grade eosinophilia (Table 131.2) during one stage of development followed by chronic low-to-moderate levels of eosinophilia. Acute schistosomiasis (Katayama syndrome)33,34 (see Chapter 122) and the larval migration stage of Ascaris35 and hookworms36 may provoke high-grade eosinophilia, which declines during the chronic stages of infection. In experimental hookworm infections, blood eosinophilia increases progressively after 2–3 weeks of infection and peaks between 5 and 9 weeks before gradually diminishing. In untreated hookworm infections, eosinophilia slowly diminishes but can persist for several years after experimental infection.37 The blood and pulmonary eosinophilia, often characterized as “transient,” associated with the pulmonary migration phases of Ascaris and hookworms typically persists for weeks or months.35,38,39 The eosinophilia with strongyloidiasis (see Chapter 117) often fluctuates over time, being high during pulmonary migration and low to moderate during chronic infection. During hyperinfection syndromes, eosinophilia can be prominent or absent, influenced by host characteristics and suppression due to corticosteroids or concomitant bacterial infection.40 The dramatic eosinophilia seen with acute trichinellosis usually disappears when a fibrous capsule forms around the larvae in muscles (except with the nonencapsulating species Trichinella pseudospiralis, which may cause a prolonged eosinophilic myositis41) (see Chapter 110). Parasites encysted in tissues, such as echinococcal cysts and cysticercosis, and physically isolated from the host by cyst walls, typically cause no eosinophilia unless disruption of the barrier allows leakage of antigen-rich material. Intermittent leakage of fluids from echinococcal cysts can transiently stimulate increases in blood eosinophilia and elicit allergic (urticaria, bronchospasm) or anaphylactic reactions.42,43 Breaching of these barriers or disintegration with death of the parasite can lead to intense tissue reactions, increased eosinophilia, and acute symptoms in the host. Adult parasites that live and migrate in tissues, such as Loa loa and Gnathostoma spinigerum, provide an ongoing stimulus to eosinophils.24,44 Many filarial infections cause persistent eosinophilia (see Chapters 104 through 107). The magnitude of eosinophilic responses and the presence and intensity of other symptoms may vary greatly depending on age at first exposure, the immunologic state of the host, and the number and timing of subsequent exposures. Temporary residents and long-term residents of endemic regions have different patterns of response to a number of helminths.23,45,46 Loiasis (see Chapter 105) in temporary residents of endemic regions is characterized by immunologic hyperresponsiveness, high-grade eosinophilia, and more severe symptoms not seen in longterm residents of the same area.24 Short-term residents are more likely to have Calabar swelling and less likely to have detectable microfilaremia than the native population.24,46 Among persons diagnosed with filariasis in the GeoSentinel Surveillance Network, those born and raised in filariaendemic regions were 2.5 times as likely to be clinically asymptomatic compared with those from nonendemic areas.47 With schistosome infections, previously unexposed and nonimmune persons, and not long-term residents of endemic areas, may experience the Katayama syndrome in acute schistosomal infection.33,34

Eosinophilia Chapter 131

Box 131.4  Factors that Lower Eosinophil Counts

EPIDEMIOLOGY Published studies provide guidance to the infections more commonly associated with eosinophilia in persons who have visited or resided in tropical regions.48 Amongst 261 patients with eosinophilia (defined as >0.5 × 109/mL) evaluated in London (Hospital for Tropical Diseases), with a standard protocol and all diagnostic studies done at one laboratory, a diagnosis related to tropical exposures was made in 64%, with the most common diagnoses being schistosomiasis (33%), strongyloidiasis (25%), hookworm (5.3%), onchocerciasis (4.2%), loiasis (2.3%), ascariasis (2.3%), and trichuriasis (2.3%).49 More than one parasitic infection was found in 17%. The predominant diagnoses varied by geographic areas of exposure. Schulte et al. in Germany found blood eosinophilia (defined as ≥8%) in 4.8% of 14 298 patients who were evaluated after return from developing countries.50 The majority (73.6%) were born in Europe, the

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Table 131.1  Helminthic Infections and Eosinophilia Parasitic Infection

Main Body Site

Diagnosis

Angiostrongyliasis cantonensis Angiostrongyliasis costaricensis Anisakiasis Ascariasis Capillariasis hepatica Capillariasis philippinensis Clonorchiasis Coenurosis Cutaneous larva migrans Cysticercosis Dicrocoeliasis Dirofilariasis Dracunculiasis Echinococcosis Echinostomiasis Enterobiasis Fascioliasis Fasciolopsiasis Filariasis Lymphatic: Wuchereria, Brugia Loiasis M. ozzardi M. perstans M. streptocerca Onchocerciasis Tropical pulmonary eosinophilia Gnathostomiasis Heterophyiasis Hookworm Hymenolepiasis Metagonimiasis Opisthorchiasis Paragonimiasis Schistosomiasis Schistosomal dermatitis S. haematobium S. intercalatum S. japonicum S. mansoni Sparganosis Strongyloidiasis Trichinellosis Trichostrongyliasis Trichuriasis Visceral larva migrans T. canis, T. cati Bayliscaris procyonis

CNS GI GI GI, lung (larvae) Hepatobiliary GI Hepatobiliary Many, including CNS Skin Many, including CNS, soft tissues GI Lung, subcutaneous Soft tissues, skin Liver, lung, CNS, other GI GI, perianal Hepatobiliary GI

Serology,a rarely larvae in CSF Tissue biopsy (ileum, colon) Parasite via endoscopy or biopsy; serologya Eggs in stool; larvae in sputum or BAL for early infection Biopsy of liver Eggs, larvae, adults: stool, duodenal or jejunal aspirate, biopsy Stool examination, duodenal aspirate; serology Parasite in tissue Clinical Tissue imaging, serology Eggs in stool, duodenal aspirate, bile Morphology in biopsied tissue Morphology of worm Morphology in tissue or aspirate; serology Eggs in stool Eggs or worms in perianal area Eggs in stool, duodenal aspirate, serologya Eggs in stool

Blood, lymphatics Subcutaneous, eye Blood, skin, body cavities Blood, body cavities Skin, subcutaneous tissues Skin, subcutaneous tissues, eye Lung Subcutaneous, other tissues GI Skin (transient), GI, lung (larvae) GI GI Hepatobiliary Lung, CNS, subcutaneous

Microfilariae in blood; worm in tissue, serologya Removed worm; microfilariae in blood, serologya Microfilariae: blood, skin biopsy Microfilariae: blood, fluids; adult in tissue Microfilariae: skin biopsy, snips; adult in tissue Microfilariae: skin snips, blood, eyes; adults: tissue, serologya Serology Morphology in surgical specimen, serologya Eggs in stool Eggs in stool Eggs in stool Eggs in stool Eggs in stool, duodenal aspirate, bile Eggs: sputum, BAL, feces, pleural fluid

Skin Urinary tract, rarely CNS Liver, GI (venules of bowel), rarely CNS Liver, GI (venules of bowel), rarely CNS Liver, GI (venules of bowel), rarely CNS Subcutaneous, multiple sites GI, lung (larvae), skin (episodic) GI (early), muscle, CNS GI GI

Clinical Eggs in urine, tissue biopsy; serology Eggs in stool, biopsy Eggs in stool, biopsy; serology Eggs in stool, biopsy; serology Parasite in surgical specimen Larvae in stool, duodenal aspirate; serology Muscle biopsy; serology Eggs in stool or duodenal aspirate Eggs in stool, worms on protoscopy

Liver, eye, lung (larvae) CNS, eye, other

Larvae in tissue; serology Identification of larva

CNS, central nervous system; CSF, cerebrospinal fluid; GI, gastrointestinal; BAL, bronchoalveolar lavage. a Serologic tests may be available in some specialist laboratories.

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median duration of travel was 35 days, and one-third were asymptomatic. A definite diagnosis was made in 36%. In 18.9%, a specific helminth infection was found, with the most common helminth infections being schistosomiasis, hookworm, cutaneous larva migrans, strongyloidiasis, filariasis, and ascariasis. Travelers to Africa were more likely to have eosinophilia than those who visited the Indian subcontinent and Latin America. In returned travelers with higher levels of eosinophils (>16%),

a helminth infection was found in 46.6%. In Belgium, of more than 8000 consultations for tropical diseases, 378 patients were found to have eosinophil levels greater than 450/µL. Specific diagnoses were made in 170 patients, and parasites were detected 107 times, most commonly intestinal helminths and filarial and schistosomal species. When serologic tests and the Mazzotti test (see Chapter 106) were added as diagnostic criteria, intestinal helminthic infections were diagnosed in 19%, filarial infections

Box 131.5  Eosinophilia and Remote Tropical or Other Exposuresa

Parasite/Disease

Magnitude

Angiostrongyliasis Ascariasis Clonorchiasis

High early Moderate to high (larval migration) High early

Fascioliasis

High early

Fasciolopsiasis Filarial infections Loiasis

High early

Clonorchiasis Coenurosis Cysticercosis Echinococcosis Fascioliasis (rarely >10 years) Gnathostomiasis Hookworm (rarely 6 years or longer) Hymenolepiasis Loiasis Mansonelliasis

Mansonellosis Onchocerciasis

High Can be high Can be high

Tropical pulmonary eosinophilia Gnathostomiasis

High

Hookworm

High during larval migration High early High in early infection High during early infection High during larval migration High during acute infection

Opisthorchiasis Paragonimiasis Schistosomiasis Strongyloidiasis Trichinellosis

Visceral larva migrans

May be high

High to moderate

Comment Often absent with adult worms May wax and wane in chronic infection May wax and wane in chronic infection

Especially in expatriates Normal eosinophil counts in up to 30%

Wax and wane in chronic infection Persistent low to moderate Low or absent late Low or absent in chronic infection Low to moderate in chronic infection Absent late, except with nonencapsulating species May persist months or longer

in 13%, and schistosomal infections in 10% of patients.51 Among 2224 asymptomatic newly arrived refugees evaluated in Boston, 12% were found to have eosinophilia. Of the 45% who had serologic testing for specific pathogens, 39% were positive for Strongyloides, 22% for schistosomiasis, and 51% for filariasis.52 For African immigrants in Spain with eosinophilia (27% of 788 immigrants) who underwent parasitologic and serologic testing within 6 months of arrival, 75% were asymptomatic.53 A final diagnosis was made in 77% (55% had one parasitic infection, 14% had 2, and 7% had 3 or more), with the most common identified infections being filariae (30%), schistosomiasis (17%), and hookworms (17%). Predominant parasites varied by country of origin. A limitation of this study was the lack of good materials for the serologic diagnosis of Strongyloides. In Indochinese refugees with persistent eosinophilia whose initial comprehensive screening failed to reveal a cause for the eosin­ ophilia, the most commonly implicated infections after further investigation were hookworm (55%) and Strongyloides (38%) infections.54 As noted above, the prevalence of various helminthic causes of eosin­ ophilia will vary from one population to another depending on their geographic origins, their duration of residence, and their exposurerelated activities. Although eosinophilia is a useful marker that suggests the possible presence of certain parasitic infections, many patients with these infections do not have eosinophilia when evaluated.55 The absence of eosinophilia does not exclude the possibility of schistosomiasis, strongyloidiasis, and other infections commonly associated with eosinophilia. In a study of 1107 travelers with schistosomiasis seen in the United Kingdom, eosinophilia was present in only 44%,56 and in another study 45% of 92 returned travelers with schistosomiasis had eosinophilia.50 When persons are known to have had intense exposure to parasites, such as schistosomes,

