Parasites of North American Freshwater Fishes [Second Edition] 9781501735059

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Library of Congress Cataloging-in-Publication Data Hoffman, Glenn L. Parasites of North American freshwater fishes / Glenn L. Hoffman ; with a foreword by Ernest H. Williams, Jr. — 2nd ed. p. cm. Includes bibliographical references (p. ) and index. ISBN 0-8014-3409-2 (cloth : alk. paper) 1. Freshwater fishes—Parasites—North America. I. Title. SH175.H6 1998 571.9'991776—dc21 98-49407 Copyright © 1967 by The Regents of the University of California Copyright © 1999 by Cornell University All rights reserved. Except for brief quotations in a review, this book, or parts thereof, must not be reproduced in any form with¬ out permission in writing from the publisher. For information, address Cornell University Press, Sage House, 512 East State Street, Ithaca, New York 14850. First edition published 1967 by University of California Press. Second edition published 1999 by Cornell University Press. Printed in the United States of America. Color plates printed in Hong Kong. Cornell University Press strives to use environmentally respon¬ sible suppliers and materials to the fullest extent possible in the publishing of its books. Such materials include vegetable-based, low-VOC inks and acid-free papers that are recycled, totally chlorine-free, or partly composed of nonwood fibers. Cloth printing

10

987654321

This book is dedicated to the memory of My wife, Carolyn Wilson Hoffman, who typed and edited the manuscript Our son, James Wilson Hoffman, who at the age of 22 was killed by a drunk driver Professor Luther Owen Nolf, my mentor at the University of Iowa during 1942 and 1946-1950 Dr. Stanislaus F. Snieszko, my director, colleague, and friend from 1958 to 1972 at the Eastern Fish Disease Laboratory, U.S. Department of the Interior, Kearneysville, West Virginia and Ernest J. and Viola M. Hoffman, my parents, who were my role models

Contents Foreword by Ernest H. Williams Jr.

ix

Preface to the Second Edition

xiii

Preface to the First Edition

xv

Abbreviations

xvii

Phylum Nemathelminthes: Class Nematoda

254

Phylum Acanthocephala

285

Phylum Annelida: Class Hirudinea (Leeches)

301

Phylum Arthropoda: Class Crustacea: Subclass Branchiura, Orders Copepoda and Isopoda

310

Introduction

1

Public Health Aspects of Fish Parasites

4

Methods

6

Miscellaneous Parasites and Other Pathogens, Including Turbellaria

332

Some North American Fish Parasites, Listed by Location in the Fish

9

Fish Parasites Found in or on Other Animals

339

Predators

340

Fish and Parasite Checklist

341

Miscellaneous Fishes: Parasites of Ornamental, Tropical, and Brackish Water to Marine Fishes

403

Chemotherapy and Prophylaxis of Parasitic Diseases of Fishes

409

Brief Descriptions of the Groups of Fish Parasites

12

Algae and Fungi (Kingdoms Protista and Fungi)

14

Subkingdom Protozoa (Kingdom Protista)

21

Phylum Coelenterata (Cnidaria)

92

Phylum Platyhelminthes: Class Monogenea

95

Phylum Platyhelminthes: Class Trematoda (Digenea): Adults

1 39

Phylum Platyhelminthes: Class Trematoda (Digenea): Larval Forms

185

Phylum Platyhelminthes: Class Cestoidea

21 7

References

415

Glossary of Fish Parasitology Terms

511

Index

527

The color plates follow page 284.

VII

Foreword I am deeply honored to present this revision of the "bible" of freshwater fish parasitology. In a way, I rep¬ resent those who used the original 1967 edition as grad¬ uate students, as professors for a text in their own courses, and as authors for published articles in the field. All of us have referred to it thousands of times (my copy of the first edition is spineless and threadbare) and have eagerly awaited this second edition. We express our gratitude to the author for this wonderfully useful text and reference book. This volume was and is a remarkable accomplishment. The first edition spawned and inspired countless stud¬ ies. Parasites of North American Freshwater Fishes has been and will continue to be a major force advancing fresh¬ water fish parasitology. This second edition includes information from the lit¬ erature through 1992, a 25-year period since the first edi¬ tion. Publication was slightly delayed due to editorial changes, the necessity of obtaining additional financial support, and the medical problems of the author. The changes improved the volume, outside support was obtained, and the author's indomitable spirit overcame any physical frailties. It was well worth the wait. A few years of delay is insignificant for an important guidebook that will be used for decades to come. The revision spans a period of high research pro¬ ductivity and great expansion of our knowledge in the study of freshwater fish parasites. Dr. Hoffman has melded this often fractured, contradictory, and widely scattered information and, with his unique breadth of knowledge and experience, has clarified and syn¬ thesized it into an outstanding guidebook. Many new parasites of North American freshwater fishes have been found and described since the first edition. A new edition was essential to keep pace with all these changes.

This new information combined with current meth¬ ods to study parasites (electron microscopy, molecular biology) and advanced data analysis (cladistic analysis, new computer statistical analysis) are effecting monu¬ mental changes in the higher classification of many parasites. Current usage employs a somewhat awkward compromise between the "traditional" and "new" par¬ asite taxonomic treatments, such as I followed in three recent fish parasite books (Bunkley-Williams and Williams, 1994, 1995; Williams and Bunkley-Williams, 1996). Dr. Hoffman's book lingers with the traditional categories to maintain a continuity in the classifica¬ tions that have been relatively stable for more than 100 years. This preserves the historic nomenclature and allows students to more easily access the majority of fish parasite literature written before the advent of new tax¬ onomic treatments and by those scientists who do not follow these systems. Everyone knows the old classifi¬ cations, and just as one is reluctant to discard comfort¬ able old shoes, there is still some resistance to change. For example, the concepts of "fungi" and "protozoa" are still important, even if fragments of both of these for¬ mer categories have been dispersed as separate phyla in different kingdoms. The drastic new arrangement of flatworms is probably correct but has not stabilized suf¬ ficiently to be accepted by everyone. It remains in flux as new discoveries are made and is also clumsy and overly fragmented with an incredible maze of unfamil¬ iar subgroupings. The exact taxonomic status of crus¬ taceans is in question. Tongueworms (Pentastomida) may soon be grouped with fish lice (Branchiura), and myxozoans with sea jellies (Cnidaria). The traditional cat¬ egories are still useful, however. Dr. Hoffman uses these older and more familiar categories to group similar¬ appearing organisms together for ease of identification, explanation, and comparison, in ways that are more IX

X

Foreword

Table 1. Higher Classification of the Parasites of North American Freshwater Fishes Contemporary or Future Classification Kingdom Prokaryotae—bacteria Phylum Cyanobacteria—blue-green algae/bacteria Kingdom Protista—one-celled organisms Subkingdom Protozoa—animal-like protistans Phylum Sarcomastigophora—amoebas and flagellates Phylum Apicomplexa—coccidians and gregarines Phylum Ciliophora—ciliates, suctorans, etc. Phylum Cilioshoraciliates, suctorans, etc. Phylum Chlorophyta—green algae Phylum Oomycota—saprolegnia fungus and allies Kingdom Fungi—fungi Phylum Microsporida(?)—microsporidians Phylum Zygomycota Phylum Deuteromycota Kingdom undetermined—Ichthyophonus fungus and allies Kingdom Animalia—multicellular animals Phylum Cmdaria—sea jellies, corals, and allies Class Myxozoa—myxosporidians Class Hydrozoa—sturgeon hydroid, etc. Phylum Platyhelminthes—flatworms Superclass Rhabditophora—fish-associated turbellarians Superclass Cercomeria Class Udonellidea—ectoparasitic worm Class Cercomeridea Subclass Trematoda Infraclass Aspidobothrea—soleworms Infraclass Digenea—flukes Superfamily Didymozoidea—tissue flukes Subclass Cercomeromorphae Infraclass Monogenea—gillworms — Order Dactylogyridea—simple gillworms Order Gyrodactylidea—live-bearing gillworms Order Capsalidea—capsalids Order Mazocraeidea—polys Infraclass Cestodaria Cohort Cestoda—tapeworms Subcohort Amphilinidea Subcohort Eucestoda Phylum Nematoda—roundworms Phylum Nematomorpha—horsehair worms Phylum Acanthocephala—spiny-headed worms Phylum Annelida—segmented worms Class Hirudinida—leaches Subclass Acanthobdellida—primitive leech Subclass Hirudinea—true leech Phylum Arthropoda Subphylum Crustacea—crustaceans Class Maxillopoda Subclass Copepoda—copepods Subclass Branchiura—fish lice and tongueworms

Class Malacostraca Order Isopoda—isopods Subphylum Chelicerata Class Arachnids—spiders and allies Order Acari—mites and ticks Phylum Mollusca—sea shells and allies Class Bivalvia—pelecypods or bivalves Order Unionida—freshwater mussels Order Veneroida—fingernail & peaclams Phylum Vertebrata—vertebrates Class Agnatha—jawless fishes Order Petromyzontiformes—lamphreys

Traditional Classification

Taxonomy of Group

—

Kingdom Protista —

Phylum Protozoa Phyla Rhizopoda & Mastigophora Group Sporozoa

Lorn and Dykova 1992

— — —

Kingdom Fungi — Phylum Protozoa — Deuteromycetes Kingdom Fungi or higher taxon unknown — Phylum Coelemterata Phylum Myxospora (Protozoa) — — Turbellaria

Rand 1996 Rand 1996 not established

Ragan et al. 1996 not established

not established Brooks and McLennan 1993

Class Monogenea Class Trematoda Subclass Aspidocotylea Subclass Digenea Subclass Didymozoidea Class Monogenea Subclass Monopisthocotylea Order Dactylogyrida Family Gyrodactylidae Family Capsalidae Subclass Polyopisthocotylida — Class Cestoidea Subclass Cestoidaria Subclass Eucestoda Phylum Nemathelminthes —

(status in question)

Khalil et al. 1994

Anderson et al. 1974-1983 Amin 1985d

—

Class Hirudinea Order Acanthobdella — —

Class Crustacea — Order Copepoda — Phylum Pentastomida Pentastomid larvae

Bowman and Abele 1982

not established

— —

Arachnida — —

Turgeon et al. 1998

—

Glochidia Sphaeriid clams —

— —

Robins et al. 1991 Hardistyand Potter 1971

Foreword

intelligible to a popular audience. For example, microsporidians superficially look like myxosporidians, despite the fact that one is apparently a fungus and the other a sea jelly. Dr. Hoffman used the traditional, better known tax¬ onomic treatments to produce a reference tool that would be more user friendly and enduring. Many of the new, highly technical classifications are intelligible only to specialized taxonomists and will probably soon change. The contemporary, if often controversial, higher taxonomy is presented in Table 1. Traditional categories or groups used in this book can be found in the center column and common group names in the left column. The right column cites references containing more com¬ plete new classifications of most groups. Some lack a uni¬ fied reference appropriate to fish parasites. For those unfamiliar with classification, Margulis and Schwartz's Five Kingdoms (1988) is a useful and entertaining introduction and a popular guide to the higher classification of life on Earth, although it is a bit dated as their five kingdoms now number at least seven. Dr. Hoffman's distinguished career spans over 50 years. He became the leading expert on the parasites of North American freshwater fishes during a series of research appointments. He received a B.A. degree from the University of Iowa in 1942 and then served as Lieu¬ tenant and Laboratory Director for the U.S. Army San¬ itary Corps until 1946. For 2 months of this service he worked at Dr. Jonas Salk's virus research laboratory in Munich. He received his Ph.D. degree in 1950 from the University of Iowa and became an Assistant Professor at the University of North Dakota Medical School Micro¬ biology Department, where he received an Outstanding Teacher award. In 1958, he was appointed as a Research Parasitologist at the National Fish Health Research Lab¬ oratory (then called the Eastern Fish Disease Laboratory) in Leetown, West Virginia. In 1974, he transferred to the U.S. Lish and Wildlife Service Fish Farming Experimen¬ tal Station at Stuttgart, Arkansas. He retired from the Fish and Wildlife Service in late 1985, but his fish parasitol¬ ogy work continues (Overstreet and Meyer 1986). The breadth and influence of his publications, which include more than 200 scientific articles and four books, have been profound. He has been instrumental in pro¬ ducing English translations of many parasite books and articles. He has served or continues to serve on various scientific committees and the editorial boards of nine scientific journals. His original research involved a variety of fish para¬ sites and diseases but focused on flukes and their life cycles, particularly those that encysted as metacercaria in fishes. By 1959, whirling disease had become a major threat to salmonid culture in the United States, and naturally, the federal government called on its leading expert to save the day. From that time to 1974, Dr. Hoff¬ man conducted extensive investigations detailing the biology and control of this disease. As other diseases threatened fish culture in the United States, he investi¬ gated and provided the information to solve these prob¬

XI

lems. The Asian tapeworm's threat to the U.S. baitfish industry is a prominent example of this work. Many of these dangerous epizootics invaded from overseas, and Dr. Hoffman thus became an expert on exotic diseases and the prevention of introductions. His review articles on parasites of exotic fishes, strigeid flukes, and whirling disease, on the treatment and prevention of disease, and on other fish diseases have been widely used by sci¬ entists. Meanwhile his numerous popular circulars and Fish Health Leaflets have explained fish diseases to the general public. Dr. Hoffman has often freely given his expert advice and encouragement and transmitted his enthusiasm about parasites. He has presented his work at more than 100 symposia and meetings around the world and has conducted collaborative research with numerous fish dis¬ ease professionals in the United States and more than 20 other countries. The fish health training courses he con¬ ducted and his many other endeavors have helped develop, mold, and inspire a generation of fish health specialists and fish parasitologists worldwide. Many of the existing programs in these fields were either created or fomented by his actions. Dr. Hoffman has received numerous awards and hon¬ ors. He was one of the first honorees enshrined in the National Lish Culture Hall of Lame in Spearfish, South Dakota, and he also received Distinguished Service Awards from the Wildlife Disease Association (1974), the American Lisheries Society (AES) Lish Health Section (1982), and the AFS Fish Culture Section (1985). The fol¬ lowing parasites were named in his honor: Biacetabulum hoffmani Mackiewicz, 1972 (tapeworm); Cercaria hoffmanensis Brooks, 1948 (fluke); Eimeria hoffmani Molnar and Hanek, 1974 (coccidian); Gyrodactylus hoffmani Wellborn and Rogers, 1967 (gillworm); Myxidium hoffmani Jayasri, 1981 (myxosporidian); Myxosoma hoffmani Meglitsch, 1963 (myxosporidian); Neascus hoffmani Pandey, 1973 (fluke); Thqcamoeba hoffmani Sawyer, Hnath, and Conrad, 1974 (amoeba); Trichodina hoff¬ mani Wellborn, 1967 (ciliate). We should be grateful that the always generous and genteel Dr. Hoffman took time from his retirement to update this important book. Everyone interested in fishes, from scientists to aquarium hobbyists, must have a copy. Fishery laboratories will need both shelf and field copies of this hands-on text. Ernest H. Williams Jr.

Selected additional literature: R. C. Anderson et al. (1974-1983), Bowman and Abele (1982), Brooks and McLennan (1993), Bunkley-Williams and Williams (1994-1995), Hardisty and Potter (1971), Khalil et al. (1994), Overstreet and Meyer (1986), Ragan et al. (1996), Rand (1996), Turgeon et al. (1988), E. H. Williams and Bunkley-Williams (1996).

Preface to the Second Edition To the best of my knowledge, this edition reviews and cites practically all the literature on North American freshwater fish parasites as well as related foreign species, from the time of Linnaeus through 1992. It should be useful to anyone interested in the subject. I have, per¬ haps uniquely, included literature on foreign freshwa¬ ter fish parasites that, because of related hosts, are likely to occur in North America. This edition may therefore be helpful in foreign countries, particularly in Europe, because of related hosts and parasites. Much credit for this edition belongs to the many people who have helped me, directly or indirectly, since the first edition. T. Sawyer, Royal Oak, MD, prepared the section on free-living amoebae, some of which may be fish pathogens. V. Sprague, University of Maryland, Natural Resources Institute, assisted me with the genera Dermocystidium and Ichthyophonus. J. Mackiewicz, State Uni¬ versity of New York, College of Education, Albany, provided the caryophyllaeid cestode key. D. J. Klemm, EPA, Cincinnati, OH, and the late M. C. Meyer, University of Maine, gave me much help with the leeches. E. H. Williams and L. Bunkley-Williams, University of Puerto Rico, Mayagiiez, P. R., prepared most of the section on isopods. R. Lichtenfels, USDA Parasitology Lab, Beltsville, MD, kindly loaned specimens for comparison. Over the past thirty years, I have been indebted to J. Lom, Ceske Budejovice, Czech Republic, for much guid¬ ance and information on the protozoa, and to the great Russian parasitologist, O. N. Bauer, St. Petersburg, for help with circumpolar and cosmopolitan fish parasites. C. D. Becker, M. Beverley-Burton (Canada), and W. Yasutake have aided me with counsel, collaboration, and specimens that have helped, directly or indirectly, in the preparation of this edition.

During my involvement with fish disease diagnosis as a teacher, diagnostician, and researcher, the follow¬ ing fish health biologists supplied me with specimens and/or shared their experiences with me—to them I am indeed grateful: D. E. Anderson, the late J. E. Camper, the late R. P. Dexter, J. R. Hnath, R.W. Horner, J. A. Hutcheson, J. O'Grodnick, H. M. Jackson, P. W. Janeke, E. L. Leek, J. C. Lientz, R. D. Major, A. J. Mitchell (my last assistant), L. Pettijohn, R. G. Piper, J. H. Schachte, C. E. Smith, V. Ch. Suppes, W. "Gib" Taylor, P. G. Walker, J. W. Warren, T. L. Wellborn, and W. G. Yoder. Several of these have advanced to researcher status in their own right, or into administration. My wife, Carolyn, not only typed the entire manu¬ script but gave editorial assistance. She also prepared most of the drawings in the first edition, many of which have been used again in this volume. Joyce Mann, Vi Catrow, and Lora McKenzie, Techni¬ cal Information Service, National Fish Health Laboratory, Kearneysville, WV, have provided invaluable library assistance. R. Putz, Barbara Rinehart, and Andrea Finger of the Freshwater Institute, Shepherdstown, WV, kindly inserted reviewer suggestions and prepared the final laser copy of the manuscript. I am particularly grateful to five highly regarded fish health specialists whose review advice has confirmed and strengthened the quality of this second edition: Paul Bowser, Professor of Aquatic Animal Health at Cornell University's College of Veterinary Medicine; Ernest H. Williams, Professor of marine Sciences at the University of Puerto Rico and Associate Editor of the Journal of Aquatic Animal Health; Professor Margaret S. Ewing of Oklahoma State University; W. A. Rogers of the Depart¬ ment of Fisheries and Allied Aquacultures in fish health, and laboratory director, now retired from the U.S. Fish xiii

XIV

Preface to the Second Edition

and Wildlife Service, National Fisheries Research Center, LaCrosse, WI. The science editors of Cornell University Press, Robb Reavill, Peter Prescott, and Helene Maddux, graciously provided counsel on the format of the book and provided suggestions for the final manuscript copy. Financial assistance was provided by the Conserva¬ tion Fund of the Freshwater Institute, the Fish Health Section of the American Fisheries Society, the American Society of Parasitologists, the Colorado Aquaculture Association, the Rocky Plains Fish Health Group, the Pacific Northwest Fish Health Protection Committee, Omar Amin, and Larry Roberts.

During my fifty-year career in fish parasitology, remarkable progress has been made. It is now gener¬ ally easier to identify the parasites—including new species; the effects of some of the more prevalent ones have been elucidated; some harmful species can now be controlled; and progress has been made in molec¬ ular techniques. Progress made in the former Soviet Union, which shares some species and many close rel¬ atives in North America, has also helped American fish parasitologists. GLENN L. HOFFMAN

Kearneysville, West Virginia

Preface to the First Edition Much credit belongs to the many people who have helped me, directly or indirectly, during the past 23 years. My major professor, the late L. O. Nolf, University of Iowa, encouraged my interest in fish parasites when I was an undergraduate in 1942. He steered me in the proper methods of parasite identification and in a method of cataloguing the literature on fish parasites that was scat¬ tered in many journals and miscellaneous publications. This book, 23 years later, is the first step in an attempt to bring that literature together in one publication. The late E. B. Speaker and the late Robert Cooper, Iowa State Conservation Commission, assisted me by pro¬ viding part-time employment during my undergraduate studies and before I entered the Armed Forces during World War II. During this time they encouraged my interest in fish parasitology. The late Chester Herrick, University of Wisconsin and the Wisconsin Conservation Department, provided working facilities for me at the Woodruff Biological Sta¬ tion, where I was able to do my first fish-parasite life his¬ tory studies (Posthodiplostomum minimum). Drs. Bangham, Fischthal, Hunter, and the late H. J. Van Cleave have given me much counsel in the study of fish parasites. They also laid the groundwork for fish parasitology in the United States (see References). The late H. S. Davis, U.S. Fish and Wildlife Service, paved the way for government participation in fish par¬ asite research at Feetown. George R. FaRue, U.S. Animal Disease and Parasite Research Branch, has given me much kindly counsel on strigeoids of fish during the past fifteen years. Dr. John Mackiewicz, State University of New York, College of Education, generously helped with the

caryophyllaeid cestode section. Marvin C. Meyer, Uni¬ versity of Maine, gave me much help with the leeches. Victor Sprague, University of Maryland, Natural Resources Institute, assisted me with the genera Dermocystidimn and Ichthyophonus. The following have aided me with counsel or speci¬ mens that helped directly or indirectly in the prepara¬ tion of this book: C. D. Becker, A. Bradford, W. Bullock, R. M. Cable, M. B. Chitwood, G. Dubois, J. E. Hall, S. H. Hopkins, E. J. Hugghins, R. R. Kudo, J. Fom, A. McIntosh, G. Malmberg, F. P. Meyer, J. D. Mizelle, J. F. Mueller, P. Osborn, F. Sogandares-Bernal, H. W. Stunkard, F. J. Thomas, J. H. Wales, R. A. Wardle, and S. Yamaguti. Paul Thompson, Director, Division of Fishery Research, U.S. Bureau of Sport Fisheries, and S. F. Snieszko, Director, Eastern Fish Disease Faboratory, have made it possible for me to continue and bring this work to com¬ pletion. Their understanding of the need for such a book for students and researchers is much appreciated. My wife, Carolyn, has graciously prepared most of the drawings, most of which were traced from photocopies of the originals. Robert Putz, my associate, has very ably carried on our laboratory work during the preparation of the book. Without the typing and library assistance of Juanita Collis, Bonnie Knott, Millicent Quimby, Mary Ann Strider, and Florence Wright of this laboratory, the fin¬ ished copy would not have been possible. Ernest Callenbach, University of California Press, and Genevieve Rogers are to be commended on their gra¬ cious and efficient handling of the many editorial details. GLENN L. HOFFMAN

Kearneysville, West Virginia, 1967

XV

Abbreviations The following are abbreviations used throughout the book.

States of the United States

AL AK AR AZ CA CO CT DE DC FL GA HI IA ID IL IN KS KY LA MA ME MD MI MN MO MS

Alabama Alaska Arkansas Arizona California Colorado Connecticut Delaware District of Columbia Florida Georgia Hawaii Iowa Idaho Illinois Indiana Kansas Kentucky Louisiana Massachusetts Maine Maryland Michigan Minnesota Missouri Mississippi

Canadian Provinces

MT NC ND NE NH NJ NM NV NY OH OK OR PA PR RI SC SD TN TX UT VA VT WA WI WV WY

Montana North Carolina North Dakota Nebraska New Hampshire New Jersey New Mexico Nevada New York Ohio Oklahoma Oregon Pennsylvania Puerto Rico Rhode Island South Carolina South Dakota Tennessee Texas Utah Virginia Vermont Washington Wisconsin West Virginia Wyoming

Alta. B.C. Lab. Man. M.P. N.B. Nfdl. N.S. N.W.T. Ont. P.E.I. Que. Sask. Y.T.

Alberta British Columbia Labrador Manitoba Maritime Provinces New Brunswick Newfoundland Nova Scotia Northwest Territories Ontario Prince Edward Island Quebec Saskatchewan Yukon Territory

Parasites of

North American Freshwater Fishes

Introduction The importance of fish parasites is related directly to the importance of the fishes they may affect. As our world becomes more populated, all foodstuffs, including fish, become increasingly valuable. It is well known that fishes are an excellent source of complete protein con¬ taining little saturated fat. Fishes provide an important recreational asset, both for sport fishing and as one of the attractions of nature. Observing live fishes, both in nature and in the display aquarium, is enjoyable to young and old alike. Chubb (1965), in the introduction to a fish parasite survey in England, clearly described the dynamics of fish parasitology: In natural populations of plants and animals par¬ asites are always present. The parasites are normally in a complex dynamic equilibrium with the freeliving communities of plants and animals. Fishes are the apex of the predator-prey pyramid within fresh waters, and therefore tend to be infected by a considerable range of parasites, which may occur in large numbers. This is the normal condition found in any natural environment. However, if some unusual event occurs in the environment, of natural or human origin, the equilibrium between host and parasite may be disturbed, and an (epizootic) of one or more species of parasites may occur. Regulating mech¬ anisms in the environment soon come into play, and a new equilibrium will be established; but in the intervening period, there may be a serious loss of fishes. It is thus important from an economic point of view, for fishing as an amenity, or for fish farming, that we have a knowledge of the occurrence of par¬

asites on our freshwater and marine fishes. Once we have a sound background knowledge, it may at least be possible to avoid some undesirable human interference in natural waters, and even to control some of the more harmful parasites. Unfor¬ tunately, for really effective control the fullest possible details of the biology of the parasites are required. Currently, we have very little information on the biology, or indeed of the occurrence, of the parasites of either freshwater or marine fishes of the British Isles. More recently, Lester (1984) reviewed methods for studying the effect of parasites in feral fishes. Before the treatment or control of fish parasitic dis¬ eases can be achieved, the study of fish parasites should follow a logical pattern: 1. Identify the parasite. 2. Obtain a thorough knowledge of its life history, which may be simple or very complicated. 3. Learn the ecological requirements of the parasite, such as host specificity, optimum temperature, pH, nutrition, and other metabolic requirements. 4. Map the geographic range of the parasite. 5. Determine the effect of immunologic mechanisms of the host on the parasite, or vice versa. 6. Study control and treatment methods. This manual is intended to be an aid to the identifi¬ cation of freshwater fish parasites of North America. Little work has been reported from Alaska and Mexico, so those areas are not well represented. Most of the keys are for generic identification. The known species of each genus are listed along with reference citations for literature pertinent to species identification, life cycles, and sometimes control. 1

2

Introduction

All families and genera are listed in alphabetical order. For the highest taxa, refer to Margulis and Schwartz (1988). Most of the identification keys are set up in couplets of contrasting characteristics, but a few are set up in triplets and quadruplets to save space. Page numbers are given in the keys only when the reference is not to a nearby page. In many cases, parasite-host records are insufficient to determine host specificity; such lack of records can be very misleading. Parasites of brackish water fishes and anadromous fishes are included because these fishes are sometimes found in freshwater. Euryhaline species are represented by Ergasilus lizae (copepod), Glugea hertwigi (Protozoa), and Ichthyophonus hoferi (fungus). The gen¬ era Gyrodactylus (Monogenea), Oodinium (Protozoa), and Trichodina (Protozoa) and some of the myxosporideans have representatives in both fresh and marine water, but they are not euryhaline. Foreign fish parasites are included only when the host exists on both continents, or if there is a probability that the parasites will also be present in North America; this includes some aquarium fishes such as tropicals, koi carp, and goldfish. International contact can be made in the International Symposium of Fish Parasitology (Color Fig. 1). Scope. In most fish disease research laboratories, the work on diseases is divided into virology, bacteriology, parasitology, and histopathology. Individuals in diag¬ nostic laboratories (hatchery biologists, etc.), however, usually handle all aspects, including environmental parameters. Importance offish parasitology. No attempt is made here to place a monetary value on the damage done to fishes by parasites. In fish hatcheries this could probably be done with some accuracy when fishes are either killed by parasites or are so adversely affected that they do not grow or reproduce properly. One infection, whirling disease of trouts, caused by Myxobolus cerebralis, sometimes results in such spectacular deformities in surviving fishes that the fishes cannot be marketed or used for stocking purposes. Another disease, caused by Ichthyophonus hoferi, occasionally brings similar crippling. In nature it has been impossible to determine the dam¬ age done by parasites, except in serious fish kills such as those caused by Ichthyophthirius multifUiis and Lernaea cyprinacea. Each true fish parasite uses the fish for its home and food, and the total damage is relative to the numbers of parasites present. If few parasites are present, the fish is usually not visibly impaired; if large numbers are present, the fish may be killed. Some parasites effect much mechanical damage by migrating through tis¬ sues and may also cause extensive tissue proliferation, which impairs growth and reproductive processes. The pathology is well covered by Dogiel et al. (1958, p. 84), Ferguson (1989), Ribelin and Migaki (1975), and Roberts (1978). For identification of various helminths in tissue

sections, see Chitwood and Lichtenfels (1972). For comparisons of parasites of humans in tissue sec¬ tions, see Kenney (1973). For histology of various fishes, see Anderson and Mitchum (1974): trout; Grizzle and Rogers (1976): channel catfish; Groman (1982): striped bass; and Yasutake and Wales (1983): salmonids. Prevention, treatment, and eradication. One of the most important factors leading to the control of fish par¬ asites is the study of the ecology of the parasite. Such information usually leads to a weak link by which con¬ trol may most easily be effected. Ecology of fish parasites is discussed by Dogiel et al. (1958, p. 1), O. N. Bauer et al. (1959), Esch (1971), and Kennedy (1981). Preventing the introduction of dangerous parasites into hatcheries or lakes and streams is far more effective than the chemical treatment of parasitized fishes. In a hatchery or other body of water that does not have serious parasites, any new stock should be carefully examined and rejected, or treated if parasitized. Chemical treatment of externally parasitized fishes is usually successful, but internal parasites are more diffi¬ cult to treat (see under Chemotherapy and Prophy¬ laxis). More research is needed in this field because of the rapidly growing importance of fish culture. Eradication of external parasites in hatcheries is pos¬ sible but is usually only temporary if there are infected fish somewhere in the water supply. Properly designed springwater supply units or wells will never be a source of parasites, and if no parasites are brought in with trans¬ ferred fish, the water will contain no parasites. Some internal infections, such as whirling disease of trouts, however, are almost impossible to eradicate because of their resistant spores and complicated life cycles. In this book the most important parasites are covered suf¬ ficiently for research background, but those of unknown importance are usually described only for identifica¬ tion purposes. Having been an excellent classical student, Carolyn has corrected the Latin endings on many of the species names. The literature has been surveyed as thoroughly as pos¬ sible through 1988, with reference to major works through 1992. The outstanding aids to the bibliographic work were the Index—Catalogue of Medical and Veterinary Zoology (U.S. Department of Agriculture), parts 1-18 and supplements 1-24; Helminthological Abstracts and Protozoological Abstracts of the Commonwealth Agri¬ cultural Bureau; Biological Abstracts; and Publications on Fish Parasites and Diseases, 330 B.C.-A.D. 1923 (McGre¬ gor, 1963). The leading contributor to freshwater fish par¬ asitology of North America, because of his numerous surveys, is the late R. V. Bangham, formerly of College of Wooster, Wooster, Ohio. Some of the most helpful major works on general fish parasitology are the following: O. N. Bauer et al. (1959): pond fishes of the former Soviet Union; O. N. Bauer

Introduction

(1984, 1987a, 1987b): major keys to identification, for¬ mer Soviet Union; O. N. Bauer et al. (1977): former Soviet Union; Bykhovskaya-Pavlovskaya et al. (1962): keys to identification, former Soviet Union; Dogiel et al. (1958): former Soviet Union; Ergens and Lom (1970): the Czech Republic and Slovakia; G. L. Hoffman (1973a): lab¬ oratory fishes; Hoffman and Schubert (1984): exotic parasites; Izyumova (1977): reservoirs; Lom (1983): First International Fish Parasitology Symposium; McDonald and Margolis (1995): Canada (not included in this book); Lom and Dykova (1992): Protozoa; Margolis and Arthur

3

(1979): Canada; Margolis and Kabata (1984-1989): Canada; Mitchum (1995): Wyoming (not included in this book); Moles (1982): Alaska; Molnar (1987): Second International Fish Parasitology Symposium; Moore et al. (1984): parasites in fish farming; Reichenbach-Klinke et al. (1980): English translation of the diseases of fishes (Germany); Schaperclaus (1954, 1986): general fish dis¬ eases and parasites; Schubert (1977): aquarium fishes; Schubert (1987): primarily aquarium fishes; Untergasser (1989): primarily aquarium fishes; Wyatt and Economan (1981): upper midwest, USA.

