978-0-12-031751-6

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Advances in PARASITOLOGY

VOLUME 51

Editorial Board M. Coluzzi, Director, Istituto de Parassitologia, Università Degli Studi di Roma ‘La Sapienza’, P. le A. Moro 5, 00185 Roma, Italy C. Combes, Laboratoire de Biologie Animale, Université de Perpignan, Centre do Biologie et d’Ecologie Tropicale et Méditerranéenne, Avenue de Villeneuve, 66860 Perpignan Cedex, France D.D. Despommier, Division of Tropical Medicine and Environmental Sciences, Department of Microbiology, Columbia University, 630 West 168th Street, New York, NY 10032, USA J.J. Shaw, Instituto de Ciências Biomédicas, Universidade de São Paulo, av. Prof Lineu Prestes 1374, 05508-900, Cidade Universitária, São Paulo, SP, Brazil K. Tanabe, Laboratory of Biology, Osaka Institute of Technology, 5-16-1 Ohmiya Asahi-Ku, Osaka, 535, Japan P. Wenk, Falkenweg 69, D-72076 Tübingen, Germany

Advances in PARASITOLOGY Edited by

J.R. BAKER Royal Society of Tropical Medicine and Hygiene, London, England

R. MULLER London School of Hygiene and Tropical Medicine, London, England and

D. ROLLINSON The Natural History Museum, London, England VOLUME 51

Amsterdam Boston London New York Oxford Paris San Diego San Francisco Singapore Sydney Tokyo

This book is printed on acid-free paper. Copyright 2002, Elsevier Science Ltd. All Rights Reserved. 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. Academic Press An Imprint of Elsevier Science 84 Theobald’s Road, London WC1X 8RR, UK http://www.academicpress.com Academic Press An Imprint of Elsevier Science 525 B Street, Suite 1900, San Diego, California 92101-4495, USA http://www.academicpress.com ISBN 0-12-031751-6 A catalogue record for this book is available from the British Library

Typeset by M Rules, London, UK Printed and bound in Great Britain by MPG Books, Bodmin, Cornwall 02 03 04 05 06 07 MP 9 8 7 6 5 4 3 2 1

CONTRIBUTORS TO VOLUME 51 B. FRIED, Department of Biology, Lafayette College, Easton, Pennsylvania 18042, USA A. HEMPHILL, Institute of Parasitology, University of Berne, LänggassStrasse 122, CH-3012 Berne, Switzerland. Email: [email protected] K. KITA, Department of Biomedical Chemistry, Graduate School of Medicine, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 1130033, Japan D.A. MAYER, Department of Surgery, New York Medical College, Valhalla, New York 10595, USA M. SILES-LUCAS, Unidad de Parasitologia, Facultad de Farmacia, Universidad de Salamanca, Avenida del Campo Charro sn, 37007, Salamanca, Spain. Email: [email protected] S. TAKAMIYA, Department of Parasitology, School of Medicine, Juntendo University, Japan

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PREFACE

As this volume was going to press, we were saddened to hear of the death, at the age of 88, of Professor W. H. Russell Lumsden. Russell succeeded Ben Dawes, the founder, as senior editor of Advances in Parasitology in 1978 (Vol. 16), assisted by two of us (J. B. and R. M.), and continued in that role until 1982 (Vol. 20). He subsequently continued to serve as an editorial board member until prevented by illness a few years ago. Russell had a distinguished and varied scientific career. He qualified initially as a zoologist in Glasgow in 1935, and subsequently in medicine in 1938. He was awarded a DSc by the University of Glasgow in 1956 and an MD in 1975. Russell served in the British Army during the 1939–45 war, latterly as commanding officer of No. 3 Malaria Field Laboratory. Subsequently he specialized in virology, joining the staff of the Yellow Fever Research Institute (later the East African Virus Research Institute) at Entebbe, Uganda in 1947. He then transferred his interests to parasitic protozoa, becoming director of the East African Trypanosomiasis Research Organization in 1957 (where one of us, J. B., was working). From there he moved to the UK in 1963 and, after five years in the University of Edinburgh, he became Professor of Medical Protozoology in the University of London (at the London School of Hygiene and Tropical Medicine and Hygiene) in 1968. During that time he co-wrote (with W. J. Herbert and G. J. C. McNeillage) Techniques with Trypanosomes (Churchill Livingstone, 1973) and edited (with David Evans) the major two-volume reference work Biology of the Kinetoplastida (Academic Press, 1976–79). He retired from the Chair in 1979, becoming Professor Emeritus and returning to his native habitat (Scotland) to live in Edinburgh accompanied by his wife Pamela, who had supported him throughout his career and who survives him. Russell’s retirement was far from idle. He pursued a variety of interests until ill health forced him to relinquish many activities,

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including a study of the design and construction of Zambian arrowheads, a subject on which he published. The publisher’s announcement of Russell’s retirement from the editorship in Vol. 20 (p. vii) included the words ‘We . . . have found him always a delight to work with, constructive, helpful and responsive . . . .’ Those of us who were fortunate enough to work with Russell in this and other ways wholeheartedly support this assessment, wishing to add only a tribute to his, often slightly mischievous, sense of humour. We shall miss him greatly as colleague and friend. In the opening review of this volume, David Mayer of the New York Medical College and Bernard Fried of Lafayette College, Pennsylvania, USA have reviewed recent developments in the study of both protozoan and helminthic human parasitic diseases in which surgical intervention may be required. The introduction of minimally invasive surgery in the last ten years has revolutionized such interventions with much smaller incisions and consequent reductions in post-operative recovery times. The authors have also reviewed new information on other aspects of the infections such as epidemiology, pathogenesis, diagnosis, treatment and prevention. Kiyoshi Kita and Shinzaburo Takamiya (University of Tokyo, Japan) provide a detailed account of electron transfer complexes in the mitochondria of Ascaris and other parasitic worms in the second review. The authors point out that parasites have exploited unique energy metabolic pathways as adaptations to their habitats within the host. Respiratory systems of parasites tend to show a greater diversity in electron pathways than those of host animals. The review is timely as there have been recent developments in our understanding of the respiratory chain concerning both the molecular structure of the biochemical components and the changes that occur during the parasite’s life-cycle. A. suum could prove to be a useful model to study regulation of transcription by the oxygen level in the environment. And finally, Mar Siles-Lucas (University of Salamanca, Spain) and Andrew Hemphill (University of Berne, Switzerland) have provided an overview of the place of laboratory models, both in vivo and in vitro, in investigating the host–parasite relationships of cestodes. Most studies have involved genera of the greatest medical and veterinary importance, such as Echinococcus, Taenia and Spirometra, but there have also been models to maintain different lifecycle stages of Hymenolepis and Mesocestoides. These models have been used for many studies on the development, morphology and metabolism and gene expression of the cestodes, on drug screening against them, and on the pathology and immunology of the infections. John Baker Ralph Muller David Rollinson

CONTENTS CONTRIBUTORS TO VOLUME 51 . . . . . . . . . . . . . . . . . . . . . . v PREFACE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vii

Aspects of Human Parasites in which Surgical Intervention May Be Important D.A. Meyer and B. Fried

1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . Trypanosoma cruzi . . . . . . . . . . . . . . . . . . . . Entamoeba histolytica . . . . . . . . . . . . . . . . . . Leishmania donovani . . . . . . . . . . . . . . . . . . Strongyloides stercoralis . . . . . . . . . . . . . . . . . Taenia solium . . . . . . . . . . . . . . . . . . . . . . Schistosoma mansoni . . . . . . . . . . . . . . . . . . Paragonimus westermani . . . . . . . . . . . . . . . . Echinococcus granulosus . . . . . . . . . . . . . . . . Ascaris lumbricoides . . . . . . . . . . . . . . . . . . . Fasciola hepatica . . . . . . . . . . . . . . . . . . . . Surgical Treatment for Parasitic Infections Not Covered in the Text . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . .

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3 3 5 11 17 24 29 36 48 53 62 70

. . . . . 76 . . . . . 79

Electron-transfer Complexes in Ascaris Mitochondria K. Kita and S.Takamiya

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96

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CONTENTS

2. 3. 4. 5. 6. 7. 8. 9.

Energy Metabolism of Parasitic Helminths . . . . . . . Developmental Changes in the Respiratory Chain . . . . NADH-Fumarate Reductase System . . . . . . . . . . . NADH-dependent 2-Methyl Branched-chain Enoyl-CoA Reductase System . . . . . . . . . . . . . . . . . . . . Role of Rhodoquinone in Anaerobic Respiration . . . . Evolution of the Parasite Electron-transport System . . Heterogeneity in Helminth Mitochondria . . . . . . . . Conclusions and Perspectives . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . 97 . . . . . 100 . . . . . 107 . . . . . . .

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. . . . . . .

115 116 120 122 123 124 124

Cestode Parasites: Application of In Vivo and In Vitro Models for Studies on the Host–Parasite Relationship M. Siles-Lucas and A. Hemphill

Abstract . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . Laboratory Models for Studies on Echinococcus spp. . Laboratory Models for Studies on Taenia spp. . . . . Hymenolepis spp.: Versatile Cestode Parasite Models . Mesocestoides spp. As Experimental Models to Study Cestode Biology . . . . . . . . . . . . . . . . . . . . 6. Experimental Investigations on Spirometra spp. . . . . 7. Conclusions . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . 1. 2. 3. 4. 5.

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134 134 136 153 164

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180 186 193 194 194

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231

Aspects of Human Parasites in which Surgical Intervention May Be Important David A. Mayer1 and Bernard Fried2 1Department

of Surgery, New York Medical College, Valhalla, New York 10595, USA; 2Department of Biology, Lafayette College, Easton, Pennsylvania 18042, USA

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Trypanosoma cruzi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. American trypanosomiasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Entamoeba histolytica . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Amebiasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.7. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Leishmania donovani . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Leishmaniasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Strongyloides stercoralis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Strongyloidiasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ADVANCES IN PARASITOLOGY VOL 51 0065–308X $30.00

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5.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Taenia solium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. Taeniasis and cysticercosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Schistosoma mansoni . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. Schistosomiasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Paragonimus westermani . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2. Paragonimiasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Echinococcus granulosus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2. Echinococcosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ascaris lumbricoides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2. Ascariasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fasciola hepatica . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.1. Case report . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2. Fascioliasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3. Epidemiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4. Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.5. Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6. Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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11.7. Prevention . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 12. Surgical Treatment for Parasitic Infections Not Covered in the Text . . . . . 76 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79

ABSTRACT

Until recently, physicians and surgeons in developed countries only occasionally encountered patients with parasitic protozoan and helminthic infections. High-speed travel, immigration and the popularity of the tropics as vacation areas have increased the number of people at risk for parasitic disease. This chapter examines the significant literature on a select number of protozoan and helminthic parasites for which surgical intervention is important in the diagnosis, treatment or cure of the disease. Although traditional surgical approaches are covered, emphasis is placed on recent advances in the areas of transplantation and minimally invasive surgery. Combining the disciplines of parasitology and surgery, this chapter covers three protozoan and seven helminthic parasites for which surgery is a valid treatment option based on the frequency of cases reported in the literature. Following coverage of the selected parasites, a table is included listing additional helminths for which surgery contributes to patient management. Physicians in the USA, UK, and Europe need to be more aware of the presentation and treatment of parasitic infections. It is our sincere hope that this review accomplishes that goal, and ultimately benefits the patients we serve.

1. INTRODUCTION

The purpose of this chapter is to examine a select number of protozoan and helminth parasites for which surgical intervention is important in the diagnosis, treatment, or cure of the parasite. The review originally started out with the title ‘Surgical Intervention in Parasitic Diseases’. As the text developed, we realized the need to include more information about the parasites covered, i.e., case history, parasite transmission, epidemiology, life cycle, pathogenesis, clinical manifestation, and treatment other than surgical. With the change in emphasis of the text for the parasites covered, evolved the new title, ‘Aspects of Human Parasites in which Surgical Intervention May Be Important.’ After completion of coverage of the selected parasites, we added a table at the end of the chapter listing additional helminths for which surgery is an option in diagnosis, treatment, or cure. Two biomedical scientists with diverse backgrounds have written this

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review. One (DAM) is a general and vascular surgeon, who developed an interest in clinical parasitology as a surgical resident at Cornell University from 1973 to 1978. The second (BF) is an experimental parasitologist, who became interested in medical parasitology during his tenure as a Louisiana State University Medical Fellow in the Central American tropics in 1967. Because numerous medical and surgical terms are used in the text, we have provided definitions (in parentheses) of some of the more esoteric terms the first time they are used. These definitions should help parasitologists not familiar with the nuances of medical and surgical terminology. Protozoan and helminthic infections are of enormous importance worldwide, and the number of people infected or at risk of infection is overwhelming. Until recently, most physicians and surgeons in developed countries only occasionally encountered patients with such infections. At present, with increased mobility of large segments of the world’s population, the popularity of the tropics as vacation areas, and the high speed of transportation, physicians in the USA, UK, and Europe need to be more aware of parasitic infections. The situation has also been exacerbated by the emergence of immunocompromise, whether by AIDS, helminthic or protozoan infections, chemotherapy, or immunosuppressive drugs. A main emphasis of this chapter is on surgical intervention in parasitic diseases. It is important to remember that during the past approximately 10 years, surgery has undergone a revolution. In most instances, the patient is no longer forced to undergo painful operations done through large, slow-healing incisions. The era of minimally invasive surgery is here. The surgeon works with small incisions using video imaging through scopes placed in body cavities. Patients usually have minimal postoperative discomfort, with relatively quick recovery time. In response to parasitic infections for which surgery is a treatment option, minimally invasive procedures such as laser photocoagulation, laparoscopy, thoracoscopy, endoscopy, and image-guided needle aspiration are being used to treat and cure parasitic infections. Our review considers ten parasites (protozoans and helminths) for which surgery is a valid treatment option based on the frequency of cases reported in the literature. Each section begins with a case report, followed by a discussion of parasite transmission, epidemiology, life cycle, pathogenesis, clinical manifestation, and treatment, especially surgical. Some topics that relate to surgical intervention include liver transplantation for alveolar echinococcosis, heart transplantation for chronic Chagas’ disease, endoscopic management of biliary liver flukes, laparoscopic treatment of ascariasis, and percutaneous drainage of amebic liver abscess. When we completed coverage of the ten selected parasites, we realized there were other parasites for which surgical management is a treatment option. Information on surgical treatment of these parasites is considered in Table 1 at the end of the chapter.