Onchocerciasis Opisthorchiasis (up to 10 years) Paragonimiasis Schistosomiasis Sparganosis (at least 9 years) Strongyloidiasis (autoinfection enables persistence for decades) Tropical pulmonary eosinophilia (filarial) Visceral larva migrans

Eosinophilia Chapter 131

Table 131.2  Common Parasitic Causes of Marked Eosinophilia

a Parasitic infections that can be associated with eosinophilia 10 years or more after exposures in tropical areas. Eosinophilia may wax and wane.

screening tests for these infections are warranted in the absence of eosinophilia or specific symptoms.34,55 Many helminthic infections eliciting eosinophilia persist for years or even decades (Box 131.5). In contrast to persons who have not left temperate areas, persons with tropical exposures are more likely to have an infection as the cause of eosinophilia. Although infections associated with eosinophilia, such as visceral larva migrans, trichinellosis, or even strongyloidiasis, can be acquired in temperate regions, these infections account for a small percentage of cases of eosinophilia in nontravelers. Eosinophilia in most temperate regions is more likely to be caused by a process other than infection. As a rule of thumb, one should think first of parasitic infections in persons with eosinophilia and tropical exposures and look first for nonparasitic causes in persons who have always lived in temperate climates. Many infections associated with eosinophilia are seen primarily in persons with prolonged exposures. In general, cysticercosis, onchocerciasis, loiasis, lymphatic filariasis, paragonimiasis, and clonorchiasis are rare in persons with brief exposures. However, infections have been documented in persons with less than 1 month of exposure.57 In the GeoSentinel study, among travelers from nonendemic areas who acquired filarial infections, 30% reported trips of 31 days or less.47 Development of acute schistosomiasis after a single brief exposure to infested water has been described repeatedly. Because of differences in the immunologic response to parasites, a nonimmune visitor to an endemic region may develop severe symptoms and more marked eosinophilia with a small parasite burden relative to that in residents of the region.

COMMON CAUSES Strongyloides Among the helminths, the principal parasite that needs to be considered is Strongyloides since it frequently elicits eosinophilia of varying magnitudes; is often difficult to detect on stool examination; can persist for decades, even without causing major symptoms; and, importantly, can cause a disseminated, often fatal, disease (hyperinfection syndrome) in patients unsuspectingly given immunosuppressive corticosteroids (see Chapter 117).40 The capacity of Strongyloides larvae to mature within the host into filariform larvae which invade the mucosa of the colon or penetrate the skin (usually in the perianal region) means that ongoing reinfection enables the parasite to persist for the lifetime of the host. In a study from Toronto General Hospital, of 51 consecutive individuals with S. stercoralis larvae documented in fecal specimens (and no other parasites found), 83% had eosinophilia (absolute eosinophil count >400 cells/µL) with a mean eosinophil count of 890 cells/µL.58 Infection was longstanding in many; 22% had immigrated to Canada more than 10 years before diagnosis. Among 76 patients with strongyloidiasis, 25% were asymptomatic, 42% had GI symptoms, and 22% had skin complaints (primarily urticaria or pruritus).58 In a study from the United Kingdom, eosinophilia was present in 88% of travelers and 76% of immigrants with Strongyloides larvae detectable on fecal examination.59 While low-level or varying

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eosinophilia may be the only clue to strongyloidiasis,60 at times the magnitude of hypereosinophilia suggests a hypereosinophilic syndrome.61 The importance of specifically considering Strongyloides infections, especially in those with eosinophilia, is based on the additional capacity of this infection to develop into disseminated, potentially fatal, disease (hyperinfection syndrome) if patients subsequently receive corticosteroids or become immunocompromised.40 Thus, clinicians should carefully assess the probability of past exposure to Strongyloides in persons who will be given steroids or other immunosuppressive agents or who develop diseases associated with immunosuppression.40 Cases of hyperinfection and dissemination have been reported more than 50 years after the last known exposure in endemic areas.62 Patients with hyperinfection and dissemination may present with acute respiratory failure and Gramnegative sepsis.63 Rarely, disseminated strongyloidiasis has been reported in apparently immunocompetent persons.64 Persons infected with human T-cell lymphotropic virus type I (HTLV-1) have an impaired immune response to Strongyloides that increases the likelihood of hyperinfection and slows clearance of parasites after treatment with ivermectin compared with persons not infected with HTLV-1.65 Although certain groups of immunocompromised patients (especially those with hematologic malignancies, or on steroids, or with severe malnutrition) appear to be at increased risk for the Strongyloides hyperinfection syndrome,66 this has not been a frequently reported complication in human immunodeficiency virus serotype 1 (HIV-1)-infected persons in geographic areas where both HIV-1 and Strongyloides infections are common. Because fecal examinations are insensitive, enzyme-linked immunosorbent assay serology is useful in detecting strongyloidiasis even when fecal examinations are unrevealing, though sensitivity of serologic tests is lower in immunocompromised individuals.54

Schistosomes Eosinophilia frequently accompanies schistosomiasis (see Chapter 122). Travelers who have visited endemic areas may present with acute schi­ stosomiasis, subacute symptoms, or even without symptoms.67 In one series of returned travelers with acute schistosomiasis, 73% had eosin­ ophilia, 43% had respiratory symptoms, and 45% had skin changes at some point in the clinical course.34 Many reports document sporadic cases and clusters of nonimmune persons with acute schistosomiasis after brief water exposures in endemic regions. Typical findings are spiking fevers, sweating, diarrhea, skin findings (urticarial rash and angioedema) and dry cough that can persist for a month or longer. Pulmonary symptoms were noted in 8 of 60 nonimmune travelers seen in Israel with acute schistosomiasis, occurring 3–6 weeks after exposure.68 Chest radiographs showed multiple small nodules and diffuse interstitial infiltrate.68 Among 31 patients with acute schistosomiasis mansoni in Brazil, all had eosinophilia, 90% had fever, 94% had diffuse abdominal pain, 81% had cough, 52% had dyspnea, and liver enzymes were elevated in 38%.69 Other symptomatic presentations of schistosomiasis in travelers can include hematuria, hematospermia, or, uncommonly, transverse myelitis.70–72 Infections can occur after a brief stay;73 but for many, schistosomiasis develops only after longer-term exposures in endemic regions. A study of expatriates and visitors to Malawi found that serologic evidence of current or past schistosome infection increased directly with the length of stay in Malawi.74 Seroprevalence was 48% in people resident 4 years or longer, in contrast to 11% for those resident 1 year or less. Recreational fresh-water exposure in Lake Malawi was an important source of infection.74

Filariae

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Eosinophilia is common in infections caused by filarial parasites (see Chapters 104–106), which differ by geographic regions and vector insects. For lymphatic filariasis, early manifestations may include filarial fevers with lymphangitis.75,76 Among 271 travelers and immigrants seen in GeoSentinel clinics who were diagnosed with filarial infections, 75% were acquired in sub-Saharan Africa and 10% in South America.47 The

most commonly diagnosed infections were onchocerciasis (37%), loiasis (25%), bancroftian filariasis (25%), and other filarial species in 6%. Twothirds of those with onchocerciasis sought care for symptoms within 1 month of return.

Human Immunodeficiency Virus Type 1 Occasional reports have noted increased levels of eosinophils in HIV-1infected persons.77 In a study of 855 HIV-infected persons in New York examined over 4 years, however, no single, consistent cause of eosinophilia could be identified.77 Increased eosinophil levels appeared to result from a relative preservation of the eosinophil cell line (while other cell lines declined as the CD4 count decreased) in some patients and in others from the presence of truly increased eosinophils.77 Increased eo­sinophil levels in HIV-infected subjects were more commonly asso­ ciated with rashes, including eosinophilic folliculitis,78 atopic dermatitis, and prurigo nodularis. One study suggested no extensive evaluation for eosinophilia was warranted.79 Although some HIV-infected patients with eosinophilia have a definable cause, such as adverse drug reactions or adrenal insufficiency due to cytomegalovirus and other infections, in most instances no obvious cause can be identified, indicating that HIV infection may be associated at times with increased levels of eosinophils.

COMMON SYNDROMES Pulmonary Eosinophilic lung diseases are a heterogeneous group of disorders characterized by a common presence of increased eosinophils in inflammatory infiltrates in the airways or parenchyma of the lungs. The clinical presentation of pulmonary eosinophilia usually consists of symptoms referable to the respiratory system accompanied by an abnormal chest radiograph and blood, sputum or bronchoalveolar lavage eosinophilia. While pathogenic mechanisms underlying many of the disorders are undefined, we classify pulmonary eosinophilias based on recognized etiologic agents and distinct clinical and pathologic patterns2 (Box 131.6). Several helminthic infections that can cause eosinophilic lung diseases can be categorized based on the behavior of the parasites.

Transpulmonary Passage of Helminth Larvae This category, the true Löffler’s syndrome, arises from reactions elicited by larvae that pass through the lungs as part of the parasite’s initial developmental cycle in the human. For three helminthic intestinal parasites (Ascaris, hookworms, and Strongyloides), infecting larvae pass through the

Box 131.6  Pulmonary Eosinophilia 1. Drug- and toxin-induced eosinophilic lung diseases 2. Helminthic infection-related eosinophilic lung diseases Transpulmonary passage of larvae (Löffler’s syndrome): Ascaris, hookworm, Strongyloides Pulmonary parenchymal invasion: mostly helminths, paragonimiasis, echinococcosis Heavy hematogenous seeding with helminths: trichinellosis, disseminated strongyloidiasis, cutaneous and visceral larva migrans, schistosomiasis

Tropical pulmonary eosinophilia: filaria 3. Fungal-related eosinophilic lung diseases Allergic bronchopulmonary aspergillosis 4. Chronic eosinophilic pneumonia 5. Acute eosinophilic pneumonia 6. Churg–Strauss syndrome – vasculitis 7. Other: neoplasia, hypereosinophilic syndromes, bronchocentric granulomatosis, sarcoidosis

Pulmonary Parenchymal Invasion with Helminths In contrast to the parasites that transit through the lungs, a few helminths, such as Paragonimus lung flukes (see Chapter 123) and echinococcal species (see Chapter 120), have a predilection to localize within the pulmonary parenchyma, and these may elicit eosinophil-enriched inflammatory reactions.81 Paragonimus larvae undergo maturation in the lungs. Larval flukes can leave hemorrhagic, necrotic tracts; their eggs provoke a granulomatous response. Because the chronic cavitary lesions of paragonimiasis can mimic tuberculosis, and many adults from Southeast Asia have positive tuberculin tests, patients may be misdiagnosed as having tuberculosis.82 Echinococcal infection can also involve the lungs, though encysted parasites typically do not elicit a peripheral eosinophilia.