Public Health Aspects of Fish Parasites As Parasites of Humans Some larval parasites of North American fish can infect humans. So far, there have been few reports of serious cases in this country, but very serious cases, including fatalities, have been reported from Europe, Asia, and the Philippine Islands (Acha and Szyfres, 1987; Arambulo, 1982; Cross et al., 1978; Ghittino, 1972; ReichenbachKlinke, 1975; Vik, 1964c). In all cases, however, thorough cooking will kill the parasites. The following parasites have been reported from humans in North America or are potentially dangerous because of their lack of host specificity. Some foreign references are given for guidance.

Trematodes Heterophyids usually lack strong host specificity, and many of the heterophyid metacercariae will likely develop in the intestine of humans—e.g., Africa et al. (1936) found that the ova of heterophyids became trapped in the intestinal villi of humans, after which the ova were forced into the circulatory system and filtered out in various organs, including vital organs, some¬ times causing death. Cryptocotyle lingua has infected people in northern Europe (Christensen and Roth, 1949), and Haplorchis pumilio of farmed Tilapia in Africa should be avoided (Sommerville, 1982a). Welberry and Pacetti (1954) reported Heterophyes heteropliyes eggs from a child from Florida. Hutton (1957), however, noted that identification of the eggs is not possible: he obtained adult Phagicola, not Heterophyes, from lab ani¬ mals that were fed the same species of fish from Florida. Thus, the Heterophyes record may be erroneous. Font et al. (1984) also reported on Phagicola as a pathogen in humans. Clinostomum complanatum, which usually lives 4

in the mouth of herons and has larvae in fishes, might migrate into the trachea of humans; a related species in India, Israel, and Japan has been so reported (Cameron, 1945; Witenberg, 1944). The opisthorchids, Metorchis and Opisthorchis, exist on this continent and are serious liver parasites of humans in Asia. Plagiorchis has been reported from people in the Philippines (Africa et al., 1936). Nanophyetus sahnincola has been reported from people in Russia (Chandler, 1955). Humans have been experimentally infected with Apophallus donicus (Niemi and Macy, 1974). Cercariae of Diplostomum spathaceum (adult in gulls, diplostomulum in fish eye lens, and cercariae in Lymnaea snails) dropped onto the eyes of small rabbits penetrated through the cornea, crossed the anterior chamber, and entered the lens, but died after two to three weeks. However, when dropped onto the eyes of adult rabbits and cold-stored, enucleated human eyes, they failed to penetrate (Fester and Freeman, 1976). In England, D. spathaceum was found in lens cataracts of a 5-month-old child and a 55-year-old fisherman (Ashton et al., 1969).

Cestodes Diphyllobothrium latum will develop in humans and is present in the north central United States, Alaska, and Canada (Ruttenber et al., 1984). Diphyllobothrium pacificum, a marine cestode, has been found in humans in Peru (Baer, 1969), and Schistocephalus solidus in Alaska (Rausch et al., 1967).

Nematodes The very serious kidney worm, Dioctophyme renale, will develop in human (Gutierrez et al., 1989). Farval migrans

Public Health Aspects of Fish Parasites

from Anisakis-type larvae cause an acute abdominal syndrome in humans (Bier et al., 1987; Cheng, 1976; Chitwood, 1970; Dailey et al., 1981; Oishi and Hiraoki, 1971; Oishi et al., 1974). Margolis (1977a) reviewed the public health aspects of "codworm" infections. This parasite is usually in marine fishes. Eustrongylides sp. ingested in raw bait fish in two cases caused serious abdominal disease requiring surgery (Anon., 1982; Lichtenfels and Stroup, 1985; S. Miller, 1982), was coughed up in one case (Abram and Lichtenfels, 1974), and in another case caused eustrongylidiasis after ingestion of raw fish sushi (Wittner et al., 1989). In the Philippines, people have suffered severe enteritis from Capillaria philippinensis from raw freshwater fish (Cross et al., 1978). In Mexico, there has been an increase in Gnathostoma spinigenun infections from eating marinated fresh¬ water fish (Akester, 1988). All of these infections are acquired by ingesting raw or undercooked fish. Some are acquired accidentally while tasting fish preparations before cooking, or while cleaning fish. All of the parasites, however, are killed by thorough cooking, hot smoking, or freezing. Although

5

pickling kills some parasites, Hutton and SogandaresBernal (1959b) found that freezing, hot smoking (4 hr at 100°-125°C), but not brine preparations, killed the metacercariae of Mesostephanus appendiculatoides.

Crustacea Argulus (fish louse) has been found in the human eye (Hargis 1958a), but this must be highly unusual.

As Esthetically Undesirable Parasites Most fish parasites, even if eaten raw, will not develop in humans, but some species are unsightly. In two instances, such diseased fishes have been banned from interstate commerce by the U.S. Food and Drug Admin¬ istration. Unsightly but dead Phocanema (codworm) are occasionally found in ocean fish fillets. Many fishes are discarded by fishermen because of unsightly parasites, but none are harmful to people if the fishes are thoroughly cooked.

Methods Examination of Fresh Fishes After collection from the wild or from culture facilities, fishes must be kept in well-aerated containers at the appropriate temperature. They should be handled humanely and killed with anesthesia. If all parasites are to be examined, the fish must be dead no longer than a few minutes, for some of the exter¬ nal protozoa leave the dead fish. Although not as satis¬ factory, in some cases preserved or frozen carcasses can be used (Pence et al., 1988). For methods on transport¬ ing and submitting samples, see Mitchell and Hoffman (1987). Some of the larger parasites can be seen with the naked eye, and almost all other parasites, or their cysts, can be seen with a good 10 to 30x dissection microscope. Some protozoa (blood, intestinal, and some external forms) can be seen only with the compound micro¬ scope. One of the most satisfactory routines is the fol¬ lowing: 1. Kill the fish by pithing or with anesthetics in the water. 2. If it is small, examine the entire fish submerged in water in a Petri dish, using a 10 and 30x dissection microscope. If the fish is too large, remove the fins and examine them as above. 3. Examine the fins or mucus scrapings from the dor¬ solateral portion of the fish for small external protozoa, using 100 and 450x magnification. 4. Remove the gills, submerge in water, and examine with 10 and lOOx dissection microscope. 5. Open the fish and remove a drop or two of blood from the heart, dilute about 50:50 with normal physio¬ 6

logical saline, and examine at lOOx for motile Trypanoplasma and Trypanosoma. At this time or before the gills are removed, prepare blood smears to be stained for blood sporozoa. 6. Remove the viscera. If the fish is small, this should be done in saline under the dissection microscope; organs should be teased apart with small forceps. Remove a drop or two from the intestine and examine for proto¬ zoa (Hexamita, Schizamoeba, and Eimeria). 7. Carefully remove the gall and urinary bladders with fine forceps and examine for sporozoa and ciliates. Always squash a small piece of kidney under a cover slip and examine for Myxosporidea and Trypanoplasma. 8. Remove the gastrointestinal tract. If the fish is small, the tract should be opened under the dissection scope. If it is larger, scrape the wall, dilute the contents with saline, shake, and allow the parasites to settle out in a conical or cylindrical container. After 5 to 10 minutes, aspirate the fluid, leaving the parasites on the bottom; repeat until clean and pour into a small Petri dish for examination under the dissection microscope. 9. Some of the body musculature should be teased apart carefully under observation with a dissection microscope. Larval worms, sometimes encysted, are usu¬ ally easily seen, but a few are very small and can be seen only with higher magnification. Many trematode and nematode larvae can be recovered by the digestion tech¬ nique (see below). 10. Remove and examine the eyes and brain. Cut the head lengthwise to remove the brain, and examine the inside of the mouth and esophagus in water under the dissection microscope.

Fish Parasite Fixation and Preservation

11. Concentration methods for myxosporidans are sometimes very helpful; for digestion techniques, see Markiw and Wolf (1974); for plankton centrifuge method, see O'Grodnick (1975) and L. G. Mitchell, Kroll, and Seymour (1983), the latter for Myxobolus species other than M. cerebralis. 12. All ectoparasites should be studied alive in chlorinefree tap water, and most internal parasites in physio¬ logical saline; 0.8-0.9% sodium chloride is adequate for temporary use (rainbow trout blood, e.g., is almost iso¬ tonic with mammalian blood; others may be different).

Freeing Intestinal Nematodes from Mucus Trematodes and cestodes are usually easily freed from mucus, but longer nematodes often become entangled in it. Shaking the nematodes for a few minutes in digest fluid (see below) usually frees them, but they will be harmed if left in the solution for very long. A good way to clean intestinal worms is as follows: slit open the intes¬ tine, place it with its contents in a physiological saline solution in a jar, shake for a minute, allow the con¬ tents to settle, and then pour off the upper, clearer por¬ tion of the liquid. Repeat as needed until the parasites are relatively clean. Examine a portion of the concentrate in a Petri dish under a dissecting microscope, usually with a strong incident light and a dark background.

Helminth Eggs, Helminth Spores, and Fecal Examination Sometimes lookalike helminth species have eggs that are recognizably different. One should therefore examine the eggs carefully. Occasionally, individual hosts cannot be sacrificed, but coprology can be done almost as with mammals. Special equipment, such as a fecal catheter with a blunt syringe, a funnel with a screen, and a rub¬ ber body sheath, can sometimes be used to obtain fecal samples (Mamedov and Mamedov, 1975; Moser and Sakanari, 1985).

Digestion Technique If trematode and nematode larvae are scarce or needed in large numbers, some species of trematodes and all lar¬ val nematodes can be concentrated by removing the fish tissue by digestion. Other species of trematodes, however, and all larval cestodes, however, are rapidly destroyed by this method. It is useful for Ichthyophonus hoferi and probably other spore-forming species, and modifica¬ tions have been used for myxosporidans (Markiw and Wolf, 1974).

7

Dissolve 0.5% of commercial pepsin in water con¬ taining 0.5% hydrochloric acid. Cut the fish into very small pieces or grind in a hand-powered kitchen food grinder; individual organs can be digested separately if necessary. Add approximately 1 g of fish to 20 ml of solu¬ tion, place in a jar containing a few glass marbles to aid mixing, and digest at 37°-39°C for one to two hours. Pro¬ longed digestion will destroy some parasites that survive the two hour method, although some nematodes such as anisakids can survive 24 hours of such treatment (Jackson et al., 1981). Continuous agitation in a water bath shaker gives the best results, but the solution can be mixed in an incubator or stationary water bath, using a magnetic stirrer, compressed air agitation, or even shaking by hand. The digested material should be strained through a wire tea strainer to remove bones and undigested par¬ ticles. Allow the digest to stand for about 15 minutes; the helminths will fall to the bottom and the supernatant can be aspirated. Add saline and repeat until clean. For counting purposes only, water instead of saline may be used. Pour the concentrate into a small Petri dish and examine under the dissection microscope.

Fish Parasite Fixation and Preservation Many parasites contract badly when placed in cold fix¬ ative; therefore, the following methods should be used. These methods, however, may not work well on all species, and the worker may want to try other procedures. I. Protozoa

A. Myxosporidea (often large white cysts) Cut out the cyst with enough adjacent tissue to prevent rupturing of the cyst. Place in a small vial of 10% formalin. B. Trophozoites of motile forms (external or inter¬ nal) 1. Place as many protozoa as possible on a microscope slide in one drop of water, add one drop of PVA-AFA (polyvinyl alcoholacetic acid formalin alcohol) fixative adhesive, mix thoroughly, spread over about half of the slide, and allow to dry for future staining. Wet fixation on a slide yields better results (con¬ sult books for techniques). 2. Place as many protozoa as possible in a small vial of 10% formalin. Protozoa usually shrink greatly. II. Trematodes

Most trematodes contract greatly if killed in cold fixative, so they should be dropped into a small vial of hot (85°-90°C) Bouin's fixative, 10% for¬ malin, or AFA. Replace the next day with 70% alco¬ hol. The hot water method is useful for mass fixing of platyhelminthes (Sinclair and John,

8

Methods

1973). Those that are too thick must be flattened under a cover slip, with the fixative drawn under the cover slip using absorbent paper; allow for distortion. For monogenea, drop infected gills or fins into 10% formalin. (See also methods in the section on monogenetic trematodes.) III. Cestodes

Much the same as for trematodes. Usually kill cestodes in 80°C water and stored in buffered 10% formalin. IV. Nematodes

Because of the impervious cuticle, it is difficult to prepare stained permanent mounts; smaller worms, however, can be mounted in glycerine jelly and per¬ haps other semisolid material. Most workers make temporary mounts of larger nematodes in lactophenol or alcohol-phenol for clearing; the speci¬ mens can be returned to alcohol after use. V. Acanthocephala

Much the same as for nematodes. If proboscis is retracted, place worm in distilled water in refigerator until proboscis everts, sometimes overnight. Before fixing in hot fixative, the cuticle must be pricked to avoid bubbles forming in the tissue. VI. Leeches

Similar to trematodes, but some are too thick for staining and must be flattened between slides and then fixed with 10% buffered formalin. (See section on leeches.) VII. Copepods (fish lice)

If easily detached (e.g., Arguhis), drop individuals into a small vial of 70% alcohol. If not easily detached, cut out a small piece of tissue contain¬ ing the parasite(s) and drop into a small vial of 70% alcohol, or carefully remove the tissue from the par¬ asite before preserving.

Permanent Preparation of Parasites

Nuclear detail shows up well with iron hema¬ toxylin staining following osmic acid fixation. Myxosporidean spores are best studied fresh but can be affixed to slides, fixed while still wet, and stained with Giemsa's. The vegetative stages are best studied in cross sections stained with hema¬ toxylin and eosin. If spores are present, Giemsa's (usually the Giemsa method of May-Griinwald) is also recommended. Two general methods for Protozoa may be found in Kirby (1950) and Lee et al. (1985). II. Trematodes

Several stains yield good results, but since worms vary considerably, no one stain is recommended. Stains most widely used are Semichon's and borax carmine, counterstained with fast green; Harris' or Heidenhain's hematoxylin; hematin; coelestin blue B; and chlorazol black E. The last is selective for some of the chitinous hard parts of Monogenea. Most of the staining involves overstaining and then destaining under observation until the desired result is attained. Some species stain more readily than others; it is difficult to recommend exact staining times. Be certain that specimens stained with hematoxylin are neutralized adequately before mounting, or they may fade. Meyer and Olsen (1975) has a good section on helminth techniques. (See also methods in the section on monogenetic trematodes.) III. Cestodes

Much the same as for trematodes. Usually kill cestodes in 80°C water and store in buffered 10% formalin. IV. Nematodes

Because of the impervious cuticle, it is difficult to prepare stained permanent mounts; smaller worms, however, can be mounted in glycerine jelly and per¬ haps other semisolid material. Most workers make temporary mounts of larger nematodes in lacto-phenol or alcohol-phenol for clearing; the specimens can be returned to alcohol after use. V. Acanthocephala

Only a brief outline of methods used is given here. The techniques can be found in books on general parasitol¬ ogy and histologic technique. I. Protozoa

Blood flagellates are found more easily alive in wet mounts, but all blood protozoa should be spread in a thin smear on a microslide and stained with Giemsa's stain. The stain and wash should be buffered at about pH 7. Ciliate morphology, including the trichodinids, is best studied with silver impregnation methods (Arthur and Lorn, 1984: dry smears; Augustin et al., 1984; Fernandez-G., 1976; Hazen et al., 1976: dry smears; Ng and Nelsen, 1977; Lorn, 1958: dry smears; Raabe, 1959: dry smears; Wilbert, 1975).

Much the same as for nematodes. If proboscis is retracted, place worm in distilled water in refrig¬ erator until proboscis everts, sometimes overnight. Before fixing in hot fixative, the cuticle must be pricked to avoid bubbles forming in the tissue. VI. Leeches

Similar to trematodes, but some are too thick for staining and must be flattened between slides and then fixed with 10% buffered formalin. (See section on leeches.) VII. Copepods

Store in alcohol and study as temporary mounts (see Harding, 1950). Small Arguhis can be cleared and mounted permanently, usually using commercial mounting rings to hold up the cover slip.

Some North American Fish Parasites Listed by Location in the Fish This section is included as an aid to identification because some parasites are found only in very selective organs or tissues. Some of these are probably rare records. Some foreign reports are cited for examples and in cases of infected imported fish. For references, see specific parasite section.

i Eggs Fungi: Saprolegnia and relatives. Protozoa: Epistylis reported from catfish eggs (Well¬ born, pers. comm., 1964); Carchesium on wall¬ eye and trout eggs; Pleistophora variae in golden shiner eggs; P. sulci in Polyodon spathula eggs; Thelohania baueri in Pungitius pungitius eggs, for¬ mer Soviet Union. Coelenterata: Hydra sometimes attack eggs and fry. Polypodium in Acipenser and Polyodon eggs. Turbellaria: Planaria sometimes attack eggs. Hirudinea: Leeches may attack eggs. Insecta: Predators, including caddis fly larvae. 2. Barbels

Protozoa: Henneguya sp. in Ictalurus nebulosus; Ichthyophthirius occasionally. Trematoda: Gyrodactylus spp. 3. Body Skin, Fin Skin, and Adipose Fin

Fungi: See external fungi, including Exophiala pisciphila. Protozoa: See ectoparasitic protozoa. Henneguya in adipose fin of Ictalurus punctatus. Monogenea: Gyrodactylus, but usually not other monogeneans. Trematoda: Metacercariae of many species. Nematoda: Undescribed larval nematode in skin nodules of ictalurids, eastern half of USA.

Arthropoda: Larval mites, former Soviet Union; Argulus; Lernaea; occasionally Salmincola. 4. Nares (Nasal Fossae)

Protozoa: Apiosoma sp., Europe; Amphileptus, Chilodonella, Myxobolus, Tetrahymena, Trichodina, Trichodinella, former Soviet Union (Yukhimenko, 1972; Kostenko and Schmalgauzen, 1983). Monogenea: Aplodiscus nasalis in Hypentelium etowanum; Cleidodiscus monticelli; Pellucidhaptor catostomi; P. nasalis; P. pricei. Nematoda: Pbilometra in bluegills and largemouth black bass. Copepoda: Ergasilus megaceros in fallfish and catfish; Ergasilus rhinos in centrarchids; Gamidactylus, Gaminispatulus, Gaminispinus, Lernaea, Paragasilus, Salmincola, Amazon River, Brazil. 5. Lateral Line Pits

Monogenea: Gyrodactylus cryptarum, Europe (Malmberg, 1970); Pellucidhaptor planacrus (Leiby et al., 1972; G. L. Hoffman, unpub., USFWS, 1980); Urocleidus adspectus (Mueller, 1936c). 6. Gills

Protozoa: Ambiphrya, Amphileptus, Chilodonella, Cryptobia, Dennocystidium, Epistylis, Ichthyobodo (Costia), Ichthyophthirius, Microspora, Myxosporea, Piscinoodinium, trichodinids, Trichophrya in or on gill arches and filaments. Monogenea: Some spp. of Dactylogyrus, Gyrodacty¬ lus, ancyrocephalids, polyopisthocotylids. Trematoda: Certain metacercariae; ova becoming miracidia of Sanguinicola. Copepoda: Adheres, Ergasilus, Lepeophtheirus, Ler¬ naea, Salmincola. Arthropoda: Larval mites. 9

Some North American Fish Parasites Listed by Location in the Fish

10

7. Branchial Cavity

Accidentally: Any parasites found on the gills. Trematoda: Syncoelium. Arthropoda: Larval mites. 8. Mouth

Protozoa: Apiosoma, Europe; Myxosporea. Monogenea: Dactylogyrus capoetobramae, former Soviet Union. Trematoda: Metacercariae of Petasiger sp. (experi¬ mentally); Leuceruthrus. Nematoda: Philometra nodulosa in suckers and buf¬ falo fishes. Copepoda: Lemaea cyprinacea, Salmincola. 9. Blood

Protozoa: Trypanosoma (Cryptobia), Trypanoplasma free; Babesiosoma, Dactylosoma, Haemogregarina in red blood cells; rarely Kucioa; Sphaerospora trophozoites (Csaba bodies) free in carp and other cyprinids, Europe. Trematoda: Sanguinicola in blood vessels, including gill vessels, also some migrating forms. Nematoda: Philometra sanguinea in caudal fin blood vessels of goldfish; P. obturans in gill vessels of pike, former Soviet Union. 10. Esophagus

Trematoda: Azygia, Cotylaspis, Derogenes, Halipegus, Proterometra. Arthropoda: Larval mites. 11. Stomach

Protozoa: Schizamoeba spp. Monogenea: Enterogyrus spp. (freshwater, tropi¬ cal); Enterogyrus sp. in foregut of Pomacanthus pant (Cone et al., 1987); Etroplus spp. (tropical, marine). Trematoda: Allocreadium, Aponeurus, Azygia, Caecincola, Centrovarium, Derogenes, Genolinea, Hemiurus, Leuceruthrus. Nematoda: Haplonema. 12. Intestine and Pyloric Ceca

Protozoa: Hexamita trophozoites and cysts in lumen; Schizamoeba cysts in fingerling trout; Eimeria, Goussia in goldfish, other cyprinids, trout. Monogenea: Enterogyrus cichlidarum in Tilapia, North Africa. 13. Feces

Coprological examination for protozoa spores and helminth eggs can be helpful (Tesarcik, 1971a; Reichenbach-Klinke, 1980; Mamedov and Mamedov, 1975; Moser and Sakanari, 1985). 14. Swim Bladder

Protozoa: Goussia cichlidarum in cyprinids and cichlids, including Tilapia, Israel. Trematoda: Acetodextra in Ictalurus punctatus. Nematoda: Cystidicola spp. in salmonids; Huffmanella huffmanella in Lepomis cyanellus; Pseudocapillaria tomentosa in Tinea tinea, Europe.

15. Body Cavity

Fungi: Ochroconis humicola in coho salmon; O. tshawytscha in chinook salmon; Phoma herbarum in northwest salmonids. Protozoa: Many Myxosporea; rarely Microsporea; Goussia spp. Trematoda: Many metacercarial species, including Ornithodiplostomum, white grub (Posthodiplostomum); adult Paurorhynchus (bucephalid), adult Acetodextra (in catfish). Cestoidea: Larval dilepids, Diphyllobothrium, Haplobothrium, Ligula, Proteocephalus, Schistocephalus, Triaenophorus. Acanthocephala: Larvae of Echinorhynchus salmonis, Leptorhynchoides thecatus, Pomphorhynchus bulbocolli. Nematoda: Adult Philonema, adult Pseudocapillaria (Europe); many larval species. Copepoda: Rarely Lemaea in small fishes. 16. Gall Bladder

Protozoa: Myxosporea; Hexamita(7), Europe. Trematoda: Crepidostomum cooperi; C. farionis; Dero¬ genes sp.; Plagioporus sinitsini; Prosthenhystera sp.; Pseudochaetosoma, Europe. Cestoidea: Eubothrium salvelini; larval Dilepidae (armed pleurocercoids). Nematoda: Rhabdochona sp.; occasionally Capillaria catostomi. 17. Hepatic Bile Duct

Trematoda: Phyllodistomum spp. 18. Kidneys

Fungi: Ichthyophonus hoferi in many fishes; Ochro¬ conis spp. in West Coast coho and chinook salmon. Protozoa: Myxosporea in tubules; sometimes inter¬ stitial, sometimes cysts. Trematoda: Metacercariae of Nanophyetus salmincola, Posthodiplostomum minimum centrarchi, probably others; adult Phyllodistomum in renal tubules and ureters. 19. Ureters

Protozoa: Vauchomia spp. in Esox spp. Trematoda: Adult Phyllodistomum. 20. Urinary Bladder

Protozoa: Myxosporea; Vauchomia (trichodinid) in Esox spp. Trematoda: Acolpenteron (Monogenea); Phyllodis¬ tomum. 21. Ovaries

Protozoa: Henneguya oviperda in Esox lucius, Europe; Pleistophora ovariae in golden shiners; Thelohania baueri in Gasterosteus aculeatus ova, former Soviet Union (Shulman, 1984). Coelenterata: Polypodium hydriforme in sturgeons and paddlefish (Hoffman et al., 1974; Raikova et al., 1979; M. L. Kent, pers. comm., Nanaimo,

Some North American Fish Parasites Listed by Location in the Fish

B.C., 1984) from California, Michigan, Missouri, former Soviet Union; Hydra sometimes attack fry. Trematoda: Acetodextra ameiuri. Nematoda: Philonema spp. in salmonids. 22. Testes

Protozoa: Hexamita (E. Moore, 1925). 23. Head Sinuses

Nematoda: Philometra sp. 24. Eyes

Protozoa: Henneguya episclera in Lepomis gibbosus; H. zikaweiensis in Carassius aurahis, China; Myxobolus corneas in cornea of Lepomis macrochirus; M. hoffmani in sclera of Pimephales promelas. Trematoda: Diplostomulum scheuringi in vitreous chamber; D. spathaceum in lens; probably other species of Diplostomulum, with some in retina; Omithodiplostomum metacercariae rarely in eye. Nematoda: Philometroides sp. in eye orbit of south¬ eastern centrarchids. 25.Cartilage

Protozoa: Henneguya brachyura in fin ray of Notropis spp.; H. schizura in sclera of Esox lucius; Hen¬ neguya sp. in branchial arch of Potnoxis spp.; M. cartilaginis in centrarchids; Myxobolus cerebralis in salmonids; M. hoffmani in sclera of eye of Pimephales promelas; M. scleropercae in cartilagi¬ nous sclera of eye of perch; 13 other species

11

listed in G. L. Hoffman (1989). Trematoda: Galactosomum metacercariae in cranium of anchovies and yellowtail, Japan; undescribed metacercariae in fin rays and gill arches of trop¬ ical fishes. 26. Nervous System

Fungi: Ichthyophonus hoferi in brain. Protozoa: Mesencephalicus in brain of Cyprinus carpio, Europe; Myxobolus cerebralis affects CNS, though parasite is in cartilage; Myxobolus hendricksoni in brain of Pimephales promelas; Myx¬ obolus arcticus, M. kisutchi, M. neurobius in CNS of salmonids. Monogenea: Some species in lateral line pits. Trematoda: Diplostomulum, Euhaplorchis Ornithodiplostomulum, Parastictodora, metacercariae on brain; Psilostomum metacercariae in lateral line canal. 27. Muscle and Body Connective Tissue

Protozoa: Many Myxosporea; some Microsporea; Sarcocystis (may be in error). Trematoda: Many metacercarial species, including Clinostomum (yellow grub) and Neascus spp. (blackspot). Cestoidea: Larval Diphyllobothrium, Triaenophorus. Acanthocephala: Some larval forms. Nematoda: Larval Eustrongylides.

Brief Descriptions of the Groups of Fish Parasites The following list is intended as an aid to identifica¬ tion. It may be difficult to place a parasite in its proper phylum, for some larval forms and at least two adults do not exhibit some of the typical morphological characteristics. 1. Algae (Kingdom Protista): Unicellular, 10-20 pm in diameter; green pigment. Usually in clumps as small cysts in gills, eye orbits, occasionally in viscera of small trop¬ ical fishes. 2. Fungi (Kingdom Fungi): Usually filamentous, about 20 pm in diameter, nonseptate. Ichthyophonus often occurring as spheres up to 100 pm in diameter, but some showing hyphalike outgrowths. Filamentous bac¬ teria (Myxobacteria) and actinomycetes only 1 to 3 pm in diameter. 3. Protozoa (Kingdom Protista): Commonly called single-celled animals. Amoebae, ciliates, and flagellates easily recognized, but vegetative stages of some sporozoa are not. Developing Myxosporea syncytial rather than truly single-celled, but with distinctive spores. The remaining groups are in the kingdom Animalia. 4. Coelenterata: One species, Polypodium hydriforme, found in ova of sturgeon and paddlefish. Hydro may attack very small fishes. 5. Monogenea and Aspidocotylea (Phylum Platyhelminthes): Body flattened dorsoventrally; true suck¬ ers absent; mouth usually opening into muscular pharynx; posterior organ of attachment (haptor) bear¬ ing chitinoid hooks or clamps; external parasites, except some species found in mouth, stomach, and intestine, and one found in urinary bladder. 12

6. Trematoda (Phylum Platyhelminthes): Digenetic; body flattened dorsoventrally; oral and ventral suckers present except in gasterostomes and Sanguinicola; only a weak oral sucker in Nematobothrium. Young metacercariae possibly resembling adults or cercariae; some developing metacercariae undergoing a "reorganiza¬ tion" stage, in which they appear inflated and may not resemble typical trematode. 7. Cestoidea (Phylum Platyhelminthes): Body flat¬ tened dorsoventrally; some adults segmented; head (scolex) typically bearring suckers, hooks, or suctorial grooves, occasionally no organs of attachment. Larvae containing microscopically conspicuous calcareous con¬ cretions. 8. Nematoda (Phylum Nemathelminthes): Body cylin¬ drical with rigid cuticle, one or both ends attenuated; no organs of attachment. 9. Acanthocephala (Phylum Acanthocephala): Body cylindrical, sometimes slightly flattened; spectacular hook-bearing eversible proboscis present. 10. Gordiacea (Phylum Nematomorpha): Not true fish parasites; may be recovered from stomach and body cavity of fishes; parasites of insects; resembling nema¬ todes except for extreme length; both ends blunt rather than attenuated; body surface ornamented. 11. Turbellaria (Phylum Platyhelminthes): Small cil¬ iated flatworms with eye spots, sometimes attacking fish in municipal aquariums, probably mostly marine. 12. Hirudinea (Leeches) (Phylum Annelida): Some flattened dorsoventrally, some only slightly; body seg¬ mented; anterior and posterior suckers larger or smaller than body diameter; external parasites.

Brief Descriptions of the Groups of Fish Parasites

13. Phyla Copepoda, Branchiura, and Isopoda: Louse¬ like (Argiilus), wormlike (Lemaea), or grublike (Salmincola, Ergasilus); external parasites. 14. Glochidia (Larvae of Pelecypod Clams): Larval clams encysted in fins and gills; bivalve shell, usually armed with hooks. 15. Linguatulid Larvae (Phylum Arthropoda, Order Pentastomida): Larvae of this reptilian parasite are

13

sometimes found in fishes in the southeastern United States. 16. Larval Mites (Phylum Arthropoda): Rarely found encapsulated in skin, gills, and esophagus of fishes. 17. Insect Predators (Phylum Insecta). 18. Plant Seeds, Pollen, and Diatoms (Plant King¬ dom): Occasionally caught in the mouth or gills of fishes.

Algae and Fungi (Kingdoms Protista and Fungi)

Algae Algae as well as protozoa are in the kingdom Protista. Algal parasites of North American freshwater fishes are probably rare; there are few published records of them. For a general account of the algae, refer to Bold and Wynne (1985). A dinoflagellate, Piscinoodinium pillulare (Oodinium limneticum Jacobs, 1946), was described from fishes in Minnesota. This species is discussed under protozoa. A filamentous green alga, Cladopliora sp., has been described from the opercula of black bass (Microptems sp.) (Vinyard, 1955). The alga probably started growing on bone that was exposed after spawning activity. I have seen similar growth on the opercula of rainbow trout that had been kept in concrete ponds for two or three years. Miller and Ballantine (1974) described opercular algal growth on Tilapia aurea cultured in seawater. Stigeoclonium (filamentous) and a species of Chlorococcales (unicellular) were found growing subepidermally within the nasal capsule and below the frontal bones of a kissing gourami (Helostoma temmincki) (Nigrelli et al., 1958). The latter alga grew visibly in three months after being injected subdermally into a young gourami. A unicellular alga was reported, but not named, from the gills and surprisingly in the visceral organs of a green swordtail (Xiphoplwnis helleri) and a kissing gourami from a tropical fish farm in Florida (G. L. Hoffman et ah, 1960). The fish farm was losing many fishes; extensive invasion by the alga was probably a major cause of mortality. Nine cysts, 0.29-1.16 mm in diameter, of Chlorella sp. (Chlorophyta, or green alga) were found in the eye

14

orbit of a bluegill (G. L. Hoffman et ah, 1965). The inte¬ rior of each cyst was filled predominantly with Chlorella (Fig. 1), a unicellular, round, smooth alga further char¬ acterized by having a single, cup-shaped green chloroplast and usually lacking pyrenoids. Multiplication of Chlorella is by autospores—2, 4, 8, or 16 of which are formed within a cell and liberated by the rupture of the mother cell wall (Fig. lc,d). Individual cells observed in the bluegill were 7 to 10 pm in diameter. The cysts contained a lesser amount of blue-green algal growth, apparently Phormidium mucicola. Two years later, simi¬ lar cysts were found in the gills of a bluegill from the same pond. These were successfully transferred to other fish by injection. Myxonema tenue, a filamentous green alga, was found growing on Haplochilus latipes in Japan (Minakata, 1908). Other European algae of fishes are discussed in Schaperclaus (1954, 1986). Parasitic algae, Cladopliora sp. and Chlorella sp., were found invading common carp by Edwards (1978), who also reviewed parasitic algae in fishes. Achloric unicellular algae of the genus Prototheca have caused disease in fishes (R. D. Baker, 1971; Gentles and Bond, 1977) and other animals, including humans (Kaplan, 1977). For many years there have been scattered references to a small, nondescript, fish disease-causing alga known as Mucophihis cyprini (mucophilosis) (Lucky, 1970; Sapozhnikov et al., 1974), but Molnar and Boros (1981) showed with electron microscopy that these mucophilus cysts contain rickettsia- or chlamydia-like organisms. I am unaware of a satisfactory treatment of algal dis¬ eases of fishes.