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2. TRYPANOSOMA CRUZI 2.1. Case report

A 44-year-old woman from Ecuador who immigrated to the USA at age 21, presented with a mild stroke caused by embolization of clot from an aneurysm of the left ventricle of the heart. She had depressed left ventricular function with marked cardiomegaly. No coronary artery disease was found on angiogram, and endomyocardial biopsies showed diffuse myocardial fibrosis. No parasites were observed on microscopic examination. The diagnosis of Chagas’ cardiomyopathy was confirmed by a positive immunoprecipitation and complement fixation test. The patient experienced progressive deterioration of cardiac function, accompanied by severe mitral and tricuspid valvular regurgitation. Refractory ventricular tachycardia and intractable congestive heart failure made heart transplantation her only chance for survival. Orthotopic heart transplantation was performed. In this method, the donor heart is sutured in place in the anatomic bed of the explanted heart, as opposed to the heterotopic transplant, in which the new heart is placed on top of the diseased heart, which is left in situ. The patient was immunosuppressed over the eight post-transplant months with a multidrug regimen consisting of cyclosporin, steroids, azathioprine and OKT3 (a monoclonal antibody against human CD3 T-cell antigen). The goal of immunosuppression is to allow proper healing while inducing tolerance to the allograft. Cyclosporin (2–8 mg kg–1 day–1) is the cornerstone of all regimens, acting by inhibiting production of interleukin-2 and attenuating cytotoxic Tlymphocytes. Steroids (methylprednisolone 0.15–0.2 mg kg–1 day–1) and azathioprine (2 mg kg–1), when added to cyclosporin, are known to increase cardiac transplant survival to 85–90% at 1 year (Gay, 1995). Additionally, OKT3 was administered at 5 mg day–1, both for immunosuppression and to enable the use of lower steroid dosages, thereby reducing side effects. Although no preoperative therapy for Chagas’ disease was given, nifurtimox (120 mg day–1 for 3 months, followed by 150 mg on alternating days) was administered postoperatively to suppress potential recurrence of Trypanosoma cruzi, with plans to continue the drug indefinitely. The patient was alive and well 6 years after transplant, and remained free of reactivation of Chagas’ disease. Pathology of the explanted heart revealed diffuse endocardial interstitial fibrosis. Lymphocytic, plasmocytic, and eosinophilic infiltrates of heart muscle indicated acute and chronic myocarditis. Again, no parasites were seen microscopically (Blanche et al., 1995).

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2.2. American Trypanosomiasis

Chagas’ disease, also called American trypanosomiasis to differentiate it from its African counterpart, is caused by infection with T. cruzi. This flagellate protozoan is an anthropozoonotic parasite infecting many vertebrates. Triatome or reduvid bugs are the vectors. In 1909, a Brazilian medical student, Carlos Chagas, discovered the etiology of the disease that now bears his name. Human infection starts with an acute phase of high parasitemia lasting several weeks, followed by a long chronic phase of positive serology and low parasitemia. The severity of the disease increases as one moves south from Texas to Brazil. Years after acute infection, 20–30% of patients develop chronic cardiomyopathy, 10% develop intestinal megasyndromes (pathologic dilatation of the esophagus or colon), and a small percentage develop peripheral nerve involvement. The polymorph T. cruzi has an indirect life cycle. The insect vectors, reduvid bugs, are obligate blood feeders measuring 5–45 mm in length. By ingesting blood from infected human or animal reservoirs, they become infected with circulating trypomastigotes. In the bug, these transform into flagellated epimastigotes, 20 µm long, with an anterior kinetoplast near the nucleus. In the midgut of the bug, they divide by binary fission. The daughter epimastigotes migrate to the hindgut to become metacyclic trypomastigotes. Humans are infected when the feeding triatome bug defecates metacyclic trypomastigotes onto the skin near the bite site. Often aided by scratching by the human victim, the trypomastigotes penetrate the slightest break in the cutaneous barrier to infect tissue histiocytes. Here, nests of intracellular oval amastigotes, 3 µm in diameter, multiply by binary fission. Daughter amastigotes form trypomastigotes, which are released upon rupture of the host cell to enter the bloodstream. The trypomastigotes are flagellated, 15–20 µm long, with a large posterior kinetoplast. Especially infective to muscle and nerve cells, they use receptors to attach and enter other host cells to again form pseudocysts of amastigotes. The time from penetration of a trypanomastigote into a host cell to the cell’s eventual rupture is approximately 5 days. The insect vector feeding on the blood of the infected vertebrate host completes the cycle (Magill and Reed, 2000).

2.3. Epidemiology

The majority of cases of human Chagas’ disease are reported from Brazil, but Bolivia has the highest (20%) seropositive rate. American trypanosomiasis is prevalent from the southwestern USA as far south as Argentina and Chile. Pathogenicity of T. cruzi isolates is greater in Brazil than in North or Central America, with megasyndromes (pathologic dilatation) of the gut and

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cardiomyopathy being more common. In Latin America, 16–18 million people are seropositive, or about 4–5% of the population. An estimated 100 000 seropositive persons are currently living in the USA. T. cruzi has been identified in more than 100 mammalian species, but the sylvatic cycle is an important facilitator of human infection by domestic animals such as dogs lying around the house at night, when reduvid bugs feed. Of the many vector species, Triatoma infestans is predominant due to its predilection for living in wall cracks in primitive housing and its preference for human blood. Although direct bug bites account for the majority of cases of Chagas’ disease, additional modes of transmission include contaminated blood transfusions, maternal to fetal infection, accidental ingestion of contaminated food, and laboratory accidents (Magill and Reed, 2000).