Heavy Hematogenous Seeding with Helminths This disease category includes the eosinophilic pulmonary responses elicited by helminthic larvae or eggs that are carried into the lungs hema­ togenously in an aberrant fashion. Thus, in contrast to the “normal” transpulmonary migration noted previously, etiologic helminths include abnormal numbers of nonhuman hookworms or ascarids causing cutaneous or visceral larva migrans,83,84 abnormal numbers of hematogenous larvae in heavy trichinellosis infections,81 and abnormal spread following chemotherapy of schistosomal parasites via collateral vessels into the lungs.81,85 Also included in this category is disseminated strongyloidiasis, which develops when the Strongyloides autoinfection cycle becomes unbridled, often in association with corticosteroid drug administration (see Chapter 117). Large numbers of larvae traverse the lungs eliciting pulmonary findings. Adult parasites can develop in the bronchial tree, causing bronchospasm mimicking asthma.86 In disseminated strongyloidiasis, filariform larvae can be found in many sites, including the stool,

sputum, and bronchoalveolar lavage. The usual eosinophilia of strongyloidiasis can be suppressed in disseminated disease because of concomitant pyogenic infection or steroid administration.

Tropical Pulmonary Eosinophilia Tropical pulmonary eosinophilia (TPE) results from a distinct immune response to the normally bloodborne microfilarial stages of lymphatic filariae, Wuchereria bancrofti, and, less commonly, Brugia malayi (see Chapter 104).87–89 TPE is prevalent in regions where these filariae are endemic, and residents or immigrants from these regions are those who may experience TPE. Males are affected more often than females in a ratio of about 4 : 1. Dyspnea, cough, wheezing (especially at night), and chest discomfort may be accompanied by weight loss and malaise. Infection is commonly misdiagnosed, with asthma being a common diagnosis.89,90 In addition to blood eosinophilia, features specific to TPE include high levels of serum IgE and antifilarial antibodies. Total white blood cell count is typically elevated and blood eosinophilia may be extremely high. Bloodstream microfilariae are almost never found. Abnormalities on chest radiographs may be subtle and include diffuse miliary lesions 1–3 mm in size, patchy consolidations, cavitation, reticulonodular infiltrates of the lower lung zones, and an interstitial nodular pattern.91 Without treatment, TPE can lead to progressive fibrosis and chronic respiratory compromise.89 TPE is an immunologically mediated disorder with microfilariae trapped in the lungs. Microfilariae are detected in inflammatory foci in biopsies from the lung, liver, and lymph nodes. A related, nonfilarial syndrome of TPE of uncertain cause has been recognized.92

Eosinophilia Chapter 131

lungs, entering via the bloodstream, penetrating into alveoli, and then ascending the airway to transit down the esophagus into the small bowel. Löffler, who first described this syndrome of migratory pulmonary infiltrates and blood eosinophilia in Swiss patients, subsequently implicated Ascaris infection, acquired from the use of contaminated human feces as fertilizer, as the cause of this syndrome.80 Ascaris is especially capable of eliciting eosinophilic inflammatory responses and is prevalent in regions where human feces contaminate soil or are used as fertilizer (see Chapter 115). Although infecting hookworm (see Chapter 116) and Strongyloides (see Chapter 117) larvae likewise traverse the lungs, these larvae rarely elicit symptoms of pulmonary eosinophilia in natural or experimental infections.81 Since Löffler’s syndrome occurs only when developing larvae are transiting the lungs, about 9–12 days after ingestion of Ascaris eggs, this cause of pulmonary eosinophilia should be considered only in those with a recent exposure to Ascaris eggs (see Chapter 115). In symptomatic patients, common complaints are an irritating, nonproductive cough and burning substernal discomfort, and over half of patients have rales and wheezing. Acute symptoms generally subside within 5–10 days. Chest radiographs show unilateral or bilateral nonsegmental densities with indefinite borders ranging in size from several millimeters to several centimeters. Infiltrates are generally transient and migratory and clear over 1 or more weeks. Blood eosinophilia increases after several days of symptoms and resolves over many weeks. Ascaris pneumonia is diagnosed at the time of pneumonic involvement only by detecting Ascaris larvae in respiratory secretions or gastric aspirates.81 At least 40 days must elapse before the larvae responsible for pulmonary infiltrates have matured sufficiently to produce eggs detectable in the stool. Negative stool examinations during or soon after an episode of pneumonitis do not exclude Ascaris as the cause, nor do positive stool examinations for Ascaris eggs during the stage of pulmonary involvement establish the cause, as these eggs reflect infection acquired 2–24 months earlier.

Other Pulmonary Eosinophilias Cases of acute eosinophilic pneumonia occurred in US military per­ sonnel deployed in or near Iraq in 2003–2004.93 The etiology was not determined; an association with recent initiation of cigarette smoking was noted. Among the drugs reported to cause pulmonary eosinophilia are ones sometimes taken for malaria prophylaxis and hence pertinent in patients who will have had tropical exposures. These include pyrimethamine, and dapsone.94,95 Eosinophilic pleural effusions (Box 131.7) can have multiple causes.96 In the setting of someone with potential infectious exposures, several helminthic infections, including toxocariasis,97 TPE,98 paragonimiasis,99 loiasis,100 and anisakiasis,101 can cause eosinophilic pleural effusions.

Abdominal Pain or Diarrhea The gut is the primary or sole residence for many helminthic parasites. Symptoms from these eoosinophilia-associated parasitic infections (Box 131.8) may result from several mechanisms, including attachment of the Box 131.7  Eosinophilic Pleural Effusions Helminthic Infections Anisakiasis (rare) Echinococcosis Gnathostomiasis Loiasis Paragonimiasis Strongyloidiasis (disseminated infection; may also find larvae in fluid) Toxocariasis Trichinellosis (early weeks of infection) Tropical pulmonary eosinophilia

Other Infections Coccidioidomycosis Tuberculosis Other Causes Hemothorax Hypersensitivity reactions, including drug reactions Malignancy Pneumothorax, including from thoracentesis Pulmonary infarct Rheumatologic Diseases

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Box 131.8  Parasites Causing Eosinophilia and Abdominal Pain or Diarrhea Helminths Ancylostoma caninum (eosinophilic enteritis) Anisakis spp. and other genera (anisakiasis) Angiostrongylus costaricensis Ascaris lumbricoides Capillaria philippinensis Clonorchis sinensis Echinococcus Enterobius vermicularis (eosinophilic colitis) Fasciola hepatica Fasciolopsis buski Gnathostoma Hookworm (heavy infection) Schistosomes Strongyloides Trichinella Toxocara canis, T. cati, other (visceral larva migrans)

Protozoa Dientamoeba fragilis Isospora belli Other Causes Eosinophilic gastroenteritis Dermatitis herpetiformis Churg–Strauss syndrome (vasculitis) Regional enteritis Ulcerative colitis Lymphoma Solid tumors Drug reaction (e.g., eosinophilic colitis from naproxen) Hypereosinophilic syndromes Allergy (e.g., cow’s milk, soy protein in infants)

parasite to, or penetration of, the gut mucosa, causing irritation and inflammation; migration through the wall or to other adjacent sites (such as biliary and pancreatic ducts); intraluminal obstruction by a bolus of worms; and entry into the appendix, among others. The canine hookworm (Ancyclostoma caninum) has been associated with a syndrome of abdominal pain, sometimes acute and severe, and peripheral eosinophilia.102 Larvae of the pinworm Enterobius vermicularis (see Chapter 113) have also been reported to cause eosinophilic colitis and enteritis.103 Capillariasis (Capillaria philippinensis) (see Chapter 112) can be associated with chronic, severe diarrhea and intestinal tissue infiltration of eosinophils.104 Helminth-elicited disease must be distinguished from often idiopathic eosinophilic gastroenteritis.105,106

Table 131.3  Helminths Associated with Eosinophilia and Skin and Soft-tissue Changes Parasite/Disease

Manifestation

Ascariasis Coenurosis Cutaneous larva migransa Cysticercosis Dracunculiasis

Urticaria Subcutaneous nodules, usually solitary Characteristic serpiginous lesions Rubbery, painless cysts; often multiple Papules, vesicles; rupture and discharge larvae Soft, subcutaneous cysts, varying sizes; urticaria Urticaria; rare painful or itchy subcutaneous nodules

Echinococcosis Fascioliasis Filariasis   Brugia zoonotic filariae   Zoonotic dirofilariasis   Loiasis   Onchocerciasis   Wuchereria bancrofti

  Brugia malayi, B. timori

Gnathostomiasis

Hookworm Paragonimiasis Schistosomiasis Sparganosis Strongyloidiasis

Skin

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Intermittent and migratory lesions of the skin and subcutaneous tissues can reflect migration of parasites in human tissues (Table 131.3) (see Chapter 132). Skin lesions associated with gnathostomiasis (see Chapter 112) may begin as early as 3–4 weeks after ingestion of the parasite or may be delayed until months or even years later. The third-stage larvae cause localized swellings that typically last 1–2 weeks and are associated with edema, pain, itching, and variable erythema. Swellings may recur off and on for 10–12 years. Because a single gnathostome can cause symptoms, cases have occurred after a brief stay in an endemic area or after eating foods, usually raw fish, from those areas. Disease has been most often reported from Southeast Asia. Increases in cases in Mexico, Peru, and Ecuador have been noted.107,108 In addition to causing cutaneous disease, the worm can also migrate to tissues throughout the body, causing pulmonary, gastrointestinal, genitourinary, central nervous system, ocular, and other localized disease.44 Loiasis, another disease causing migratory lesions and eosinophilia, follows exposures to Loa loa in westcentral Africa (see Chapter 105).57 Symptoms typically do not appear until at least 4 months after exposure and can first appear more than 5 years after exposure. Classic findings are localized areas of angioedema (Calabar swellings), which may be warm, red, itchy, or painful,24,46 usually disappear within a few days, and may recur multiple times per year. A pruritic skin rash is a common feature of onchocerciasis (see Chapter 106). Swelling of an extremity should suggest obstruction of lymphatics caused by one of the lymphatic filarial parasites. Patients may experience recurrent lymphangitis (often with retrograde progression) lasting 3–7 days, which may be associated with fever.76 Orchitis and epididymitis are

Trichinellosis Visceral larva migrans a

Local lymphadenopathy Subcutaneous nodule Pruritic 5–10-cm Calabar swellings; urticaria; papulovesicular lesions Subcutaneous nodules; papules and severe itching; urticaria; lizard skin Recurrent retrograde lymphangitis and lymphadenitis; scrotal mass; hydrocele, elephantiasis Recurrent retrograde lymphangitis and lymphadenitis; scrotal mass; hydrocele, elephantiasis Urticarial, edematous, recurrent, migratory subcutaneous swellings; creeping eruption; panniculitis Urticaria; itchy maculopapular rash; may be vesicular Urticaria (early); subcutaneous nodules Urticaria, itchy maculopapules (early); papules (ectopic egg deposition) Edematous, painful migratory swellings secondary to worm migration Itchy papular and migratory serpiginous lesions at points of penetration and dermal migration (larva currens); urticaria; papulovesicular lesions; petechiae and purpuric lesions in hyperinfection syndrome Urticaria; periorbital edema; splinter hemorrhages Urticaria; nodules

Eosinophilia usually low-grade or absent.

also characteristic features. Although eosinophilia can occur in patients with cutaneous larva migrans caused by the hookworm Ancylostoma braziliense and related nematodes, in one series only 20% had at least 7% blood eosinophilia.109 Penetration of schistosomal cercariae is typically followed by the development of an itchy, erythematous, papular rash.110 Exposures to avian schistosomes elicit a similar skin reaction (swimmer’s itch).110,111