Fungi

15

cases it would be wise to seek the aid of a mycologist. Some species must be cultured for identification.

Fungi Fungi are nonmotile, filamentous organims that lack chlorophyll; the assimilative phase consists of a true plasmodium or mycelium, or rarely of separate unicellular, independent cells, not amoeboid, and at no time unit¬ ing as a plasmodium-like structure. Only those fungi asso¬ ciated with North American freshwater fishes will be considered; the serious student of aquatic fungi, includ¬ ing fungi of fishes, should consult Alderman (1982), Neish and Hughes (1980), Olah and Farkas (1978), Spar¬ row (1960, 1973), or Srivastava (1980, 1987). In many

Class Saprolegniales Mycelium, if present, usually continuous in active assim¬ ilative phase (nonseptate); if lacking, reproduction not by budding; perfect (sexual) stage usually represented by oospores or zygospores; imperfect (asexual) stage by sporangiospores or modified sporangia, less commonly by true conidia. (text continues on page 17)

-1. ChlorellaChlorella sp. from eye orbit of bluegill (from G. L. Hoffman, et al., 1965): a, cyst containing many algal cells; b, single cell showing cup-shaped chloroplast and starch bodies; c and d, dividing stages showing mother cell wall.

sporangium and zoospores

2. Saprolegnia FIG. 2. Saprolegnia sp.: top, sporangium with many flagellated zoospores; middle, an old sporangium with a new one proliferating into it, a characteristic of the genus; bottom, sex organs, antheridium on left, oogonium on right, usually found only on special media. FIG. 3. Large lesions of Saprolegnia sp. on Micropterus dolomieui (from G. L. Hoffman and Meyer, 1974).

-FungiFIG. 4. Exophiala pisciphilus from culture. Conidia in slimy mass at apex of phialide (reprinted by permission from McGinnis and Ajello, 1974, Mycologia 66: 518-520, copyright The New York Botanical Garden). FIG. 5. Ochronis humicola conidia from slide culture (from Ajello et al., 1977, reprinted by permission of Kluwer Academic Publishers, The Netherlands). FIG. 6. Phoma herbarum hyaline conidia from slide culture (from Ross et al., 1975). FIG. 7. Ichthyophonus hoferi brain infection of Poeciliopsis occidentalis. Note exophthalmia. FIG. 8. /. Iwferi in brain of P. occidentalis, sagittal section. FIG. 9.1. hoferi spores from brain of P. occidentalis. FIG. 10. 7. hoferi branching body from viscera of Oncorhynchus mykiss.

Fungi

17

12. Unknown -FungiFIG. 11. Ichthyophonus hoferi spores from Oncorhynchus mykiss showing variation in size and "hyphal proliferation." FIG. 12. Bodies of variable sizes from unknown fungi resembling Ichthyophonus hoferi, in gills of Notemigonus crysoleucas, AR. FIG. 13. Branchiomyces demigrans from gills of Esox Indus showing burst cell tube with "escaping" large and small spores and other proliferated tubes (redrawn from Wundsch, 1930). FIG. 14. Branchiomyces sanguinis in gill capillary (photo by the late Pietro Ghittino, Turin, Italy).

Genus Saprolegnia and Related Genera (Figs. 2, 3) Saprolegnia species are usually implicated in fungus dis¬ eases of fish and fish eggs, although Achlya, Aphanomyces, Leptomitus, and Pythium have also been reported. These fungus diseases of fishes are often considered primary or secondary invaders following injury; once they start growing on a fish, however, the lesions usually con¬ tinue to enlarge and may cause death unless medication is provided. Fungi often attack dead fish eggs and soon encompass adjacent live eggs, which are attacked and killed. Fungi thus constitute one of the most important egg diseases. These fungi grow on many types of decay¬ ing organic matter and are widespread in nature. — Ref. Dick (1973), Scott (1964), Scott and O'Bier (1962), Scott and Warren (1964), Seymour (1970), Willoughby (1978, 1985). Morphology, Identification, and Life Cycle The presence of fungus on fish or fish eggs is indicated by a white cottony growth that consists of a mass (mycelium) of nonseptate filaments (hyphae), each of which is about 20 pm in diameter. Under low magnifi¬ cation, the filaments of older infections may be seen ter¬ minating in clublike enlargements that contain the flagellated zoospores (Fig. 2). These zoospores eventually escape and infect other fishes or eggs. Fish eggs. During incubation, some eggs die and may soon be invaded by a fungus. Surrounding eggs then

become covered by the mycelium and eventually die. Unless control measures are used, the ever-expanding growth will kill virtually every egg. Kanouse (1932) found circumstantial evidence that, under some condi¬ tions (probably crowding), fungus filaments can pene¬ trate living eggs. Under experimental conditions, living eggs that were in no way crowded were not invaded. H. S. Davis (1953) found no evidence that Saprolegnia can develop on normal eggs unless foreign organic matter is present. Fish. Injuries produced by spawning activity and other trauma, or lesions caused by other infections, are often attacked (G. L. Hoffman, 1949; Scott, 1964; Scott and O'Bier, 1962; Scott and Warren, 1964; Vishniac and Nigrelli, 1957). Conditions such as holding warm-water fishes in cold water during the summer and debilitation caused by other factors probably render fishes suscep¬ tible to attack by a fungus. Fungi that invade under these conditions belong to the Phycomycetes and are typified by a nonseptate mycelium. Phycomycetes belonging to the Saprolegnia family (Saprolegniaceae) are characterized by having club-shaped sporangia containing zoospores (Fig. 2) and round oogonia (sexual reproductive structures). The latter can be obtained only by culturing on special material (Bailey, 1983a, 1983b; Cooke, 1954; G. L. Hoff¬ man, 1949; Neish, 1975; Scott and O'Bier, 1962; Scott et al., 1963; Willoughby, 1985). Species can be identified only after studying these structures. The other genera mentioned above can be likewise identified, but Scott (1964) and Scott and O'Bier (1962) should be consulted.

Algae and Fungi

18

Depending on the temperature, 24 to 48 hours are required to complete a life cycle from reproductive spore to mycelium to reproductive spore. Transmission and Pathogenicity

Phycomycetes grow on many types of decomposing organic matter, and the resulting asexual reproductive spores are ubiquitous in natural waters. Dead eggs and fish are generally susceptible to inva¬ sion by fungi. Under favorable conditions, healthy eggs resist fungus invasion, and at times dead eggs do not suc¬ cumb to penetration for many days after turning white. Infertile eggs can remain resistant to fungal invasion for a month or longer when incubated in the presence of Saprolegnia and its relatives. Treatment is discussed by Alderman (1982a, 1985); Bailey (1983a, 1983b), G. L. Hoffman (1963), G. L. Hoff¬ man and Meyer (1974), Scott and Warren (1964), Willoughby (1985). Recently, it has been shown that bac¬ teria may have antagonistic action against Saprolegnia and other fungi on fishes (Haitai and Willoughby, 1988; Olah and Farkas, 1978; Willoughby, 1983). Geographical Range

Saprolegnia and its relatives are present throughout North America. There are no known seasonal restrictions to infesta¬ tion of fish eggs by Saprolegniaceae, but fishes are more likely to be affected in early spring (temperate zone) and after spawning activity.

Deuteromycetes (Fungi Imperfecti)

These are forms for which no sexual reproduction has been found—hence "imperfect."

Blastomycetes (Yeasts) Candida spp. (yeast) have been found in fishes that were fed rations containing large numbers of yeasts. Reports have come from the United States, Japan, and the for¬ mer Soviet Union, but true pathogenicity seems to be controversial (Neish and Hughes, 1980). There have been reports of yeast in fish disease from the United States, England, and the former Soviet Union.

Hyphomycetes Characterized by segmented hyphae that are either ster¬ ile or bear conidia (asexual spores) not contained in a dis¬ crete fructification. Thought to occur in fishes when present in large numbers in the feed or elsewhere in the environment. Not true parasites of fish; nevertheless, able

to cause disease and sometimes high mortality. Identi¬ fication possible by studying conidia in cultures. ■ Exophialapisciphilus McGinnis and Ajello, 1974 (Fig. 4). Originally identified as E. salmonis by Fijan (1969) from material collected during an epizootic that occurred among channel catfish in Alabama. In cultures, conidia (asexual spores) are nonseptate, 2-3 x 3-5 pm, subglobose to ovoid. Fishes show round to irregular exter¬ nal ulcers 2-25 mm in diameter, soft nodules in the viscera, and an exudate. Ictalurus punctatus, L catus, and Lepomis macrochirus have been affected. Exophiala pisciphila was recently found in a cranial infection of Salmo salar in Australia (Langdon and McDonald, 1987). ■ E. salmonis Carmichael, 1966. Originally described from cerebral mycetomas from Salmo clarki, Alta. In culture, conidia are typically singly septate and cylin¬ drical, about 3 x 5-8 pm, and quite distinct from E. pisciphila. Exophiala salmonis has also been found in visceral mycetomas in Salmo salar in Scotland (Richards et al., 1978), ME, and N.B. (Otis et al., 1985).

Genus Ochroconis de Floog and von Arx, 1973 (Syn. Scolecobasidium, in part) (Fig. 5) In culture, Ochroconis spp. produce an ellipsoidal to cylindrical sympodioconidium with a broadly rounded tip and a conspicuous hilum (narrowing) at the conidial base, which marks the point of attachment to a denticle on the conidiogenous cell. ■ Ochroconis humicola (Barron and Busch, 1962) de Hoog and von Arx, 1973: redescription of the genus; Ajello, et al., (1977): pathogen of rainbow trout, surface lesions on kidney, other internal organs, TN; Elkan and Philpot (1973): pathogen of frogs, England; Ross and Yasutake (1973); pathogen of young coho salmon, mostly in the kidney, WA. Culture at 25°C on potato dextrose agar or Sabourand's dextrose agar, etc., where the dematiaceous hyphae bear conidiophores, usually unbranched, 1.5-2.5 x 5.0-30.0 pm in size. Pale to olive brown; two-celled; finely echinulate conidia developing sympodially from the condiophores. Conidia oblong and rounded at both ends, with a truncated protuberant hilum. Conidia 2.5-4.0 x 6.0-11.5 pm. ■ Ochroconus tshawytschae (Doty and Slater, 1946) McGinnis and Ajello, 1974 (Syns., according to McGin¬ nis and Ajello, 1975: Heterosporiwn tshawytschae Doty and Slater, 1946; Scolecbasidium macrosponnn Roy et al., 1962; S. variabile Barron and Busch, 1963): infections mostly in posterior kidney, Oncorhynchus tshawytschae, CA. In culture at 25°C, conidiogenous cells sympodial, conidia septate (1-3), conidia 3.5 (2.2-3.6) x 13.6 (9.0-19.8) pm.

Fungi

Coelomycetes Fungi imperfecti with conidia produced in some type of fructification—pycnidia, acervuli, and stromata. ■ Phoma herbarum (Westendorp, 1852) Boerema, 1970 (Fig. 6): isolated from soil, water, milk, butter, paint; Ross et al., (1975): found as a visceral disease agent primar¬ ily in fry and fingerlings of Oncorhynchus kitsutch, O. tshawytscha, O. mykiss (Salmo gairdneri) from 10 national fish hatcheries in the Pacific Northwest, USA. In culture, aseptate hyaline conidia borne in dark pycnidia.

Higher Taxon Unknown Genus Ichthyophonus

19

have been obtained (McVicar, 1982; Okamota et al., 1984). Spores have been examined using electron microscopy (Paperna, 1986). The organism is commonly found in the kidney, spleen, liver, heart, stomach, intestine, visceral serosa, peritoneal exudate, gills, and brain. In the only severe epizootic recorded of rainbow trout, the spores were numerous in the brain and its cavities, as well as in the musculature. Central nervous system involvement appar¬ ently resulted in partial denervation of the skeletal mus¬ culature, which caused spinal curvature. In one interesting epizootic, many spores were found only in the brain and spinal column of the endangered Gila topminnow, Poeciliopsis occidental^, in NM. The brains were so severely infected that obvious exophthal¬ mia resulted (Figs. 7-9) (G. E. Hoffman, unpub., 1982).

(Figs. 7-11) Diagnosis

■ Ichthyophonus hoferi Plehn and Mulsow, 1911 (Syn. Ichthyosporidium hoferi). This fish fungus was consid¬ ered to belong to the Phycomycetes by Sproston (1944). Others have avoided trying to place the parasite because it is not a typical member of any class of fungi. Sprague (pers. comm., Univ. of Maryland, 1965) states that the generic name Ichthyophonus has priority over Ichthyo¬ sporidium, which is a protozoan. McVicar (1982) reviewed the genus. Ichthyophonus hoferi and Ichthyosporidium sp. have been reported from Atlantic Ocean fishes (cf. Sindermann and Scattergood, 1954), West Coast marine fishes (Hendricks, 1972), Australian Atlantic fishes (Machado-Cruz, 1960), golden shiner gills (C. Wilson, 1989), European freshwater fishes (cf. Schaperclaus, 1954), aquarium fishes (Reichenbach-Klinke, 1957), North American rainbow trout (J. D. Erickson, 1965; Gustafson and Rucker, 1956; Ross and Parisot, 1958), and amphibia (Elkan and Reichenbach-Klinke, 1974). It is not known whether these records concern one species or several. ■ Ichthyophonus (Ichthyosporidium) in rainbow trout in North America was first reported by Rucker and Gustaf¬ son (1953) from three localities in western WA. Ross and Parisot (1958) reported it from hatcheries adjacent to the Snake River in south central ID. The parasite may have spread where hatchery personnel fed trout uncooked trash fish and the raw viscera of dressed trout. If it is true that the trash fish in the Snake River watershed are infected, it is probable that the parasite has become well-established in freshwater. Morphology and Life Cycle

The stage usually seen is the resting spore; spherical, 10-200 pm in diameter, granular, opaque to translucent, with relatively thick walls. Germinating spores are usu¬ ally not seen in fresh material, but after death of the host the spores tend to germinate, producing single then branched hyphae (McVicar, 1982). Successful cultures

Apparently no other spore-forming parasite of fishes produces spores with such extreme variation in size (10-200 pm). The internal portion of the spore appears granular and amorphous in light microscopy. Nuclei can be seen only with special staining or electron microscopy. The spores are usually numerous enough for diagnosis, but digest techniques can be used in cases of low inten¬ sity (Ghittino, 1978). Pathology

Most of the spores become encapsulated in small host cysts or granulomas, which have been studied in detail by McVicar and McLay (1985). Control

No chemotherapy is known. Prophylaxis consists of sanitation and not feeding raw fish products. ■ Ichthyophonus intestinalis (Basidiobolus intestinalis Leger and Hesse, 1923). This intestinal form reported from Salmo trutta of Europe resembles I. hoferi, but the spores are much smaller. The late Ed Dunbar and I saw it in 1960 from the intestine of Salvelinus fontinalis from the Berlin, New Hampshire, National Fish Hatchery. It is not considered pathogenic.

Ichthyophonus-like "thing" (Fig. 12)

During my tenure (1974-1985) at the USFWS laboratory in Stuttgart, AR, Fred Meyer, Drew Mitchell, and I saw variably sized, encapsulated spheres containing appar¬ ently amorphous material in the gills of Pimephales pronielas and Carassius auratus. We could never relate them to anything else, except possibly Ichthyophonus, epitheliocystis, or lymphocystis.

20

Algae and Fungi

Genus Branchiomyces (Figs. 13, 14)

■ Branchiomyces sanguinis Plehn, 1912. The nonseptate filaments, 9-15 pm in diameter, of this fungus invade the blood capillaries of the gills and eventually produce masses of spores 5-8 pm in diameter. These masses, often forming branchial arrangements, are very obvious in gill squash preparations (Fig. 13). Because of resultant necro¬ sis, the disease has been named “gill rot" or “foul gill." This species has been reported in Europe from ten fam¬ ilies of fishes, including Salmonidae (Neish and Hughes, 1980). It has also caused epizootics of eels, Anguilla japonica, in Taiwan (Chien et al., 1979), and Japan (Egusa and Ohiwa, 1972), and in Channa marulius in India (Srivastava and Srivastava, 1976). In the United States it has been found in Centrarchidae: Lepomis gibbosus, L. macrochirus (Wolke, 1975); Micropterus dolomieui (Rogers, pers. comm., Auburn Univ., 1973); M. salmoides (F. Meyer and Robinson, 1973); Cyprinidae: Carassius auratus, Notemigonus crysoleucas (G. L. Hoffman and Mitchell, 1978, 1980); Plumb, pers. comm., Auburn Univ., 1981); Percichthyidae: Morone saxatilis (F. Meyer and Robinson, 1973); Percidae: Stizostedion vitreum (Horner, 1985); Poeciliidae: Poecilia reticulata (G. L. Hoffman, unpub., USFWS, 1972); Siluridae: Ictalurus punctatus (Plumb and Rogers, 1979). Thus, either this parasite has a natural worldwide distribution in temperate-subtropical areas or it has been transferred with fish shipments. Epizootiology

This prolific organism has little host specificity and exists widely in freshwaters, but it does not become

apparent until the water warms (20°C or higher) and becomes enriched with organic matter from decom¬ posing plants, fish food, etc. During an epizootic it kills about 30% of the fishes, with the remainder making a complete recovery. It is neither highly contagious to fishes in adjacent ponds nor usually recurrent in the same pond—a disease thus requiring a rare combination of numerous factors. Epizootics also occur in the wild. Grimaldi (1971) reported a massive kill (20-50 tons) of Albumus alborella (Cyprinidae) due to Branchiomyces sp., probably B. sanguinis, in Lake Lugano in northern Italy, with lesser kills in two nearby lakes. The epizootics were associated with warm temperatures (above 20°C) and algal blooms. Giussani and Ghittino (1980) reported a similar natural epizootic in a different lake in northwest Italy; many Scardinius erythrophthalmus and Tinea tinea were killed. Additional information can be found in O. N. Bauer et al. (1969) and Schaperclaus (1986). ■ Branchiomyces demigrans Wundsch, 1929. This species differs from B. sanguinis in that there is little develop¬ ment in the gill capillaries, which contain only a few thick fungal filaments. There is, however, striking growth of the fungus on the exterior of the gill tissue. Exten¬ sive necrosis of gill tissue does not occur (Wundsch, 1930). The hyphal walls are much thicker (0.5-0.7 pm vs. 0.2 pm for B. sanguinis) and the spores are larger (12-17 pm vs. 5-9 pm for B. sanguinis) (Neish and Hughes, 1980). Prior to 1973, B. demigrans had been reported only from European Esox Indus and Tinea tinea, but in 1973 there was an epizootic of B. demigrans that killed 20,000-25,000 E. Indus, 14 to 24 inches long, in the Fox River, WI (Warren, 1973).

Subkingdom

Protozoa (Kingdom Protista) (Included in the Eukaryotic Protists or Microbes by some modern protozoologists)

■ Key to the Groups of Protozoa . . .22 Phylum Rhizopoda

.22

Subphylum Amoebozoa.22 Order Unknown .23 Genus Schizamoeba.23 Facultative Parasitic Amoebae . . .23 Laboratory Cultivation and Identification of Free-living and Facultative Parasitic Amoebae.23 Order Amoebida .25 Genus Acanthamoeba .25 Genus Paramoeba.25 Genus Thecamoeba.25 Order Schizopyrenida .25 Genus Naegleria .25 Genus Vahlkampfia .25 Phylum Mastigophora

.25

■ Key to the Genera of Mastigophora .26 Class Dinoflagellida.26 Genus Amyloodinium.26 Genus Crepidoodinium.26 Genus Piscinoodinium.29 Class Euglenida .29 Genus Euglenosoma.29 Class Kinetoplastida .29 Genus Cryptobia .29 Genus Ichthyobodo .30 Genus Lamellasoma.30 Genus Trypanoplasma .30 Genus Trypanosoma.31 Order Proteromonadida .31 Genus Karatomorpha.31 Class Diplomonadida.32 Genus Hexamita .32 Genus Spironucleus .32 Subphylum Opalinata .32 Genus Protoopalina.33 Subclass Suctoria.33

Phylum Ciliophora.33 Subclass Suctoria.33 Genus Trichophrya.33 ■ Key to the Genera of Ciliophora of Fishes.34 Nonsuctorian Ciliates .40 Subclass Trichostomatia.40 Genus Balantidium.40 Subclass Heterotrichia.40 Genus Nyctotherus.40 Subclass Hymenostomata .40 Genus Ichthyophthirius.40 Genus Ophryoglena.41 Genus Tetrahymena.42 Subclass Phyllopharyngia.42 Genus Chilodonella .42 Class Litostomatea .43 Genus Amphileptus .43 Subclass Peritrichia .43 Suborder 1. Sessilina .43 ■ Key to the Genera of Sessile Peritrichs .43 Genus Ambiphrya .44 Genus Apiosoma .44 Genus Carchesium.44 Genus Epistylis.44 Genus Heteropolaria.45 Genus Zoothamnium.45 Suborder 2. Mobilina.45 Family Trichodinidae .45 ■ Key to the Genera of Urceolarids of Fishes . . . .46 Subgenus Foliella.46 Genus Paratrichodina.46 Genus Trichodina.46 Genus Trichodinella .48 Genus Tripartiella.49 Genus Vauchomia .49 Group Sporozoa

.49

■ Key to the Groups of Sporozoa

. .49

Phylum Apicomplexa

.50

Subclass Coccidia.50 Order Eucoccidiida.50 Suborder Eimeriina.50 ■ Key to the Genera of Eimeriina of Fishes .50 Genus Calyptospora.50 Genus Eimeria.50 Genus Epieimeria.53 Genus Goussia.54 Genus Haemogregarina ... .54 Family Sarcocystidae .55 Genus Sarcocystis.55 Family Dactylosomatidae . . . .55 Genus Babesiosoma.55 Genus Dactylosoma.55 ■ Key to Sporozoa Other Than Coccidia and Haemosporidia Found in Fishes.55 Phylum Myxozoa: Class Myxosporea.55 Order Bivalvulida.56 Suborder Platysporina . . . .56 Family Myxobolidae.56 Most Likely Sites of Myxobolus Species .56 ■ Key to the Genera of Myxosporea of North American Freshwater Fishes.57 Genus Dicauda .64 Genus Facieplatycauda.64 Genus Fienneguya .64 Genus Myxobolus.66 Genus Thelohanellus .72 Genus Unicauda .-..73 Suborder Variisporina.73 Family Ceratomyxidae.73 Genus Ceratomyxa.73 Family Chloromyxidae.73 Genus Agarella .73 Genus Caudomyxum .74 Genus Chloromyxum .74

(continued) 21

22

Subkingdom Protozoa

Family Myxidiidae . Genus Myxidium . Genus Zschokkella. Family Ortholineidae. Genus Neomyxobolus. Family Parvicapsulidae. Genus Parvicapsula . Family Sphaerosporidae . . . . Genus Acauda. Genus Hoferellus . Genus Myxobilatus. Genus Sphaerospora. Proliferative Kidney Disease (PKD) of Salmonids . . Genus Wardia .

.75 .75 .77 .77 .77 .77 .77 77 .78 .78 .78 .79 .80 .80

Order Multivalvulida . Family Kudoidae. Genus Kudoa. Family Pentacapsulidae . . Genus Pentacapsula ....

. . . . .

. . . . .

.80 .80 .80 .81 .81

Class Actinosporea . Order Actinomyxida . Family Triactinomyxidae Genus Triactinomyxon . .

. . . .

. . . .

.81 .81 .81 .81

Phylum Microspora. . . .81 Class Microsporea. . . .82 Order Pleistophoridida .... . . .82 Family Pleistophoridae . . . . . .82

At present, protozoa probably cause more disease in fish culture than any other type of animal parasite. In some instances, perhaps under environmental circum¬ stances unfavorable to the fish, large populations of external protozoa appear on the body and gills of the fish. When present in small numbers, these protozoa usually produce no obvious damage, but in large numbers they greatly impair the epithelium, particularly of the gills. Some of the protozoa actually feed on the cells and mucus of the fish; others that attach to the fish but do not feed on it may injure the host by blockage, especially to the gills. One external protozoan, Ichthyophthirius multifiliis, produces severe devastation by burrowing under the epithelium. Ichthyobodo (Costia) and some species of Trichodina cause "blue slime" of the body. The possible damage done by internal protozoan fish parasites has not been adequately studied. Myxosporea have been found in most organs of fishes and can be very harmful. The blood flagellate, Trypanoplasma (Cryptobia), and the sporozoan, Haemogregarina, occur in the blood and are transmitted by leeches. Few protozoans have been found in the intestine; these include one fla¬ gellate, Hexamita (Octomitus), one cililate, Protoopalina, one amoeba, Schizamoeba, two sporozoans, Eimeria and Goussia, and one ciliate, Protoopalina. Fishes in nature are infected with a great variety of pro¬ tozoa. Disease resulting from these infections has not been reported often, for several possible reasons: (1) certain stages of the parasites are dispersed in a large vol¬ ume of water, and fishes are therefore not heavily par¬ asitized, (2) the parasites, even if present in large numbers, do little harm, or (3) the most severely affected fishes die, or are weakened, then killed by natural preda¬ tors, and therefore not seen. The most comprehensive works that include proto¬ zoa in fish culture are Bykhovskaya-Pavlovskaya (1962), H. S. Davis (1953), Dogiel et al. (1958), Ergens and Lorn (1970), Jahn et al. (1979), Lorn and Dykova (1992), Schaperclaus (1986), and Shulman (1984). For a key to protozoa of North American fishes, see Wellborn and Rogers (1966). For general descriptions of protozoa, as well as of specific organisms, refer to Kudo (1971) and Lee et al. (1985).

Genus Enterocytozoon .82 ■ Key to the Genera of Microsporidians in Fishes 82 Genus Glugea .86 Genus Heterosporis .87 Genus Loma .87 Genus Pleistophora.88 Genus Thelohania .89 Collective Group Microsporidium .89 Sporozoa of Uncertain Classification .89 Genus Dermocystidium.89

Key to the Groups of Protozoa (See also Wellborn and Rogers, 1966) Trophozoite with pseudopodia (forms cysts in intestine) .Subphylum Amoebozoa, P- 22 Trophozoite with flagellum or flagella .Phylum Mastigophora, P- 25 Trophozoite with cilia.Phylum Ciliophora, p. 33 Trophozoite with tentacles .... See no. 1 of Ciliophora, p. 33 Usually lacking cell organs of locomotion, but some myxosporidean trophozoites possessing very sluggish pseudopodia; producing spores .Unofficial group Sporozoa, p. 49

For technique in studying protozoa, refer to the sec¬ tion on methods (Chemotherapy and Prophylaxis). The classification of the Society of Protozoology (Lee et al., 1985) has been followed wherever possible. Certain protozoa from other parts of the world are included for reference, in the event that related forms are discovered in North America.

Phylum Rhizopoda This phylum was proposed by Honigberg and Balamuth (1963) and, in spite of unwieldy proportions and dubi¬ ous relationships, has been tentatively accepted by protist taxonomists. The grouping is based primarily on Pascher (1918), who hypothesized that all amoeboid organisms evolved from a flagellate by loss of flagella. So this group tentatively contains subphyla Sarcodina, Mastigophora, and Opalinata.

Subphylum Amoebozoa Liihe, 1913 The members of this group formerly called Sarcodina do not possess a thick pellicle and are capable of forming

Subphylum Amoebozoa

pseudopodia. The presence of pseudopodia is one of the most useful criteria for their identification. The trophozoites of some Myxosporidea have sluggish pseudopodia. Sarcodina that are parasitic in the intestinal tract usually feed on bacteria and other organic matter. Those parasitic on the gills of fishes probably feed on bacteria and organic matter trapped between the gill lamellae. The life cycle of the species in the stomach of trout involves a trophozoite that produces cysts that are passed out with the host feces. The new host is infected when the cysts are accidentally ingested. Those sarcodinids that develop on fish gills probably also produce cysts, but these species have not been well studied. The organism causing proliferative kidney disease (PKD) was long thought to be an amoeba but is now listed among the Myxosporea as an uncertain species. For a complete description of the Sarcodina, see Kudo (1971, p. 496) and Lee et al. (1985).

Order Unknown Genus Schizamoeba Davis, 1926 (Fig. 15)

Trophozoite nucleus vesicular, without endosome, but with large, discoid granules along nuclear membrane; one to many nuclei present; cyst nuclei with large endosome. One species. ■ Schizamoeba salmonis Davis, 1926. Trophozoite in mucus of stomach of trout; cysts from intestine seen more often. Common in young hatchery trout but apparently not pathogenic (H. S. Davis, 1926, 1946, p. 53).

Facultative Parasitic Amoebae Human amoebiasis, which is caused by usually freeliving amoebae that become pathogens after accidental contact with vulnerable tissues, has been known for some time (Martinez, 1985; Visvesvara, 1980). Acanthamoeba, for example, has recently been associated with keratitis of contact lens users (Newton et al., 1986). Another, Naegleria, has been known for some time as a cause of encephalitis. Thus, it is not surprising that sim¬ ilar amoebae become pathogenic to fishes under con¬ ducive environmental conditions. Few fish disease workers have had experience in identifying these amoe¬ bae; which may thus be more common than the reports indicate. For methods, see Cursons et al. (1979), Kent et al., (1988), and Page (1966). For culture methods and a key to species, see Page (1988). Laboratory Cultivation and Identification of Free-Living and Facultative Parasitic Amoebae (Courtesy of T. K. Sawyer, Rescon Associates, Inc., Royal Oak, MD 21662)

23

Recent work on free-living amoebae as potential pathogens has shown that some of them are capable of causing systemic infections as well as occurring externally on body surfaces. In contrast to obligate parasites, some amoebae are "opportunistic" pathogens that thrive on standard laboratory liquid and agar media. Species that form characteristic cysts may often be easily identified, whereas others may require attempts to obtain pure clonal strains and studies on stained specimens. Such methods are presently available, and excellent descrip¬ tions of many genera and species have appeared in recent years. Although attempts to culture fish tissues and body fluids for protozoa may not always be successful when parasitic species are present, such techniques are increasingly effective with "opportunistic" species. Free-living freshwater or soil amoebae are often pres¬ ent on the gills of fishes suffering from respiratory stress (Daoust and Ferguson, 1985; Kubota and Kamata, 1971; T. K. Sawyer et al., 1974, 1975). The first detailed description and identification of amoebae associated with gill disease was published by Chatton (1909), who described Amoeba (= Vahlkampfia) mucicola from marine fishes. Chatton (1909), and later Kent et al. (1988), who identified Paramoeba pemaquidensis Page, 1970, from the gills of coho salmon, Onchorhynchus kisutch, reared in seawater, have published descriptive accounts sufficient to justify amoeba identifications to genus and species. In contrast to most parasitic protozoans, which are readily identified in histologic sections, free-living amoe¬ bae usually do not retain characteristic locomotive, feeding, and resting features after fixation and staining in or on host tissues. In spite of this limitation, T. K. Sawyer et al. (1974) described Thecamoeba hoffmani from sectioned gills of salmonid fishes on the basis of obvi¬ ous ectoplasmic ridges and folds characteristic of the genus. Daoust and Ferguson (1985) recognized the prob¬ lems associated with amoeba identification in the absence of cultures of living specimens and did not attempt to provide a generic name for amoebae associated with proliferative gill disease of rainbow trout. These authors published excellent photographs of intact gills using both scanning electron and brightfield microscopy. Their scanning photomicrographs showed massive gill fouling by bacteria and amoebae, whereas histologic sections showed severe gill pathology but little evidence of fouling microorganisms. Fishes showing evidence of respiratory stress, espe¬ cially under conditions of poor water quality, unsea¬ sonably warm weather, or overcrowding, should be examined for evidence of severe gill disease. Recom¬ mended methods are as follows: 1. Examine small fragments of gill tissue for evi¬ dence of debris, diatoms, protozoans, and exces¬ sive mucus secretion. 2. Make preliminary microscopic observations with hanging drop preparations or by examining

Subkingdom Protozoa

24

3.