2.4. Pathogenesis

The acute phase of human infection is characterized by rapid parasite multiplication inside cells, with amastigotes having a predilection for the heart, brain, and liver. With advancing infection, inflammatory lymphocytes and plasma cells appear, and parasites are harder to find. The heart becomes enlarged and dilated, replete with nests of amastigotes encased in pseudocysts prior to rupture of myofibrils. Chagas’ disease is characterized by one of the most severe forms of myocarditis, and congestive heart failure leads to organ congestion and fluid retention. Focal myocardial inflammation and scarring can also destroy the cardiac conducting system. The brain and meninges can become inflamed, and lesions of the esophagus and gut occur in muscle layers and nerve plexi. Since the reduvid bugs have an affinity for biting the face, an early indication of acute infection is unilateral orbital edema from rubbing bug feces in the eye, known as Romana’s sign. In the chronic phase, the early severe myocarditis can cause Chagas’ cardiomyopathy. The organ becomes thin walled, lacking in tone, with fibrosis of the chambers and dilated valve rings. Electrocardiogram findings of extra ventricular beats and heart block often occur early, with eventual congestive heart failure, left ventricular aneurysms, and pulmonary or systemic emboli from intracardiac clot. Megasyndromes also occur late in the chronic phase consisting of a dilated esophagus or sigmoid colon. The cause is parasite destruction of the submucosal parasympathetic ganglion cells leading to difficulty swallowing (megaesophagus) or constipation (megacolon). Megaesophagus is graded according to degree of esophageal dilatation: grade 1: 10 cm. Pressure at the gastroesophageal junction is increased, with no relaxation upon swallowing as normally occurs. Reflux of gastric secretions with their aspiration into the tracheobronchial tree often

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lead to secondary pneumonia, exacerbating wasting from malnutrition. Megacolon is evident with progressive constipation and abdominal distension. The sigmoid colon may twist, called a volvulus, resulting in a surgical emergency. Fecaloma, or impaction, may also acutely obstruct affected patients (Kirchoff, 1996).

2.5. Diagnosis

Parasites detected in tissues, or the presence of antibodies to T. cruzi in the blood, constitute a positive diagnosis. During the acute phase, fresh smears of unstained blood or buffy coat are examined for motile trypomastigotes. If unsuccessful, polymerase chain reaction (PCR) assays for T. cruzi nuclear and kinetoplast DNA can be tried. Xenodiagnosis, where uninfected laboratorybred reduvid bugs are fed on the patient, may be helpful. A month later, the bug feces should show flagellates if the patient is seropositive. In the chronic phase, serologic tests are preferable since parasitemia is low. Antibody detection is possible with indirect hemagglutination assay (IHA), immunofluorescence test (IF), complement fixation test (CF), and enzymelinked immunosorbent assay (ELISA). In all these methods, a lysate of culture-derived epimastigotes is the source of capture antigen. Negative seroconversion is believed to represent parasitologic cure (Markell et al., 1999b).

2.6. Treatment

The role of heart transplantation in end-stage Chagas’ cardiomyopathy is not well defined. There are relatively few publications regarding this controversial issue (Milei et al., 1992; Bocchi et al., 1994; Blanche et al., 1995; de Carvalho et al., 1996). The fear is that drugs used to prevent rejection, by suppressing the immune system, may lead to recurrence of T. cruzi infection with resultant damage to the allografted heart. The actual operation required to transplant a heart is well established, with centers reporting 1-year survival of over 90% and 5-year survival of over 80%. Generally, cardiac transplantation is limited to patients with terminal heart disease who cannot achieve palliation or prolongation of life with conventional medical or surgical therapy. Active infection has been considered a contraindication because of the probability of dissemination with immunosuppression (Gay, 1995). There may be 75 000 undiagnosed cases of Chagas’ cardiomyopathy in the USA today. Leiby et al. (2000) raised the concern that the emigration of several million people from T. cruzi endemic areas into the USA has caused this increased prevalence. In their study of 11 430 cardiac surgery patients, enzyme immunoassay and radioimmunoprecipitation analysis of postoperative blood

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samples showed six positive tests (0.05%). Overall, 2.7% of Hispanic patients tested positive. Interestingly the diagnosis of Chagas’ disease was not considered preoperatively in any of the six seropositive cases. Leiby et al. (2000) concluded that T. cruzi is an underdiagnosed cause of cardiac disease in North America, especially in regard to those patients born in endemic areas. In patients with Chagas’ cardiomyopathy, heart failure, left ventricular aneurysm, or global dysfunction (weak contraction of the entire heart muscle) are ominous signs. The life expectancy from the start of heart failure is 6–13 months. Therefore, the role of cardiac transplantation in this disease is an issue of great relevance (Milei et al., 1992). Libow et al. (1991) showed that T. cruzi replicates actively both in vitro and in vivo in the presence of immunosuppressive drugs. Therefore, organ transplantation in patients with Chagas’ disease would appear problematic, since the obligatory immunosuppressive agents might trigger a resurgence of the parasite. Nonetheless, the overall experience with kidney transplantation in Chagas’ disease is a positive one (Luders et al., 1992). Similarly, American trypanosomiasis is not considered an absolute contraindication to cardiac transplantation. Bocchi et al. (1994) reported heart transplantation in 18 patients in Brazil, with T. cruzi reactivation in 13 (72%). In another Brazilian study, de Carvalho et al. (1996) found a lower incidence, with parasitemia present in three of ten cases, yet no allograft involvement. Fever, subcutaneous nodules, and myocarditis of the transplanted hearts were common findings. Although all patients in these studies were successfully treated with benzonidazole (5–10 mg kg–1 day–1 for 30–60 days), this agent may have been responsible for increased incidence of lymphoproliferative disease. De Carvalho et al. (1996) presented recommendations not to use prophylactic drug therapy since they believe disease reactivation is not prevented. Blanche et al. (1995), in a USA study, continued nifurtimox therapy (8–10 mg kg–1 day–1) permanently posttransplant with no reactivation. Almeida et al. (1996) confirmed the efficacy of the less toxic anti-gout drug allopurinol in controlling post-transplant Chagas’ parasitemia and myocarditis. In addition, although clearly immunosuppression is needed for the survival of the allografted heart, there is evidence that reducing the dosage of ciclosporin may decrease T. cruzi disease recurrence and improve survival. The authors cited above confirm that heart transplantation is a reasonable option for terminal patients with advanced Chagas’ cardiomyopathy. Owing to small numbers of cases, there are no clear guidelines for postoperative drug prophylaxis against T. cruzi, but there is consensus that lower doses of immunosuppressives enhance survival in these patients. The available literature demonstrates encouraging survival of approximately 75% at 1 year, and up to 65% at 10 years after transplant. Chronic Chagas’ gastrointestinal disease or megasyndrome differs from T. cruzi cardiac involvement in that treatment algorithms are both well studied

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and standardized. Denervation in the gut underlies the pathology, and affects both sympathetic and parasympathetic pathways. A loss of neurons in the submucosal Meissner and myenteric Auerbach plexi occurs, together with preganglionic lesions in vagal innervation. In a study of 1600 patients, de Rezende (1979) found the lifetime risk of esophageal dysfunction in T. cruzi-infected persons is about 30%, megaesophagus usually occurring many years after initial infection. Esophageal dilatation and wall thickening make swallowing difficult. Manometry (pressure readings obtained using a swallowed intraesophageal probe) shows a failure to relax the lower esophageal sphincter, similar to the acquired disease achalasia (a similar but nonparasitic condition, in which a high-pressure lower esophageal zone leads to massive esophageal dilatation). In most cases of Chagas’ megaesophagus, initial treatment is by balloon dilatation of the lower esophageal sphincter. Surgery is reserved for those patients who fail repeated balloon dilatations. Pinotti et al. (1993) found good long-term results in 95% of 722 cases treated with surgery to incise the muscular lower esophageal wall at the narrowed area (cardiomyotomy). Recently this has been accomplished by laparoscopy, thus sparing the patient a major incision with its inherent slow recovery. Laparoscopic myotomy may become the procedure of choice for Chagastic megaesophagus. After the esophagus, the colon is next most often involved, especially the lower left side or rectosigmoid. As aganglionosis progresses, the entire colon can be involved, causing constipation, cramps, and abdominal distension. Early stages can be managed with high-fiber diets and laxatives, and even occasional manual disimpactions. Complications of toxic megacolon and twisting of the dilated sigmoid colon (volvulus) require emergency operations. Failures of conservative therapy may also need surgical treatment. Cutait and Cutait (1991) noted various surgical procedures that have been applied to Chagastic megacolon. All of them include resection of the sigmoid colon as well as a portion of the rectum. If a part of the rectum is not removed, the portion of proximal bowel brought down and sutured to the rectum may become dilated, leading to a recurrence of megacolon.