Hepatobiliary Helminths that cause eosinophilia and hepatic dysfunction (Box 131.9) include several parasites that reside in the biliary tree and cause localized inflammatory changes and can obstruct the bile or pancreatic ducts, leading to a clinical picture that can mimic acute cholangitis or cholecystitis (see Chapter 123). Ascaris adult worms can wander from their usual habitat within the gut lumen to enter pancreatic or biliary ducts causing

Box 131.11  Causes of Eosinophilia in Cerebrospinal Fluid

Angiostrongyliasis cantonensis (infrequent hepatic involvement) Ascariasis (mechanical obstruction of bile ducts by migrating worm) Capillariasis due to Capillaria hepatica Clonorchiasis Dicrocoeliasis Echinococcosis (eosinophilia may be absent with intact cysts; liver function may remain normal despite multiple cysts)

Helminths

Fascioliasis Metorchis conjunctus Opisthorchiasis Schistosomiasis Strongyloidiasis (with hyperinfection) Taeniasis (Taenia saginata; rare obstruction of bile ducts) Visceral larva migrans (toxocariasis)

Box 131.10  Helminthic Infections Causing Eosinophilia and Prominent Fevera Clonorchiasis Fascioliasis Gnathostomiasis (early, episodic) Onchocerciasis Schistosomiasis (Katayama syndrome)

Trichinellosis Visceral larva migrans (toxocariasis)

Nematode (Roundworm) Infections with Migrating Larvae Inherently Neurotropic Angiostrongylus cantonensis Gnathostoma spinigerum Baylisascaris procyonis Cestode (Tapeworm) Infection with Cysts Developing in the Central Nervous System Cysticercosis Echinococcosis Coenurosis Trematode (Fluke) Infection with Ectopic Central Nervous System Localization Paragonimus westermani Schistosomiasis Fascioliasis

Miscellaneous Infectious Causes Coccidioidomycosis Cryptococcus neoformans Toxocariasis (Toxocara canis) Trichinellosis Myiasis (larvae of cattle botflies) Noninfectious Causes Ventriculoperitoneal shunts Leukemia or lymphoma with central nervous system involvement (Hodgkin’s) Drug hypersensitivity reactions: nonsteroidal anti-inflammatory agents, antibiotics (e.g., sulfonamides, ciprofloxacin, vancomycin), phenytoin; myelography contrast agents Hypereosinophilic syndromes Sarcoidosis

Eosinophilia Chapter 131

Box 131.9  Infections Causing Eosinophilia and Hepatic Dysfunction

a

Acute, early Ascaris, hookworm, or Strongyloides may be associated with fever.

Table 131.4  Examples of Drug-associated Reactions with Eosinophilia acute symptoms (see Chapter 115). In fascioliasis, young flukes penetrate the capsule of the liver and migrate through liver parenchyma causing local necrosis and inflammation before they enter the bile ducts, where they mature and produce eggs that are released into bile and passed in feces.112,113 Eosinophilia may wax and wane when infection becomes chronic. Clonorchiasis, endemic in eastern and southeastern Asia, and opisthorchiasis, acquired mainly in Southeast Asia, are other parasites that reside in bile ducts (see Chapter 123). Among 17 immigrants who were found to have ova of Opisthorchis spp. or Clonorchis sinensis in fecal specimens when evaluated in the US Midwest, 88% had eosinophilia (defined as absolute eosinophil counts >500/µL). Clinical symptoms included vague right upper abdominal discomfort; 24% of those infected had been in the United States at least 5 years.114

Drug Reactions

Examples

Pulmonary infiltrates Pleuropulmonary Interstitial nephritis Necrotizing myocarditis Hepatitis Hypersensitivity vasculitis Gastroenterocolitis Asthma, nasal polyps DRESS Asymptomatic

NSAIDs Dantrolene Semisynthetic penicillins, cephalosporins Ranitidine Semisynthetic penicillins, tetracyclines Allopurinol, phenytoin NSAIDs Aspirin Antiepileptics Ampicillin, penicillins, cephalosporins

NSAIDs, nonsteroidal anti-inflammatory drugs; DRESS, drug-induced rash, eosinophilia, and systemic symptoms.

Fever Only a few helminthic infections cause eosinophilia and prominent fevers (Box 131.10). Fever and eosinophilia can be seen in acute schistosomiasis,33,115 visceral larva migrans,116 trichinellosis,117 fascioliasis,112 gnathostomiasis,44 and some other infections. In many instances, focal findings (e.g., localized subcutaneous swellings, muscle tenderness, lymphangitis, cough and pulmonary infiltrates, abdominal pain, and others) provide diagnostic clues. The combination of exposure history (when, where, and what activities) along with clinical findings can help the clinician to focus on more likely diagnoses.

Central Nervous System Findings Larvae of Angiostrongylus cantonensis migrate to the brain, spinal cord, and eye, where the larvae and young adults provoke an intense inflammatory response and cause the clinical picture of eosinophilic meningoencephalitis (see Chapter 111). A number of other parasites or their eggs can be associated with an eosinophilic tissue response and with variable eosinophils in the spinal fluid. Cysticercosis, which commonly involves the central nervous system, typically causes focal neurologic findings, including seizures (see Chapter 119). The cerebrospinal fluid may be acellular and peripheral eosinophilia absent. In addition to helminthic parasites, other infections, drug reactions, and other processes can induce an eosinophilic meningitis118,119 (Box 131.11).

NONINFECTIOUS DISEASES Many processes (Box 131.1) associated with eosinophilia may need to be considered if helminthic infections do not explain a patient’s eosinophilia. Drug hypersensitivity reactions should always be considered as a potential cause of eosinophilia, even in persons who have visited or lived in tropical regions. The presence of a characteristic rash may suggest a hypersensitivity reaction, but drug-associated eosinophilia can occur in the absence of skin changes. In addition to peripheral eosinophilia, drug reactions may be associated with pathologic changes and dysfunction of other tissues and organs (Table 131.4). Any of these findings may predominate, or a combination of several may be seen at the same time or in staggered development. Some medication-related eosinophilic responses (e.g., Stevens–Johnson syndrome, acute anaphylaxis, and drug-induced rash, eosinophilia, and systemic symptoms (DRESS)) can be life-threatening and preclude continued use of or rechallenge with candidate offending medications. Because of the striking blood or tissue eosinophilia, some drug reactions can mimic parasitic infections.

Evaluation of Patients with Eosinophilia Several key concepts are relevant to the evaluation of a patient with eosinophilia after tropical exposures:

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• Multiple parasitic infections may be present. • Fecal excretion of eggs may be in small numbers or intermittent. • Examination of stool for eggs is useless during the prepatent period. Symptoms such as fever and eosinophilia may be present for 2 weeks or more before the products of the adult parasite (e.g., eggs or larvae) are passed in the stool. • Eosinophilia is most prominent with tissue-dwelling helminths, and many of these never enter the intestinal tract. Stool examinations will not detect tissue-dwelling helminths or exclude their presence as a cause of eosinophilia. • Serologic tests may be negative early in the symptomatic period. It is necessary to have some idea of the possible infections that might be present in order to focus the evaluation and determine which tests to do and which tissues to examine. Even if the patient is entirely asymptomatic and the physical examination is unremarkable, the presence of eosinophilia should lead to a workup to identify treatable diseases that could cause future problems. Eosinophilia, however, need not be present.55 Knowledge of the geographic distribution of various parasites and the characteristic time interval between exposure and onset of symptoms and the presence of diagnostic forms of parasites or eggs in tissue or fluids is an essential part of the evaluation (Box 131.12). Even when sensitive and specific serologic tests are available, they may be negative early in the course of infection. In a study of patients with acute schistosomiasis, serologic results were negative in 3 of 8 persons tested during the first week of illness.33 If suspicion is high, serologic tests should be repeated if initially negative. During the prepatent period, serologic studies may be the only way to confirm the diagnosis.120 Timing of obtaining blood to search for microfilariae needs to be done cognizant of the potential periodicity exhibited by the species and strains of filariae (see Chapters 104 and 105). In general, it is preferable to diagnose infection by identification of eggs, larvae, adults, or other parasitic stages or products in tissues, fluids, or stool. Sometimes diagnostic material is unavailable or its retrieval would entail serious risk to the patient (e.g., brain biopsy, liver biopsy, major surgical intervention). When parasites, eggs, or other products are unavailable, serologic tests are useful in some infections. Problems with serologic tests may include lack of sensitivity, availability, standardization and quality control, and low specificity. Cross-reactivity to related parasites is common for many of the tests. Acid-fast techniques stain schistosome eggs and hooklets of Echinococcus granulosus.

RESPONSE TO TREATMENT Several key points should be kept in mind when treating patients with helminthic infections: • Treatment may not be curative; efficacy may vary depending on the stage of the parasite. • Symptoms may worsen, usually only transiently, with therapy. • The level of eosinophilia may be a poor predictor of whether treatment has cured infection. • Eosinophilia may rise following initiation of treatment which elicited killing of helminths • Relapses can occur late. Even when parasitic infections are recognized and treated, anthelminthic drugs may fail to cure infection.

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Box 131.12  Eggs or Parasites in Tissues or Fluids In Urine Filariasis (microfilariae, e.g., Wuchereria bancrofti, Loa loa) Gnathostomiasis (parasite passed in urine; rare) Schistosome eggs (Schistosoma haematobium) Strongyloidiasis (larvae, rare) In Sputum Ascariasis (larvae during early migration) Echinococcosis (fragments of larvae and free scolices after rupture of cyst into bronchus: rare) Gnathostomiasis (rarely, coughed-up worm) Hookworm (larvae, early) Paragonimiasis (eggs in sputum) Schistosomiasis (eggs) Strongyloidiasis (filariform larvae, and, rarely, eggs and adults in sputum in hyperinfection syndrome) In Pleural Fluid Paragonimiasis (eggs) Strongyloidiasis (filariform larvae)

On Blood Smears Microfilariae of filariasis (e.g., W. bancrofti, Brugia malayi, L. loa, Mansonella perstans, M. ozzardi) Trichinellosis (rarely observed on blood smears) In Cerebrospinal Fluid Angiostrongyliasis due to Angiostrongylus cantonensis (larvae rarely in cerebrospinal fluid) On Perirectal Skin (Tape Test) Pinworm eggs Tapeworm proglottids In Fecesa Strongyloidiasis (20–30% positive with a single examination; 60–70% with three or more; 40–90% positive on duodenal sample or endoscopic aspirate) Several intestinal nematodes, trematodes, and cestodes

a Eggs and parasites are not found in feces in patients with many helminthic infections, such as echinococcosis, toxocaral visceral larva migrans, and trichinellosis; stool examinations are often negative at the time of earliest symptoms in patients with many infections, including schistosomiasis, ascariasis, and hookworm infections.

A temporary rise in eosinophilia after treatment has been observed in a number of helminthic infections.121,122 The magnitude of the posttreatment eosinophilia correlates with the parasite burden prior to therapy.123 This may be associated with other symptoms, such as pruritus, dermatitis, arthralgia, and myalgia in patients with loiasis.124 Patients with schistosomiasis may develop respiratory symptoms and new pulmonary infiltrates in association with therapy.85,122,123 Severe symptoms have been reported when treatment is given during the early Katayama syndrome.121 These worsening symptoms and the rise in eosinophils may reflect the host immune response to antigens released or exposed from dead or dying larvae or worms. In patients with loiasis treated with diethylcarbamizine, levels of eosinophils usually returned to normal within 6 months.124 This occurred even in patients who had late relapses of infection.124 Thus, normalization of levels of eosinophils after treatment does not necessarily indicate that infection has been eradicated. Conversely, however, the persistence of eosinophilia likely indicates that infections persist. Box 131.13 outlines an approach to the evaluation of the patient with eosinophilia.