4.

5. 6.

culture dishes containing filtered water under an inverted microscope. Centrifuge culture fluids containing amoebae, and transfer the resulting pellet to freshly poured agar plates. Pour 1.5% non-nutrient agar into sterile plates, allow to harden, and streak with Escherichia coli or Klebsiella pneumoniae to provide the food source for the amoebae. Low nutrient agar may be prepared by adding 0.1 g malt extract and 0.1 g yeast extract/liter to the culture medium. Inoculate the culture plates with a centrifuged pellet or with small fragments of suspected tissue. Store cultures in covered containers lined with moist paper towels, and examine at frequent intervals.

Most soil amoebae, including Acanthamoeba, Hart¬ manella, Naegleria, and Vahlkampfia, form resistant cysts that aid in making preliminary identification to genus. Amoebae that do not form cysts should be subcultured to fresh plates at frequent intervals. Consult Page (1976) for information on culture methods, culture media and procedures for maintaining different genera of freeliving amoebae. An extensive listing of culture media, and special culture conditions for algae and protozoa are available from the American Type Culture Collection, 12301 Parklawn Drive, Rockville, MD 20852. Amoebae not readily identified on the basis of gross characteristics should be cloned and studied in both living and stained conditions. Clones may be prepared by streaking small blocks of agar with cultured amoebae across the surface of a fresh plate, making daily obser¬ vations, and marking the location of individual cysts or trophozoites. Second-step cultures can be made by trans¬ ferring the cyst or single colony of amoebae and repeat¬ ing the procedure five to six times in succession. Clones may also be obtained by diluting amoebae or cysts in liq¬ uid to concentrations of one or two organisms per ml and preparing 10 cultures, each inoculated with 0.1 ml of the 1.0 ml suspension. Cloning procedures can some¬ times be improved by inoculating plates at tempera¬ tures slightly lower than those that encourage rapid growth. Some species of Hartmanella, Naegleria, Vahlkampfia, etc., yield cysts after only several days of growth at ambient temperatures and may be more difficult to clone than those with slower growth characteristics. Some amoebae are affected by light passing through the microscope. In such cases they may contract, detach from the cover glasses used for preparing hanging drops, or fail to move in locomotive patterns useful for diag¬ nosis. Other amoebae, e.g., eruptive species such as Naegleria and Tetramitus, must be studied for their abil¬ ity to transform to amoeboflagellates; this will distinguish them from Vahlkampfia, which does not transform. Staining procedures to determine patterns of nuclear division are important to distinguish certain hartmanellids and vahlkampfids. The nuclear membrane and

nucleus disappear early in the mitotic cycle in eumitotic amoebae such as Acanthamoeba and Hartmanella, whereas both structures are retained throughout cell division in Naegleria, Vahlkampfia, and other vahlkampfid-like amoe¬ bae. Amoebae that are allowed to attach on slides or cover glasses prior to staining often show typical locomotive forms that are otherwise inhibited in specimens studied live under the microscope. Amoebae may be stained using most of the standard techniques described in books or laboratory manuals. Drops of water or liquid culture approximately 1 cm or less should be placed on slides or cover glasses, inocu¬ lated with small blocks of agar with amoebae facing down, and incubated for one hour or more in a moist chamber. Slides are more convenient to use than cover glasses, but the latter provide a smaller area to be scanned under the microscope. Most fixatives are suit¬ able, but the use of Nissenbaum's as described by Page (1976) yields fixed preparations in five seconds or less. Staining may be carried out with most described his¬ tological procedures, but the Feulgen procedure for DNA and Kernechtrot (nuclear red) as reported by Page (1976) and others, are adequate for most purposes. Either procedure will provide excellent nuclear and morphological details, especially when studied or pho¬ tographed with phase contrast optics. Formulas for Nissenbaum's fixing solution and Kernechtrot are pro¬ vided in the section on methods (Chemotherapy and Prophylaxis). Methods for cultivation and identification of freeliving amoebae from gills also are suitable for testing internal organs for the same or related species. Taylor (1977) cultured a species of Acanthamoeba from the spleen of red-eye bass, Microptenis coosae, and showed that the strain could be used to establish laboratory infections in experimental fishes. The strain isolated by Taylor (1977) is readily maintained on agar media and has been deposited at the American Type Culture Collection as strain ATCC #30807. Franke and Mackiewicz (1982) cultured species of Acanthamoeba and Naegleria from the intestinal contents of freshwater fishes and found that A. polyphaga successfully infected goldfish, Carassius auratus. Studies reported by Taylor (1977) and Franke and Mackiewicz (1982) suggest that future attempts to cul¬ ture organs other than gills may lead to the identifica¬ tion and description of more amoebae affecting fish health. T. K. Sawyer et al. (1978) described Vexillifera bacillipedes Page, 1969 from rainbow trout reared in hatch¬ eries in Italy. The amoeba, originally thought to cause PKD (proliferative kidney disease), grew well on agar media but did not form cysts and required frequent subculture; it was thought to be a contaminant from hatchery water and unrelated to kidney disease in this host. Attempts to culture nonprasitic or nonpathogenic amoebae such as Schizamoeba salmonis, originally described by H. S. Davis (1926, 1953), may lead to reports on other genera and species of well-known amoebae, and to descriptions of new ones. Nash et al.

Phylum Mastigophora

(1988) described a probable systemic amoebic infection in catfish, Siluris glanix L., in Europe. The authors pro¬ vided excellent illustrations of probable amoebae in sectioned tissue but did not determine whether the organisms were free-living or parasitic.

25

■ Thecamoeba hoffmani Sawyer, Hnath, and Conrad, 1974: on gill lamellae, Oncorhynchus kisutch, O. mykiss, O. tshawytsclia, MI, OR; smaller than most Thecamoeba, 22-42 pm in length during locomotion in stained sec¬ tions; pronounced longitudinal ridges or folds present in hyaline ectoplasm.

Order Amoebida Kent, 1880

Order Schizopyrenida

Naked, usually uninucleate (some multinucleate); with mitochondria; no flagellate stage.

Singh, 1952 Fimaciform (slug-shaped), more or less cylindroid, monopodial; clear, hemispherical, eruptive, with anterior bulges; typically uninucleate; nuclear division promitotic; temporary flagellated stages common; cysts variable.

Genus Acanthamoeba Volkonsky, 1931—emended Page, 1967 (No fig.)

Oval to triangulate, irregular; several to many slender, pointed pseudopodia, often from clear margin and body surface; extranuclear centrospheres; cysts polyhedral or convex; endocysts polygonal to stellate; with opercula. ■ Acanthamoeba polyphaga Franke and Mackiewicz (1982): cultured from intestinal contents of Catostomus commersoni, Notropis comutus, and experimentally infected Carassius auratus, NJ; Taylor (1977): on gills and in spleen, blood, urinary bladder, gall bladder, Carassius aureus, Micorpterus coosae, M. salmoides, Morone saxatilis, Oncorhynchus mykiss, Oreochromis aurea (Tilapia), south¬ eastern USA. ■ Acanthamoeba sp.(?). Nash et al. (1988): histopathology of a possible Acanthamoeba in Silums glanis, Germany.

Genus Naegleria Alexeiff, 1912—emended Calkins, 1913 (No fig.)

Flagellated stage lacking cytostome; two anterior fla¬ gella present; snail-like movement with lobopodia; vesic¬ ular nucleus with large endosome; contractile vacuole conspicuous; cysts uninucleate. ■ Naegleria sp. Taylor (1977): gills, Oreochromis nilotica (Tilapia), AT.

Genus Vahlkampfia Chatton and Lalung-Bonnaire, 1912 —emended Page, 1967 (No fig.)

Genus Paramoebo Schaudin, 1896 (No fig.)

Radiate body, 20-30 pm, pseudopods clear, conical, 8-90 pm long; nucleus spherical, 10-12 pm, endosome 5-6 pm, surrounded by granules; paranuclear body pres¬ ent; usually parasitic in crabs. ■ Par amoeba pemaquidensis Page, 1970: on gills of Onco¬ rhynchus kisutch reared in net pens in seawater, WA; included here because of the host's anadromous nature; typical of Paramoeba spp., the parasite had a Feulgenpositive paranuclear body adjacent to the nucleus; float¬ ing and transitional forms had digitiform pseudopodia; surface filaments were present on the plasmalemma; they measured 24-30 pm directly from gills; amoebae disappeared when fishes were held in reduced salinity. v

Genus Thecamoeba Shaeffer, 1926 (Fig. 16)

Usually large, up to 180 pm; usually oblong to triangu¬ late; uninucleate to multinucleate; dorsum wrinkled and/or ridged.

No flagellated stage; small amoebae; vesicular nucleus with large endosome and peripheral chromatin; snaillike movement with one broad pseudopodium; fresh¬ water or parasitic. ■ Vahlkampfia spp. Taylor (1977): gills and body mucus, Hypophthalmichthys molotrix, Micropterus salmoides, Ore¬ ochromis aureus, O. nilotica, Salmo trutta, AF, AR, FF, TN.

Phylum Mastigophora Diesing, 1866 (Flagellata)

The parasitic protozoa of this group possess one to sev¬ eral flagella that are usually visible, except on the attached forms of Ichthyobodo, Oodinium, and Euglenosoma. Some forms ingest food through the cytostome, and some absorb food in solution through their pellicles. Most species have one nucleus, but Hexamita spp. have two. The nucleus is vesicular and contains an endo¬ some. Contractile vacuoles are present in some species. Asexual reproduction is by longitudinal fission, and trypanosomes in culture divide to produce "rosettes"

26

Subkingdom Protozoa

that are composed of many individuals and eventually separate. — Ref. Kudo (1971, p. 303); Becker (1977); Lee et al. (1985).

Key to the Genera of Mastigophora

Flagella do not show on forms attached to fish. Comprises many free-living species, mostly marine; at least 55 species are parasitic in or on invertebrates. Three genera have been found on fishes. The following is based on the review of Lorn, 1981.

Genus Amyloodinium Brown and Hovasse, 1946

1. With chromatophores.Order Dinoflagellida 2 1. Without chromatophores. . . Order Zoomastigophorea 3 2. (1) With two flagella in free-swimming stage, one transverse; parasitic stage without flagella, but basal filipodia about 50 pm long .(Fig. 17) (Oodinium)

Piscinoodinium

2. (1) With two flagella in free-swimming stage, none transverse; globular on gills of crappies; one species.(Fig. 18)

Euglenosoma

3. (1) With one or two flagella.Order Kinetoplastida 4 3. (1) With three to eight flagella, which may not show on some attached forms . . . Order Polymastigina 8 4. (3) With one flagellum

.... Family Trypanosomatidae 5

4. (3) With two flagella.6 5. (4) In blood.(Fig. 24)

Trypanosoma

5. (4) On gills; several rows of small refringent rods running parallel to body axis. . . (Fig. 22)

Lamellasoma

6. (4) Trypanosome-like, but with two flagella, one anterior, and one posterior; in blood .(Fig. 23)

Trypanoplasma

6. (4) Not trypansosome-like, body more cylindrical;

(Syn. Oodinium Chatton, 1912, in part) (No fig.)

Ectoparasitic on gills and body of many marine fishes. Included here because of prevalence in municipal aquar¬ iums, etc. Trophont possessing attachment disc with very short peduncle; periphery of the disc radiating into long, filiform projections embedded deeply in host's epithelial cells. Special tentacle-like movable organ, the stompode, extending with the peduncle of the attach¬ ment disc from the basal end of the parasite; stompode possibly functioning in food intake and/or injection of lytic material into the host cells. No chloroplasts. Size of trophont up to 15 pm. Up to 256 gymnospores pro¬ duced in the palmella (tomont) stage; gymnospore with stigma (eye spot). ■ Amyloodinium ocellatum (Brown, 1931) Brown and Hovasse, 1946. As above. Bower (1987): standardized propagation method; Cheung et al. (1981a): scanning electron microscopy of development; Cheung et al. (1981b): in the kidney and other internal organs, Anisotremus virginicus.

one anterior flagellum, one trailing flagellum; on gills... 7

Genus Crepidoodinium

7. (6) With rod-shaped blepharoplast; body triangular; movement jerky.Cryptobia

agitans

7. (6) With round or oval blepharoplast; body elongate, movement not jerky.(Fig. 19)

Cryptobia branchialis

8. (3) With one nucleus and two anterior flagella; latter may not show on attached forms .(Figs. 20, 21) (Costia)

Ichthyobodo

8. (3) With two nuclei and eight flagella; in intestine of fish 9 9. (83) With movable posterior appendages.Urophagus 9. (83) Without movable posterior appendages; from salmonids, perhaps others .(Fig. 25) (Octomitus)

Hexamita

Class Dinoflagellida Butschli, 1885 With characteristics of the order (see Kudo, 1971, p. 370). The most striking features are the presence of yel¬ low, brown, and green chromatophores; a posterior fla¬ gellum; and a "girdle" with a transverse flagellum.

Lorn and Lawler, 1981 (see Lorn, 1981) (Syn. Oodinium Chatton, 1912, in part) (No fig.)

Ectoparasitic on gills of estuarine and marine fishes of Cyprinodontidae. Trophont attached by its basal end, flat¬ tened into a broad holdfast bearing "rhizoids" with fingerlike tips that closely adhere to host cell mem¬ branes without penetrating host cell. No stompode (foot mouth). Well-developed chloroplasts present. Nucleus with large interphasic chromosomes. Up to 670 pm in length. Up to 2048 gymnospores produced in the palmella (tomont) stage; gymnospore without stigma (eye spot). ■ Crepidoodinium cyprinodontinn (Lawler, 1967) Lorn, 1981 (Syn. Oodinium cyprinodontinn Lawler, 1967): Cyprinodon variegatus, Fundulus heteroclitus, F. magalis, Luanda parva, brackish water, VA. Lawler (1968): F. luciae; Lorn and Lawler (1973): electron microscopy of attachment; Lom (1981): cytoplasm curiously spongy, unlike Pisci¬ noodinium and other fish-invading dinoflagellates; also unlike other fish dinoflagellates, this parasite does not (text continues on page 29)

17. Piscinoodiniutn

15. Schizantoeba

Sarcodines -and flagellatesFIG. 15. Schizantoeba salmonis: a, stained trophozoite; b, stained cyst (from Davis, 1926). FIG. 16. Thecatnoeba hoffmani in histologic section of gill of Oncorhynchus kisutch, phase contrast (from T. K. Sawyer et al., 1974, by permission of the University of Wisconsin Press). FIG. 17. Piscinoodinium limneticum, parasite form (from Jacobs, 1946). FIG. 18. Euglenosoma branchialis: a, free form; b, attached form (from Jacobs, 1946). FIG. 19. Cryptobia branchialis (from Ergens and Lorn, 1970, by permission of the Czecho¬ slovak Academy of Sciences). FIG. 20. Ichtliyobocio necator: a, dorsal view; b, lateral view; c, attached (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences).

20. Ichthyobodo

25. Hexamita

E

n

o VO

■ If

27. Protoopalina

26. Spironucleus

-Flagellates and ciliatesFIG. 21. Ichthyobocio necator attached to fish skin (histologic section by the late H. S. Davis, USFWS). FIG. 22. Lamellasoma bacillaria (from H. S. Davis, 1947). FIG. 23. Trypanoplasma salmositica, from stained blood smear (from Katz, 1951). FIG. 24. Trypanosoma percae canadense (from Fantham et al., 1942). FIG. 25. Hexamita salmonis: a, schematic, f = flagellar pocket, o = orifice of flagellar pocket, pf = posterior flagellum (courtesy of Ergens and Lorn, 1970, by permission of the Czechosolovak Academy of Sciences); b, cyst; c, phase contrast, anterior flagella not showing (photo by the late Pietro Ghittino, Turin, Italy). FIG. 26. Spiromicleus elegans: pf = posterior flagellum (courtesy of Ergens and Lorn, 1970). FIG. 27. Protoopalina symphysodonis, schematic of body form, nucleus, and cilia from a living specimen (courtesy of Foissner et al., 1979).

Class Kinetoplastida

directly destroy host cells, so appears to be a symbiont rather than a true parasite; E. H. Williams (1972): Cyprin-

Class Kinetoplastida

odon variegatus, Fundulus similis, Gulf of Mexico.

Honigberg, 1963

29

—emended Vickerman, 1976

Genus Piscinoodinium Lorn, 1981 (Syn. Oodinium Chatton, 1912, in part) (Fig. 1 7) Ectoparasitic on gills and skin of freshwater fishes; rarely subcuticular. Trophont 12-96 pm in greatest diameter; numerous rodlike organelles, rhizocysts, radiating from

With characteristics of the order (see Levine et al., 1980). Plastic forms with one or two flagella.

Genus Cryptobia Leidy, 1846 (Fig. 19)

attachment disc, which possesses a very short peduncle; rhizocysts penetrating into and firmly embedded in the

Elongate to triangular protozoa with two flagella, one

epithelial cells of the host. Without stompode (foot

running backward along the body, adhering to the sur¬

mouth). Well-developed chloroplasts and starch grains present; theca (shell) without plates; mucocysts present

face, and extending posteriorly from the body; undula¬

subthecally. Palmella, without common envelope, pro¬ ducing up to 256 flagellated gymnospores without stigma. ■ Piscinoodinium limneticum (Jacobs, 1946) Lorn, 1981: skin of freshwater aquarium fishes; 12-96 pm in great¬ est diameter, MN; G. L. Hoffman (1959c): causes disease at aquarium fish hatcheries, FL. ■ P. pillulare (Schaperclaus, 1954) Lorn, 1981: on feral

directed forward and the other originating anteriorly but

tions of the flagellum not discernible on body surface, as in Trypanoplasma. Kinetoplast oblong and slender; Fib¬ ril formation (aciculum) present at anterior end. Most species on invertebrates, but two species found on fishes. Life cycle usually completed on the gills through simple division. ■ Cryptobia agitans (Colponema agitans H. S. Davis, 1947): 6-8 pm long, 3-4.5 pm wide; pyriform to oval, but often appearing triangular; trailing flagellum about 10 pm long; jerky movement unlike other species and

Carassius carassius Schaperclaus, 1951; Geus (1960): experimentally infected Cyprinus carpio, Leucaspis delineatus, Tinea tinea, and larval amphibians Amblystonia mexicanum, Rana temporaria, R. arvalis; Lorn and Schu¬

macrochirus, WV. Chen (1956b): description, Aristichthys nobilis, Hypophthalmichthys molitrix, China; G. L. Hoff¬

bert (1983): electron microscopy of the trophont;

man (unpub. USFWS, 1975-1979): on grass carp and

Reichenbach-Klinke and Braun (1968): disease in Oncorhynchus mykiss, Germany; Schubert (1974): general

goldfish, probably not pathogenic; Molnar (1979a): A.

discussion, 20-70 pm in greatest diameter.

■ C. branchialis Nie, 1955 (Bodomonas concava H. S.

possibly species specific; on gills, Pomoxis spp., Lepomis

nobilis, H. molitrix, Czech Republic and Slovakia. Davis, 1947): larger than C. agitans, body elongated,

Class Euglenida

blunt anteriorly but tapering posteriorly, body 12-23 pm long and 4-5 pm broad; free part of trailing fla¬

Butschli, 1884

gellum as long as anterior flagellum; movement not jerky, as in C. agitans. Brugerolle et al. (1979): cellular

With characteristics of the order (see p. 293 in Kudo, 1954). Main characteristics are the presence of chromatophores, paramylon body, and anterior stigma. The status is not clear in the one species described from fishes.

Genus Euglenosoma Davis, 1947 (Fig. 18)

structures of Ichthyobodo, Cryptobia, Trypanoplasma; Bykhovskaya-Pavlovskaya et al. (1962): gills of grass carp, crucian carp, former Soviet Union; Chen (1956a): description, grass carp, China; H. S. Davis (1947): as B.

concava on Lepomis macrochirus, Pomoxis spp, WV; Ergens and Lorn (1970): description, Acerina cernua, Carassius carassius, Tinea tinea; G. L. Hoffman (1978d): Carassius auratus, Ictalurus punctatus, correct name, AR; G. L. Hoffman (unpub., USFWS 1975-1985): gold¬ fish, golden shiners, channel catfish, many occur¬

Body euglenoid, long, spindle-shaped, tapering to sharp

rences, AR; Lorn (1980): electron microscopy showing

point posteriorly; body plastic; anterior end spatulate

intake of bacteria, not fish cells, indicating ectocommensal behavior; Lorn et al. (1976): Cyprinus carpio, Bul¬

and flexible; ventral groove and cytostome present; two equal flagella arising from cytostome; body twisted spirally.

garia; Molnar (1979a): C. carpio, grass carp, Hungary; Molnar, Hanek, and Fernando (1974): Ictalurus nebu-

■ Euglenosoma branchialis H. S. Davis, 1947: reported

losus, Ont.

once; on gills, Pomoxis annularis, P. sparoides, WV; 20-30 pm long x 4-5 pm wide.

■ C. carassii (Trypanoplasma carassii Swezy, 1915): prob¬ ably Cryptobia agitans or very similar.

30

Subkingdom Protozoa

Genus Ichthyobodo Pinto, 1928 (sometimes erroneously spelled Ictyobodo) (Syn. Costia) (Figs. 20, 21)

Flagellate with flat body; two trailing flagella originat¬ ing from infusorial cavity on ventral side of body. ■ Ichthyobodo necator (Henneguy, 1884; Joyon and Lom, 1969) Lom and Dykova, 1992 (Syn. Costia necatrix Henneguy, 1884; Costia pyriformis Davis, 1943): freeswimming, with flat, moderately asymmetrical, oval body measuring about 10 x 5 pm. Infusorial cavity opening on ventral part of body, from which two par¬ allel flagella of unequal lengths extend. Ventral surface recessed into broad groove in which both flagella move. Centrally located nucleus round, 2.5 pm, containing large nucleolus (1.5 pm). Quadriflagellar specimens in incip¬ ient stages prior to division. When feeding, organism adheres tightly to an epithelial cell, penetrates with protruding cytostomatic channel, and ingests suctorially. Lacks host specificity, even infesting amphibian larvae; very pathogenic. Marine strains reported (Ellis and Wootten, 1978). — Ref. Joyon and Lom (1966); Lom and Dykova (1992); Schubert (1966): ultrastructure.

Genus Lamellasoma Davis, 1947 (Fig. 22)

Resembling trypanosomes; body flattened, with bluntly rounded anterior end, but wider near middle. Long fla¬ gellum thicker at base, tapering gradually toward tip. Sev¬ eral rows of small refringent rods mnning parallel to body axis; small contractile vacuole in anterior end. Body 15-29 pm long x 3-4 pm wide; flagellum up to 30 pm; nucleus located about one-third of body length from anterior end, rounded or oval, with several large, periph¬ eral masses of chromatin; rounded karyosome, sometimes present. Blepharoplast anterior to nucleus, staining intensely. ■ Lamellasoma bacillaria Davis, 1947: on gills, Pomoxis annularis, P. sparoides, WV.

Genus Trypanoplasma Laveran and Mesnil, 1901 (Syn. Cryptobia Leidy, 1846, in part) (Fig. 23)

Biflagellate, trypanosome-like; one flagellum free, other marking outer margin of undulating membrane, kinetoplast (parabasal body) elongated. Parasitic in blood of fishes; fish leech is vector and intermediate host. Path¬ ogenicity not well understood but considered serious in some western trout hatcheries.

Woo and Wehnert (1983) suggest that Trypanoplasma salmositica belongs to the genus Cryptobia. Lom (1979), however, distinguishes the blood biflagellates of fishes ('Trypanoplasma) from ectoparasitic forms (Cryptobia); that designation is used here. — Ref. Becker (1977); Kudo (1971, p. 425); Lom (1979): review.

■ Trypanoplasma acipenseris Ioff, Lewaschow, and Boschenko, 1926: Acipenser mthenus, former Soviet Union. ■ T. borelli Laveran and Mesnil, 1902: Leuciscus erythrophthalmus, France; Brugerolle et al. (1979): cellular detail; Lom et al. (1986): experimentally severe patho¬ genicity, Carassius carpio fingerlings, Czech Republic and Slovakia. ■ T. carassii (Kaschkovsky, 1974) Lom, 1979: Carassius auratus, Siberia, Russia. This may be T. carassii Swezy, 1915; See also Swezy, 1919. ■ T. cataractae (Putz, 1972) Lom, 1979: Campostoma anomalum, Exoglossum maxilingua, Rhinichthys atratulus, R. cataractae, WV. ■ T. catostomi (Bower and Woo, 1977a): Catostomus commersonii, Ont.; Bower and Woo (1977b): develop¬ ment; Bower and Woo (1977c): experimental host speci¬ ficity; Muzzal (1979): incidence of 67%, NH. ■ T. cyprini Plehn, 1903: Cyprinus carpio, Germany. ■ T. guemeyorum Minchin, 1909: Esox Indus, England; Laird (1961): E. Indus, Coregonus clupeaformis, Salvelinus namaycush, northern Canada. ■ T. iubilans Nohynkova, 1984: intestinal parasite, but invading visceral organs; Dykova and Lom (1985): histopathology in pet cichlids, Herichthys cyanoguttaum, Cichlasoma meeki; is truly pathogenic. ■ T. lynchi Katz, 1951: see T. salmositica. ■ T. makeevi Akhmerov, 1959 (see BykhovskayaPavlovskaya, 1962, p. 19): Oncorhynchns gorbuscha, O. keta, East Asia. ■ T. markewitschi Schapoval, 1953 (see Schapoval, 1954): Anguilla anguilla, former Soviet Union. ■ T. salmositica Katz, 1951 (Syn. Cryptobia lynchi, C. borelli Laveran and Mesnil, 1901, in part): Oncorhynchns kisutch, U.S. Pacific Coast; Arai and Mudry (1983): O. tshawytscha, B.C.; C. Becker (1964): salmon, O. mykiss, Cottus spp., Gasterosteus aculeatus, Prosopium williamsoni, Rhinichthys cataractae, CA, OR, WA; C. Becker (1980): Acipenser transmontanus, Catostomus columbianus, C. macrocheilus, Cottus asper, C. bairdi, C. beldingi, C. confusus, Ptychocheilus oregonensis, WA; C. Becker and Katz (1965a, 1966, 1969): Cottus rhotheus, O. kisutsch, vector Piscicola salmositica, WA; Cottus aleuticus, O. gorbuscha, R. cataractae, refractive granule; Bower and Margolis (1983): direct transmission; Bower and Margolis (1984a):

Order Proteromonadida

centrifuge for detection, difference in host susceptibil¬ ity; Bower and Margolis (1984b): geographical distribu¬ tion, B.C.; Bower and Margolis (1985a): experimental infection, effect of temperature, salinity, host, O. tshawytscha most susceptible; Davison and Breese (1954): found in Oregon salmon only in November-February; Hoskins et al. (1976): coho salmon, B.C.; Jones and Woo (1987): immune response; Katz et al. (1961): count¬ ing technique; Katz et al. (1966): first record in O. nerka; Laidley et al. (1988): "stress response"; Lowe-Jinde (1979): experimentally in O. mykiss, pathogenesis; Lowe-Jinde (1980): observations on infected O. mykiss; Lowe-Jinde and Niimi (1983): none in 131 fishes, including salmonids, Lake Ontario; Paterson and Woo (1983) elec¬ tron micrography; Thomas and Woo (1988): mecha¬ nism of anemia in O. mykiss; Wehnert and Woo (1980): host specificity; Woo (1978): division process; Woo (1979): experimental infection; Woo and Wehnert (1983): direct transmission; Woo and Wehnert (1986): effect of hypoxia on infection; Woo et al. (1987): cortisol increases susceptibility. ■ T. tincae Schaperclaus, 1954: Tinea tinea, Germany. ■ T. truttae Brumpt, 1906: Salmo trutta, France. ■ T. valentini Gauthier, 1920: Salmo trutta, France. ■ Trypanoplasma sp. Wolf (pers. comm., State Dept. Fish and Game, Sacramento, CA, 1960): Dorosoma petenense and salmonids.

Genus Trypanosoma Gruby, 1841 (Fig. 24) Body leaflike; single nucleus; kinetoplast present, from which single flagellum arises; basal portion of flagellum forming undulating membrane extending along one side of body. Parasitic in blood of mammals, birds, rep¬ tiles, amphibia, fishes. Leeces are vectors for fish and amphibian forms. Pathogenicity in fishes not well known. Life cycle: trypanosomes picked up by leech with blood meal; appear in crop on second day; multiply rapidly; become crithidia in four to six days. At seven days, become slender metacyclic trypanosomes and migrate back to proboscis, where they are expelled during next blood meal (Qadri, 1962a). For methods, see C. Becker (1977) and Daly and DeGiusti (1971).

31

■ T. granulosum Laveran and Mesnil, 1909 (see Shulman, 1984, p. 28): Anguilla fluviatilis, France. ■ T. lotai Smirnova, 1970: Lota lota, former Soviet Union. ■ T. myoxocephala Fantham, Porter, and Richardson, 1942: in blood, Myoxocephalus octodecimspinosus, Que.; 42-49 pm long. ■ T. occidentalis Becker, 1967a: in blood, Cottusgulosus, C. rhotheus, Gasterosteus aculeatus, WA, 40-55 pm long. ■ T. percae canadense, Fantham, Porter, and Richard¬ son, 1942: in blood, Perea flavescens, Que.; Kennedy (1974): Perea fluviatilis, England; 45-50 pm long. ■ T. pingi Chen and Hsieh, 1964: Carassius auratus, China. ■ T. rebeloi Dias, 1955: Oreochromis mossambica. ■ T. remaki Laveran and Mesnil, 1902. Kudo (1921): Esox reticulahis, NY; Kennedy (1974): Esox lucius, England; Molnar (1979a): E. lucius, Hungary; 24-30 pm long. ■ T. serranoi Dias, 1955: Oreochromis mossambica (Tilapia m.), Africa. ■ T. shulmani Khaybulyayev, 1971 (see Shulman, 1984, p. 16): Esox lucius, former Soviet Union. ■ T. tincae Laveran and Mesnil, 1904. Kennedy (1974): Tinea tinea, England; Molnar (1979a): T. tinea, Hungary. ■ T. winchesiense Qadri, 1962a: Cyprinus carpio, England. ■ T. zilli Leger, 1914: Tilapia lata, Africa. ■ Trypanosoma sp. Fantham and Porter (1947): Salvelinus fontinalis, Que.; Strout (pers. comm., Univ. of New Hampshire, 1961): Esox niger, NH.

Order Proteromonadida Grasse, 1952 —emended Vickerman, 1976 One or two pairs of heterodynamic flagella present, without paraxial rods; single mitochondrion present, distant from kinetosomes, curving around nucleus, not extending the length of body; without Feulgenpositive kinetoplast. Golgi apparatus encircling band¬ shaped rhizoplast, passing from kinetosomes near sur¬ face of nucleus to mitochondrion; cysts present; parasitic.

— Ref. C. Becker (1977), Kudo (1971, p. 409), Lom (1979). ■ Trypanosoma catostomi Daly and DeGiusti, 1971: in blood, Catostomus commersoni, MI, 38-57 pm long; Bower and Woo (1979): C. commersoni prevalence, Ont. ■ T. ctenopharyngodoni Chen and Hsieh, 1964: Ctenopharyngodon idella, China. ■ T. danilewskyi Laveran and Mesnil, 1904 (see Shulman, 1984, p. 19): Cyprinus carpio, Europe.

Genus Karatomorpha Dobell, 1909 ■ Karatomorpha spp. Bykhovskaya-Pavlovskaya (1962), Shulman (1984): in intestine, Amur River fishes, Russia. Included here because I have seen flagellates that were not Hexamita or Spironucleus in the intestine of gold¬ fish and other pet fishes.

32

Subkingdom Protozoa

Class Diplomonadida Wenyon, 1926 —emended. Brugerolle, 1975 Usually with six anterior flagella, two nuclei, one or two karyomastigonts; cysts present; free-living or para¬ sitic (see Kudo, 1971, p. 438, and Lee et al., 1985).