2.7. Prevention

Currently, in the absence of a vaccine or risk-free drug treatment, Chagas’ disease must be controlled by preventing transmission. Efforts include elimination of domiciliated reduvid bugs in housing, better screening of contaminated blood in blood banks, and closer monitoring of maternal transmission (Magill and Reed, 2000).

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3. ENTAMOEBA HISTOLYTICA 3.1. Case Report

A 44-year-old man presented with a 2-day history of difficulty in breathing. Hospital two-dimensional cardiac echo (ultrasound) revealed fluid in the pericardial sac. Cardiac tamponade (fluid compressing the heart and restricting normal cardiac filling and contractility) warranted emergency operative decompression by pericardiectomy (removal of a portion of the pericardium). Upon opening the pericardial sac, the surgeons found chocolate-colored sterile pus containing Entamoeba histolytica trophozoites. An abdominal computed tomography(CT) scan identified a hypodense liver abscess in the left lobe. The diagnosis of amebic liver abscess with perforation into the pericardium was confirmed by a high serum titer of antiamebic antibodies on hemagglutination testing. The patient was treated with 2 weeks of metronidazole therapy (500 mg intravenously every 6 h), and was subsequently discharged in good condition (Chao et al., 1998).

3.2. Amebiasis

Jackson and Gathiram (2000) defined amebiasis as an infection caused by the ameba E. histolytica, a tissue-invasive pathogen that grows at 37°C in vitro producing quadrinucleated cysts 10–15 µm in diameter. The disease in humans ranges from subclinical to deep tissue invasion. This spectrum includes patients with acute life-threatening amebic dysentery to chronic asymptomatic cystpassers, the latter group comprising some 90% of those infected. Symptomatic or asymptomatic intestinal infection may be followed by the sequelae of tissue invasion: amebic liver abscess, pulmonary amebiasis, amebic pericarditis, ameboma (a granulomatous mass lesion of the cecum or rectosigmoid often mistaken for colon cancer), or stricture of the bowel. Fedor Losch first discovered E. histolytica in 1875, describing motile trophozoites in the stool of a Russian worker with dysentery. Interestingly he did not implicate the amebae as causative agents in the patient’s infection, believing them to be simply facilitators of a primary bacterial inflammatory process. In 1928, Emile Brumpt theorized that E. histolytica consisted of two separate species, E. dispar and E. dysenteriae (Jackson and Gathiram, 2000). E. dispar was nonpathogenic and geographically confined mainly to the temperate zones, whereas E. dysenteriae was responsible for invasive amebiasis and was more prevalent in the tropics. Brumpt’s work was largely ignored over the subsequent 50 years, but its validity is now recognized. It is currently believed that the morphologically identical E. histolytica and E. dispar are two

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distinct species, with the latter being a nonpathogenic gut commensal. Isoenzyme analysis, especially phosphoglucomutase and hexokinase, allows consistent identification of these two species.

3.3. Epidemiology

The life cycle of E. histolytica has three stages: trophozoite, precyst, and cyst. The cysts, whether uninucleated or quadrinucleated, are the infectious agents. They must be orally ingested by the human host through fecal contamination of water, food, or fingers. The cysts pass into the small intestine, where excystation occurs releasing metacystic quadrinucleated amebae. Cytoplasmic division occurs, producing eight metacystic uninucleated trophozoites from each quadrinucleated ameba. The infective trophozoites are the actively growing and multiplying stage and contain pseudopodia. They make their way down the bowel until an area suitable for invasion is found. The cecum is frequently preferred, where the trophozoites rapidly multiply by binary fission, invading the bowel wall using lytic and physical means. The trophozoites may metastasize through the portal system to the liver and other sites. As stool passes to the left colon and rectum, trophozoites are carried along, and eliminate their food vacuoles and other inclusions to become precysts. After the cyst wall thickens, the uninucleated cysts become mature quadrinucleated cysts, which are passed rectally to await transmission to the next host. In instances of profuse diarrhea, trophozoites are shed rectally before encystation can occur. Maturation into cysts outside the body has been described under certain favorable circumstances (Jackson and Gathiram, 2000). Invasive infection and severe disease caused by E. histolytica is more prevalent in the tropics, but amebiasis is found worldwide. Poor sanitation and nutritional deficiencies aid spread. Walsh (1986) estimated that 10% of the world’s population harbors the parasite. Endemic areas include West and South Africa, Mexico, western South America, the Middle East, and Southeast Asia. The 3% figure used to estimate the number of people in the USA who pass amebic cysts or trophozoites in their stools is misleading in the sense that institutionalized patients, homosexuals and Native Americans on reservation land tend to have much higher infectivity. Amebic cysts are killed by boiling and exposure to temperatures over 40°C or under 5°C. The cysts are hardy, however, and are not affected by the chlorine levels present in the water supply. They may remain viable for up to a month in raw sewage and natural water at 4°C. Cysts can live up to 48 h on food products at 20–25°C. Although animals like monkeys, dogs and pigs can serve as animal reservoirs, they have little impact on human transmission. Infection mainly occurs

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through direct person-to-person contact, or through contaminated food or water. In areas where tainted water or sewage is used to grow or freshen vegetables, or where human excrement serves as fertilizer, the ingestion of leafy or root salads becomes hazardous. Although females are infected as frequently as males, invasive amebiasis is much more common in males. Host genetic factors appear to have less influence on infectivity than do socioeconomic ones (Jackson and Gathiram, 2000).