Eosinophil Determinations Confirm that the eosinophil count is elevated Estimate the absolute blood eosinophil count: normal, ≤350 eosinophils/µL; elevated, >450 eosinophils/µL Categorize eosinophilia as low (3000/µL) (see Table 131.2) Medical History Eosinophil History Inquire from history or medical records if eosinophil counts were previously normal or elevated, and if so for how long Medication History Review drug exposure history for recent or current drugs that may be associated with eosinophilia. Discontinue any drugs that are commonly associated with eosinophilia and make a complete list of other medications, vitamins, supplements, or herbal preparations that the patient is taking. If patient is taking drugs that have been associated with eosinophilia, assess for associated serious organ involvement. Note any history of allergies to drugs or other agents Disease History Review the medical history for diseases or disorders typically associated with eosinophilia (see Box 131.1). Given their prevalence, allergic and atopic disorders should be noted, although new onset of some allergic manifestations (e.g., urticaria, bronchospasm, wheezing) may be secondary to helminthic infections Geographic History Review history of past residence and travel in other countries or regions. Relevant are time periods extending even decades before. Especially relevant are exposures in tropical regions and those with poor sanitation. Note places, dates, and duration of exposures. Some helminths have discrete geographic distributions (e.g., Clonorchis in the Far East; Angiostrongylus cantonensis, a cause of eosinophilic meningitis, principally, but not exclusively, in the Pacific Basin; Loa loa in central-west Africa; Onchocerca volvulus in equatorial Africa and elevated regions in Central America). Even nontropical geographic histories are relevant to some helminths and other agents, e.g., the southwestern United States for coccidioidomycosis. Exposures in sheep-rearing areas are pertinent to Echinococcus granulosus Activity History Review occupational and recreational exposures. Histories of swimming in or contact with fresh water in areas where schistosomiasis occurs are pertinent. Contact with fresh or salt water followed by a rash on water-exposed skin suggests schistosome dermatitis (avian schistosome species). Skin contact with soil potentially contaminated with human or dog feces, walking barefoot, or occupational (military or job-related) exposures are pertinent to the acquisition of cutaneous larva migrans, hookworm, and Strongyloides infections

Other Epidemiologic History Helpful in the evaluation is knowledge of whether the patient traveled with others and whether others developed similar illnesses – relevant to common-source water exposures (e.g., schistosomiasis) or foodborne illness (e.g., trichinellosis). In concert with geographic exposures, histories of exposures to insect vectors are pertinent (e.g., filarial infections) General History Do a careful review of systems for any current or recent symptoms, including a history of fevers (see Box 131.10), skin lesions (see Table 131.3) that may have cleared, and gastrointestinal illnesses (see Box 131.8)

Eosinophilia Chapter 131

Box 131.13  Evaluation of the Patient with Eosinophilia

Physical Examination Do a careful physical examination, paying close attention to skin and soft tissue for nodules or masses Initial Laboratory Evaluation The presence of specific clinical symptoms and physical findings, as well as other information from the history, may begin to direct laboratory testing As needed, check routine studies to assess organ involvement (e.g., liver function tests, renal function tests, urinalysis, serum troponin for cardaic involvement, chest radiograph (CXR)) or high-resolution chest computed tomography (CT) Further Diagnostic Evaluations Symptomatic Patient with Localizing Findings The evaluation will be guided by the nature of the historical information and results from the physical examination and initial laboratory tests based on the focal findings. Skin: skin snips, skin biopsy, excision of mass Central Nervous System CT and magnetic resonance imaging (MRI); cerebrospinal fluid examination (cell counts, differential, protein, glucose; larvae) Pulmonary Findings or Abnormal CXR Sputum examination (routine, acid-fast bacteria, ova and parasites (O&P)), CT or MRI to define lesions better, including those not seen on CXR Stool O&P; urine O&P; rectal biopsy; other tests (see Box 131.12) Eosinophilia in a Patient without Other Findings Stools for O&P (×3 as needed) Strongyloides serology (if stools negative), toxocara serology Check serologic studies for schistosomiasis, filariasis, and onchocerciasis if patient may have been exposed

Dietary History Dietary histories are pertinent to several helminthic infections, including anisakiasis (raw fish), fish tapeworm (fish), Nanophyetus salmincola (salmon), Taenia solium (pork), T. saginata (beef), fascioliasis (watercress), fasciolopsiasis (water chestnut), gnathostomiasis (fresh-water fish, eels, frogs, snakes, and poultry and pigs fed on fish), Angiostrongylus cantonensis (land snails or slugs, fresh-water shrimp, crabs, some marine fish), and trichinellosis (pork, boar, bear, horse, walrus, warthog)

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CHAPTER 132 

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Cutaneous Lesions Steven D. Mawhorter • David L. Longworth

INTRODUCTION Cutaneous lesions can be cardinal manifestations of diverse infectious diseases wherever they are acquired. Notably, in travelers returning from tropical regions, skin lesions are one of the top five medical concerns.1–5 The top three dermatologic diagnoses in travelers are cutaneous larva migrans (CLM), bacterial infections (cellulitis, abscess, and pyoderma), and arthropod/insect-related lesions.5–7 The majority of these lesions develop prior to return home and rarely require hospitalization, although often lead to medical evaluation.4,5 The skin represents the largest organ of the body and the most accessible to direct examination and observation. As a result, it is often a sentinel for many infectious diseases by being the primary site of involvement; the entry site of parasitic or bacterial pathogens; or by demonstrating lesions that result from toxin-, inflammatory-, or vascular-mediated changes associated with infection.8 Specific pathogens and the clinical manifestations of diseases commonly found in people residing in, or traveling to, tropical areas are discussed in detail in pathogen-specific chapters. This chapter considers the pathophysiology of tropical and parasite-associated skin reactions, and focuses on the approach to the patient who presents with cutaneous lesions, to assist in the generation of a differential diagnosis.

THE SKIN AS TARGET ORGAN – PATHOGENETIC MECHANISMS

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Although many pathogens can produce cutaneous manifestations, the skin has a limited number of responses to inflammatory or infectious processes. In general, the pathogenesis of skin manifestations of infectious diseases can be considered under six broad categories. The first involves bloodborne dissemination of the actual infectious agent.9,10 This category can be further subdivided into two areas based on whether the etiologic agent engages in direct interaction with the skin and dermal appendages to produce cutaneous manifestations or mediates the interaction through cellular, humoral, or other immune mechanisms. Examples of organisms that directly involve the skin include Neisseria meningitidis, Rickettsia spp., endocarditis-related bacterial embolization (e.g., Janeway lesions, Osler’s nodes), Pseudomonas spp. and other Gram-negative ecthyma lesions, Bacillus anthracis, and Candida spp. In this setting, Gram stain of the skin lesion may reveal the offending organism.11,12 Viral pathogens, such as varicella-zoster virus and many enteroviruses, may elicit cutaneous findings following viremic seeding of the skin.10 Examples of immune reactions involving the skin to bloodborne organisms include Salmonella spp. (rose spots), N. meningitidis and N. gonorrhoeae, measles, and rubella.10 In these cases, cutaneous manifestations are typically due to local cutaneous immune vasculitis or dermal immune responses, including, but not limited to, immune complex deposition. Since the skin changes are largely immune complex-mediated, direct smears of the lesions are unlikely to reveal a pathogen. Many diseases associated with the bacter-

emic or viremic pathogenesis of this first category of cutaneous involvement are severe and occasionally life-threatening.10 The second category involves local reactions to parasites that gain access to the body elsewhere and migrate to the skin but are not bloodborne. They can elicit a direct or indirect immunologic response to produce the cutaneous changes. This category includes agents such as the filarial parasites Onchocerca volvulus (see Chapter 106) and Loa loa (see Chapter 105).13,14 The nodules of onchocerciasis and loiasis often contain adult worms (direct mechanism). However, generalized pruritus and maculopapular eruptions appear to be mediated by immune reactions to dermal-based microfilariae (indirect mechanism).15 Other helminthic examples include the cutaneous nodules of Dracunculus medinensis (see Chapter 108) and Spirometra spp. (sparganosis, see Chapter 121).13,16 A third category involves cutaneous immune reactions to direct penetration of the organism, which may or may not proceed to the further development of active infection. Examples are primarily parasitic, including human hookworm “ground itch,” which occurs at the site of primary penetration.17 When nonhuman helminths, such as feline or canine hookworms, penetrate the skin, they are unable to complete their life cycle and migrate through the epidermis with characteristic serpiginous tracks, called CLM, created by the worm and the local host immune reaction (see Chapter 109). “Swimmer’s itch,” caused by avian schistosome cercariae penetrating the skin of sensitized people, is another example of an abortive helminthic infection.18,19 The immunologic nature of the response (immunoglobulin E (IgE)-mediated histamine response in this case) is shown by the absent or minimal reaction in previously unexposed people.20,21 Similar pruritic reactions occur in human schistosomiasis at the time of cercarial penetration (see Chapter 122).19 Myiasis and arthropod infestations, such as scabies and lice, are further examples in this category.22–25 Arthropod bite and sting reactions are similar in that they usually represent a local immune reaction to deposited salivary contents that many vectors use to facilitate their blood meal. Mosquito salivaspecific IgE antibodies have been detected in humans.26,27 The fourth pathogenetic category involves dissemination to the skin of toxins produced by infectious agents.9 The causative microorganisms are typically localized and distant from the skin. Examples are toxic shock syndrome, streptococcal scarlet fever, and staphylococcal scalded skin syndrome (see Chapter 30).9,28 Depending on the agent involved and the host response, diffuse erythema or vesiculobullous lesions can be seen, at times with associated hemodynamic instability. The fifth category relates to mechanical disruption of normal cutaneous homeostatic mechanisms resulting in pathologic changes in the skin and its supporting structures. Lymphatic filariasis, with its inflammatory and physical distortion of the normal lymphatic structures, is a good example of this category.29,30 The resulting lymphedema and increased susceptibility to bacterial infection contribute to skin changes that can occur over time. Similarly, lesions that form ulcers significantly increase the likelihood of secondary bacterial infection, usually prevented by an intact dermal–epidermal layer. This is often compounded by the limited

Figure 132.1  Erythema nodosum.

(Courtesy of Kenneth J. Tomecki, MD, Cleveland, OH.)