Genus Hexamita Dujardin, 1841 (Syn. Octomitus Prowazek) (Fig. 25)

Pyriform; 8-14 pm long by 6-10 pm wide; two sym¬ metrically arranged oval nuclei present in anterior end; six anterior and two posterior flagella present. Freeliving in polluted waters; parasitic in intestine of amphib¬ ians and fishes; sometimes systemic. One species in fishes (Kudo, 1971, p. 449); review of diplomastigine fla¬ gellates from the intestine of fishes (Kulda and Lorn, 1964a). ■ Hexamita intestinalis truttae (Dujardin, 1841) W. Schmidt, 1919: in intestine of Salmo trutta and frogs, and in rectum of Moteila, Europe; 5-15 pm by 2-7 pm; Kulda and Lorn 1964a); may be synonymous with H. salmonis, below. ■ H. salmonis (Moore, 1923) Kudo, 1954: in intestine, young trout and salmon, sometimes pathogenic; 6-12 pm long by 3.5-5 pm wide; Arthur et al. (1976): Coregonus clupeaformis, Lotalota, Prosopium cylindricum, Salvelinus namaycush, Thymallus arcticus, western Canada; Ferguson (1979): electron microscopy and mortality; Hare and Frantsi (1974): Gasterosteus aculeatus, P.E.I.; H. Jackson (La Crosse, WI, unpub. res., 1984): growth in tissue culture of ovarian fluid, brown trout, Atlantic salmon, Chinook salmon, Duluth, MN; M. Kent (pers. comm., Nanaimo, B.C., 1991): in intestine and blood, pen-reared salmon, B.C.; Kulda and Lorn (1964a): differention of H. salmonis from Spironucleus elegans; Lester (1975): first report of H. salmonis in Gasterosteus aculea¬ tus, Vancouver, B.C.; Lester (1975): Lota lota, B.C.; Li and Desser (1985b): in intestine and gall bladder, Notemigonuscrysoleucas, Notropis cornutus, N. heterolepis, Semotilusatromaculatus, Algonquin Park, Ont.; McElwain and Post (1968): Cyzine for trout hexamitiasis, cultured in new born calf serum medium; M'Gonigle (1941): Salvelimts fontinalis, maritime hatcheries; Naich and Bennett (1987): caused significant disease, England; Richardson (1935, 1936): S. fontinalis, Que.; Rothenbacher and Bohl (1975): review of pathogenicity, therapy; Sano (1970): description of cysts, histopathology, Japan; Sanzin (1965): no intracellular stage, ecology, taxonomy and variabil¬ ity, Germany; Tesarcik and Rehulka (1973): in air blad¬ der, Oncorhynchus mykiss, Czech Republic and Slovakia; Uzmann and Hayduk (1963b): culture method, 6-12 pm x 3.5-5 pm; Uzmann and Jesse (1963a), Uzmann et al.

(1965): pathogenicity; Yasutake et al. (1961): chemo¬ therapy. ■ Hexamita sp. Gratzek (pers. comm., Univ. of Georgia, 1989): common in small discus and angel fish, where it is usually pathogenic, cured by Metronidazole. ■ Hexamita sp. Hanek and Molnar (1974): intestine, Catostomus catostomus, Que.; Hoskins and Hulstein (1977): Oncorhynchus keta, O. tshawytscha, B.C. ■ Hexamita sp. as hyperparasite, D. Huffman (pers. comm., San Marcos, TX, 1975) Spinitectus (nematode); Hunninen and Wichterman (1938): Deropristis inflata (trematode).

Genus Spironucleus Lavier, 1936 (Fig. 26)

Similar to Hexamita, but with the two nuclei oblong and curved in spirally; body more elongated. In the rectum of amphibians and intestine of fishes; only one species known to be parasitic. ■ Spironucleus elegans Lavier, 1936. Ergens and Lorn (1970): in aquarium fishes, capable of causing serious dis¬ ease leading to death; Giavenni (1981): Pterophyllum scalare, northern Italy; Kulda and Lorn (1964a, 1964b): P. scalare, differentiation from Hexamita; Molnar (1973): caused disease in grass carps, Barbus sp., angel fish; Mol¬ nar et al. (1974): Micropterus salmoides, Notropis het¬ erolexis, Perea flavescens, Pimephales notatus, Ont., first record in North America. ■ S. sytnphysodotiis (in Schubert, 1977, p. 688): in intes¬ tine, Pterophyllum scalare, Synnphysodon discus, other cichlids; controversial Lochkrankheit (hole-in-head) disease discussed as related to hexamitids. ■ Spironucleus spp.(?) and Hexamita spp. There have been many unpublished accounts of apparently un¬ described hexamitids from the intestine, gallbladder and other viscera, and the hole-in-head disease of S. discus. Definitive identification is lacking. The follow¬ ing accounts may be related: H. Ferguson and Moccia (1980): hexamitid flagellates recovered from the viscera of morbid Siamese fighting fish (Betta spendens); Herkner (1970): hexamitid illustrated as cause of hole-in-head disease may be Spironucleus, rather than Hexamita.

Subphylum Opalinata Lee, Hutner, and Bovee, 1985 This confusing group has characteristics mostly of Ciliophora but also of Mastigophora and Sarcodina. They are cylindrical or spindle-shaped, circular in cross section. The kinetosomes are unlike those of true ciliates in that there is a submarginal falx (sickle-shaped) composed of linear rows of closely set kinetosomes, with their

Subclass Suctoria

cilia extending along the anterior curved margin (Wessenberg, 1978). Gametes are flagellated. Usually commen¬ sals in the alimentary tract of amphibians, but two species found in marine fishes and one in freshwater fishes (Foissner et al., 1979).

Genus Protoopalina Metcalf, 1923 (Fig. 27)

■ Protoopalina symphysodonis Foissner, Schubert, and Wilbert, 1979: in rectum, freshwater Symphysodon aequifasciata; 97 (45-140) pm x 10 (6-15) pm; spectac¬ ularly pointed "tail"; most infected fishes died. Gratzek (pers. comm., Univ. of Georgia, 1989): Symphysodon dis¬ cus, concomitant with Hexamita sp., Protoopalina appar¬ ently not pathogenic.

Subclass 1. Suctoria Claparede and Lachmann, 1858 Adult stages sessile, without cilia, but with suctorial tentacles.

Phylum Ciliophora Doflein, 1901 Possessing cilia or compound ciliary organelles typically in at least one stage of life cycle; subpellicular infraciliature (ciliary rows) present even when cilia absent. Two types of nuclei present, usually one macro- and one micronucleus. Typically dividing by transverse binary fission, but also by budding and multiple fission. Con¬ tractile vacuole usually present. Most species free-living, but many commensal and some truly parasitic. — Ref. Corliss (1979); Ergens and Lorn (1970, p. 99); Fernandez-G (1976): silver stain; Foissner (1976): dry silver stain; G. L. Hoffman (1978a); Lee et al., (1985, p. 393).

See the Key to the Genera of Ciliophora of Fishes on p. 34.

Subclass Suctoria Genus Trichophrya Claparede and Lachmann, 1858 (Fig. 28)

Body small, round, elongated; 30-40 pm; without stalk; suctorial tentacles in fascicles, not branching, often irregularly arranged around periphery, pre-division stages sometimes showing two tufts on temporary ends of body. With simple or multiple endogenous budding; attached on invertebrates and vertebrates. Small strains probably host specific, otherwise fish host specificity apparently lacking.

33

— Ref. Culbertson and Hull (1962): all described species syn. of T. piscium; Kudo (1971, p. 1043).

■ Trichophrya ictaluri H. S. Davis, 1942: on gills, Ictalunis punctatus, IA. ■ T. liaohoensis Chen, 1984: on nine fish species, no dif¬ ferentiation from T. piscium, China. ■ T. micropteri H. S. Davis, 1942: on gills, Micropterus dolomieui, WV; on gills, Notropis cornutus, Ont. ■ T. piscium (Biitschli, 1889) Culbertson and Hull, 1962 (Syn. according to Culbertson and Hall, 1962: T. ictaluri Davis, 1947; T. intermedia Prost, 1952; T. micropteri Davis, 1942; Trichophrya sp. from Salvelinus fontinalis H. S. Davis, 1942; and probably T. sinensis Chen, 1955). According to Shulman (1984), species is known as Caprinianapiscium (Biitschli, 1889) Jankowski, 1973, in the former Soviet Union. T. piscium or Tricophrya sp. reported on the gills of Anguilla sp., Catostomus catostomus, Coregonus albula, Ctenopharyngodon idella, Cyprinus carpio, Esox Indus, Ictalurus nebulosus, I. punctatus, Lepomis humilis, Micropterus dolomieui, Morone chrysops, M. saxatilis, Notiopis atherinoides, N. cornutus, Oncorhynchus clarki, O. keta, O. kisutch, O. mykiss (Saltno gairdneri), O. nerka, Perea fluviatilis, Salmo salar, S. trutta, Salvelinus fontinalis, S. namaycush from Man., Nfdl., N.S., Ont., England, Finland, Italy, former Soviet Union, IA, MD, NJ, TX, VT, WA, WV, WY. — Ref. Becker (1967b), Bogdanova (1967), Calenius (1980b), Dechtiar (1972a), Egusa and Ahmed (1970), Hare and Frantsi (1974), Heckmann and Carroll, (1985), Heckmann and Ching (1987), Kennedy (1974), Li and Desser (1985a), Lorn et al. (1976), Muzzal and Peebles (1987), Nepszy (1988), Paperna and Zwerner (1974), Sandeman and Pippy (1967), Scheubel (1973), Wiles and Cone (1986). For comparison, see Morgensen and Butler (1984): cytology of T. rotunda.

This species has been reviewed by Culbertson and Hull (1962), who placed the above-listed four species in syn¬ onymy with it. They misquote two species: T. ictalurus Davis, 1942 is T. ictaluri Davis, 1947; and T. salvelinus Davis, 1942 is Tricophtya sp. Davis, 1942. They comment that T. piscium is a symbiont, probably feeding on fish ectoparasites. Tricophrya has been found on fish gills that were not heavily parasitized by other ectoparasites, so it must have been feeding on fish mucus or other fish material. Kozicka (1966) believes it to be pathogenic because she found fish red blood cells in it. Lorn (1971a) indicates it is a commensal, because electron microscopy revealed no fish tissue in the tentacles. U.S. hatchery biol¬ ogists H. S. Davis (1942) and F. Meyer (pers. comm., USFWS, 1964) believe it contributes to morbidity when present in large numbers. H. S. Davis (1942) found T. micropteri only on Micropterus dolomieui at Leetown, WV. It was not present on the closely related M. salmoides or any other fish at the hatchery. F. Meyer (pers. comm., USFWS, 1964) found a Tricophrya sp. only on Lepomis humilis, although it was in a pond containing catfish,

Subkingdom Protozoa

34

Key to the Genera of Ciliophora of Fishes 1. Cilia present throughout trophic life; no tentacles .Nonsuctorian Ciliates 2

9. (3) Attached; usually lacking body cilia, though telotroch (larva) possessing posterior ring of cilia .Suborder Sessilina 10

1. Cilia only when young; adult with feeding tentacles.Subclass I Suctoria (Fig. 28)

Trichophrya

9. (3) Free-swimming; low barrel- to saucer-shaped, with highly developed attaching organellae on posterior

2 (1) Without adoral (posterior) .

zone of membranellae.Subclass III Holotrichia 6

end consisting of circle of posterior cilia and chitinoid ring attachment disc with radially arranged teeth

2 (1) With adoral zone of membranellare.3 .

3. (2) Adoral zone of membranellae winding

(denticular ring) .Suborder Mobilina, Family Urceolariidae 14

counterclockwise; external parasites .Subclass IV Peritrichia 9

10. (9) Without stalk.Family Scyphidiidae 11 10. (9) With stalk (resembling

3. (2) Adoral zone of membranellae winding clockwise to cytostome; large, oval, intestinal parasites; usually active, cilia obvious; rare in fishes.Subclass II Spirotrichia 4 4. (3) Peristome (mouth funnel) sunk in funnel-like

Vorticella).12

11. (10) Cylindrical; posterior end attached, usually by widened scopula; body usually cross-striated; nucleus ribbon-shaped; adoral membrane small .(Fig. 42)

hollow at anterior end, sometimes difficult to see; 70-80 pm long.(Fig. 29)

Balantidium

11. (10) Elongate, pear-shaped; macronucleus a solid inverted cone; scopula small.(Fig. 43)

4. (3) Peristome not sunk in funnel-like hollow at anterior end.5

Epistylis and (Figs. 45-48) Heteropolaria

12. (10) Stalk contractile.13

parallel to body axis, then turning toward

13. (12) Entire colony contracting or

right in front of cytostome (mouth); up to 150 pm long.(no fig.)

Apiosoma (Glossatella)

12. (10) Stalk noncontractile .(Fig. 44)

5. (4) Peristome starting at anterior end, running

Ambiphrya (Scyphidia)

Nyctotherus

6. (2) Cytostome in peristome, with

expanding simultaneously.(no fig.)

Zoothamnium

13 (12) Colony not contracting simultaneously;

inconspicuous membrane . . . Order Hymenostomatida 7

6. (2) Cytostome on body surface or in peristome, without membrane or strong cilia.Order Gymnostomatida 8 7. (6) Cytostome near anterior end, but difficult to determine

recorded from pikeperch egg disease, also

Carchesium

found on trout eggs.(no fig.) 14. (9) With incomplete turn of adoral ciliary spiral;

anterior projection of tooth present; gill parasites,

in large specimens; body oval to round; ciliation uniform;

usually "cupped" over gill lamellae, dying soon

pellicle longitudinally striated; mature macronucleus

after removal from fish.15

horseshoe-shaped; multiplication occurring within cyst;

14. (9) With one or more complete turns of adoral ciliary

subepithelial parasite; trophonts varying from 50 to

spiral; no anterior projection of tooth present;

1000 pm in diameter

parasitic on body, gills, or in urinary bladder.16

.(Figs. 30-33)

Ichthyophthirius multifiliis

7. (6) Cytostome near anterior end of body, but apparent because ciliature longer than body cilia; body pear-shaped to ovate; 25 to 31 meridional ciliary rows present; macronucleus round to oval; multiplication by binary

15. (14) Central ray (centripetal thorn) of denticle slender but well-developed; blade curved or rodlike .(Figs. 52, 62)

Tripartiella

15. (14) Central ray of denticle a small, insignificant crook.(Figs. 54, 61)

Tdchodinella

fission; possibly causing ulcerlike lesions from which parasites may invade deeper tissues; 30-70 pm long .(Figs. 36, 37)

Tetrahymena corlissi

8. (6) Cytostome ventral, in anterior half; dorsoventrally flattened; ovoid dorsal surface without cilia, convex; ventral surface flat, with subpellicular infraciliature (cilary rows); oral basket conspicuous; macronucleus

16. (14) Adoral ciliary spiral with fewer less than two turns, but at least one complete spiral-.17 16. (14) Adoral ciliary spiral with two or three complete turns; urinary bladder parasites.(Fig. 55)

17. (16) Ciliary girdle with supplementary circlet of cirri .(no fig.) (Cydochaeta)

Trichodina (Cyclochaeta domerguei as

rounded; 50-70 pm long .(Figs. 38-40)

described by MacLennan, 1939,

Chilodonella (Chilodon)

8 (6) Cytostome a long slit; flask-shaped; somewhat

Vauchomia

is

Trichodina domerguei nomen nudum;

.

compressed; ciliation uniform, complete; two macronuclei present; one species.(Fig. 41)

Amphileptus

cf. Lorn and Hoffman, 1964) 17. (16) Ciliary girdle without supplementary circlet of cirri.(Figs. 53, 56-60)

Trichodina

(text continues on page 40)

30. Ichthyophthirius - Ciliates FIG. 28. Trichophrya piscium: a, on gills of Ictalurus punctatus, x 160; b, attached to gills of Micropterus dolomieui (redrawn from H. S. Davis. FIG. 29. Balantidium ctenopliaryngodoni (modified from Chen, 1955). FIG. 30. Ichthyophthirius multifiliis: a, trophont (cytostome usually not seen); b, oral detail of theront (from J. L. Lee et al., 1985). FIG. 31. /. multifiliis theront showing nucleus at anterior, instead of at midbody as is shown by some authors, methyl green stain, x 400. FIG. 32. /. multifiliis trophont in wet mount showing prominent macronucleus. FIG. 33. I. multifiliis mature trophonts embedded in skin of fin. FIG. 34. Ophryoglena sp. from infected Micropterus salmoides. FIG. 35. Ophryoglena sp. causing epithelial sloughing of small M. salmoides.

Ciliates

41. Amphileptus

FIG. 36. Tetrahymena corlissi, showing membranelles, ciliary rows, macroand micronuclei, vacuole, and caudal cilium (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences). FIG. 37. T. corlissi from infected Poecilia reticulata (from G. L. Hoffman et al., 1975, J. Parasitol.). FIG. 38. Chilodonella cyprini, ventral view (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences). FIG. 39. C. cyprini, silver stain showing ciliary rows (kineties) and posterior indentation (courtesy of W. Foissner, Austria). FIG. 40. Chilodonella hexasticha, silver stain, showing fewer ciliary rows and no posterior indentation (courtesy 44. Epistylis 42. Ambiphrya of W. Foissner, Austria). FIG. 41. Amphileptus branchiarum: a, dorsal view of free-swimming stage; b, ventral view showing M-shaped buccal orifice; c, embedded stage in gill, note the two macronuclei (N) (redrawn from Wenrich, 1924). FIG. 42. Ambiphrya tholiformis, showing ribbonlike nucleus, circle of body cilia, and large scopula (from Surber, 1942). FIG. 43. Apiosoma sp., schematic: a = anterior half of body, b = posterior half of body, e = epistomal disc, i = oral funnel, ma = macronucleus, mi = micronucleus, p = striped surface of pellicle, pc = groove of pectinilar wreath, pr = peristomal lip, r = groove containing the peristomal ciliary spiral, s = scopula, v = contractile vacuole (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences). FIG. 44. Epistylis niagarae (from Bishop and Jahn, 1941). This species is not listed in text because it is found on turtles.

45. Heteropolaria

Peritrichs FIGS. 45-48. Heteropolaria colisamm. FIG. 45. Zoids, AW = anlagen of the ciliary girdle, CV = contractile vacuole, E = cytoplasmic inclusions, Ma = macronucleus, Mi = micronucleus, My15 = parts of the myoneme system, Nv = food vacuole, Pd = peristomal disc, Pk = peristomal collar (from Foissner and Schubert, 1977). FIG. 46. Zoids from a colony on Lepomis cyatiellas. FIG. 47. Telotroch, note eccentric position of scopula (S) (from Foissner and Schubert, 1977). FIG. 48. Colonies on skin, particularly the dorsal and anal fins, of Lepomis cyanellus. FIG. 49. Method for measuring the adhesive disc of urceolarids: a = diameter of denticulate ring, b = diameter of adhesive disc, o = width of peripheral membrane, p = number of rods corresponding to the width of one denticle (from Ergens and Eom, 1970, by permission of the Czechoslovak Academy of Sciences). FIG. 50. Method for measuring the denticle of trichodinids: a = overall length, b = length of inner thorn, c = width of central part, d = length of blade (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences). FIG. 51. Denticle shapes of urceolarids: a = Foliella, b = Paratrichodina, c = Trichodina, d = Tricliodinella, e = Tripartiella, f = Vauchomia (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences). FIG. 52. Example of Tripartiella: T. lata attached to gill lamella (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences).

49

51

52. Tripartiella

—--TrichodinidsFIG. 53. Trichodina triittae. aboral disc (redrawn from Mueller, 1937c). FIG. 54. Trichodinella epizootica: a, aboral view; b, lateral view (redrawn from Lorn, 1956). FIG. 55. Vauchomia nephritica: a, lateral view; b, aboral end (attachment disc) (redrawn from Mueller, 1938).

53. Trichodina

54. Trichodinella

'll a

b 55. Vauchomia

Trichodinids FIG. 56. Trichodina fultoni, daughter stage after division showing one half of the old denticles (inner ring) and a full complement of new denticles forming in the more peripheral (dark) ring, x 650 (from H. S. Davis, 1947). FIG. 57. Trichodina platyformis, x 600 (from H. S. Davis). FIGS. 58-62 are examples of silver impregnated specimens. FIG. 58. Trichodina reticulata from Carassius auratus, AR. xlOOO. Note reticulated center (photo by K. Migala, Wroclaw, Poland). FIG. 59. Trichodina nigra from Acantharchus pomohis, WV (photo byj. Lorn, Ceske Budejovice, Czech. Republic). FIG. 60. Trichodina donierguei acuta from Notemigonus crysoleucas, AR, x 1000 (photo by K. Migala, Wroclaw, Poland). FIG. 61. Trichodinella epizootica from Carassius auratus, AR, x 1500 (photo by K. Migala, Wroclaw, Poland). FIG. 62. Tripartiella bursiformis from Acantharchus pomotis, WV, x 2000 (photo by J. Lorn, Ceske Budejovice, Czech. Republic).

Subkingdom Protozoa

40

goldfish, carp, golden shiners, buffalo, largemouth bass, and green sunfish. Apparently, there are distinct phys¬

■ Nyctotherus dilleri Earl and Jimenez, 1969: intestine,

iological species or strains of Tricophrya, despite the

■ N. pangasia Tripathi, 1954a: intestine, freshwater fish,

morphological similarity.

India.

■ Trichophrya sp. H. S. Davis (1942): on gills, Salvelinus

■ Nyctotherus sp. Fantham and Porter (1947): intestine,

fontinalis.

brook trout, perch, central Europe; originally erroneously

■

reported as Balantidium sp.

Tricophrya sp. as T. catostomi, Heckmann and Carroll (1985); Catostomus commersoni, WY, parasite not prop¬ erly described, may have been T. piscium; Wiles and Cone (1986): description from C. commersoni but prob¬ ably not T. piscium; N.S.

■ Tricophrya sp. as T. clarki, Heckmann (1970); Onco-

rhynchus clarki (Salmo clarki), WY; Ph.D. thesis—new species may not be named solely in theses or abstracts; Heckmann and Ching (1987): O. clarki, Catostomus

catostomus, WY.

Cichlasoma fenestration, Mexico.

■ Nyctotherus spp. Ky (1971): comparison of known species from freshwater fishes.

Subclass Hymenostomata (Syn. Holotrichia) Usually holotrichous (uniform cilia over body); body cilia as monokinetids or dikinetids; oral cilia as right oral dikinetid of three segments at most, and usually

Nonsuctorian Ciliates

three left oral polykinetids (membranelles); caudal cilia common.

All possessing cilia or cirri in the trophozoite stage. Par¬ asitic ciliates of fishes varying from 20 pm to 1 mm in diameter. Cytostome present in all species parasitic in or

Genus Ichthyophthirius

on fishes. Cysts produced by some species.

Fouquet, 1 876

— Ref. Corliss (1959, 1961).

(Figs. 30-33, Color Fig. 2)

■ Ichthyophthirius multifiliis F. (Syn. Enchelys parasitica

Subclass Trichostomatia

Dorier, 1926 is probably juvenile stage of I. multifiliis,

Butschli, 1899

according to Schaperclaus, 1954). Commonly known as "Ich" and "whitespot," this

Genus Balantidium

species is found in the skin of many, probably most, species of freshwater fishes in the temperate zone world¬

Claparede and Lachmann, 1858 (Fig. 29)

wide. It is probably the most important freshwater fish parasite in the world. New hosts are continually being

Oval, ellipsoid to subcylindrical; cytostome and oral cav¬

reported, e.g., Giavenni (1981): ornamental fishes, Italy;

ity long, apicoventral; concrement vacuole absent; body

Mpoame (1982): seven species of African fishes. Ich has

cilia holotrichous. Endosymbionts of diverse hosts from

been reported from Australia, China, all of Europe,

insects to primates; may damage intestinal epithelium.

India, Indonesia, Iraq, North America, Peru, South Africa,

— Ref Kudo (1971, p. 880).

the former Soviet Union, and probably more. Body oval to round but very plastic; 50 pm to 1 mm

■ Balantidium ctenopharyngodoni Chen, 1955: intestine, Ctenopharyngodon idella, China; Ky (1971): comparison

ated; cytostome (oral cavity) 8-20 pm, containing larger

with other species, North Vietnam; Molnar and Reinhardt

right undulating membrane and three smaller left mem¬

(1978): intestinal lesions, C. idella, Hungary.

branelles near middle of youngest (30-50 pm) stages, but

in diameter; ciliation uniform; pellicle longitudinally stri¬

with unequal growth of ciliate producing gradual shift

Subclass Heterotrichia Genus Nyctotherus Leidy (No fig.)

forward until cytostome nearly anterior in larger (350-800 pm) individuals; cytostome very difficult to see in larger specimens; details best seen in silver-impreg¬ nated specimens. Horseshoe-shaped macronucleus vis¬ ible without staining, micronucleus adhering to it in larger specimens; macronucleus spherical in youngest

Oval, compressed; peristome beginning at anterior end,

individuals, becoming oval, then elongated, and finally

turning to right, and ending in cytostome (mouth) mid¬

horseshoe-shaped. Large size of the mature parasite and

way; cytopharynx running dorsally and posteriorly,

horseshoe-shaped macronucleus are best identifying

forming a long tube with undulating membrane. Mas¬

features. Electron micrographic descriptions given by

sive macronucleus present anteriorly; micronucleus

Chapman (1984), Chapman and Kern (1983), Kozel

present; cytopyge and contractile vacuole terminal; in colon of invertebrates, amphibia, and fishes.

(1986), McCartney et al. (1985), and Peshkov and Tikhomirova (1968).

Subclass Hymenostomata

Life cycle: after invading the fish, the organism under¬ goes very limited division (Ewing et al., 1985; Kozel, 1986), but tremendous growth of the trophont (Ewing and Kocan, 1986), occurs over 4 to 40 days, depending on temperature, with the optimum usually 18-24°C. At maturity the (now) tomont leaves the fish (Ewing and Kocan, 1987), attaches to a substrate, and becomes a cyst. Overnight, up to 1000 small (30-40 pm), oval, ciliated theronts (tomites), each with single anterior knob (per¬ foratorium), are produced from each cyst; they rupture the cyst wall, swim away randomly, and seek a fish (Lorn and Cerkasovova, 1974). Upon reaching a fish, the theront invades the epithelium (Ewing et al., 1985), or sometimes the body or gill, depending on fish species, /c/2 strain, and perhaps other factors. Various factors affecting the development of Ich are elucidated by Ewing, Blazer and Kocan (1988) and Ewing et al. (1986). Temperature greatly influences Ich development and fish infection. In the southern United States, epizootics at fish farms usually occur in early spring when the water temperature approaches 19°C. One catfish farmer reported that Ich epizootics typically came in the spring when a cold spell followed two weeks of warm weather, and when the water temperature was about 16°C. In Fin¬ land, Ich epizootics of salmon occurred at 15°C (TellervoValtonen and Keranen, 1981). In Maine, P. Walker (pers. comm., 1980) reported epizootics of Ich when the ice went out. In the former Soviet Union, Musselius (1979) reported outbreaks at 1-2°C. Herman et al. (1959) dis¬ covered physiological variants of Ich that were mainly temperature related; this variation was further elaborated on by Nigrelli et al. (1976). Two temperature strains, 22-24°C and 28-29°C, were described by Yunchis (1978). Beckert (1967) failed to culture Ich in various media, fish mucus, fish blood, carp peptone, Ordal's bacterio¬ logical medium, and tissue culture but had limited survival of five days in gill tissue medium. Current researchers maintain Ich on live fishes, transferring to nonimmune fish periodically at controlled tempera¬ tures. More precise details of the methods, involving the number of theronts for infections, can be found in Beck¬ ert and Allison (1964), Clayton and Price (1988), and Ewing, Ewing, and Kocan (1988); intraperitoneal injec¬ tion is discussed by Dickerson et al. (1985). Although it is believed that all freshwater fishes are susceptible to infection, at times some appear to have a natural resistance to Ich disease. Newly hatched bluegill fry are refractory for about one month (this needs fur¬ ther study). Paperna (pers. comm., Israel, 1972) kept various species of fish enclosed in live boxes in a pond during an epizootic of Ich, and Tilapia were not affected. M. Martin (pers. comm., Stuttgart, AR, 1975) observed that golden shiners were seldom diseased when kept in ponds of channel catfish during an Ich epizootic, but could become diseased when kept in other ponds. Polzin and Bremer (1971) in Germany found that Ich developed normally in Rutilus but remained small in Abramis, with few Ich able to leave the fish.

41

It is now common knowledge that survivors of Ich epi¬ zootics usually have developed a temporary but good immunity (Amandi and Banner, 1985; Areerat, 1974; Beckert and Allison, 1964; T. Clark et al., 1987; Hines and Spira, 1974; Parker, 1965; Purvis, 1966; Wahli and Meier, 1985). Adding immune fishes to a pond during an Ich epizootic decreases the mortality (R. Allison, 1968-1969) and intraperitoneal injection can be used to immunize fishes (Dickerson et al., 1985). Due to cross reactions, Tetrahymena or its sheared cilia can be used to immunize fishes (Goven, Dawe, and Gratzek, 1980; Wolf and Markiw, 1982); U.S. Patent #4,309,416 has been granted in the name of Gratzek, Goven, and Dawe, G. L. Hoff¬ man (pers. comm., USFWS, 1982). The sera of immune fish will agglutinate theronts in vitro. Death of the fish usually occurs during the growth period of the parasite, when much damage is done by the large, active trophonts. In severe infections the epithelium sloughs off and the fish succumbs quickly (Bauer et al., 1959; Chapman, 1984; Dogiel et al., 1958; Ventura and Paperna, 1985). Diagnosis of Ich is made simple by the large size of the motile trophont and its horseshoe-shaped macronucleus. Early stages, with smaller trophonts containing round to oval nuclei, have been confused with Amphileptus branchiarum; the latter has two spherical macronuclei, however. Checking for Ich carriers in ponds, etc., can be done by xenodiagnosis (uninfected test fish in a live box). Treatment during an epizootic is effective only if given early, because only the free-swimming tomonts and theronts are killed. In warm water ponds, it is important to treat early in the spring, probably before the water reaches 18°C. Copper sulfate or formalin are usually effective, and immunization with a sublethal dosage of Ich theronts or Tetrahymena as the antigen should be con¬ sidered. For details on method and dosage, see Treatment, p. 409. It is of interest that Allen and Avault (1970) found channel catfish to be Ich-free in brackish water cul¬ ture in the southern United States. Another interesting sidelight is the finding of Myxidium (Myxosporidea) spores in Ich (Oka, 1973). — Ref. Bauer et al. (1959), Canella and Canella (1976), H. S. Davis (1953, p. 209), Kudo (1971, p. 893), F. Meyer (1966a), Mugard (1949).

■ I. marinus Sikama, 1961. Author: this may be Cryptocaryon—marine Ich.

Genus Ophryoglena Ehrenberg (Figs. 34, 35) (Not included in key because there is only one North American report.)

Ellipsoidal to cylindrical; ends attenuated or rounded; preoral depression in the form of a "6"; tetrahymenal buccal ciliation present; refractile "watch glass" body in

Subkingdom Protozoa

42

cytoplasm near buccal cavity; macronuclei of various shapes with several endosomes; micronucleus present; fresh- or saltwater parasite. — Ref. Canella and Canella (1976): Italy; Kudo (1971, p. 34).

■ Ophryoglena sp. G. L. Hoffman (1967b): infected Cot¬ tas bairdi, Lepomis macrochirus, Semotilus atromaculatus, S. corporalis in aquariums, producing sloughing of epithe¬ lium and mortality; 150-200 x 85 pm; usually ellip¬ soidal, slightly attentuated anteriorly, but sometimes rounding up when in tissue; cytostome about 45 pm from anterior end; probably uncommon, WV.

Genus Chilodonella Strand, 1926 (Syn. Chilodon Ehrenberg, 1838) (Figs. 38-40)

Ovoid, tapered anteriad; dorsoventrally flattened; ven¬ tral surface with kineties (ciliary rows); anteriorly flattened dorsal surface with cross-row of bristles; cytostome round; oral basket conspicuous, protrusible; macronu¬ cleus rounded. On fishes and amphipods; many species. For treatment, see under Chemotherapy and Prophylaxis. — Ref. O. N. Bauer et al. (1969); Bykhovskaya-Pavlovskaya et al. (1962); H. S. Davis (1953); Kazubski and Migala (1974): descriptions; Kudo (1971, p. 582).

Genus Tetrahymena Ferguson, 1941 (Figs.

36, 37)

Pyriform; small, with uniform ciliation; 17-42 ciliary rows and two postoral rows present; preoral suture straight. Cytostome small, close to anterior end, pyriform; undulating membrane on right side and three membranellae on left side in cytostome. Macronucleus ovoid; micronucleus absent in some species. In freshwater or facultatively parasitic. — Ref Corliss (1972, 1973, 1979), Elliot (1973), Kudo (1971, p. 894).