3.4. Pathogenesis

The number of amebae ingested is directly proportional to the frequency and severity of gut lesions. The process by which trophozoites of E. histolytica adhere to the large-bowel mucosa is mediated by a lectin that is inhibited by galactose-N-acetyl-galactosamine. Adherence occurs only under conditions of low oxygen tension, as produced by a normal enteric bacterial flora. Areas of necrosis and ulceration occur as the amebae secrete proteolytic enzymes and cytokines that induce cytolysis of host mucosal cells, aided by phagocytosis (Abd-Alla et al., 1993). Flask-shaped ulcers with undermined edges are typical of amebic lesions of the cecal area, but may be disseminated throughout the colon. The center of the ulcer crater contains gray necrotic material consisting of fibrin, trophozoites and cellular detritus. The amebae can invade the submucosa entering blood vessels. Vessel thrombosis often leads to transmural infarction (death) of entire segments of bowel, accompanied by secondary bacterial infection (Ravdin, 1988). Perforation is a frequent complication of transmural (involving the entire bowel wall) intestinal amebiasis. Presentation is that of a toxic, febrile patient with increasing abdominal pain and rigidity. Abdominal distension from ileus (peritonitis-induced paralysis and swelling of the intestines) is typical. A mass may be palpable, often in the right lower quadrant or cecal area, if the greater omentum has walled off the perforation. Free air may be seen under the hemidiaphragms on upright chest X-ray or on CT scans. Toxic megacolon (pathologic septic colonic dilatation) with rectal bleeding may result, usually after several weeks of cramps, tenesmus (spasm), fever and other evidence of amebic dysentery (Abbas et al., 2000). Amebic appendicitis has been reported. Difficult to differentiate clinically from bacterial acute appendicitis, amebic appendicitis has the gray slough and ulcerations characteristic of lesions of E. histolytica (Demartines et al., 1995). Amebomas are firm inflammatory masses of the bowel wall found in about 1% of cases of colonic amebiasis. These granulomas are most common in the cecal area, and consist of fibroblasts, collagen, inflammatory cells, eosinophils, and scant amebae. When found during colonoscopy or barium enema, amebomas can be mistaken for colon cancer. Following an attack of transmural

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colitis, the inflamed bowel may heal with a fibrous amebic stricture or narrowing. The rectum and sigmoid colon are most often involved, and signs of high- or low-grade intestinal obstruction may result in constipation and abdominal pain and distension (Adams and MacLeod, 1977). Amebic liver abscess, caused by trophozoites being carried to the liver via the portal venous circulation, is five times more frequent in the right hepatic lobe than the left. Liquefactive necrosis yields gray or yellow liquid material in the center, with a fibrinous periphery where trophozoites may be found. Twenty per cent of cases of amebic liver abscess will have E. histolytica found on microscopic stool examinations, but 70% will have positive stool cultures. Patients present with right upper quadrant abdominal and right chest pain, elevated right hemidiaphragm, pleuritic pain (on taking a deep breath), chills, and fever (Akgun et al., 1999). Liver abscesses, especially those in the left lobe, may extend through the diaphragm into the pleural space. When the lung and bronchial tree are involved, the patient may cough up copious amounts of abscess material. Amebic pericarditis, as described in our case report above, also results from direct extension from a hepatic amebic abscess (Mbaye et al., 1998).

3.5. Diagnosis

The finding of trophozoites of E. histolytica in the stool is consistent with invasive amebiasis. Although E. histolytica cysts can remain viable outside the body for up to a month in raw sewage and natural surface water, trophozoites usually disappear within 30 min of passage. Commercial kits are available which contain fixatives to preserve cysts and trophozoites until they can be examined microscopically with wet mounts or iodine-stained smears. Since microscopy does not differentiate between E. histolytica and the nonpathogenic commensal E. dispar, further studies using isoenzyme electrophoresis or monoclonal antibodies in antigen-capture ELISA may be useful. Between 70 and 90% of infections are diagnosed when stool specimens are collected over 3 days or longer. Laboratories experienced in stool culture can improve this diagnostic yield. If invasive amebiasis is still suspected after negative stool examinations, serologic testing can be performed. Serum antibody detection is possible using IF, IHA, the latex agglutination test, the gel diffusion precipitation test, counterimmunoelectrophoresis, and ELISA. Seropositivity is seen in 96% of patients with amebic liver abscess, and 85% of patients with amebic dysentery. Seronegativity virtually rules out an amebic origin for the patient’s illness. Antibodies may be found in the sera of patients 6 months to 3 years following acute infections with E. histolytica (Markell et al., 1999e).

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3.6. Treatment

Before discussing the surgical treatment of invasive amebiasis, it must be mentioned that antibiotic therapy alone is generally curative in the majority of patients with invasive colonic and extracolonic disease. Metronidazole is given 500 mg intravenously every 6 h for 5–10 days or orally at 750 mg thrice daily, also for 5–10 days. Although curative in 90% of cases of invasive E. histolytica infection, metronidazole can cause nausea, headache, metallic taste, confusion, and ataxia (unsteadiness). Most seriously, psychosis or seizures mandate stopping the drug. With severe invasive amebiasis, dehydroemetine, at 1–1.5 mg kg–1 day–1 intramuscularly for 5 days, may be given together with metronidazole for a more rapid antiamebic effect. Following treatment of invasive disease, a luminal agent is given to eradicate intestinal presence of the organism. Paromomycin (30 mg kg–1 day–1 orally for 7 days) or iodoquinol (650 mg orally thrice daily for 20 days) may be used (Li and Stanley, 1996). Further treatment is indicated for the management of complications of invasive amebiasis. In a patient with invasive amebic colitis and suspected transmural involvement, antibiotics effective against enteric bacteria should be added to the antiamebic regimen. If metronidazole therapy has been started, a quinolone such as levaquin can be administered (500 mg intravenously every 24 h) to provide this coverage. Nasogastric suction, along with liberal intravenous hydration, is also required. Although small perforations of the colon that are walled off early by the greater omentum may occasionally be treated conservatively without surgery, caution is necessary when utilizing this approach. Takahashi et al. (1997), in a study of 55 Mexican patients, demonstrated that the nonsurgical treatment of fulminant amebic colitis increased mortality. Indications for emergency surgery include: toxic megacolon, perforation of amebic intestinal ulcers, gangrene of segments of colon, massive hemorrhage from transmurally involved large bowel, and intra-abdominal abscess from a colonic leak (Dautov, 1997). The goal of surgical intervention is to treat infection and stop blood loss. This generally means resection of all of the non-viable transmurally inflamed or perforated colon. No attempt at an anastomosis (connection) between the ends of resected bowel is advisable at the time of emergency operation owing to the likelihood of poor healing in the presence of inflammation and infection. It may be necessary to excise the entire abdominal colon, leaving the ileum as a skin-level stoma (a small-bowel version of a colostomy) and a closed rectal pouch in the pelvis. Once the patient has recovered, the small bowel can be reconnected to the rectum using stapling instruments in a secondary elective procedure 3–6 months later (Abbas et al., 2000) The development of amebomas or late fibrous strictures of the colon are occasional indications for elective operations, usually to differentiate them from colon cancer or to treat varying degrees of obstruction. Unlike the