Bacteria Bartonella spp. (cat-scratch disease) Brucella spp. Chlamydia trachomatis (lymphogranuloma venereum) Chlamydia psittaci (psitticosis) Francisella tularensis Streptococcal infections Yersinia spp., Campylobacter spp., Salmonella spp. (diarrheal diseases) Mycobacteria M. tuberculosis M. leprae M. marinum

A

Viruses Cytomegalovirus Epstein–Barr virus Hepatitis C virus Vaccinia (smallpox inoculation agent)

Fungi Blastomycosis Cryptococcosis Coccidioidomycosis Histoplasmosis Trichophyton: deep-seated infection Protozoa Giardia lamblia Trypanosomiasis, African Helminths Ascaris lumbricoides Filariases (especially Wuchereria bancrofti)

Cutaneous Lesions Chapter 132

Box 132.1  Differential Diagnosis of Erythema Nodosum

Drugs Noninfectious: systemic lupus erythematosus, sarcoid, pregnancy, Crohn’s disease, ulcerative colitis, Behçet’s syndrome Idiopathic (up to 40%)

Box 132.2  Causes of Erythema Multiformea

B

Figure 132.2  Erythema multiforme of the mouth (A) and hand (B). (Courtesy of Kenneth J. Tomecki, MD, Cleveland, OH.)

hygiene available in many developing countries to both residents and travelers. A sixth, less well-understood, category involves an apparently systemic immunologic pathogenesis. Examples include erythema nodosum (Fig. 132.1), classic erythema multiforme, and Stevens–Johnson syndrome (SJS) (Fig. 132.2). In some cases of erythema multiforme, herpes simplex virus or Mycoplasma pneumoniae organisms have been found in the skin of such lesions, but most cases lack evidence of demonstrable cutaneous microbial antigen localization or toxin production.31–38 Infectious, noninfectious, and idiopathic causes are seen in this category. Infectious agents associated with erythema nodosum (Box 132.1) and erythema multiforme (Box 132.2) include bacteria, mycobacteria, fungi, viruses, and helminths. In addition, many serious cutaneous manifestations due to medications are mediated through this mechanism.39–41

GENERAL APPROACH TO THE PATIENT In approaching patients with rash and tropical exposures, there are three important steps to help define the diagnostic possibilities. First, the patient’s general medical and exposure history needs to be obtained.42 Second, the rash should be accurately defined based on morphology (e.g., macule, papule, vesicle, and nodule), location, and distribution.43,44 Third, associated clinical information gathered from a complete medical history and physical examination needs to be integrated. Such information includes a medication list, any sensations associated with the rash, pigmentation, migratory nature, duration, and changes in the rash over time.

Bacteria Chlamydia spp. Proteus spp. Francisella tularensis Salmonella spp. Staphylococcus spp. Streptococci, hemolytic Vibrio spp. Yersinia spp. Mycoplasma pneumoniae Mycobacteria Mycobacterium tuberculosis Viruses Herpes simplex virus types 1 and 2 (commonly associated) Epstein–Barr virus Adenovirus Coxsackievirus (especially B5) Orf virus

Fungi Histoplasmosis Coccidioidomycosis Parasites Cutaneous larva migrans Trichomonas Drugs Nonsteroidal anti-inflammatory agents Antituberculosis drugs Sulfonamides Case Reports Mebendazole, mefloquine, albendazole Streptococcal toxic shock syndrome Idiopathic (up to 50%)

a

Important to differentiate from giant urticaria and Stevens–Johnson syndrome (erythema multiforme major).

Ancillary clinical information not directly related to the rash must also be considered, including other organ system involvement and the results of laboratory tests. Patients often present with a specific rash, which provides a starting point for the evaluation and construction of the differential diagnosis.45,46 The importance of careful characterization of the rash cannot be overemphasized. This chapter is organized to provide important general information regarding the evaluation of patients with rash who have potential infectious diseases exposures and includes tables organized by the character of the specific lesion or primary associated symptom to facilitate the practical usefulness of the information. We focus the text and tables on

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parasite-associated skin conditions and those nonparasitic conditions that are more prevalent in tropical and subtropical climates. Most of the tables based on presenting cutaneous manifestations (Tables 132.6–132.10) include information about the geographic distribution of the disease entities. They also provide acquisition and incubation information together with parasite survival times to help assess which diagnoses fit the medical history of exposure. Common associated findings are included since cutaneous findings are often not the sole problem but, rather, one aspect of a multisystem disorder. The presence or absence of specific associated findings may also help the provider to focus further evaluation by knowing what other things to look for to rule in or rule out a specific diagnostic consideration.

HISTORY A detailed history (Box 132.3) is the starting point in evaluating a patient with a cutaneous lesion who has traveled abroad. In broad terms, the

Box 132.3  Important Historical Aspects Regarding Etiologies of Rash Patient Demographics Age, sex, occupation Reason for travel (vacation versus business versus other) Medications, allergies, immunizations Pretravel evaluation (compliance with pretravel recommendations?) Medical conditions that may alter immune status Time interval from travel dates to onset of symptoms Exposure Histories Geographic location, description of location, duration of exposure Vector exposures (precautions taken?) Animal exposures (wild versus domestic) List of items purchased (e.g., animal hide rugs, nickelcontaining jewelry)

Sexual contact with new partners Parenteral exposure (e.g., vaccine or injection abroad, acupuncture, tattoos, poultices on open sores) If Immigrant Country of origin Age on arrival in developed country Frequency and duration of return visits Rash-related Information Prodromal symptoms Character, distribution, progression, including speed and physical pattern Relation to fever Previous treatment and efficacy

history has three main components. First is information about the patient, second is information about the patient’s exposure to possible pathogens, and third is a detailed history regarding the rash.47 As a general rule, travel within the past 3 months is more likely to be relevant to the diagnosis of an acute illness than more remote travel.47,48 The underlying immune status of the patient also has important implications regarding the differential diagnosis. Immunosuppressed travelers may be at increased risk of acquiring intestinal protozoa, including Giardia lamblia, Isospora belli, Entamoeba histolytica, and Cryptosporidium.49,50 Interestingly, there is little evidence that human immunodeficiency virus (HIV) infection increases the rate of helminthic infection or decreases the efficacy of treatment, with the limited exception of strongyloidiasis.51 Some infections acquired in the developing world have the potential for reactivation or dissemination years later, especially in the setting of immunosuppression. Examples include coccidioidomycosis, histoplasmosis, hepatitis B, leishmaniasis, strongyloidiasis, and Trypanosoma cruzi infection.52 Details related to the specific exposures of an individual patient are a crucial aspect of the history. A detailed dietary history is essential. Even if high-risk exposures are not identified, foodborne illnesses such as hepatitis A can be acquired from food preparers who themselves reside in high-risk areas for infections and may contaminate a traveler’s food during its preparation.53 Because sexually transmitted infections rank high on lists of disease prevalence in travelers (see Chapters 126 and 138), a sexual history is an essential part of the exposure evaluation.1,54 This should include inquiry regarding the use of condoms. Moreover, many sexually transmitted infections, such as primary HIV, syphilis, and the group of genital ulcerative conditions, have prominent cutaneous findings (Table 132.1 and Fig. 132.3D). Vector exposure related to geography, types of vectors present, and intensity of exposure all need to be considered. Many tropical diseases have very defined distributions (even to regions within countries), making geographic exposure very important. Emerging infections (e.g., dengue, chikungunya, West Nile, Zika virus) and reemerging ones (measles, Staphylococcus aureus – including methicillin-resistant S. aureus) represent challenging concerns as international travel increases exposure to these pathogens with broader geographic distributions.55–57 Of note, fever and rash are prominent presentations for most of this group of infections. Recent exposure is typically most important, especially exposure within the past 3 months.58 However, as noted in the tables, malaria, many helminths, and other tropically acquired pathogens can persist for months to years in the human host, requiring their consideration in selected clinical settings well beyond the actual time of travel.42 In such cases, the presentation is typically subacute or chronic, although it may be punctuated by acute episodic illness.

Table 132.1  Dermatologic Manifestations of Sexually Transmitted Diseases

952

Disease

Ulcer

Chancroid Donovanosis Gonorrhea Granuloma inguinale HIV Herpes simplex virus Lymphogranuloma venereum Syphilis, endemic (bejel) Syphilis, venereal

X (p) X (nt)

Yaws Scabies

X (nt)

Nodule

Maculopapular

Pigment Change

Lymphadenopathy

Other

X (p, L) If disseminated

X (nt)

X

Pseudobubo X (nt, G)

1° X (p) X (nt) X (nt) X (nt)

Occasional painful necrotic ulcer Vesicle ulcerates

X

X (p, L) ↓ (late)

X 2°

X

X X

↓ (late)

X (G) 2° (L, G)

Occasional oropharyngeal mucous patches

X (L)

X, lesion develops; G, generalized lymphadenopathy; HIV, human immunodeficiency virus; L, localized lymphadenopathy; nt, nontender; p, painful; 1°, primary disease; 2°, secondary disease; ↓, hypopigmented.

Cutaneous Lesions Chapter 132

A

B

C

D

E

Figure 132.3  (A) Tropical phagedenic ulcer. (B) Buruli ulcer. (C) Myiasis extruding larva. (D) Granuloma inguinale. (E) South American blastomycosis (paracoccidioidomycosis). ((A–C) Courtesy of Jay S. Keystone, MD. (D and E) Courtesy of R. L. Guerrant, MD, Charlottesville, VA.)

Understanding the life cycles of parasitic pathogens is important when obtaining the history, as the life cycle may predict when a traveler becomes ill. For example, Loa loa-associated symptoms often require at least 6 months to appear and can first present as late as 18 months after the last possible exposure (see Chapter 105).59 Because of an autoinfective cycle, Strongyloides stercoralis infections (see Chapter 117) acquired decades earlier can result in risk to people who require immunosuppression many years after their disease exposure and acquisition.60,61 Another exposure-related issue is whether the patient is a lifelong resident of a developed country or has emigrated there recently or remotely. Expatriates are more likely to have an allergic-hypersensitivity reaction to L. loa infection compared to the local population.62 The frequency and duration of return visits to tropical areas may be very important information in creating the proper differential diagnosis. Malaria may be more frequent and severe in people who have returned to an endemic area after several years away. People traveling to visit family and relatives in malarious countries are often less rigorous about seeking pretravel advice and using prophylactic medication (see Chapter 126). Rarely, the index patient being evaluated has not had exposures in parasite endemic regions but has become infected via contacts through a friend or worker. For example, cysticercosis in Orthodox Jewish community members in New York who had not traveled outside the United States was attributable to household employees from endemic areas in Central America.63 Questioning the patient about unusual situations or risky behaviors may uncover relevant events, such as ingestion of raw seafood or questionable medical interventions. Acupuncture with unsterilized needles carries a risk of hepatitis and HIV transmission. A peculiar route of acquisition of sparganosis is seen when raw frog flesh is used as a poultice (see Chapter 121). Physicians unaware of alternative medical practices of some immigrant populations usually do not consider such a route of exposure when obtaining a history. Asking about souvenirs purchased may reveal

exposure to animal products (e.g., animal skin rugs) that carry the risk of disease transmission, such as brucellosis or anthrax, or to new materials (e.g., nickel-containing jewelry) that can result in allergic contact skin reactions.64 Occasionally, the history may help to point to an obsessional concern about the presence of parasites without other evidence to support such a diagnosis, and the possibility of delusional parasitosis should be considered (see Chapter 140).65,66 Specific questions about the rash and any associated clinical symptoms, such as fever pattern, weight loss, diarrhea, and response to any attempted treatments, are also important. The symptoms related to the rash may, at times, provide important diagnostic clues. For example, patients with myiasis (see Chapter 124), produced by cutaneous invasion by larvae of the bot fly in Latin America (Fig. 132.3C) or tumbu fly in Africa often present with papules, nodules, and subcutaneous swellings and will frequently describe a sensation of movement and intermittent pain at the site of the lesion.