■ Tetrahymena corlissi Thompson, 1955. Corliss (1970): description; G. L. Hoffman et al. (1975): epizootic of gup¬ pies (Poecilia reticulata), parasite invaded tissue and many fishes died, NC; Lynn (1975): life cycle; Thomp¬ son (1958): experimental infection; J. Wright (1981): in turbellaria. ■ T. pyriformis (Ehrenberg). Bykhovskaya-Pavlovskaya (1962): several fishes, former Soviet Union, Corliss and Daggatt (1983): redescription; Gilbert et al. (1979): use for in vitro testing of parasiticide; Goven, Dawe, and Gratzek (1980): use for immunizing catfish against Ich; Wolf and Markiw (1982): for immunizing rainbow trout against Ich; Yukhimenko (1972): in and on larvae of Ctenopharyngodon idella, Hypophthahnichthys molitrix, former Soviet Union. ■ Tetrahymena spp. H. W. Ferguson et al. (1987): cranial ulceration, Salmo salar, Scotland: Fijan (pers. comm., for¬ mer Yugoslavia, 1974): mortality of carp fingerling; Nigrelli et al. (1955): epizootic in aquaria, NY.

Subclass Phyllopharyngia Trophonts free-swimming; cilia mainly on ventrum; true cytostome present, oral region with phyllae and rod¬ shaped nematodesmata. Some species external sym¬ bionts, but at least two species highly pathogenic.

■ Chilodonella cucullulus (Muller, 1941): free-living, but may become so numerous in tropical fish culture that it appears parasitic; tending to come to the surface in huge numbers. ■ Chilodonella cyprini (Moroff, 1902) (coldwater Chil¬ odonella): on gills, skin of many fishes, Australia, China, Europe, North America; epizootics usually at 5-10°C, rarely up to 22°C; body 30-80 pm x 20-62 pm (usually about 60 x 40 pm); posterior indentation present; kineties (ciliary rows) 9-15 left, 8-13 right. Ashburner and Ehl (1973): treatment, Australia; Heckmann and Fiebelt (1970): new host, Catostomus catostomus, MT; G. F. Hoff¬ man et al. (1979): comparison with Chilodonella hexasticha, AR; Kazubski and Migala (1974): description of Chilodotiella cyprini, C. hexasticha, Poland; Krascheninnikow (1953): silver line staining, C. cyprini, C. hexasticha, Ukraine; Rydlo and Foissner (1986): taxonomy, redescrip¬ tion, and treatment of C. cyprini, C. hexasticha, Austria; Vanyatinskiy (1978): comparison of C. cyprini and C. hexa¬ sticha, former Soviet Union; Wiles et al. (1985): scanning EM, C. cyprini, C. hexasticha, N.S. ■ C. dentatus Fantham and Porter, 1947: on gills, Micropterus dolomieui, Que. ■ C. hexasticha (Kiernik, 1909) ("warmwater Chilodo¬ nella”): on gills, skin of many fishes, Bulgaria, Europe, Malaysia, Mexico, South Africa, USA, former Soviet Union,; epizootics usually at 19-26°C; body 30-60 pm x 20-44 pm (usually about 50 x 33 pm); posterior inden¬ tation absent; kineties (ciliary rows) 6-10 left, 5-7 right. G. F. Hoffman et al. (1979): comparison with C. cyprini, AR; Jimenez Guzman et al. (1988a): channel catfish, northeast Mexico; Kazubski and Migala (1974): descrip¬ tion, Poland; Lorn et al. (1976): carp, number of ciliary rows (kineties) of C. hexasticha and C. cyprini seem to over¬ lap, species determination needs more study; Paperna and van As (1983): epizootiology and pathology in tilapia, etc., South Africa; Rydlo and Foissner (1986): see C. cyprini; van As and Basson (1984): on tilapia, goldfish, South Africa; Vanyatinskiy (1978), Wiles et al. (1985): see C. cyprini.

Suborder 1. Sessilina

■ C. truttae E. Moore, 1925: goldfish, trout, NY. Author: this may be C. hexasticha; no posterior indentation; greater damage to goldfish.

Class Litostomatea Body monokinetids (ciliary rows) with tangential trans¬ verse ribbon; slightly convergent postciliary ribbon and laterally directed kinetodesmal fibril not overlapping those of adjacent kineties (ciliary rows); simple oral cilia present.

43

(scopula) of mobile species or secrete the stalk of the ses¬ sile species, (3) telotroch band, permanently ciliated on mobile species, temporarily so during dispersal of ses¬ sile species. — Ref. Calenius (1980c): staining method; Ergens and Lorn (1970, p. 110); Foissner (1976): dry silver stain; Hazen et al. (1976): fixing and staining method; G. L. Hoffman (1978a); Kudo (1971, p. 1016); Lee et al. (1985, p. 546); Lom (1966); Shulman (1984, p. 280).

Suborder 1. Sessilina

Genus Amphileptus

Kahl, 1935

Ehrenberg, 1838 (Syn. Hemiophrys Wrzesniowski, 1870) (Fig. 41, Color Fig. 3)

Flask-shaped; somewhat compressed; ciliation uniform and complete; slitlike cytostonre not reaching middle of body, without trichocyst borders; many contractile vac¬ uoles and two or more macronuclei present. —

Ref. Canella (1960). Foissner (1983), Kudo (1971, p. 852).

■ Amphileptus branchiarwn Wenrich, 1924 (Syn. A. voracus Davis, 1947; Hemiophrys branchiarum Kahl, 1931). Ergens and Lorn (1970, p. 100): Europe; Mitchell and Smith (1988): originally reported from frog tadpole gills, and has since been reported from many fishes, mostly in southern United States. Main identifying character¬ istics: free-swimming form elongated, slightly attenuated at both ends, 65-130 pm x 38-75 pm; slitlike, lateral cytostome about one-third of body length; two promi¬ nent macronuclei visible with acidic methyl green stain (differentiating species from Ich); attached form globu¬ lar in a pit, sometimes becoming completely embedded; seldom pathogenic, possibly beneficial because it some¬ times feeds on other ectoparasites. ■ A. disciformis Chen, 1956a (may be A. branchiarwn): cyprinids, China; Chen (1984, part 1): China. ■ A. macrostoma (Chen, 1955): described as Hemiophrys macrostoma in China, but Foissner (1983) created the new genus Pseudoamphileptus for it; invades Heteropolaria (Epistylis) Iwoffi on fishes and consumes all but the pel¬ licle, Austria.

Sedentary and sessile, except some with mobile telotrochs; body characteristically bell- or goblet-shaped, with stalk or holdfast at tapered end; mouthless telotroch with locomotory posterior ciliary girdle; some species colonial; adults typically filter-feeding bactivores; attached to various substrates, including fishes. — Ref. Lom (1964): description of buccal ciliary organelles; Lom (1966): review and new species; Shulman (1984, p. 281).

Key to the Genera of Sessile Peritrichs 1. Body stalked .4 1. Body not stalked.2 2. (1) Nucleus ribbon-shaped.3 2. (1) Nucleus conical or oval.(Fig. 43)

Apiosoma

3. (2) Body with permanent telotroch ciliary girdle (about midbody), scopula large.(Fig. 42)

Ambiphrya

3. (2) Body without telotroch ciliary girdle; scopula very small.(no fig.) 4. (1) Very short, single stalk.(no fig.)

Scyphidia

Rhabdostyla

4. (1) Longer, branching stalks, forming colonies.5 5. (4) Noncontractile stalks.5 5. (4) Contractile stalks.7 6. (5) Scopula of swarmer centrally located .(Fig. 44)

Subclass Peritrichia Stein, 1859 Body of three major regions: (1) oral, with prominent peristome bordered by adoral ciliature that originates in the oral cavity (infundibulum) at base of which is the cytostome; region able to contract and withdraw, (2) aboral, including kinetosomes that form the suction disc

Epistylis

6. (5) Scopula of swarmer not centrally located .(Figs. 45-48, Color Fig. 4)

Heteropolaria

7. (5) All stalks connected; entire colony contracting simultaneously.(no fig.)

Zoothamnium

7. (5) All stalks connected but contracting independently.(no fig.)

Carchesi

44

Subkingdom Protozoa

Genus Ambiphryo Raabe, 1952 (Syn. Scyphidia Dujardin, 1841, in part) (Fig. 42)

Cylindrical; posterior end (scopula) attached; body usu¬ ally cross-striated; scopula broad, flattened; ciliary girdle always present. Macronucleus usually ribbon-shaped. — Ref. H. S. Davis (1953, p. 212), Kahl (1935), Kudo (1971, p. 1022), Raabe (1952), Shulman (1984, p. 283). ■ Ambiphrya ameiuri (Thompson, Kirkegard, and Jahn,

1947): macronucleus without end branches; on gills, Ictalurus melas, I A, Scyphidia macropodia Davis, 1947, probably a synonym that was reported from Ictalurus nebulosus, I. punctatus, and accidentally on Lepomis macrochirus; Fitzgerald et al. (1982): channel catfish, scanning EM, TN; Goncharenko et al. (1985): channel catfish, former Soviet Union, Lorn and Dykova (1988): fry of Ctenopharyingodon idella, Cyprinus carpio, ciliate feeds on waterborn organic particles, but large numbers on gills of fry may injure or kill the fish, Czech Republic and Slo¬ vakia; Trombitsky (1987): fry of Ictiobus cyprinellus, com¬ mon carp, silver, bighead carp, former Soviet Union.

■ A. triangularis Li and Desser, 1985a: on fins, gills, nares, Notemigonus crysoleucas, Notropis comutus, Perea flavescens, Semotilus atromaculatus, Ont. ■ Apiosoma spp. Hanek and Molnar (1974): fins, Catostomus catostomus, Gasterosteus acideahis, Salvelinus fontinalis, Ont.; Heckmann and Farley (1973): as Glossatella sp. from Hesperoleucas symmetricus, CA; Heckmann et al. (1987): Cottus bairdi, UT; Lester (1974): Gasterosteus aadeahis, B.C.; Molnar et al. (1974): from six fish species, Ont.; van As and Basson (1984): Oncorhynchus mykiss, South Africa.

Genus Carchesium Ehrenberg, 1838 (No fig.)

Like Zoothamnium, but stalks contracting independently. — Ref. H. S. Davis (1953, p. 214); Kudo (1971, p. 1018).

■ Carchesium sp. H. S. Davis (1953): on eggs, pikeperch trout; G. L. Hoffman (pers. obs.): on skin, two moribund golden shiners, AR; Lom (1964): on tadpoles, morphol¬ ogy and morphogenesis, Czech Repbulic and Slovakia; Randall (1956): electron micrography; Zagon and Small (1970): scanning electron micrography.

■ A. micropteri Surber, 1940: see Apiosoma m. ■ A. tholiformis Surber, 1942: macronucleus with end

Genus Epistylis

branches; on gills, Micropterus dolomieui, M. salmoides, WV; Lorn and Dykova (1992): needs further study, may be A. ameiuri.

Ehrenberg, 1830

Genus Apiosoma Blanchard, 1885 (Syn. Glossatella Butschli, 1889) (Fig. 43)

Peritrich with elongated conical body, with only ante¬ rior end contractile; macronucleus conical, triangular from lateral view in aboral half of body; peristomal spi¬ ral encircling epistomal disc, making a full turn, with the remaining three-fourths encircled in the infundibulum; stalk visible as thin pad with electron microscopy, but hardly recognizable with light microscopy; found on skin, gills. Many species described in Europe, some considered harmful. Found on many fishes, including Cyprinus car¬ pio, Dalia pectoralis, Esox Indus, Gasterosteus aculeatus, grass carps, Lota lota, Perea fluviatilis, Oncorhynchus mykiss, S. trutta; undescribed species in North America. — Ref. Bykhovskaya-Pavlovskaya (1962): as Glossatella; Calenius (1980): staining method; Ergens and Lom (1970, p. 112); Kahl (1935); Lom (1966, 1973): EM of attachment; Scheubel (1973): key; Shulman (1984): many species. ■ Apiosoma micropteri (Surber, 1940) (Syn. Scyphidia m.

Surber, 1940): on gills, Micropterus dolomieui, M. salmoides, WV.

(Fig. 44)

Inverted bell form; usually on branched noncontractile stalk; often forming large colonies; in division, each newly formed organism forming its own stalk, which adheres to the common stalk; scopula of telotroch (swarmer) centrally located. Epistylids are common on fishes and sometimes con¬ tribute to morbidity via partial smothering of gills. Only Heteropolaria colisanim (see below) and H. Iwoffi have been properly identified, however. Those species on North American fishes have not been studied described; several species have been described in Europe. — Ref. Arvy et al. (1969): species descriptions; Davis (1953, p. 214); Ergens and Lom (1970, p. 114); Fischthal (1949b); Hazen et al. (1976): fixing and staining; Kahl (1935); Kudo (1954, p. 853); Lom (1973): description of attachment; Lom and Vavra (1961); Shulman (1984, p. 290).

■ Epistylis longicorpora Uyemura, 1938: from freshwater mussel; Miyazaki and Egusa (1973): bacterial secondary infections in koi carp; Sejima et al. (1973): control of E. longicorpora disease in goldfish, koi carp, Japan; Takase et al. (1973): Epistylis disease in koi carp, associated bac¬ teria. Foissner et al. (1985) suggest that E. longicorpora is Heteropolaria colisanim or closely related to it. ■ Epistj’lis spp. Arai and Mudry (1983): on gills, Cottus asper, Onchorliynchus nerka, B.C.; Arthur et al. (1976): Cothis cognatus, B.C.; Blasiola and Dempster (1975): nonfatal but debilitating to Cichlasoma; Cloutman (1975a): epi-

Family Trichodinidae

zootic on white bass, striped bass; Cloutman et al. (1978): on 8 of 22 fish species examined (may be Heteropolaris colisarum), and a different species on Lernaea (anchor parasite); Dean (1974): eight feral fish species killed by Epistylis-Aeromonas liquefaciens (red-sore) dis¬ ease, SC; Dechtiar (1972b): on Etheostoma exile, Ont.; Esch et al. (1976), Hazen et al. (1978): Aeromonas primary and Epistylis secondary in red-sore disease, NC (Foissner et al., 1985, think this is probably H.colisarum); Fischthal (1949b): on trout, darters, WI; Hazen et al. (1978): EM of lesions showing bacteria; G. F. Hoffman (pers. obs., 1959): on brook trout, NH, NC; Feitritz (1960); Feitriz and Fewis (1976): on rainbow trout, CA; R. Miller and Chapman (1976): on 15 fish species in reservoirs, NC; Mitchum (pers. comm., 1966): on brook and cutthroat trout, WY; Muzzal (1986): on brook trout, MI; Muzzal and Peebles (1987): Notropis atherinoides, MI; Muzzal and Sweet (1986): Cottus bairdi, MI; Muzzal et al. (1987): Lota lota, MI; Paperna and Zwerner (1976): Morone saxatilis, VA. ■ Epistylis spp. G. F. Hoffman (pers. obs., USFWS, AR, 1976), M. Meyer (pers. comm., USFWS, ME, 1976): on fish leeches, but not the same species as on host fish; G. F. Hoffman (pers. obs., 1979): Lernaea cyprinacea, but not the same species as on host fish, AR.

Genus Heteropolaria Foissner and Schubert, 1977 (Figs. 45-48, Color Fig. 4)

hike Epistylis, except with scopula of telotroch (swarmer) not centrally located, and the myonema of the peristomal disc with a peculiar branch. Fom and Dykova (1992) con¬ sider this genus provisional, pending further study, and suggest retention in Epistylis. ■ Heteropolaria colisarum Foissner and Schubert, 1977: from body, Colisa fasciata, aquarium fish, Germany; Foissner et al. (1985): from fins, body, Lepomis cyanellus, L. macrochirus, Ictalurus punctatus, AR, CA, ID, SC, TN; colonies from very small on scales to large (3 cm), pulvinate whitish coat on fins); stalks 12-35 pm in diameter and up to 1.18 mm in length; branching dichotomously; zoids cylindrical, 150-300 pm long x 40-60 pm in diameter; telotroch with eccentrically placed scopula. This species probably the same as Epistylis of Blasiola and Dempster (1975), Cloutman et al. (1978), Esch et al. (1976), Hazen et al. (1978), R. Miller and Chapman (1976), and Rogers (1971); and the primary or secondary cause of warm-water red-sore disease. Sogandares and Hutton (pers. comm., FF, 1958): unknown species, perhaps H. colisarum, on Mugil cephalus in fresh¬ water, FA. ■ H. lwoffi (Faure-Fremiet, 1943) Foissner and Schubert, 1977 (Syn. Epistylis lwoffi Faure-Fremiet, 1943). Cone and Odense (1987): scanning EM of H. lwoffi and Apiosoma piscicola on fry of Salvelinus fontinalis, N.S.; Foissner

45

(1983): morphology and predation by Pseudoamphileptus macrostoma, Austria; Lester (1974): on Gasterosteus aculeatus, B.C.; Lorn (1966): description and relationship to host, on many fishes, Europe; Lorn and Dykova (1992): is Epistylis Iwoff.

Genus Zoothamnium (Bory, 1824) Ehrenberg, 1838; Stein, 1854 (No fig.)

Similar to Vorticella, but colonial; all stalks connected and entire colony contracting simultaneously. Many species, some on marine fishes and shrimp. — Ref. Kudo (1971, p. 1018).

■ Zoothamnium sp. Khajuria and Pillay (1950): from grey mullet, India.

Suborder 2. Mobilina Kahl, 1933 Mobile forms, those on fish usually discoidal to bonnet¬ shaped and orally-aborally flattened, with permanently ciliated trochal (wheel-like) band (ciliary girdle); com¬ plex thigmotactic apparatus at aboral end, often with highly distinctive denticulate ring. Some species path¬ ogenic to fishes; one family.

Family Trichodinidae Raabe, 1959 With characteristics of the suborder. Six genera: Foliella, Paratrichodina, Trichodina, Trichodinella, Tripartiella, Vauchomia. Found mostly on fishes, skin or gills, but some occurring in urinary bladder as endocommensals (Lorn and Haidar, 1976). One species is found in the intestine (Cunha and Pinto, 1928), some on tadpoles, one on copepods (Migala and Grygierek, 1972), one on Gyrodactylus (Yin and Sproston, 1948), and one as an endozooic tissue parasite in eels (McArthur, 1976). Some species tend to be host specific, but others are not (Kazubski, 1965b; Lorn, 1961; Wellborn, 1967). Should be studied live, stained by silver impregnation (Arthur and Lom, 1984), and perhaps examined by scanning EM (Hamilton-Atwell and van As, 1982). Should be mea¬ sured according to Lom (1958, 1963) and Ergens and Lom (1970). Growth of skeletal parts has been described by Kazubski (1965a). The life cycle of Trichodina reticu¬ lata has been studied in detail (Ahmed, 1977), and H. S. Davis (1947) describes division of Trichodina fidtoni. Some species are highly seasonal, with definite temperature optima, and are usually more numerous in

46

Subkingdom Protozoa

cool weather (Kazubski and Migala, 1968; F. Meyer, 1970b). Under certain conditions, some species are highly pathogenic (McArdle, 1984). A suctorian, Endosphaerci engelmanni, has been found parasitic in Trichodina sphaeroidesi, infecting a marine fish (Padnos and Nigrelli, 1947). — Ref. Bykhovskaya-Pavlovskaya (1962); H. S. Davis (1947): described several species; Ergens and Lom (1970): Czech species, including methods of measurement; G. L. Hoff¬ man (1978a); Lom (1959, 1961, 1963, 1970); Shulman (1984); Wellborn (1967): North America; Wellborn and Rogers (1966).

Key to the Genera of Urceolarids of Fishes (Modified from Ergens and Lom, 1970, and Lom, 1963) (For measurement methods see Figs. 49 and 50)

1. With one or more complete turns of adoral ciliary spiral.2

Genus Paratrichodina Lom, 1963 (Fig. 51b)

Body a more or less flattened or vaulted hemisphere; ado¬ ral spiral making a turn of 150 to 280 degrees. Denticles with well-developed thorns, wedged together only by central conical parts; if anterior projection present near base of the blade, it is not inserted in notch in blade of preceding denticle. Macronucleus generally horseshoe¬ shaped. On gills, freshwater and marine fishes; two for¬ eign species parasitic in fish urinary tract. ■ Paratrichodina erectispina Lom and Haidar, 1977: on gills, Pimephales vigilax, ML Body 30 pm in diameter; adhesive disc 25 pm; denticulate ring 15 pm; 25 den¬ ticles present; 4 to 5 radial pins per denticle. ■ Paratrichodina sp. Lom and Haidar (1977): on gills, Notemigonus crysoleucas, IL; disc 12 pm; denticulate ring 9 pm; denticulate: thorns 1.5 to 2 pm, blade 2 to 3 pm, central part 1 pm; with very short anterior projection and kidney-shaped blade.

1. With less than one complete turn of adoral ciliary spiral.3 2. (1) Usually ectoparasitic; body usually saucer-shaped,

Ehrenberg, 1831 (Figs. 51c, 53, 56-60, Color Figs. 1, 5, 6)

with obvious tripartite denticle structure (blade, central conical part, inner thorn) .(Figs. 51c, 53, 56-60, Color Figs. 5, 6)

Trichodina

2. (1) Endocommensal; body hemispherical to conical in side view; tripartite denticles as above.(Figs. 5If, 55)

Vauchomia

3. (1) Denticles tripartite.4 3. (1) Denticles without inner thorn.5 4. (3) Tripartite denticles with discernible inner thorn, but denticles small and complex, with central conical parts very fine and projecting anteriorly; body dish-shaped in side view .(Figs. 51 e, 52, 62)

Tripartiella

4. (3) Tripartite denticles, but wedged together only

Paratrichodina

5. (3) Denticles with sharp anterior projection from the blade; body usually bonnet-shaped.(Figs. 5Id, 54, 61)

Body saucer-shaped to low bonnet-shaped. Aboral ciliature with three ciliary girdles: first made of short, simple cilia, closely associated with border membane; sec¬ ond a girdle of strong, long locomotor pectinelles not separated by a septum; third with tactile marginal cilia inserted on border of velum. Chitinoid ring of attach¬ ing disc with radially arranged hooked teeth, each with outer flat blade, central cone, and inner ray (thorn). Buccal ciliary spiral making between one and two com¬ plete turns. Numerous species; many pathogenic. Usu¬ ally on the gills, but in weakened fishes reproducing rapidly and possibly covering the entire surface of the fish, often leading to death of the infested fish. — Ref. see Urceolaridae, above.

by central conical parts (similar to Trichodina); body similar to Tripartiella.(Fig. 51b)

Genus Trichodina

Trichodinella

5. (3) Denticles with two anterior projections from the blade.(Fig. 51a)

Foliella

Subgenus Foliella Lom, 1959 (Fig. 51a)

Proposed as a subgenus of Trichodinella, but this has apparently been dropped. T. subtilis was described as T. (Foliella) subtilis Lom, 1959.

■ Trichodina acuta Lom, 1961: described as T. domerguei f. acuta f. n., on body, gills of fishes, including Cyprinus carpio and frog tadpoles, Czech Republic and Slvakia; Duncan (1977): Tilapia spp., Philippines; Lom (1970): C. carpio, Ictalurus nebulosus, Lepomis gibbosus, Perea fluviatilis, Czech Republic and Slovakia; Migala (1970): parasite population influenced more by light and tem¬ perature than carp density; van As and Basson (1984): C. carpio, South Africa; Vismanis et al. (1975): C. carpio, Latvia. Central clear area of adhesive disc is helpful in identification; considered pathogenic. ■ T. algonquinensis Li and Desser, 1985a: syn. of T. urinaria. ■ T. anguilli Wu, 1962. Ergens and Lom (1970): Anguilla vulgaris, Czech Republic and Slovakia; Markiewicz and Migala (1980): A. anguilla, not as pathogenic as T. fidtoni,

Family Trichodinidae

Poland; diameter 25-42 pm, adhesive disc 11-18 pm, 21-24 denticles present. ■ T. califomica H. S. Davis, 1947: on gills, body, Oncorhynchus tshawytscha, CA; Bykhovskaya-Pavlovskaya (1962): is Tripartiella californica from O. keta, O. gorbuscha, O. masu, Salvelinus leucomaenis, S. malma; Lorn (1970): is nomen dubium because silver stain was not used; Stein (1976): Salmo salar, former Soviet Union; Wellborn (1967): examination of type material. ■ T. centrostrigeata Basson, van As, and Paperna, 1983: Cyprinus carpio, Tilapia spp., South Africa. ■ T. chlorophora (Richards, 1948) Blecka and Garoian, 1972: redescription, Physa gyrina (snail), IL. ■ T. cubanensis Arthur and Lorn, 1984: from skin, Cichlasoma tetracantha, Cuba. ■ T. dallii Zhukov, 1964: Dallia pectoralis, lakes in Chukhotka, Russia; Lorn (1970): nomen dubium. ■ T. davisi Wellborn, 1967: on fins, body, gills, Morone saxatilis, NC; Lorn (1970): similar to T. funduli, T. hypselepsis, T. salmincola; Paperna and Zwerner (1976): M. saxa¬ tilis, Chesapeake Bay, VA. ■ T. discoidea H. S. Davis, 1947: on gills, Ambloplites rupestris, Ictalurus punctatus, Lepomis macrochirus, Pomoxis sparoides, I A, WV; Blecka (1972): I. punctatus, Lepomis cyanellus, southern IL; Meryman (1975): I. punctatus, IL. ■ T. domerguei (Wallengren, 1897) (Syn. Cyclochaeta d.)\ saucer-shaped (see MacLennan, 1939, for description of Cyclochaeta domerguei); Frank (1962): histopathology, Carassius auratus, Germany; Lorn and Hoffman (1964): consider species a Trichodina because marginal cilia (cirri) of many other Trichodina spp. are equally long; they consider this species nomen nudum until it can be com¬ pared with the European T. domerguei. See also Calenius (1980a): Gasterosteus aculeatus, Myxocephalus quadricornis, Finland; Kazubski and Migala (1968): seasonality on carp (good photos), Poland; Lester (1974): G. aculeatus, B.C.; Lom (1970), review of this confusing species; Lom and Stein (1966), Shtein (1976): G. aculeatus, Pungitius pungitius, White Sea, Russia. ■ T. epizootica: syn. of Trichodinella epizootica. ■ T. esocis Lom, 1961: Esox Indus, Czech Republic and Slovakia; Shulman (1984): E. Indus, Luciperca lucioperca, former Soviet Union. ■ T. fultoni H. S. Davis, 1947: on gills, Ambloplites rupestris, Ictalurus punctatus, Lepomis macrochirus, Micropterus dolomieui, M. salmoides, Oncorhynchus mykiss, WV; Calenius (1980c): Esox Indus (Syn. of T. pediculus), Fin¬ land; H. S. Davis (1953): on salamander; G. L. Hoffman and Lom (1967): Carassius auratus, Catostomus commersoni, Lepomis cyanellus, L. gibbosus, L. macrochirus, Micropterus dolomieui, M. salmoides, Notropis rubellus, N. spectrunculus, Pomoxis annularis, R. atratulus, R. cataractae, Salvelinus fontinalis, Semotilus margarita, WV; Lom (1970): Tinea tinea, Czech Republic and Slovakia; Lom

47

and Hoffman (1964): Lepomis cyanellus, M. salmoides, Rhinichthys atrahdus, WV; Markiewicz and Migala (1980): Anguilla anguilla, very pathogenic, Poland; Walker and McNeish (1981): L. aurihis, M. dolomieui, feral die-off, ME; Wellborn (1967): redescription, very large, 92-111 pm. ■ T. funduli Wellborn, 1967: body, fins, Fundulus notti AL; Lom (1970): similar to T. davisi, T. hypselepis, T. salmincola, possibly infraspecific varieties of same species. ■ T. gasterostei Shtein, 1967: Gasterosteus aculeahis, fresh¬ water lakes, former Soviet Union. ■ T. globosa Wellborn, 1967: Etheostoma radiosum, AR; Kozel and Whittaker (1982): E. caeruleum, KY. ■ T. guberleti (MacLennan, 1939) (Syn. Cyclochaeta d.)\ Richardsonius, Apocape, WA; Lom (1970): nomen dubium until verified with silver impregnation stain. ■ T. heterodentata Duncan, 1977: Tilapia mossambica, T. zilli, Trichogaster trichopterus, Philippines; HamiltonAtwell et al. (1986): scanning EM, South Africa; van As and Basson (1984): Cyprinus carpio, South Africa. ■ T. hoffmani Wellborn, 1967: Etheostoma edwini, AL, seven radial pins per denticle. ■ T. hypsilepis Wellborn, 1967: Notropis hypselepis, AL; Arthur and Lom (1984): frog tadpoles, Cuba; Lom (1970): similar to T. davisi, T. funduli, T. salmincola, possibly infraspecific varieties of the same species, 10 radial pins per denticle. ■ T. kuleminae Lom, 1970 (Syn. T. meridionals Kulemina, 1968): in nasal fossae, Europe; Lom et al. (1976): nasal fossae, Cyprinus carpio, Bulgaria. ■ T. microdenticulata Wellborn, 1967: Dorosoma petense, small, 26 pm in diameter, LA. ■ T. mutabilis Kazubski and Migala, 1968: Cyprinus car¬ pio, seasonal occurrence, variability, Poland; Chen (1956b): Ctenopharyngodon idella, Hypophthahnichthys molitrix, China; Kostenko (1971): Tinea tinea, Danube River, former Soviet Union; Lom (1970): C. carpio, Caras¬ sius auratus, review, Czech Republic and Slovakia; Lom et al. (1976): C. carpio, Bulgaria; Migala (1970): C. carpio, population influenced more by light and temperature than fish density, Poland; van As and Basson (1984); C. auratus, South Africa; Vismanis et al. (1975): C. carpio, former Soviet Union. ■ T. nigra Lom, 1961: Cobitis taenia, Czech Republic and Slovakia; Arthur and Lom (1984): on gills Abramis ballems, Radius nitilus, former Soviet Union; BykhovskayaPavlovskaya (1962): including three subspecies, former Soviet Union; Chen (1984); grass carp, China; Hoffman and Lom (1967): Acantharcus pomotis, WV; Kazubski (1967): seasonal variation, Poland; Kazubski and Migala (1968): C. carpio, seasonal variation, Poland; Lom (1970): discussion of four subspecies, Europe, former Soviet Union; Lom and Hoffman (1964): on gills, Lepomis macrochirus, Micropterus salmoides, WV; Migala (1970): population influenced more by environmental factors

48

Subkingdom Protozoa

than carp density, Poland; Mukherjee and Haidar (1982): West Bengal, India. This species, comprising four sub¬ species, is 50-90 pm in diameter, with outer blades rounded and 8-10 striations per denticle. ■ T. nobilis Chen, 1963: skin, rarely gills, Cyprinus carpio, Ctenopharyngodon idella, Hypophthalmichthys molitrix, frog tadpoles, China; Lorn et al. (1976): C. idella, C. carpio, H. molitrix, Bulgaria; Shtein (1968): Carassius auratus, C. idella, C. carpio, H. molitrix, Leuciscus waleckii, former Soviet Union. ■ T. noturi Wellborn, 1967: Notions leptocanthus, AL. ■ T. opeongoensis Li and Desser, 1985a: in ureters, uri¬ nary bladder, Notemigonus crysoleucas, Semotilus atromaculatus, Ont. ■ T. ovaliformis Chen, 1955: Ctenopharyngodon idella, China; Li and Desser (1985a): gills, Notropis comutus, Perea flavescens, Ont.; Lorn (1970): nomen dubium until silver impregnation done; Musselius (1967): Hypophthalmichthys molitrix, C. idella, former Soviet Union. ■ T. pediculus (O. F. Muller, 1786) Ehrenberg, 1838. This large species about 100 pm in diameter, has been found on many fishes, Hydra, and tadpoles in Europe and North America; H. S. Davis (1947): Muller's material on Micropterus salmoides was T. fultoni. ■ T. platyformis H. S. Davis, 1947: on gills, Margariscus margarita, Rhinichthys atronasus, WV. ■ T. reticulata Hirschmann and Partsch, 1955: Carassius auratus, Europe; Lorn and Hoffman (1964): same, WV. ■ T. hnttae Mueller, 1937c: on gills, Oncorhynchus clarki, OR; H. S. Davis (1947): O. clarki; Bogdanova and Stein (1963): salmon, Sakhalin Island, Russia. ■ T. tumefaciens H. S. Davis, 1947: on gills, Cottus bairdi, WV. ■ T. vallata H. S. Davis, 1947: on gills, Ictalurus punctatus, IA. ■ Trichodina spp.(?). Reported from Ambloplites rupestris, Campostoma anomalum, Culaea (Eucalia) inconstans, Cyprinus carpio, Esox Indus, E. masquinongy, Etheostoma nigrum, Hyborhynchus notatus, Hypentelium nigricans, Lepomis gibbosus, L. macrochirus, Micropterus dolomieui, Notions gyrinus, Perea flavescens, Percina caprodes, Poecilichtliys exilis, Salvelinus fontinalis, Semotilus atromaculatus, Stizostedion vitreum by Richardson (1938), Bangham (1944), Fantham and Porter (1947), Fischthal (1947a, 1950c), Bangham and Adams (1954), Anthony (1963), and others.