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surgical treatment of toxic transmural colitis, a primary anastomosis can generally be performed in the right or left colon in these situations without the need for a colostomy (Ito et al., 2000). Amebic liver abscess generally responds to metronidazole therapy alone. It has been reported by Li and Stanley (1996) that amebic liver abscesses resolve slowly. Even successfully treated ones may transiently enlarge over the first few weeks before regressing. Ultrasound is commonly used to follow resolution of amebic liver abscesses, and most will have disappeared within 6 months of successful treatment. Ten per cent of patients will have abnormal ultrasound findings more than a year post-therapy. Jackson and Gathiram (2000) reported that needle drainage under ultrasound radiologic guidance is appropriate for abscesses over 10 cm in diameter, which may potentially rupture. Left lobe liver abscesses should also be aspirated to prevent rupture into the pericardium. Although the majority of amebic liver abscesses under 10 cm usually respond to drug therapy alone, if pain and fever persist past 3–5 days, aspiration should be considered as an additional modality. Should the liver abscess extend into the chest, fluid in the pleural space should also be aspirated or drained with a chest tube. Pham et al. (1996) reported a study of 1512 patients with amebic liver abscess in Vietnam. Ultrasound-guided aspiration was performed in 1289 patients, with 24.9% requiring a second aspiration, and 9.4% a third. They recommended aspiration and antiamebic treatment as the standard therapy for abscesses 4–17 cm in size. Abscesses smaller than 4 cm usually resolved with drug treatment alone. Abscesses greater than 17 cm responded poorly to aspiration and were best dealt with surgically. Surgical drainage is appropriate for a large left lobe abscess that is inaccessible to percutaneous needle drainage. Abscesses complicated by rupture into the peritoneal cavity also require laparotomy (operative exploration of the peritoneal cavity) and surgical drainage, as may abscesses not responding to antiamebic therapy and needle aspiration. Moazam and Nazir (1998) noted that, unlike adults, children with complicated amebic liver abscess are best managed non-surgically. In all 48 of their patients, aged 3 weeks to 14.5 years, none required open surgical drainage. Metronidazole and ultrasound-guided needle aspiration obviated the need for surgery, even in malnourished children with ruptured amebic liver abscess. Mondragon-Sanchez et al. (1995) reported 50 consecutive patients with amebic liver abscess in Mexico City. Medical (drug) treatment alone controlled the disease in 24 patients, percutaneous drainage was necessary in 15, and surgery was needed to treat complications in 11. They concluded that multimodality treatment carried a low (2%) mortality rate. This is consistent with Akgun et al. (1999), who noted a 2.2% mortality rate for their 44 patients with amebic liver abscess similarly treated.

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3.7. Prevention

The prevention of amebic infection starts with the avoidance of fecally contaminated food and water. Visitors to endemic areas should eat only cooked foods and peeled fruit, and drink only bottled water. A prototype vaccine has raised hope that invasive amebiasis may some day be eradicated (Zhang et al., 1994). Recombinant amebic antigens have been bioengineered: 170-kDa galactose-inhibitable adhesin, serine-rich E. histolytica protein (SREHP), and 29-kDa antigen. Efficacy, in terms of protection, has been shown in experimental animal studies for a combination of SREHP and the 170-kDa antigen. Cieslak et al. (1993) reported progress in developing an oral combined typhoid fever–amebiasis vaccine, in which recombinant SREHP was inserted into avirulent Salmonella strains. Zhang and Stanley. (1995) induced IgA and IgG antiamebic antibodies in mice through oral vaccination with SREHP fused to a cholera toxin B subunit.

4. LEISHMANIA DONOVANI 4.1. Case Report

A 30-year-old Japanese woman was admitted to hospital with a fever of unknown origin. At age 28, she had been to Bangalore, India to do graduate research and had spent 2.5 years there. Soon after returning, she began to have chills and fevers up to 40°C. She had a workup at New England Medical Center in Boston, Massachusetts. Blood tests showed a mild pancytopenia (diminished erythrocytes, leukocytes and platelets), elevated liver enzymes (lactate dehydrogenase, aspartate aminotransferase, alanine aminotransferase), and an elevated sedimentation rate (a nonspecific test for inflammation or infection). Physical examination revealed hepatosplenomegaly (enlarged liver and spleen), a finding confirmed on abdominal CT scan. Although an infectious etiology was suspected due to her recent stay in India, blood cultures and smears were negative. Liver biopsy and bone marrow aspiration showed infiltration of monocytes and plasma cells. Malignant lymphoma was suspected, and she underwent a splenectomy in an attempt to diagnose the condition. No evidence of malignancy was found on pathologic examination of the spleen, and steroids (methylprednisolone 120 mg day–1) were started. Her fever went down and she returned to Japan. She was admitted to the Tokyo Medical and Dental School of Medicine for further treatment. The liver remained enlarged; blood cultures were again negative; and blood smears showed no evidence of malarial or other parasites. With no proof of malignant lymphoma, steroids were stopped. Blood tests then

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showed progressive leukopenia (low white cell counts) and predominance of monocytes. A polyclonal hypergammaglobulinemia was also noted. A bone marrow biopsy was repeated, and smears finally showed monocytes filled with the rounded amastigotes of Leishmania donovani, each parasite with the characteristic nucleus and kinetoplast. The diagnosis of visceral leishmaniasis was confirmed by culturing peripheral blood in a medium of human leukocytes. Three days later, promastigotes were found moving actively in the medium. Antileishmanial therapy was started with pentamidine isetionate intravenously at 200 mg day–1. Three days later the fever broke. A month later, the white cell count was up to normal and the monocytosis disappeared. Pentavalent sodium stibogluconate was given intravenously at 500 mg day–1 for 20 days to prevent relapse. Bone marrow reaspiration was then negative for amastigotes, and serum gammaglobulin and liver function tests improved. The patient was discharged on no medications, and remained relapse-free at 2 years follow-up (Kawakami et al., 1996).

4.2. Leishmaniasis

Magill (2000) defined leishmaniasis as a group of diverse clinical syndromes caused by the protozoan parasites of the genus Leishmania. Vectors include sandflies from the genera Phlebotomus and Lutzomyia. In 1885, parasites from lesions of cutaneous leishmaniasis were first described by Cunningham. In 1903, Leishman found the causative organisms in the spleen of a soldier who was stationed in Dum Dum, India. He used the name ‘Dumdum fever’, but the disease also became known as kala-azar, Hindi for black sickness.

4.3. Epidemiology

Visceral leishmaniasis or kala-azar is caused by L. donovani. Infection is characterized by fever, weight loss, enlargement of the liver and spleen, pancytopenia, and hypergammaglobulinemia. Old World visceral leishmaniasis is transmitted by a sandfly vector from the genus Phlebotomus. Zijlstra et al. (1995) described major epidemic areas in southern Sudan and the Ganges river basin in India. Endemic areas also include the Mediterranean basin, the Middle East, Central Asia, and northwest China. New World visceral leishmaniasis is spread by a sandfly vector from the genus Lutzomyia. Abramson et al. (1995) noted American kala-azar in northeastern Brazil, Argentina, Bolivia, Colombia, Venezuela, Central America and Mexico. The worldwide incidence of visceral leishmaniasis is estimated to be 100 000 new cases annually.