PHYSICAL EXAMINATION A complete physical examination is important since cutaneous manifestations may be only one component of a systemic disorder. The entire skin should be examined with attention to the elements listed in Box 132.4. Repeated examination over time may be necessary. The rapidity of spread of a rash can have implications regarding the aggressiveness of the evaluation. The general level of a patient’s toxicity and the nature, tempo, and pattern of the rash dictate the initial evaluation and therapeutic approach. Although only 7% of cases with petechiae and fever will turn out to be meningococcemia,67 the life-threatening nature of meningococcal disease requires an aggressive evaluation and often empirical therapy when this diagnosis is a consideration (see Chapter 24).8,9

953

Box 132.4  Important Aspects of the Physical Examination in the Patient with Rash Vital signs General appearance • Level of toxicity to establish tempo of evaluation and therapy Primary character of the rash • Petechial or hemorrhagic • Macular • Papular • Migratory • Urticarial • Nodular • Ulcerative Secondary character of the rash • Shape (linear versus circular versus irregular)

• Size • Scale on surface • Other Location of primary lesion, if any Distribution of rash Direction of spread (centripetal or centrifugal) Mucosal or conjunctival involvement Genital involvement Associated physical findings • Visible parasite at the site of the rash • Lymphadenopathy • Splenomegaly or hepatosplenomegaly

Section III

progression, the rash usually becomes petechial or purpuric or both (Fig. 132.5). Dengue fever rash often begins as a transient macular exanthem approximately 24 hours after the abrupt onset of fever (see Chapter 75). Three or 4 days later, a generalized maculopapular truncal rash appears, with centripetal spread, sparing the palms and soles (Fig. 132.6). This rash may become petechial and may even desquamate.71–73 The original and subsequent distribution of the rash, consideration of mucosal (enanthem) or genital involvement, and other associated physical findings may be helpful in establishing a diagnosis. The presence of lymphadenopathy (Box 132.5) and splenomegaly or hepatosplenomegaly Box 132.5  Differential Diagnosis of Infections with Rash and Lymphadenopathy Helminths Lymphatic filariasis Brugia malayi B. timori Wuchereria bancrofti Loiasis Mansonelliasis Onchocerciasis Opisthorchiasis Schistosomiasis (Katayama fever) Protozoa Leishmaniasis Cutaneous and mucocutaneous Visceral Toxoplasmosis

Trypanosomiasis African American Fungi Coccidioidomycosis Chromomycosis Cryptococcosis Histoplasmosis Lobomycosis Paracoccidioidomycosis Penicilliosis marneffei Sporotrichosis Bacteria Numerous Viruses Numerous, including primary human immunodeficiency virus

  

 

PRACTICE: APPROACH TO THE PATIENT IN THE TROPICS

The classification of the rash is often the focal entry point into the creation of a differential diagnosis.4,5,68 As a result, the primary and secondary characteristics of the rash should be carefully determined (Box 132.4). Some rashes are fleetingly present, such as Salmonella spp. rose spots. Other rashes may evolve in relatively unique patterns over time, such as anthrax, Rocky Mountain spotted fever (RMSF), and dengue.22,69–73 For example, cutaneously acquired anthrax (see Chapter 38) characteristically evolves from a papule to a vesicle with surrounding brawny, gelatinous edema, and occasional satellite vesicles69,70 (Fig. 132.4). The lesion then progresses to hemorrhage and necrosis, eventually turning into an eschar. RMSF skin changes are often absent in the first few days of clinical illness. Then a macular rash typically starts on the wrists and ankles with centripetal spread (see Chapter 49).22 During

Figure 132.4  Cutaneous anthrax showing characteristic central eschar with surrounding edema and vesicles. (Courtesy of Kenneth J. Tomecki, MD, Cleveland, OH.)

A

954

Figure 132.5  Early macular rash of Rocky Mountain spotted fever. Papular and petechial changes are often seen. The rash typically begins around the wrists and ankles.

B

Figure 132.6  Cutaneous manifestations of dengue. (A) Early maculopapular nonpruritic rash usually seen at the time of defervescence. (B) Late hemorrhagic and purpuric skin changes of a patient with dengue hemorrhagic fever. (Courtesy of James H. Maguire, MD, Centers for Disease Control and Prevention, Atlanta, GA.)

Characterizing the Rash Careful identification of the primary and secondary characteristics of the rash will facilitate creating a differential diagnosis, especially when integrated with the clinical history and any associated physical findings. After Box 132.6  Differential Diagnosis of Rash with Splenomegaly or Hepatosplenomegaly Organomegaly Commonly Associated Brucellosis Ehrlichiosis Epstein–Barr virus infection Glanders (Burkholderia mallei) Leishmaniasis, visceral Penicilliosis (Penicillium marneffei) Q fever Relapsing fever (Borrelia recurrentis) Schistosomiasis (Schistosoma mansoni, S. japonicum) Trypanosomiasis (Trypanosoma cruzi)

Organomegaly Occasionally Associated Cytomegalovirus infection Histoplasmosis Leptospirosis Malaria Psittacosis Rocky Mountain spotted fever (Rickettsia rickettsii) Typhoid fever Scrub typhus (R. tsutsugamushi)

defining the rash, the reader should refer to the table related to that characteristic to help with diagnosing the individual patient. The role of tropical exposures is highlighted here. Significant medical conditions with prominent cutaneous manifestations may be due to common pathogens with worldwide distribution despite exposure to specific tropical pathogens. For example, although petechiae in the recently returned febrile traveler may represent malaria, dengue, or even Lassa fever, it is still crucial to consider meningococcemia, rickettsial disease, and other diagnoses. Life-threatening diagnoses with cutaneous manifestations are seen throughout the temperate and tropical world (Table 132.3).8,9

Hemorrhage and Petechiae Hemorrhagic or petechial rashes (Table 132.4) must be considered of life-threatening importance and evaluated rapidly.8,9,22 Although many nonlethal rashes have this presentation, it is difficult, if not impossible, to rule out serious pathogens at the initial evaluation. Worldwide, meningococcemia is the most important cause of life-threatening disease presenting with fever and cutaneous hemorrhage or petechiae (see Chapter 24). Infection-mediated petechial or purpuric lesions may arise directly from infected emboli or via an indirect immunologic mechanism.

Cutaneous Lesions Chapter 132

(Box 132.6) are examples of findings that may help focus the evaluation. If a primary lesion or an inoculation site can be determined, it should be given careful attention. For example, an eschar would indicate rickettsial disease or anthrax (Table 132.2), whereas periorbital swelling in a child soon after a visit to rural Brazil would focus the evaluation on Chagas’ disease. Despite these examples in which the appearance of the rash can point to the diagnosis, the appearance of the rash is often not pathognomonic of a single disease entity.

Table 132.2  Diagnostic Considerations in the Patient with an Eschar

Organomegaly Rarely Associated Babesiosis Drug reaction Serum sickness

Pathogen

Condition (Vector)

Rickettsia africae R. akari R. australis R. conorii R. sibirica R. tsutsugamushi Bacillus anthracis Loxosceles reclusa

African tick bite fever Rickettsialpox (mite) Queensland coastal fever (tick) Mediterranean spotted fever (tick) North Asian tick typhus Scrub typhus (mite) Anthraxa Brown recluse spider bite

a

Usually transmitted by contact with infected animals or their products, rarely mechanically via biting flies or insects.

Table 132.3  Life-threatening Conditions with Cutaneous Manifestations Time to Appearance (after Onset of Illness)

Pathophysiology

Gram-negative bacteria Neisseria meningitidis Rickettsia rickettsii Gram-negative bacteria Pseudomonas spp. Gram-negative bacteria N. meningitidis N. gonorrhoeae N. meningitidis N. gonorrhoeae Salmonella spp. Salmonella spp. Staphylococcus aureus

12–36 hours 12–36 hours (later for Rickettsia)

Vascular thrombosis due to shock Vascular invasion (?) Schwartzman reaction

Days

Vascular invasion, mostly venous

3–10 days

Immune vasculitis due to immune complex deposition Immune vasculitis

Streptococcus spp. S. aureus S. aureus Streptococcus pyogenes Candida spp. Strongyloides stercoralis (hyperinfection syndrome) Viral hemorrhagic fevers

Days to weeks

Immune vasculitis Emboli from endocarditis with microabscesses Emboli from endocarditis

Minutes

Vascular toxin with peripheral vasodilation

Several days Hours to days from immunosuppression Within 3 weeks of exposure

Vascular invasion of dermis ? Larval skin penetration ? Immune vasculitis Immune vasculitis; disseminated intravascular coagulation, thrombocytopenia, decreased clotting factor synthesis

Cutaneous Manifestation

Pathogen

Peripheral gangrene Scattered, multiple petechiae and purpura Ecthyma gangrenosum Asymmetrical scattered maculopapular rash Polymorphous lesions

Rose spots Osler’s nodes; small ( female; can have 2° tetanus and gas gangrene; multiple and recurrent ulcers seen Rare form of tuberculosis

Tularemia

Unusual direct cutaneous inoculation Bite of infected arthropod or animal; contact with infected animal tissue; inhalation; ingestion of contaminated meat or water

Papule/pustule/vesicle → painful ulcer 1–6 cm diameter, circular raised edge and surrounding edema; deeply penetrating and destructive Inoculation papule can ulcerate Clinical sx reflect portal of entry; skin inoculation → local, painful ulcer followed by painful regional LN, fever; rare erythema nodosum

Disease range is asymptomatic to fatal; ulceroglandular form most common; chronic/ recurrent fevers and LN seen

Yaws

See Table 132.8 See Table 132.10

All invasive fungal infections can cause ulcerative skin lesions

Deep abscesses and sinus tracts may form

Fungal

3–5 days (range, 1–25 days)

Helminthic Dracunculiasis

See Table 132.6

Ulcer at site of worm eruption

Protozoan Amebiasis, cutaneous

Worldwide See also Table 132.8

Leishmaniasis, cutaneous and mucocutaneous

See Table 132.8

  

 

PRACTICE: APPROACH TO THE PATIENT IN THE TROPICS

Table 132.9  Differential Diagnosis of Ulcerative Skin Lesions—cont’d

Cyst ingestion in contaminated food or water; uncommon sexual transmission

1–3 weeks (range, 2 days to months)

Months to years

When cutaneous, painful, rapid-growing ulcers with necrosis

Uncommon sexual transmission: anal > vaginal intercourse

Type 1: direct contact with infected oral secretions Type 2: contact with genital secretions

2–12 days

Lifetime in human host

Grouped vesicles on erythematous base ulcerate; lesions painful; moderate to severe generalized systemic sx with 1° infection; local LN common

Recurrence due to latent virus in neural cells; autoinoculation possible

Spider bite

Immediate to hours

Bite → mild sting followed by intense pain in 2–8 hours, bullae and erythema, local ischemic necrosis, deep ulceration Occasional fever, myalgias, and generalized morbilliform rash 24–48 hours after bite

Parenteral steroid use in first 24 hours controversial; heals spontaneously though scars common Can be confused with pyoderma gangrenosum

Viral Orf (Parapoxvirus) Herpes simplex

See Table 132.8 Worldwide

Arthropod Brown recluse spider bite (Loxosceles reclusa)

?