Genus Trichodinella Sramek-Husek, 1953

(Syn. Brachyspina Raabe, 1950) (Figs. 51 d, 54, 61)

Adoral spiral making a turn of 180 to 270 degrees; den¬ ticles each possessing delicate central part, with anterior right border extending into a projection and thereby fit¬

ting into notch between central part and blade of pre¬ ceding denticle. Inner thorn not stout and straight as in Trichodina, instead forming short, delicate hook curved outward; small hook not easily distinguished in live ciliates and difficult to impregnate with silver. Macro¬ nucleus generally horse-shoe shaped. Body a more or less flattened or vaulted hemisphere. Exclusively on fish gills, usually clamped over end of lamella; on fresh¬ water, rarely marine, fishes. — Ref. Lorn and Haidar (1977).

■ Trichodinella carpi (Duncan, 1977): on gills, Cyprinus carpio, Philippines. Distinct from other Trichodinella species in complete absence of centripetal projection. Body low, hat-shaped; diameter 31 pm; adoral groove turning fewer than 180 degrees; denticular ring 8 pm; 18 denticles present, length 2.7 pm. ■ T. epizootica (Raabe, 1950) Sramek-Husek, 1953. Arthur and Lorn (1984): gills, Perea fluviatilis, Lota lota, Rybinsk, Russia; Calenius (1980a): gills, Acerina cemua, Esoc Indus, Lota lota, brackish water and freshwater; Hoffman and Migala (unpub. 1976): C. auratus, AR; Lorn (1964): buc¬ cal ciliature; Lom and Haidar (1977): on 18 fish species, including Carassius auratus, Cyprinus carpio, Esox Indus, Oncorhynchus nerka, Salmo trutta, Tinea tinea, Europe; van As and Basson (1984): C. carpio, South Africa. ■ T. minuta (Chen, 1956) Kostenko, 1969: on gills, Aristichthys nobilis, Hypophthalmichthys molitrix, China; Lom and Haidar (1977): review, body 26-35 pm, adhesive disc 11-21 pm, 23-24 denticles present, blade 4.4 pm, pos¬ sibly syn. of T. epizootica. ■ T. myakkae (Mueller, 1937) Raabe, 1950: on gills, Carpiodes carpio, Ictiobus bubalus, Micropterus salmoides, Salvelinus fontinalis, FL; Chen (1984): on seven fish species, including Carassius auratus, Cyprinus carpio; H. S. Davis (1947): IA, NY, WV; Lom and Haidar (1977): in absence of silver impregnated specimens, is consid¬ ered nomen dubium; possibly T. epizootica. ■ T. subtilis (Lom, 1959) Lom and Haidar, 1977: on Carassius carassius, Czech Republic and Slovakia; Kazubski and Migala (1968): Cyprinus carpio, seasonality and variation, Poland; Lom and Hoffman (1964): Carassius auratus, WV; Lom and Haidar (1977): on seven fish species, including C. carpio, Oncorhynchus mykiss, Eura¬ sia; Migala (1970): density of carp population has less influence than light and temperature, Poland. ■ T. symmetrica (Davis, 1947) Lom, 1959: originally described as Trichodina symmetrica from gills of Ictalurus punctatus, Rhinichthys atronasus, Semotilus margarita, IA, WV; Lom and Haidar (1977): is nomen nudum because Davis's specimens seem to be a mixture of Tripartiella and Trichodinella; see Figs. 50, 51, and 138 in H. S. Davis (1947). ■ T. tilapiae Duncan, 1977: on gills, Tilapia zilli, Philip¬ pines, most closely related to T. subtilis, but the finely drawn out central part of the denticle does not bend acutely forward.

Group Sporozoa

Genus Tripartiella

Genus Vauchomia

Lorn, 1959

Mueller, 1938

(Syn. Trichodina Ehrenberg, 1831, in part)

(Figs. 51 f, 55)

(Figs. 51 e, 52, 62)

Adoral spiral making a turn of 180 to 290 degrees. Del¬ icate central part of denticle bearing a straight thorn, directed backward in most species; blades slanted obliquely backward; narrow base by which they join cen¬ tral part of the denticle extending anteriorly in a pro¬ jection that may be short and thin or wide and kneelike; projections fitting well into corresponding notch in preceding denticle. Macronucleus generally horseshoe¬ shaped; body with the shape of a flattened hemisphere, or sacklike with a deeply vaulted adhesive disc. Exclu¬ sively on gills of freshwater fishes. — Ref Lorn (1963); Lom and Haidar (1977): synopsis.

■ Tripartiella bulbosa (H. S. Davis, 1947): on gills, Semotilus margarita, WV; Chen (1956a): four species of grass carp, China; Ivanova (1966): grass and silver carp; Lom and Haidar (1977): grass carp, Abramis brama, Hun¬ gary; Mukherjee and Haidar (1982): five species of native fishes, West Bengal, India; outer process of denticle rod¬ like, inner process short; 18-24 denticles present; body 16-27 pm in diameter and height. ■ T. bursiformis (H. S. Davis, 1947) (Syn. Trichodina b.)\ on gills, Ambloplites rupestris, WV; Chen (1984): five species of native fishes, including Aristichthys nobilis, Hypophthalmichthys molotrix, China; Hoffman and Lom (1967): Acantharcuspomotis, NC; Lom and Haidar (1977): description; much like T. bulbosa but larger and body compressed laterally. ■ T. kashkovskyi Lom and Haidar, 1977: gills, Fundulus dispar, Mississippi River, IL; differing from T. bulbosa in having longer, thinner thorns of denticles; differing from T. bursiformis in having kneelike junction in cen¬ ter of denticle and in being smaller. ■ T. lata Lom, 1963: Phoxinus laevis, Czech Republic and Slovakia; Calenius (1980a): Osmerus eperlanus, fresh¬ water, Linland; Lom (1970): Ctenopharyngodon idella, Czech Republic and Slovakia; Lom and Haidar (1977): gills, Pimephales vigilax, MI; differing from T. bulbosa and T. bursiformis in having slender, tapering, back¬ ward-slanting thorns and in having a rounded "knee" of denticle projecting forward. ■ T. symmetricus H. S. Davis 1947) (Syn. Trichodina s.): on gills, Ictalurus punctatus, Semotilus marginata, Rhinichthys atratulus, I A, WV; Li and Desser (1985b): gills, fins, skin, Perea flavescens, Semotilus atromaculatus, Ont.; Lom and Haidar (1977): nomen nudum because the original description is a mixture of Tripartiella and Trichodinella; for evidence, see H. S. Davis (1947, Figs. 50-51, 138, 139).

49

Similar to Trichodina, except with buccal ciliary spiral making two to three complete turns; parasitic in the uri¬ nary system. This genus is retained on the advice of Lom (1964), although Uzmann and Stickney (1954) recommend reduction to subgenus. ■ Vauchomia nephritica Mueller, 1938b: in urinary blad¬ der, Esox masquinongy, NY; Lom and Haidar (1976): redescription, NY. ■ V. renicola Mueller, 1932: in urinary tract, Esox spp., NY; Bangham (1944) and Fischthal (1947a): Esox mas¬ quinongy, WI; Crossman (1962): E. americanus vermiculatus, Ont.; Fantham and Porter (1947): Trichodina sp. in urinary tract, E. niger, Que.; Hunter and Rankin (1940): Esox spp., CT.; Lom and Haidar (1976): E. niger, silver stain of adoral disc, NY; Sindermann (1953): E. niger, MA.

Group Sporozoa For many years protozoa that formed spores and had no organelles of locomotion were placed in Class Sporozoa Leuckart, 1879 (see Kudo, 1971, and Lee et al., 1985). Some stages of these organisms do possess organelles of locomotion, however, and because electron microscopy has revealed some organelles useful for grouping. A new taxonomic system has therefore been devised (Lee et al. 1985; Levine et al., 1980).

Key to the Groups of Sporozoa 1. Apical complex of sporozoites visible with electron microscopy; spores usually not in cysts (Eimeria, Goussia, Haemogregarina, Babesiosoma, Dactylosoma, Sarcocystis) .(p. 50) Phylum Apicomplexa 1. Apical complex absent, spores usually in cysts that are often visible without magnification.2 2. (1) Spores oval, comparatively large (6-65 pm); one to six polar capsules usually easily seen at magnifications as low as 400x .(p. 55) Phylum MyxoZoa 2. (1) Spores oval, comparatively small (3-8 pm) .(p. 81) Phylum Microspora 2. (1) Spores spherical (3-12 pm) with large vacuoles displacing nucleus to edge (Dermocystidium) .(p. 89) Sporozoa of uncertain classification

Subkingdom Protozoa

50

Phylum Apicomplexa Levine, 1970 (in Levine, 1985) Apical complex visible with electron microscopy, gen¬ erally consisting of polar ring(s), two to eight rhoptries (electron-dense tubular, saccular, or club-shaped organ¬ elles), numerous micronemes (convoluted, elongated electron-dense organelles); conoid or subpellicular micro¬ tubules present at some stage. Microspores generally present at some stage. All species parasitic.

pendently; zygote not motile; sporozoites typically enclosed in sporocyst within oocyst; homoxenous (one host life cycle) or heteroxenous (more than one host in the life cycle). Stieda body (structure that plugs the sporocyst aperture pending sporozoite release) present or absent. — Ref Dykova and Lom (1983): list of species from fishes; Lom and Dykova (1992).

Genus Calyptospora Overstreet, Hawkins, and Fournie, 1984 (Fig. 63)

Subclass Coccidia Leuckart, 1879 Gamonts (the gamete-producing stages) ordinarily pres¬ ent; mature gamonts small, typically intracellular. Life cycle characteristically consisting of merogony (pro¬ duction of merozoites from an antecedent cell), gametogony (production of gametes), and sporogony (production of spores). Most species parasites of verte¬ brates, including fishes.

Sporocyst covered with a thin veil supported by one or more sporopodia (elongated projections of sporocyst wall); lacking Stieda body; possessing membranecovered, distinct, oblong opening, with elongated por¬ tions of rim abutted in fresh material. Suture extending a short distance distally on opposite sides before trans¬ forming into low, protruding ridges that continue around posterior end and converge or join another extension of sporocyst wall. Sporozoite developing in invertebrate digestive tract; definitive host a marine or brackish water fish. ■ Calyptospora empristica Fournie, Hawkins and Overstreet, 1985: hepatocytes and pancreatic acinar cells, Fundulus notti, MS.

Order Eucoccidiida Leger and Duboscq, 1910 (or Order Eimeriida Leger and Duboscq, 1911) Merogony present (production of merozoites from an antecedent schizont).

Suborder Eimeriina Leger, 1911 Macrogamete (single female gamete) and microgamont (cell producing many male gametes) developing inde-

Key to the Genera of Eimeriina of Fishes

■

C. funduli (Duszynski, Solangi, and Overstreet, 1979) Overstreet, Hawkins, and Fournie, 1984 (Syn. Eitneria funduli Duszynski, Solangi, and Overstreet, 1979): Fundulus grandis, MS, AL, VA; Duszynski et al. (1981): experi¬ mental infection of intermediate host, grass shrimp, Palaemonetus pugio; Dykova and Lom (1983): review; Fournie et al. (1983a): true invertebrate intermediate host; Hawkins et al. (1981): ultrastructural effect on liver of host; Hawkins et al. (1983a): ultrastructure of macrogamont; Hawkins et al. (1983b): ultrastructure of oocysts; Hawkins et al. (1984): intrahepatic stages in fishes; Hawkins et al. (1984a, 1984b): ultrastructure of parasitehost interface; Overstreet et al. (1984): Calyptospora n. g. and Calyptosporidae n. fam.; Solangi and Overstreet (1980): biology and pathogenesis; Solangi et al. (1982): effect of low temperature; Upton and Duszynski (1982): reviewed as Eimeria funduli.

Sporocysts not bivalved; with or without Stieda body .(Figs. 64, 65)

Eimeria

Genus Eimeria

Sporocysts bivalved; without Stieda body .... (Fig. 66)

Goussia

Schneider, 1875

Similar to

(Figs. 64, 65)

Eimeria, but with epicellular

merogony and gamogony; sporogony intracellular, as in Eimeria.(no fig.) Similar to

Epieimeria

Goussia, but with incomplete bivalve condition;

with anterior apical opening and a veil supported by sporopodia; requiring an invertebrate intermediate host .(Fig. 63)

Calyptospora

Mainly in cells of the digestive tract, although some¬ times found in kidneys, gonads, liver, spleen, gall blad¬ der, and even gills (£. branchiphiala). Oocyst containing four sporocysts, with two sporozoites in each; sporo¬ zoites oval to vermiculiform. Oocyst wall in fishes extremely thin, unlike in homotherms, and usually possessing no aperture (micropyle) for sporozoite

so

ss

MO

10 pm

63. Calyptospora

69. Babesiosonta

70. Dactylosoma

-Coccidia and haemosporidiaFIG. 63. Calyptospora fUnduli, diagrammatic representation of a sporulated oocyst; note unique footlike "sporopodia" on sporocysts (from Duszynski et al., 1979). FIG. 64. Eimeria oocyst, schematic: MO ,= micropyle of oocyst, MS = micropyle of sporocyst, P = microphyle end, S = sporozoite, So = oocyst wall, SS = sporocyst wall, ZO = residual body of oocyst, ZS = residual body of sporocyst (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences). • FIG. 65. Eimeria sporocyst, schematic: S = Steida body, SS = sub-Steida body (from Dykova and Lorn, 1981, J. Fish Dis., by permission of the Czechoslovak Academy of Sciences). FIG. 66. Goussia sporocyst, schematic: SL = suture line of the two valves (from Dykova and Lorn, 1981). FIG. 67. Haemogregarina catostomi, in red blood cell, N = nucleus (redrawn from C. D. Becker, 1962). FIG. 68. Sarcocystis salvelini, cross section through cysts in muscle (from Fantham and Porter, 1943). FIG. 69. Babesiosoma tetragonis, in red blood cells of Catostomus: a, typical rosette stage, b, separated merozoites, N = nucleus (drawn from Becker and Katz, 1965b). FIG. 70. Dactylosoma salvelini, in red blood cell, N = nucleus (redrawn from Fantham et al., 1942).

52

Subkingdom Protozoa

release. Sporocyst wall with Stieda body (a structure to plug the aperture for sporozoite release) at one end, usu¬ ally encircled by a prominent thickening in the wall. Depending on the species, oocyst possibly containing residual body: coarse, granular material formed by sub¬ stances left from the original division of the zygotes to form spores. Sporocysts sometimes possessing a simi¬ lar cluster of residual granules from the division of spores into two sporozoites. Life cycle: when an oocyst is swallowed by a fish, the sporozoites are released from the sporocyst and attack the cells of the intestinal epithelium; with the aid of lym¬ phocytes and blood circulation, the sporozoites may penetrate the cells of other organs. In the cell, the sporo¬ zoite grows to become an oval schizont (asexual repro¬ ductive cell); following schizogenesis, the schizont splits into many merozoites (slender vermiform cells), which penetrate the cells of adjacent tissue and there grow again and transform into schizonts. Schizogenesis may recur in this manner several times, but the number of merozoite generations is limited, depending on the species. Following the terminal merozoite cycle, gametogenesis occurs, producing macrogametocytes (female, single gamete) and microgametocytes (male) containing many microgametes; those the author has seen were biflagellated. After fertilization, the zygote forms a strong membrane and becomes an oocyst as described earlier; the sporulated oocysts leave the fish and are able to survive in the water. — Ref. Desser (1981); Ergens and Lom (1970); Kudo (1971); J. L. Lee et al. (1985); Upton et al. (1984): general. See also P. Cheng, (pers. comm., N.Y. Aquarium, 1982): stain smears with Ziehl-Neelsen stain, sporocysts red; Doran (1970): E. tenella survives freezing; Doran (1974): pan¬ creatic extract causes excystation to free sporozoites, culture of E. tenella; Dykova and Lom (1980): histopathology; Dykova and Lom (1981): classification and patho¬ genesis; Hawkins et al. (1981): EM of E. funduli; Long and Joyner (1984): problems in identifying Eimeria of birds; Molnar (1977a): methods; Molnar (1979a): experimen¬ tal infection by E. carpelli; Molnar (1981): oocyst discharge and histopathology; Paterson and Desser (1982): meth¬ ods, experimental infections, temperature dependency; Patton and Brigman (1979): excystment of E. tenella sporozoites; Santos Pinto (1956): parasitic castration by E. sardinae; Speer (1979): E. magna in cell culture; Stotish and Wang (1975): preparation and purification of mero¬ zoites of E. tenella; Upton et al. (1984): review and key to species.

■ Eimeria aculeata Jastrzebski, 1984: intestine, Gasterosteus aculeatus, Poland. ■ E. anguillae: transfer to Epieimeria by Dykova and Lom (1981).

■ E. carassiaurati Romero Rodriguez, 1978: intestine, Carassius auratus, Spain; no Stieda body, oocyst 14.8-15.2 pm x 13.3 pm, sporocyst 13.6 x 5.8 pm, causing haem¬ orrhagic enteritis. ■ E. catostomi Molnar and Hanek, 1974: in intestine, Catostomus commersoni, Hypentelium nigricans, no Stieda body; Dykova and Lom (1983): review; Upton et al. (1984): review. ■ E. chenchingensis Chen, 1984: in spleen, Carassius auratus, oocyst ovate and 21-24 x 17-19 pm, China. ■ E. cheni Shulman and Zaika, 1962 (Syn. E. intestinalis Chen, 1956): anterior intestine, Aristichthys nobilis, Hypophthalmichthys molitrix, Mylopharyngodon piceus, no Stieda body, former Soviet Union; Dykova and Lom (1983): review, China, Hungary. ■ E. cotti Gauthier, 1921: caecal opening, Cottus gobio, France; Arai and Mudry (1983): pyloric caeca, C. cognatus, B.C.; Dykova and Lom (1983): review, Stieda body present. ■ E. duszynskii Conder, Oberdorfer, and Heckmann, 1980: intestine Cottus bairdi, no Stieda body, oocysts irregular in shape, UT; Dykova and Lom (1983): review; Heckmann et al. (1987): C. bairdi, UT; Upton et al. (1984): review. ■ E. esoci Shulman and Zaika, 1962 in BykhovskayaPavlooskaya (1962): intestine, urinary bladder wall, Esox lucius, Stieda body present, former Soviet Union; Li and Desser (1985b): caecum, gall bladder, gills, intestine, heart, kidney, liver, muscle, spleen, swim bladder, Lepomis gibbosus, Ont. ■ £. etheostomae Molnar and Hanek, 1974: intestine, Etheostotna exile, E. nigrum, Ont.; Dykova and Lom (1983): review, no Stieda body; Upton et al. (1984): review. ■ E. fernandoae Molnar and Hanek, 1974: intestine Catostomus commersoni, Hypentelium nigricans, Ont.; Dykova and Lom (1983): review, no Stieda body; Upton et al. (1984): review. ■ E. freemani Molnar and Fernando, 1974: syn. of Goussia leucisci. ■ E. funduli; see Calyptospora funduli. ■ E. gasterostei (Thelohan, 1890) Doflein, 1909: liver, Gasterosteus aculeatus, France; Dykova and Lom (1983): review, no Stieda body; Jastrzebski and Pastuszko (1987): light and electron microscopy; Lester (1974): G. aculea¬ tus, B.C.; Upton et al. (1984): review. ■ E. glenorensis Molnar and Fernando, 1974: intestine, Morone americana, Ont.; Upton et al. (1984): review.

■ E. aristichthysi Lee and Chen, 1964: Aristichthys nobilis, Hypophthalmichthys molitrix, no Stieda body, China.

■ E. haichengensis Chen, 1984: intestine, Cyprinus carpio, China; Dykova and Lom (1983): review, no Stieda body.

■ E. aurati G. L. Hoffman, 1965: Carassius auratus, intes¬ tine, no Stieda body, oocysts in long, wormlike, white intestinal cast, PA.

■ £. haneki Molnar and Fernando, 1974: intestine, Culaea inconstans, Ont. Dykova and Lom (1983): review, no Stieda body.

Suborder Eimeriina

53

■ E. hofjmani Molnar and Hanek, 1974: intestine, Umbra limi, Ont.; Dykova and Lom (1983): review, no Stieda body; Upton et al. (1984): review.

■ E. sinensis Chen, 1956b: intestine, Aristychthis nobilis, Hypophthalmichthys molitrix, China; Molnar (1976a): histology.

■ E. hupensis Chen and Hsieh, 1964: intestine, Carassius auratus, China; Dykova and Lom (1983): review, no Stieda body.

■ E. spleni: syn. of Goussia degiustii.

■ E. hybognathi Molnar and Fernando, 1974: intestine, Hybognathus hankinsoni, Ont.; Dykova and Lom (1983): review, no Stieda body. ■ E. hypophthalmichthys Akhmerov, 1959: kidney, Hypophthalmichthys molitrix, former Soviet Union; Dykova and Lom (1983): review, no Stieda body. ■ E. ictaluri Molnar and Fernando, 1974: intestine, Ictaluras nebulosus, Ont.; Dykova and Lom (1983): review, no Stieda body.

■ E. tedlai Molnar and Fernando, 1974: intestine, Perea flavescens, Ont.; Dykova and Lom (1983): review, Stieda body present; Jastrzebski (1984a): P. fluviatilis, Poland. ■ E. truttae (Leger et Hesse, 1919): Stankovich, 1924: pyloric caeca, intestine, Salmo trutta, France; Awakura et al. (1983): pyloric caeca, Oncorhyjichus mason, Japan; Molnar and Hanek (1974): intestine, Salvelinus fonti¬ nalis, Que.; Upton et al. (1984): review. ■ E. zarnowski Jastrzebski, 1982: intestine, Gasterosteus aculeatus, Poland.

■ E. laureleus Molnar and Fernando, 1974: intestine, Perea flavescens, Ont.; Stieda body not described.

■ Eimeria sp. H. S. Davis (1946): intestine and pyloric caeca, Salvelinus fontinalis, VT.

■ E. liaohoensis Chen, 1984: intestine, Carassius auratus, Pseudogobio rivularis, Hypseleotis sp., China.

■ Eimeria, sp. Fantham and Porter (1947): Fundulus heterocleitis, N.S.; Duszynski et al. (1979): may be E. funduli.

■ E. micropteri Molnar and Hanek, 1974: intestine, Micropterus dolomieui, M. salmoides, Ont.; Dykova and Lom (1983): review, no Stieda body.

■ Eimeria sp. Fantham and Porter (1947): Fundulus heteroclitus, N.S.

■ E. moronei Molnar and Fernando, 1974: intestine, Morone americana, Ont.; Dykova and Lom (1983): review, Stieda body present. ■ E. newchongensis Chen, 1984: in liver, Carassius aura¬ tus, Erythroculter erythropterus, China. ■ E. ojibwana Molnar and Fernando, 1974: intestine, Coftus bairdi, Ont.; Dykova and Lom (1983): review, no Stieda body. ■ E. osmeri Molnar and Fernando, 1974: intestine, Osmerus mordax, Ont. ■ E. patersoni Lom, Desser, and Dykova, 1989: in renal tubule cells, spleen, liver parenchyma, Lepomis gibbosus, oocyst 11.9 x 10.6 pm, Ont. ■ E. percae (Dujarrie de la Riviere, 1914) Reichnow, 1921: intestine, Perea fluviatilis, former Soviet Union; Dykova and Lom (1983): review, no Stieda body; Jastrzebski (1984a): P. fluviatilis, Poland; this species included because the European P. fluviatilis and Ameri¬ can P. flavescens are so similar. ■ E. pungitii Molnar and Hanek, 1974: intestine, Pungitius pungitius, Que.; Dykova and Lom (1983): review, no Stieda body; Jastrzebski (1984a): Poland. ■ E. rouxi (Elmassian, 1909) Reichnow, 1921: intestine, Tinea tinea, France; Dykova and Lom (1983): review, no Stieda body. ■ E. salvelini Molnar and Hanek, 1974: intestine, Salvelinus fontinalis, Que.; Dykova and Lom (1983): review, Stieda body present. ■ E. saurogobii Chen, 1964: intestine, Ctenopharyngodon idella, China; Dykova and Lom (1983): review, no Stieda body.

■ Eimeria sp. Fantham and Porter (1947): intestine, Salvelinus fontinalis, Que. ■ Eimeria sp. G. L. Hoffman unpub., (1978): liver, cul¬ tured Polyodon spathula, collection by C. Suppes, MO Dep. of Conservation, specimens sent to R. Overstreet, Gulf Coast Research, Ocean Springs, MS 39564-7000, biflagellated microgametes and oocysts found in visceral ascitic fluid. ■ Eimeria sp. Li and Desser (1985b): swim bladder, Notemigonus crysoleucas, Ont. ■ Eimeria sp. Molnar et al. (1974): Etheostoma exile, Ont.

Genus Epieimeria Dykova and Lom, 1981 (No fig.)

Similar to Eimeria; tetrasporocystic; dizoic; sporocysts with Stieda body. Merogony and gamogony completed in epicellular state; sporogony intracellular. Parasites of fishes. — Ref. Dykova and Lom (1981).

■ Epieimeria anguilllae (Leger and Holland, 1922) Dykova and Lom, 1981. Dykova and Lom (1983): review; Hanek and Molnar (1974): intestine, Anguilla rostrata, Que.; Hine (1975c): Anguilla australis, A. dieffenbachii, New Zealand; Hine and Boustead (1974): New Zealand; Jas¬ trzebski (1984a): A. anguilla, Poland; Lacey and Williams (1983): A. anguilla, England; Molnar and Baska (1986): light and electron microscopy, Hungary; Molnar and Hanek (1974): A. rostrata, Que.

54

Subkingdom Protozoa

Genus Goussia Labbe, 1896 (Fig. 66)

Tetrasporocystic, dizoic; sporocysts without Stieda body; walls consisting of two valves joined meridionally. Merogony, gamogony and sporogony completed within host tissue. Bivalved sporocyst distinctive to genus. Parasites of Chondrichthyes and Teleostei. — Ref. see Eimeria; Dykova and Lom (1981): revalidation of genus Goussia.

■ Goussia acipenseris Molnar, 1986: intestine, Acipenser ruthensis, Europe. ■ G. carpelli (Leger et Stankovitch, 1921) Dykova and Lom, 1983: intestine, seven fish species, including Cypri¬ ans carpio, Hypophthalmichthys molitrix, Europe, former Soviet Union; Alvarez-Pellitero and Gonzalez-Lanza (1986) in cyprinids, host-parasite relationships, Spain; Jastrzebski (1984a): in Carassius auratus, Cyprians carpio, Gobio gobio, Poland; Kent and Hedrick (1985b): C. aura¬ tus, life cycle in tubifex and grass shrimp, CA; Lom and Dykova (1982): C. carpio, structure of sporocysts and "yel¬ low bodies," Czech Republic and Slovakia; Molnar (1981): C. carpio, oocyst rejection and histopathology, Hungary; Steinhagen and Korting (1988): C. carpio, experimental transmission, Germany. ■ G. cichlidarum Landsberg and Paperna, 1985: swim bladder of cichlids, including Oreochromis aureus, O. aureus x O. ailoticus, Tilapia zilli, Israel. ■ G. degiustii (Molnar and Fernando, 1974) Dykova and Lom, 1983 (Syn. Eimeria degiustii Molnar and Fer¬ nando, 1974; E. spleai Degiusti and Gnath, 1968 = aomea nudum): spleen, Campostoma aaomalum, Notropis corautus, MI; Li and Desser (1985a): spleen, gall bladder, intestine, liver, N. corautus, N. heterolepis, Semotilus atromaculatus, Ont.; Lom (1971b): EM of spore envelope; Lom, Desser, and Dykova (1989): in spleen, kidney, pancreatic islets, swim bladder, N. corautus, description of oocysts and sporocysts, Ont.; Molnar and Fernando (1974): spleen, N. corautus, Pimephales promelas, Ont.; R. Walker (pers. comm., Rensselaer Polytechnic Institute, 1983): spleen, Notropis hudsoaius, NY.

Lepomis gibbosus, Perea flavescens, Semotilus atromacu¬ latus, Ont. ■ G. leucisci (Shulman and Zaika, 1964) (Syn. Eimeria freemaai Molnar and Fernando, 1974; Eimeria leucisci Shul¬ man and Zaika, 1964): in kidney tubule, gall bladder, no Stieda body, European cyprinids; in kidney, Notropis cor¬ autus, oocyst 23 x 17 pm, Ont.; Lom, Desser, and Dykova (1989): kidney interstitium, lumen of renal tubules, N. corautus, Ont. ■ G. aotemigoaica Li and Desser, 1985a: kidney, spleen, swim bladder, ureters, Notemigonus crysoleucas, Ont. ■ G. uotropicum Li and Desser, 1985: intestine, Notropis cormttus, Ont. ■ G. subepithelialis (Moroff et Fiebiger, 1905) Dykova and Lom, 1983 (Syn. Eimeria subepithelialis Moroff et Fiebiger, 1905): intestine, Cypriaus carpio and probably Tinea tiaca, Europe; Molnar (1982): intestinal nodular coccidiosis of T. tiaca, but differing from that caused by C. carpio, Hungary. ■ G. vargai Molnar, 1986: intestine, Acipenser ruthenus, Hungary. ■ Goussia spp. Li and Desser (1985a): Goussia sp. "a" and "b" in swim bladder of N. corautus, Ont.

Genus Haemogregarina Danilewsky, 1885 (Fig. 67, Color Fig. 16)

Schizogony occurring in blood cells of vertebrates; cyto¬ plasm and nuclei staining readily; developmental stages relatively large, ovoid to slender vermiform; one end sometimes clubbed, the other sometimes attenuated, but both rounded; larger ones sometimes reflexed around nucleus of red blood cell. Sexual reproduction occuring when gametocytes taken into gut of leech; microgametocyte developing two to four microgametes; sporo¬ zoites forming without production of spores. Usually studied in stained blood smears of fishes or frogs. — Ref. Becker (1970): review of fish hematozoa; Kudo (1971, p. 706).

■ G. iroquoiaa (Molnar and Fernando, 1974) Paterson and Desser, 1984: intestine, Notropis corautus, Ont.; Mol¬ nar and Hanek (1974): Nocomis biguttatus, Notropis het¬ erolepis, N. rubellus, Pimephales aotatus, Rhiaichthys atratulus, Semotilus atromaculatus, Ont.; Molnar, et al. (1974): Notropis biguttatus, N. corautus, N. heterolepis, P. aotatus, P. promelas, Ont.; Paterson and Desser (1981a): experimental infection, P. promelas, EM of microgametes; Paterson and Desser (1981b): EM of macrogametes, P. promelas; Paterson and Desser (1982): experimental infection and development; Paterson and Desser (1984): EM of fertilization and sporulation, transfer to Goussia.

■ Haemogregarina sp. Boggs and Myers (1968): Ictalurus punctatus, 17% infected, VA.

■ G. laruelensis (Molnar and Fernando, 1974) Li and Desser, 1985a: intestine, caecum, gall bladder, liver,

■ Haemogregarina sp. Fantham and Porter (1947): Salveli¬ nus fontinalis, Que.

■ Haemogregarina catostomi C. D. Becker, 1962: in red blood cells, Catostomus macrocheilus, WA; C. D. Becker (1980): C. macrocheilus, C. columbianus, WA. ■ H. cyprini Smirnova, 1971: a. sp. and effect on carp; Lom et al. (1976): Cypriaus carpio, Europe. ■ H. irkaluppiki M. Laird, 1961; Salvelinus alpinus, north¬ ern Canada; Arai and Mudry (1983): S. malma, Canada; Arthur et al. (1976): Coregonus clupeaformis, Prosopium cylindraceum, B.C.

Phylum Myxozoa

■ Haemogregarina sp. Heckmann (1971b): Onchorhynchus clarki (=Salmo clarki), Prosopium williamsoni, but not other spp., WY; Heckmann and Ching (1987): O. clarki (=S. clarki), WY. ■ Haemogregarina sp. McCarthy (1974): Salmo salar smolts, England.

Family Sarcocystidae Asexual development in prey host; sexual development in predator host. Sarcocystis reported once from fishes; Toxoplasma /Isospora not in fishes.

Genus Sarcocystis Lankaster (Fig. 68) Muscle parasites of vertebrates; spores found in opaque white cysts (Meischer's tubes), cylindrical to ovoid. When mature, banana-shaped spores filling cyst; spores containing nucleus and granules. Many species. ■ Sarcocystis salvelini Fantham and Porter, 1943: in muscle, Salvelinus fontinalis, Que. This has not been reported for 46 years, so it may be in error.