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Cutaneous leishmaniasis infection is identified by nodular and ulcerative skin lesions. Similar to the visceral form, it is transmitted by sandfly vectors, and is prevalent in both the Old and New World geographic distributions. Human disease is a rural zoonosis. Inhabitants of villages near infected rodents have prevalence rates approaching 100%. Tourists, military personnel, hunters, and others entering endemic ecosystems are at risk for infection (Samady and Schwartz, 1997). The life cycle of Leishmania parasites starts when promastigotes in the proboscis of a female sandfly are introduced through the skin into a vertebrate host during blood feeding. Promastigotes measure 15–20 µm in length and have a single free flagellum. A large central nucleus and a kinetoplast are present. The promastigotes invade the reticuloendothelial system, and transform intracellularly into amastigotes. Amastigotes are round or oval, measure 2–5 µm in diameter, and contain a large nucleus and kinetoplast. They multiply within phagolysosomes by binary fission, and invade further reticuloendothelial cells. When sandflies feed on infected vertebrates, parasitized cells are ingested, and the amastigotes transform into promastigotes. The promastigotes multiply in the gut of the vector and migrate to the proboscis to complete the life cycle (Magill, 2000). Reservoir hosts include dogs, foxes and rodents such as rats and gerbils. The sandfly vectors are 1.5–2.5 mm in size, have V-shaped wings, and move with a characteristic hopping motion. Sandfly saliva has a strong vasodilatory peptide known as maxadilin, which correlates with the course of the disease in humans. Warburg et al. (1994) noted that Costa Rican sandflies have small amounts of salivary maxadilin. There, visceral leishmaniasis is rare, but cutaneous leishmaniasis is common. Costa Rican sandfly bites induce little erythema (redness), but enhance cutaneous proliferation of parasites. In contrast, Brazilian sandflies have high levels of salivary maxadilin. In Brazil, visceral leishmaniasis is prevalent, but the cutaneous form rare. Sandfly bites in Brazil produce marked erythema, but cutaneous infection is not facilitated.

4.4. Pathogenesis

Berman (1997) described the pathogenesis of leishmaniasis. For the visceral form, promastigotes inoculated by the sandfly bite enter reticuloendothelial cells, where multiplication takes place. A granuloma forms at the bite site, filled with amastigotes and inflammatory cells. Parasites spread first to lymph nodes, and then by hematogenous pathways to the liver, spleen and bone marrow. The subsequent course can be a granulomatous-cell-mediated immunity type, which results in mild, self-limited disease, or a more virulent course, leading to the visceral leishmaniasis syndrome. This latter group of

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patients develops splenomegaly from hyperplasia of reticuloendothelial cells filled with Leishmania. The liver is also enlarged, with Küpfer cells replete with amastigotes. Macrophages containing parasites are found in the bone marrow. The patient with visceral leishmaniasis is weak, febrile, emaciated, and has hepatosplenomegaly and prominent lymph nodes. Alvar et al. (1997) noted the association between human immunodeficiency virus (HIV) infection and leishmaniasis. Visceral leishmaniasis is a significant opportunistic infection in AIDS patients when CD4 T-lymphocyte counts fall below 50 cells mm–3. Involvement of the gastrointestinal tract is more common in these patients, with macrophages containing parasites being found from esophagus to rectum. High parasite burdens are also seen, often with negative serologic tests making diagnosis difficult. Treatment must not only be focused on eradicating the Leishmania, but also on increasing the CD4 T-lymphocyte count with highly active antiretroviral therapy (HAART). Visceral leishmaniasis is also found as an opportunistic infection with renal and heart transplantation, as well as in immunosuppressed patients undergoing cancer chemotherapy or chronic corticosteroid use. Ramesh and Mukherjee (1995) reviewed post-kala-azar dermal leishmaniasis, and noted that it was first discovered in India, where up to 20% of patients with visceral leishmaniasis developed the characteristic nodules around the mouth, face, trunk, and extremities. In Sudan, the incidence is over 50%. The skin lesions appear a few months to 10 years after apparent successful treatment of kala-azar. Histologically, the lesions are granulomas containing L. donovani and inflammatory cells. The skin lesions may persist for up to 20 years, and afflicted patients pose a public health problem as persistent reservoirs of infection. Magill (2000) described cutaneous leishmaniasis as nodular and ulcerative skin lesions caused by L. tropica, L. major, L. aethiopica and L. infantum in the Old World, and L. mexicana, and L. braziliensis in the New World. Some colorful names for cutaneous leishmaniasis include: oriental sore, Baghdad boil, Dehli boil, aleppo evil, chiclero ulcer, forest yaws, and bay sore. The condition results from a cell-mediated response at the inoculation site, where parasites enter macrophages. Lymphocytes, mononuclear cells and plasma cells surround the macrophages in a granulomatous reaction. Once the parasite count diminishes, epithelioid and giant cells appear. Tissue necrosis occurs next, and the lesions eventually heal by fibrosis. The incubation period for cutaneous leishmaniasis is 2–8 weeks, but may be up to 3 years in rare cases. Multiple lesions are common, and more than 100 may be seen on the same patient. Lesions are most commonly found on exposed skin surfaces: face, hands, arms, and legs. Lymphatic spread, seen with L. major, is evidenced by a linear array of skin nodules leading to groups of enlarged lymph nodes.

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Diffuse cutaneous leishmaniasis, a result of L. aethiopica, is a cell-mediated immune response to the parasites, leading to multiple, large plaque-like nodules covering the face, nose, limbs, and buttocks. The appearance has been likened to leprosy, and coexisting visceral disease is uncommon. Bomfin et al. (1996) noted low interferon gamma (IFN-γ) and interleukin-10 expression, explaining the inability of diffuse cutaneous leishmaniasis patients to mount an effective anti-Leishmania immune response that would lead to clinical improvement. Leishmaniasis recidivans, found primarily in Iraq and Iran, is another unusual chronic form of cutaneous leishmaniasis, which can persist for 20–40 years, with lesions beginning on the face and progressing relentlessly (Grevelink and Lerner, 1996). Three per cent of persons infected with L. braziliensis develop mucosal disease. Mucocutaneous leishmaniasis is more prevalent in the southern latitudes, often appearing months to years after the initial cutaneous infection. Progressive mutilating destruction of the nasal septum, followed by the palate, lips, pharynx, and larynx, can cause death from aspiration or malnutrition. Susceptibility to mucosal disease has been correlated with regulatory polymorphisms leading to high levels of tumor necrosis factor alpha (TNF-α) in Venezuelan patients (Marsden, 1986).

4.5. Diagnosis

In endemic areas, visceral leishmaniasis is reliably diagnosed on clinical grounds, and confirmed by response to antileishmanial therapy. Parasitologic diagnosis is always preferable, especially if the clinical picture is less certain. Splenic aspiration is the most definitive method of confirming the diagnosis. Chulay and Bryceson (1983) reported 3000 splenic aspirates in 400 cases, with three patients dying in shock within 24 h of aspiration, presumably from splenic laceration from the needle. Another four patients had intraabdominal bleeding which resolved spontaneously. Because of the risk of significant morbidity and mortality, splenic aspiration is not recommended for the soft spleen of acute disease, or for the patient with abnormal clotting parameters or low platelet count (