Myiasis

See Table 132.7 See Table 132.7

Tungiasis

968

sx, symptoms; GI, gastrointestinal; LN, lymphadenopathy; CNS, central nervous system; CXR, chest X-ray film. a Several rickettsial infections are associated with an ulcerative eschar; see Table 132.2.

Entity

Distribution

Acquisition

Incubation

Chromomycosis (several agents)

Worldwide, especially tropics and subtropics

Percutaneous inoculation of nonintact skin

Weeks to months

Entomophthoromycosis (Basidiobolus baptosporus)

Tropical Africa, rarely elsewhere

Percutaneous inoculation of nonintact skin

Lobomycosis (Loboa loboi)

Tropical Central and South America

Mycetoma Mixed bacterial and fungal infection; begins as subcutaneous nodule

Organism Survival

Common Associated Findings

Additional Information

Years

Nodule I ++++

?

Years

Percutaneous inoculation of nonintact skin

1–2 years

Years

Worldwide (especially tropics)

Percutaneous inoculation

Weeks to months

Years (up to 25 years untreated)

Pityriasis (tinea) versicolor (Malassezia furfur)

Worldwide

Direct inoculation of fungal elements

?

?

Sporotrichosis (Sporothrix schenckii)

Worldwide

Direct percutaneous inoculation

3–8 weeks

Months to years

Tinea imbricata (Trichophyton concentricum)

Asia; South Pacific islands; Mexico, Central and South America

Direct contact with infected person

10 days

Lifetime of host

Painless subcutaneous nodule → expanding woody swelling Ulcer Nodule None I ++++ Starts as painless, small mobile nodules Ulcer Nodule S I + ++++ Granules may be discharged from wound; sinus tracts common; nodules slowly enlarge and become phlegmonous Ulcer Nodule S I ++ +++ Circumscribed hypo- or hyper- (brown) pigmented scaly lesions; usually upper chest and back Nodule Ulcer None None Papule → painless; nodule → may ulcerate; secondary nodules along lymphatic channels Ulcer Nodule S I ++ ++++ Concentric rings 0.3–1.25 cm apart; no systemic sx except pruritus Nodule Ulcer None None

Lymphatics involved, (e.g., elephantiasis and lymphedema); starts as local papule → painless, irregular, papule → nodule Rare systemic sx or dissemination

Primary Skin Infections Ulcer S +++

Cutaneous Lesions Chapter 132

Table 132.10  Differential Diagnosis of Nodular and Ulcerative Fungal Infections

Autoinoculation seen; histopathology characteristic

Systemic sx usually absent; male:female 4 : 1; local bone invasion with cortical erosion, lytic lesions; tendons spared; path: granuloma around purulent center Superficial stratum corneum infection; enzyme of fungus produces pigment changes 25% extracutaneous lung, bone, joints

Hypopigmentation >hyperpigmentation; covers up to 70% of body

Primary Nasal and Oropharyngeal Infections Entomophthoromycosis (Conidiobolus coronatus)

Tropical Africa; rarely elsewhere

Inhalation of spore usually on to nasal turbinate

?

Years

Rhinosporidiosis (Rhinosporidium seeberi)

Worldwide (sporadic); endemic foci in India and Sri Lanka

Direct inoculation of fungus

Weeks to months

Years to decades

Painless indurated mass → local insidious spread Ulcer Nodule None I ++++ Characteristic, friable, vascular sessile growth or polyps on mucosal surfaces; lesions painless Ulcer Nodule None I ++++

Male : female 8 : 1

Treatment surgical, but recurrence common

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Table 132.10  Differential Diagnosis of Nodular and Ulcerative Fungal Infections—cont’d Entity

Organism Survival

Distribution

Acquisition

Incubation

Common Associated Findings

North America (most); sporadic in Central and South America, Africa, Poland, India, Middle East Southwest United States, Mexico, Central and South America

Inhalation of infectious conidia; rare percutaneous inoculation; rare STD from men with GU infection

1–2 months inhaled; 2 weeks cutaneous

~15 years in human host

Nodule I +++

Inhalation of infectious arthroconidia; rare cutaneous inoculation

2 weeks

Years

Cryptococcosis (Cryptococcus neoformans)

Worldwide

Inhalation of infectious conidia

?

?

Histoplasmosis (Histoplasma capsulatum)

Worldwide (but focal and sporadic)

Inhalation of infectious spores

2 weeks

Years

Histoplasmosis, African (Histoplasma capsulatum, var. duboisii)

Tropical Africa

Uncertain; suspect inhalation of infected spores

?

Years

Paracoccidioidomycosis (Paracoccidioides brasiliensis)

Central and South America

Inhalation of infectious spores

Months to years

Years to decades

Penicilliosis (Penicillium marneffei)

East and Southeast Asia

Inhalation/ ingestion of infectious spores; rare percutaneous inoculation

9 days to 1 month

Years

1° disease; fever, cough, chest pain, malaise; hypersensitivity reactions (15–20%): erythema nodosum, generalized erythema, maculopapular rash, arthralgias, urticaria, rare erythema multiforme Nodule Ulcer + + Mostly pulmonary with no sx; skin lesions 10% (hematogenous papule → nodule → ulcer) Ulcer Nodule S I ++ ++ Mostly pulmonary with no sx; skin changes (erythema nodosum) seen with reactions to 1° infection Ulcer Nodule I I Oropharynx Often oropharynx + + Common painless skin lesions from skin and underlying bony disease Ulcer Nodule I I +++ +++ Initial respiratory infection usually no sx; common mucocutaneous oropharyngeal lesions due to hematogenous dissemination (60%) Ulcer Nodule I I ++ +++ Nodules are subcutaneous abscesses; ulcers from necrosis of nodules/ papules Ulcer Nodule S I ++ +++

Additional Information

Primary Respiratory Infections Blastomycosis (Blastomyces dermatitidis)

Coccidioidomycosis (Coccidioides immitis)

Ulcer S ++

Most hematogenous to skin; 20–80% of patients disseminate to skin, bone, GU tract (male > female), CNS; male : female 3–9 : 1

Extrapulmonary disease rare ( male

Antimalarials Chloroquine

Mefloquine Proguanil

Primaquine Doxycycline

Quinine

Halofantrine

Pyrimethamine use Renal failure

Female > male 2 of 6 patients receiving concurrent chloroquine

Incidence 0.3–2.3%; black race; family member with history of pruritus posttreatment Family member with history of pruritus posttreatment

Antidiarrheals Pepto-Bismol Trimethoprim– sulfamethoxazole

  

 

PRACTICE: APPROACH TO THE PATIENT IN THE TROPICS

Table 132.13  Cutaneous Reactions to Travel Medications

Ciprofloxacin

Ofloxacin

Metronidazole Furazolidone

Salicylate hypersensitivity, erythematous rash, bismuth reaction, transient darkening of tongue Diffuse maculopapular rash, allergic/toxic dermatitis Rare: erythema multiforme Exfoliative eczema exanthems Stevens–Johnson syndrome TEN Generalized skin rash Rare: pruritus, urticaria, photosensitivity, flushing, angioedema, hyperpigmentation, erythema nodosum, fixed drug eruption, pustulosis, TEN Generalized skin rash, pruritus, eczema Rare: hypersensitivity vasculitis, phototoxic dermatitis, photo-onycholysis Hypersensitivity reaction, flushing, ± erythematous rash Rare: pityriasis rosea, fixed drug eruption Maculopapular eruptions, vesicular morbilliform, hypersensitivity reaction, pruritic rash, contact dermatitis, erythema multiforme

Overall incidence 2.2%; HIV/AIDS higher incidence ∼1/200 000 doses ∼1/200 000 doses Incidence of 1.1%

Incidence up to 14% of patients

Alcohol ingestion 0.5% incidence rate overall (based on evaluation of 10 433 patients)

Anthelminthics Albendazole

Hair loss (reversible) Rash and pruritus Stevens–Johnson syndrome Diethylcarbamazine Hypersensitivity reaction (Mazzotti reaction) Ivermectin Rash and pruritus Biting arthropod repellent Diethyltoluamide Rare: local irritant reaction, scarring bullous dermatitis

High-dose chronic therapy

Onchocerciasis >> Loiasis > other filariae

HIV/AIDS, human immunodeficiency virus/acquired immunodeficiency syndrome. TEN, toxic epidermal necrolysis. (Data from references 105–117, 119–123, 125–134.)

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SECTION III: PRACTICE: APPROACH TO THE PATIENT IN THE TROPICS

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CHAPTER 133  Hepatobiliary Disease Mohammad S. Khuroo • Mehnaaz S. Khuroo

INTRODUCTION

DISEASES CAUSED BY HEPATITIS VIRUSES

A broad group of infections affect hepatobiliary organs in the tropical countries. These include infections prevalent in nontropical zones and those restricted to tropical zones of the world (Table 133.1). Amongst the hepatitis viruses, hepatitis E virus (HEV) has the most significant palpable impact.1 A number of nonviral infectious agents predominantly prevalent in tropical countries affect the liver and biliary tract as primary targets.2 Commonest amongst these are amebiasis, echinococcosis, schistosomiasis and hepatobiliary and pancreatic ascariasis (HBPA). Tuberculosis and the acquired immunodeficiency syndrome (AIDS) are endemic in tropics and can involve both the liver and/or biliary tree.3,4 Several viral and nonviral tropical infections such as malaria, typhoid, dengue fever, and leptospirosis may affect the heptobiliary system as a part of multiorgan involvement.5 Hepatobiliary infections prevalent in tropical countries have a special predilection and distinct pattern for expatriate Western travelers to such countries.6,7

Acute Sporadic Viral Hepatitis

DIAGNOSTIC CONSIDERATIONS Methodology to be followed for proper evaluation of hepatobiliary diseases in tropics is no different than that employed in nontropical countries.8 This includes a proper history, thorough physical examination, routine blood counts and serum chemistries including bilirubin, alanine aminotransferase (ALT), aspartate aminotransferase (AST), and alkaline phosphatase (ALP). Abnormal liver tests can be classified into definite patterns: (1) hepatocellular (elevated bilirubin; elevated ALT/AST >500 IU; ALT>AST; normal to 4 times normal elevation of ALP). Each pattern has different clinical connotations and defines corresponding further workup. Hepatobiliary ultrasound with Doppler is a valued complement to clinical examination for hepatobiliary diseases and is strongly recommended as a part of primary evaluation. Ultrasound with Doppler is useful for detecting gallstones, thickened gallbladder, dilated bile ducts, cysts and masses within the liver, periportal fibrosis, hepatosplenomegaly, and evidence of portal hypertension.9 Based on findings, further cost-effective, evidencebased laboratory tests, imaging tools such as computed tomography (CT), magnetic resonance imaging (MRI), magnetic resonance cholangiopancreatography (MRCP), endoscopic retrograde cholangiopancreatography (ERCP), or other invasive tests such as liver biopsy or guided fine needle aspiration biopsies may need to be employed.

Viral hepatitis (see Chapters 64–66) is highly endemic in tropical countries and is a cause of significant morbidity and mortality. Etiology of acute sporadic hepatitis in these countries is different than in the West. Around half of sporadic viral hepatitis in the tropics is related to HEV. Hepatitis B virus (HBV), hepatitis C virus (HCV), and non-A-E hepatitis constitute around 22%, 9%, and 25% of patients, respectively. Hepatitis A virus (HAV), a dominant cause of acute hepatitis in the West, is an uncommon (