Family Dactylosomatidae (Jakowska and Nigrelli, 1955) Becker, 1970 Parasitic in erythrocytes and sometimes in other circu¬ lating cells and fixed cells. Pyriform, round, rod-shaped, or amoeboid; without conoid, oocysts, spores, pseudo¬ cysts, or flagella; with polar ring and rhoptries; loco¬ motion by body flexion, gliding; heteroxenous, with merogony in vertebrate host and sporogony in inver¬ tebrate host; sporozoites with single-membraned wall; vectors of species in fishes unknown. — Ref. J. L. Lee et al. (1985), Levine (1961).

55

Genus Dactylosoma Labbe, 1894 (Fig. 70) Small parasites, not filling red blood cells. Cytoplasm and nucleus staining faintly, with no pigment; 4 to 16 nuclei present; gametocytes with karyosomes; 4 to 16 merozoites in fanlike arrangement. Invertebrate host unknown. ■ Dactylosoma marine Hoare, 1930. J. R. Baker (1960): Tilapia spp., East Africa. ■ D. salvelini Fantham et al., 1942: in red blood cells, brook trout, Que.; Fantham and Porter (1947): same.

Key to Sporozoa Other Than Coccidia and Haemosporidia Found in Fishes 1. Spores large (6-65 pm); polar capsules usually easily seen.Phylum Myxozoa 2 1. Spores about 10 pm; no polar capsules; spherical spore with large vacuole .(Sporozoa of uncertain classification)

Dermocystidium 1. Spores small (3-6 pm); no polar capsule; polar filament usually not easily visible, but demonstrated by extrusion and electron microscopy.Phylum Microspora 2. (1) In fishes, rarely amphibia; spores usually with two valves (like a clam shell) but rarely with up to eight valves; spore with one to eight, usually two, polar capsules.Class Myxosporea 2. (1) In invertebrates; included here because one has been reported as part of the life cycle of a myxosporea n, Myxobolus cerebral is .Class Actinosporea

Phylum Myxozoa Class Myxosporea Butschli, 1881

Genus Babesiosoma Jakowska and Nigrelli, 1956 (Fig. 69, Color Fig. 15) Small parasites, not filling red blood cells. Cytoplasm and nucleus staining faintly, with no pigment; cytoplasm less granular but more vacuolated than Dactylosoma; one to four Babesia-like nuclei present; no definite karyosome. Reproducing by schizogony, fission, and budding. Four or fewer merozoites arranged in rosette or cross. Inver¬ tebrate host unknown. ■ Babesiosoma tetragonis C. D. Becker and Katz, 1965b: Catostomus sp., CA, similar to B. marine of East Africa (see J. R. Baker, 1960), but with fan-shaped schizont.

Myxosporea are usually seen when the parasite is in the spore stage, at which time an opaque white cyst con¬ taining spores may be seen with the naked eye. Identi¬ fication is typically based on spore morphology. The histozoic species usually form cysts, sometimes very large, but species that develop in the gall bladder, kid¬ ney lobules, and urinary bladder do not form such cysts, and the trophozoite is the stage usually detected; iden¬ tification is still based mainly on the spore, however. The resistant proteinaceous spore wall is usually bivalve but rarely has up to eight valves. Fom and Vavra (1963a) demonstrated mucous envelopes on the spores; species differences were easily recognized. Each spore contains

56

Subkingdom Protozoa

one or two sporoplasms and one to four (and rarely more) polar capsules containing coiled filaments. Con¬ sult Lorn and Arthur (1989) for guidelines on describing new species or reviewing species for clarification. Spore measurements are important in species descrip¬ tions and later in identification (Fig. 71). Spores fixed in formalin, etc., however, shrink significantly. Such mea¬ surements should therefore be taken also on fresh spores that have been temporarily stored at room or refrigera¬ tor temperature, or more permanently stored in the deep freeze (Long and Meglitsch, 1970). As with many groups of fish parasites, some species of myxosporeans are highly pathogenic and others have not been proven harmful. In fish culture, Ceratomyxa shastn (salmon), Henneguya spp. (catfish), Hoferellus carassi (goldfish), Myxobolus argenteus (golden shiner), M. cerebrnlis (salmonids), and M. notemigoni (golden shiner) have been proven pathogenic. Species of questionable pathogenicity include Henneguya zscbokkei (salmon), Myxobolus arcticus (salmonids), M. cartilaginis (centrarchids), M. corneus (bluegill), M. hoffmani (fathead min¬ now), M. neurobius (salmonids), and M. squamae (salmon). It is seldom possible to assess parasite damage to feral fishes, but many myxosporeans produce no detectible damage in the wild. The life cycles of myxosporidans have been difficult to study because the direct, fish-to-fish cycles are unclear. Early studies indicated that Myxidium, Chloromyxum, and Leptotheca would infect fishes directly (see Kudo, 1930a, p. 313). Fish-to-fish transmission with spores of Ceratomyxa shasta, for example, was not possible, although susceptible salmon were easily infected when held for as little as 30 minutes in enzootic water (Bartholomew et al., 1989b; K. A. Johnson et al., 1979; K. A. Johnson, 1980). The infective stage has not been identified but will not pass through a 14 pm-pore filter. It was believed for many years that spores of Myxobolus cerebrnlis "aged" for 3Vi months were infective for trout (G. L. Hoffman and Putz, 1969; Prihoda, 1978, 1983; Uspenskaya, 1957, 1978), but Wolf and Markiw (1984) demonstrated that Tubifex is the obligate intermediate host for a Triactinomyxon stage of Myxobolus. Several researchers are working to clarify this life cycle, and some have presented corroborating evidence with other Myxobolus species. Myxosporean life cycles have been reviewed by Lorn and Dykova (1992). Development in the fish host has been studied in many myxosporean species. After reaching the target organ, the invasive sporoplasm grows into a trophozoite, whose nucleus divides repeatedly. Some nuclei become surrounded by dense cytoplasm and become sporonts; each sporont grows, with its nucleus dividing, to become the sporoblast, producing two to many spores, depend¬ ing on the species. Many species continue to produce spores within the host tissue envelope until a visible cyst is produced. Lumen-dwelling forms become large tropho¬ zoites, which produce one to many spores within the

trophozoite. Some species, particularly those that develop in the kidneys, do not regularly produce spores and are thus difficult to identify (e.g., Hoferellus carassii and Sphaerospora cyprini). Mature spores leave the fish in excreta, in the case of coelozoic species; some escape when cysts near the surface of the fish burst; and those located in tissues escape upon death and decay of the host or during digestion by fish-eating animals, in whose feces they pass into the water. In some histozoic species, e.g., Myxobolus cartilaginis and M. cerebrnlis, some spores are "forced" through tissue and escape through the gills or in the feces (G. L. Hoffman, Putz, and Dunbar, 1965; Uspenskaya, 1957). — Ref. Bykhowskaya-Pavlovskaya (1962): key; Kudo (1920): synopsis and keys; Kudo (1930a): methods; Kudo (1971): text; Lom and Arthur (1989): standardization of species descriptions; Lom and Dykova (1992); L. G. Mitchell (1977): review; Shulman (1966, 1984): synopsis.

See the Key to the Genera of Myxosporea of North American Freshwater Fishes on p. 57.

Order Bivalvulida Suborder Platysporina Kudo, 1919 Polar capsules (usually two, rarely one) opening at apex (anterior) of the spore, lying solely in the sutural plane of a bilaterally symmetrical spore; width usually greater than thickness. Generally histozoic parasites of freshwater fishes, producing large polysporous trophozoites (visible "cysts").

Family Myxobolidae Thelohan, 1892 Spores flattened parallel to the straight suture line; suture forming an elevated ridge and valves possibly drawn out into long projections ("tails"). One of the two polar capsules sometimes smaller; absent in two genera. Most species with an iodinophilous vacuole. Most Likely Sites of Myxobolus Species

Because of the large number of Myxobolus species, many have similar spores, and identification is sometimes dif¬ ficult. Some species tend to be site specific in the fish; others do not. In some cases, expecially when a cyst has ruptured, spores may be relocated by the movement of tissue, sometimes far removed from the cyst site; e.g., spores of the cartilaginophilous M. cerebrnlis have been found in intestinal contents. Possibly some spores are forced into the vascular system and are moved by blood flow. Host specificity is also variable; nevertheless, site and host records may still aid in identification. The follow¬ ing list of organ sites is intended to help steer the worker toward the proper species.

Family Myxobolidae

57

Key to the Genera of Myxosporea of North American Freshwater Fishes (Following the classification of Lom and Noble, 1984) 1. With two valves (shells) and two polar capsules

8. (6) Spores spherical or elongated parallel to suture line.

(rarely one: Thelohanellus), one capsule opening

Suture line appearing to pass between the polar

in each of the valves.Order Bivalvulida 2

capsules if two polar capsules present.9

1. With three or more valves and a polar capsule opening in each.Order Multivalvulida 16

8. (6) Spores spherical to irregularly ellipsoid, bilaterally symmetrical along straight suture line; two polar

2. (1) Spores flattened parallel to the straight suture line,

capsules set widely apart in sutural plane;

which forms an elevated ridge; projections possibly

coelozoic in marine and freshwater fishes

extending from the valves .Suborder Platysporina, Family Myxobolidae 3 2. (1) Spores not flattened parallel to the suture line.Suborder Variisporina 6

.Family Ortholineidae (Fig. 104) Neomyxobolus 9. (8) Spores spherical to elongate, bisected by straight meridional suture; four polar capsules present .Family Chloromyxidae 10 9. (8) Similar to no. 9, above, but with two

Suborder Platysporina 3. (2) Spores ellipsoid, ovoid or rounded in valvular view; two polar capsules present, usually pyriform; most species with iodinophilous vacuole, but some, formerly Myxosoma, without; no caudal appendages

(Myxosoma in part).(Figs. 78-90, 96) Myxobolus 3. (2) With caudal appendages or only one polar capsule . . 4 4. (3) With caudal appendages and two polar capsules.... 5 4. (3) With no caudal appendages, one polar capsule.(Fig. 97) Thelohanellus 5. (4) With two caudal appendages (tails), each a continuation of one valve.(Figs. 76, 77) Henneguya 5. (4) With one caudal appendage that is not an extension of the valves.(Fig. 98) Unicauda 5. (4) Similar to Unicauda, but having two, sometimes three, caudal appendages extending in opposite directions. . . . (Figs. 72-74) Dicauda 5. (4) With single flat appendage, otherwise similar to Myxobolus.(Fig. 75) Facieplatycauda

Suborder Variisporina 6. (2) Spores spindle-shaped, sigmoid or crescentic; two polar capsules present, one in each end of spore; longitudinal suture line straight, curved, or sigmoid .Family Myxidiidae 7 6. (2) Spores variously shaped; suture line usually appearing to pass between the two polar capsules.8 7. (6) Spores usually fusiform, straight, slightly crescentic, or sigmoid.(Figs. 91, 102) Myxidlum 7. (6) Spores ellipsoidal in suture view and slightly bent or semicircular in valvular view, with rounded or bluntly pointed ends; polar capsules opening slightly subterminally, both located to side .... (Fig. 103) Zschokkella

polar capsules in one end.11 10. (9) Spores spherical.(Fig. 101) Chloromyxum 10. (9) Spores elongated, with two "tails" .(no fig.) Caudomyxum 10. (9) Spores elongated, with four "tails" . (Fig. 100) Agarella 11. (9) Spores symmetrical; suture line appearing to pass between polar capsules.Family Sphaerosporidae 12 11. (9) Spores asymmetrical, thin-walled, elongated roughly in sutural plane; valves unequal, separated by a curved suture line .Family Parvicapsulidae (Fig. 105) Parvicapsula 12. (11) Spores spherical.(Fig. 110) Sphaerospora 12. (11) Spores not spherical .13 13. (12) Spores in the shape of an isosceles triangle.(Fig. Ill) V/ardia 13. (12) Spores oval to elongated.14 14. (13) Spores anteriorly pointed, miter-like, or rounded in valvular view, with filaments at posterior end .(Mltraspora in part) (Figs. 92-95, 108, 109, 115) Hoferellus 14. (13) Spores elongated, oval, without filaments at posterior end.15 15. (14) Shell valves, usually with fine ridges extending posteriorly in two caudal appendages; resembling

Henneguya, but with suture line passing between polar capsules.(Fig. 107) Myxobilatus 15. (14) As with Myxobilatus, but with no caudal appendages.(Fig. 106) Acauda

Order Multivalvulida 16. (1) Spores with four polar capsules.(Fig. 112) Kudoa 16. (1) Spores with five polar capsules. (Fig. 113) Pentacapsula

8. (6) Spores considerably elongated perpendicular to straight central transverse suture .Family Ceratomyxidae (Fig. 99) Ceratomyxa

(text continues on page 62)

T

TL

-71. Methods for measuring myxosporean spores of various generaMyxobolus in frontal (a) and sutural (b) or side views; Henneguya in frontal (c) and side (d) views; Myxidium in frontal (e) and side (f) views; Chloromyxum in side or sutural (g) and frontal (h) views; Kudoa in apical (i) and one of the possible side (j) views, which is the diagonal one. Measurement of the polar capsule is indicated in (a) and (b); L = length of spore, W = width of the spore, T = thickness of the spore. In spores with caudal appendages, such as in Heimeguya: AL = length of the caudal appendage, TL = total length of the spore (from Lorn and Arthur, 1989, J. Fish Dis., by permission of the Czechoslovak Academy of Sciences).

75.

Facieplatycauda

73.

Dicauda

76.

Henneguya

suture line polar capsule old nuclei polar filament sporoplasm iodinophilous vacuole

78.

Myxobolus

-Myxosporea-

FIG. 72. Dicauda atherinoidi, subdermal cysts in Notropis atherinoides (from G. L. Hoffman and Walker, 1978). FIG. 73. D. atherinoidi, mature spore (from Hoffman and Walker, 1978). FIG. 74. D. atherinoidi, young spore with tails half grown from G. L. Hoffman and Walker, 1978. FIG. 75. Facieplatycauda pratti, line diagrams of two spores: a = front view, b = side view (from Wyatt, 1979, by permission of J. Protozool.). FIG. 76. Henneguya spp.: a, Henneguya zschokkei cysts in muscle of Prosopium williamsoni (courtesy of L. Mitchell, Iowa State Univ.); b, Henneguya doori, spore (from Guilford, 1963). FIG. 77. Henneguya exilis, infection in gills of Ictalurus punctatus, cysts appear as dark elongate masses between lamellae. FIG. 78. Myxobolus pfeifferi, schematic of iodine-stained spore showing dark iodinophilous vacuole; European sp. given as example (from Keysselitz, in Kudo, 1930). FIG. 79. Myxobolus notemigoni, whitish cysts under scales of Notemigonus crysoleucas. FIG. 80. Myxobolus cerebralis, multinucleate trophozoite in cartilage of Oncorhynchus mykiss, tissue section, hematoxylin and eosin stain (photo by G. L. Hoffman). FIG. 81. Myxobolus scleropercae, cysts in eye orbit of Perea flavescens, specimen from J. Schachte, NY (photo by G. L. Hoffman).

-Myxosporea-

FIG. 82. Myxobolus arcticus from central nervous system of Oncorhynchus nerka, B.C. (photomicrograph courtesy of L. Margolis). FIG. 83. M. cartilaginis from cartilage of Lepomis macrochirus, WV (photomicrograph from Lorn and Hoffman, 1971, J. Parasitol.). FIG. 84. M. cerebralis from cartilage of Oncorhynchus mykiss, WV (photomicrograph from Lorn and Hoffman, 1971). FIG. 85. M. cartilaginis spores in large lesion in cartilage of Lepomis macrochirus, WV (from G. Hoffman, 1973a). FIG. 86. M. cerebralis spores in tissue section, methylene blue stain (courtesy of the late P. Ghittino, Turin, Italy). FIG. 87. M. cerebralis spores with extruded polar filaments. FIG. 88. Misshapen head of O. mykiss, including short opercula caused by M. cerebralis infection, WV (from G. L. Hoffman, 1973a). FIG. 89. M. cerebralis spore in India ink to show the mucous envelope impenetrated by ink particles (from Lorn and Hoffman, 1971, /. Parasitol.). FIG. 90. M. corneas cysts in eye of Lepomis macrochirus, IL (specimen courtesy of R. Horner, IL). FIG. 91. Myxiiiium sp. spore from eel, New Zealand: note striation ridges. Scanning EM, x 8200 (courtesy of P. M. Hine, New Zealand). FIG. 92. Hoferellus carassii infection in Carassius auratus, MO. Note the enlarged polycystic kidney (from G. L. Hoffman, 1981). FIG. 93. H. carassii, unilateral infection in C. auratus kidney. FIG. 94. H. carassii, binucleate trophozoites from polycystic kidney infection (from G. L. Hoffman, 1981). FIG. 95. PKD binucleate trophozoite, presumably Hoferellus sp., in kidney of Oncorhynchus mykiss.

a 96. Myxobolus

97. Thelohanellus

99. Ceratomyxa

103. Zschokkella

101. Chloromyxum

100. Agarella

TT S

a

10 |jm

b

104. Neomyxobolus

105. Parvicapsula

106. Acauda

- Myxosporea FIG. 96. Myxobolus ovalis, spore; a, front view; b, side view (redrawn from H. S. Davis, 1953). FIG. 97. Thelohanellus notatus, spore (redrawn from Kudo, 1934). FIG. 98. Unicauda crassicauda, spore (redrawn from Kudo, 1934). FIG. 99. Ceratomyxa shasta, spore (redrawn from E. R. Noble, 1950). FIG. 100. Agarella gracilis, spore (redrawn from Dunkerly, 1915). FIG. 101. Chloromyxum gibbosum, spore; a, front view; b, side view (redrawn from Herrick, 1941). FIG. 102. Myxidium kudoi, spore (redrawn from Meglitsch, 1937). FIG. 103. Zschokkella salvelini, spore (redrawn from Fantham et al., 1939). FIG. 104. Neomyxobolus ophiocephalus, spore, shown in end (a) and front (b) views (redrawn from Chen and Hsieh, 1960). FIG. 105. Parvicapsula sp., spore, from kidney of Oncorhynchus kisutch, WA (redrawn from G. L. Hoffman, 1981). FIG. 106. Acauda elongata, spore. FIG. 107. Myxobilatus caudalis, spore: a, front view; b, side view (redrawn from Davis, 1944).

—107. Myxobilatus

62

Subkingdom Protozoa

Myxosporea -and Actinosporea-

FIG. 108. Hoferellus carassii, typical spore from Carassius auratus in sutural view (from Lorn, 1986). For life cycle diagram, see Fig. 115. FIG. 109. H. cyprini from Cyprimis carpio, sutural view (from Lorn, 1986). FIG. 110. Sphaerospom carassii, spore in sutural view (redrawn from Kudo, 1920). FIG. 111. Wardia ovinocua spore (redrawn from Kudo, 1920). FIG. 112. Kiuioa clupeidae, spore, a marine species given here as an example (redrawn from Meglitsch, 1947). FIG. 113. Pentacapsula muscularis, anterior view showing all five polar capsules (redrawn from Cheung et al., 1983, by permission of Blackwell Scientific Publishers). FIG. 114. Triactinomyxon sp., containing many spherical sporozoites in sporoplasm, from Tubifex. Freehand sketch from description of

Triactinomyxon dubium.

Bone: see cartilage. Brain: accmthogobi, arciticus, bilineatum, cerebralis (acci¬ dentally), cyprinicola, drjagini, enceplmlicus, farionis, hendricksoni, heterolepis, inaequus, kisutchi, muelleri, neurobius, neurophila, schuberti. Cartilage (sometimes locked in bone): cartilaginis, cerebralis, encephalica, eucalii, hoffmani, hyborhynchi, morone, mievoleonensis, petruschewskii, scleropercae. Connective tissue (usually subcutaneous): argentius, capsulatus, catostomi, kostiri, robustum, subtecalis. Everywhere (or seems so!): carassii, circidis, ellipsoides, krokhini, lepomicus, musculi, nemachili, pseudodispar, schuberti. Eyes: acanthogobi, corneus, couesi, heterolepis, hoffmani, magnus, orbitalis, scleropercae, spatulus, talievi. Fins (or base of fins): achmerovi, aureatus, congesticus, inomatus, percae.

Frog: at least one species found in frogs. Gall bladder: bubalis, osbumi, sparoidis. Gills: acuta, angustus, aniscapsularis, anura, aureatus, branchialis, cuneata, cyprini, dechtiari, discrepans, dispar, dujardini, endovasa, funduli, gibbosus, globosus, ibericus, insidiosus, iowanensis, koi, micro¬ cystis, muelleri, musseliusae, okobojiensis, ovalis, pavlovskii, pellucides, pendula, pseudokoi, rotundum, rotundas, symmetricus, transovalis. Heart: cordis, cyprinicola, dogieli, paralintoni. Intestinal wall: artus, intestinalis, okobojiensis, paroidis, poecilichthidis. Kidneys: agolis, albovia, kosloffi, transversalis. Liver: grandis, notropis. Mesentery: achmerovi, microthecum, osbumi. Metacercarial cysts (Uvulifer ambloplites): uvuliferensis. Mouth: dentium, macrocapsularis, pharyngeus.

Family Myxobolidae

63

FIG. 115. Life cycle of Hoferellus carassii (from Ahmed, 1973): 1, amoebula; 2, zygote 3-5; young trophozoites; 3a-c, plasmotomy; 6, trophozoite just free in the lumen; 7, young polytrophozoite; 8, polytrophozoite with a gemma and a vacuole; 9, polytrophozoite forming pansporoblast; 10, large polytrophozoite forming spore cells; 11, polysporoblast with immature spore and spore cells; d-f, development of spore from spore cell nuclei; 12, polysporoblast with fully formed spores; 13, a mature spore; 14, germination.

Muscle: catostomi, funduli, homeospora, insidiosus, multiplicata, obliquus, orbiculatus, subcircularis, teres, transovalis, transversalis, wellerae. Peritoneum: magnaspherus. Scales, in or beneath: barbi, dijagini, hudsonius, indicum, mrigala, notemigoni, sphericum, squamae, squamalis, squamosus, transovalis, vastus. Skin: bellus, bibullatum, commersoni, compressus, conspicuus, cyprini, dermatobium, ellipticoides, gravidus, homeospora, lintoni, moxostomi, mutabilis, nodosus,

notropis, ovatus, ovoidalis, procerum, pseudokoi, rhinichthidis, robustum, salmonis, squamae. Spinal cord: dijagini, farionis, kisutchi, neurobius. Spleen: agolus, albovia, brachyspora, equatorialis, galilaeus, israelensis. Testes: diaphana. Tumor: diaphana. Viscera: amurensis, exiguus, grandis, heterospora, ibericus, media, mesentericus, muelleri, notropis, parellipticoides, pfrille, scardini.

64

Subkingdom Protozoa

Genus Dicauda Hoffman and Walker, 1978 (Figs. 72-74, Color Fig. 11)

Mature spores ovoid, flattened; two polar capsules pres¬ ent; sporoplasm with small iodinophilous vacuole; elon¬ gated “tails" extending from sutural groove, apparently related to the mucoid envelope of Myxobolus and not an extension of the spore wall. No typical mucoid envelope seen in India ink preparations. Close relative of Myxobo¬ lus and Unicauda. ■ Dicauda atherinoidi G. L. Hoffman and Walker, 1978 (Figs. 72-74): primarily dermis, feral Notropis atheriuoides, pathogenicity negligible except in heavily infected fishes, MI, NY.

Genus Facieplatycoudo Wyatt, 1979 (Fig. 75)

This is a provisional genus because Lorn and Noble (1984) consider it a synonym of Myxobolus. However, it is retained here to aid in identification. It differs from Myxobolus only in having an unusual flat extension of the two valves in face view. ■ Facieplatycauda pratti Wyatt, 1979: kidney, Catostomus luxatus, OR.

Genus Henneguya (Thelohan, 1892) Davis, 1944 (Figs. 76, 77)

Spores rounded; ellipsoid or spindle-shaped in valvular view; biconcave in sutural view. Each valve continuing as a caudal projection; both projections sometimes apposed. Shell valves smooth. Two polar capsules pres¬ ent, sometimes very elongated; binucleate sporoplasm usually with iodinophilous vacuole; trophozoites com¬ monly large, polysporous with pansporoblast forma¬ tion. Histozoic in freshwater, sometimes marine, fishes. — Ref. H. S. Davis (1944a), Kudo (1920, 1934, 1971), Lorn and Noble (1984).

■ Henneguya acuta Bond, 1939b: gills, Esox masquinongy, NY; Dechtiar (1972a): same, Lake of the Woods, Ont.; Nepszy (1988): same, Lake Ontario, Ont. ■ H. adiposa Minchew, 1977: adipose fin, Ictalurus punctatus, MS, cysts to 0.5-1.5 mm; Current (1979): ultra¬ structure and sporogenesis, NE; Hoffman (unpub., 1980): AR. ■ H. ameiurensis Nigrelli and Smith, 1940: barbels, Ictalurus nebulosus, NY. ■ H. amiae Fantham, Porter, and Richardson, 1940: gills, Ami a calva, Canada. ■ H. cerebralis Pronina, 1972. Pronina and Pronin (1985): head, Thymallus arcticus, cytochemical and histopatho-

logical changes, former Soviet Union, (see O. N. Bauer, 1984). ■ H. cutanea Dogiel and Petruschewsky, 1933. Lorn et al. (1976): in several fishes including Cyprinus carpio, Czech Republic and Uzbekistan. ■ H. cyprini Lorn et al., 1976: found in Cyprinus carpio by Kohn (1935) and Jirovec (1942), Czech Republic and Slovakia. ■ H. diversus Minchew, 1977: carbuncle-like lesions at bases of barbel and pectoral fins of Ictalurus punctatus, MS. ■ H. doneci Shulman, 1962. Shulman (1966): gills, Carassius auratus, Amur River, Russia. ■ H. doori Guilford, 1963: gill filaments, Perea flavescens, WI; Cone (1979a): same, Lake Erie, Ont.; Nepszy (1988): same, Lake Huron, Lake Erie, Lake Ontario, Ont. ■ H. episclera Minchew and Sleight, 1977: sclera of eye, Lepomis gibbosus, RI; G. L. Hoffman (unpub., USFWS, 1958): same, L. macrochirus, Fishing Creek, MD, collec¬ tion of S. F. Snieszko. ■ H. esociis Fantham, Porter, and Richardson, 1939: gills, Esox niger, Que. ■ H. exilis Kudo, 1929: gills, Ictalurus melas, I. puncta¬ tus, IL; J. C. Baker and Crites (1976): gills, I. punctatus, western Lake Erie, USA; Current and Janovy (1976): ultrastructure of interlamellar form, I. punctatus, NE; Cur¬ rent and Janovy (1977): ultrastructure of sporogenesis; Current and Janovy (1978): comparative ultrastructure of interlamellar and intralamellar forms; Dechtiar (1972a): gills, I. nebulosus, I. punctatus, Lake Erie, Ont.; Duhamel et al. (1986): histopathology of granulomatous branchitis, I. punctatus, CA; Guilford (1965): gills, 1. melas, Lake Michigan; G. L. Hoffman (1979): species identification and use of plankton centrifuge for con¬ centrating spores, AR; Long and Meglitsch (1970): pre¬ treatment of spores before measuring; McCraren et al. (1975): histopathology of the various types of H. "exilis", lesions in I. punctatus; F. Meyer (1970a): description of inter- and intralamellar forms of H. "exilis" infections, AR; Minchew (1977): gills, I. punctatus, MS, comparison of six species from I. punctatus, including five new species and three from gills (H. exilis, H. longicauda, H. postexilis); L. G. Mitchell (1978b): gills, I. punctatus, Des Moines River, 1A; Molnar et al. (1974): gills, I. nebu¬ losus, Ont.; Rice and Jahn (1943): IA; Segovia Salinas (1985): intralamellar form in cultured I. punctatus, Mon¬ terrey, Mexico; C. E. Smith and Inslee (1980): histopathol¬ ogy of interlamellar Henneguya sp. in I. punctatus. A very damaging branchitis of I. punctatus known as proliferative gill disease (PGD) and hamburger gills was attributed, from 1969 to 1986, to interlamellar H. "exilis." Recent findings indicate that this disease is caused by the trophozoite of an unknown myxosporidan, possibly Sphaerospora sp. transmitted by the oligochaete Dero (Bowser and Conroy, 1985; MacMillan et al., 1988). This question apparently has not been resolved.

Family Myxobolidae

■ H. fontinalis Fantham, Porter, and Richardson, 1939: fin cyst, Salvelinus fontinalis, Que.; Fantham and Porter (1948): same, Que. ■ H. fontinalis var. notropis Fantham, Porter, and Richard¬ son, 1939: skin, Notropis cornutus, N. heterolepis, Que. ■ H. gambusi Parker, Spall, and Warner, 1971: skin cysts, Gambusia affinis, pathogenic, OK. ■ H. gurleyi Kudo, 1920: base of spines, Ictalurus melas, IA.

65

■ H. salmincola Ward, 1919: probably a syn. H. zschokkei. ■ H. salmonis Fantham, Porter, and Richardson, 1939: subcutaneous tissue, Salmo salar, Canada. ■ H. schizura (Gurley) Labbe, 1899. Shulman (1966): eye muscles, sclera, vitreous humor, Esox lucius, Europe, USA, former Soviet Union. ■ H. schulmani Ki and Ha, 1971: gills, Cyprinus carpio, northern Vietnam.

■ H. limatula Meglitsch, 1937: gall bladder, Ictalurus furcatus, IL; Guilford (1965): I. melas, MI.

■ H. theca Kent and Hoffman, 1984: brain, Eigemannia viriscens (tropical aquarium fish), spore with capsule, South America.

■ H. lobosa (Cohn, 1895). Shulman (1966): usually in gills, several fishes, including Esox Indus, Europe, former Soviet Union.

■ H. umbri Guilford, 1965: gills, Umber limi, Lake Michigan.

■ H. longicauda Minchew, 1977: in lamellar capillaries and interlamellar regions, cysts up to 370 x 130 pm, total spore length 108 pm; associated with H. postexilis, the intralamellar form, Ictalurus punctatus, MS. ■ H. magna Rice and Jahn, 1943: gills, Roccus chrysops, IA. ■ H. nigris Bond, 1939b: gills, Esox niger, MD, E. masquinongy, NY. ■ H. niisslini Schuberg and Schroder, 1905. Kudo (1920): at base of dorsal fin, Salmo trutta, Germany. ■ H. oviperda (Cohn, 1895). Shulman (1966): in ovaries, usually not in ova, occasionally in testes and intestinal wall, Esox lucius, Lucioperca lucioperca, Europe; E. H. Davies (1968): E. lucius, England; Kennedy (1974): Eng¬ land; Uspenskaya (1982): cyst ultrastructure, St. Peters¬ burg, Russia; Uspenskaya (1988): ultrastructure of intracellular plasmodia. ■ H. pellis Minchew, 1977: cysts 1-2 mm, in skin of Ictalurus furcatus; with body size 13 pm, this species is the smallest of the complex of catfish Henneguya spp., AL. ■ H. pinnae Schubert, 1967: in fins, Ctenopoma kingsleyae (tropical aquarium fish), Germany; Schubert (1968): ultractructure. ■ H. postexilis Minchew, 1977: interlamellar cysts, Ictalu¬ rus punctatus, the smallest total length (42-62 pm) of the H. "exilis" complex in catfish; C. E. Smith and Inslee (1980): histopathology of severe gill disease. ■ H. psorospermica Thelohan, 1895. Shulman (1966): gills, muscles, walls of buccal cavity, intestine of several fishes, including Coregonus autumnalis, Esox lucius, Lota lota, Europe; Dykova and Lorn (1978b): gill arteries, E. lucius, histopathology, Czech Republic and Slovakia; Lom and Vavra (1963b): ultrastructure of development of polar filaments, Czech Republic and Slovakia. ■ H. pungitii Hanek and Molnar, 1974: kidney, Gasterosteus aculeatus, Pungitius pungitius, Metamek River, Que. ■ H. renicola Schurmans-Stekhoven, 1920: kidney, Pun¬ gitius pungitius, Netherlands.

■ H. zikaweiensis Sikama, 1938. Shulman (1966): gills, skin, cornea, Carassius auratus, Yangtse River, China; Amur River, Russia. ■ H. zschokkei (Gurley, 1893) Doflein, 1901 (Syn. H. salmincola Ward, 1919, according to Shulman, 1966). Shulman (1966): gills, subcutaneous muscle of many fishes, including Coregonus autumnalis, C. lavaretus, C. nasum, C. sardinella, Esox lucius, Lota lota, Oncorhychus keta, O. kisutch, O. mykiss, O. nerka, Prosopium cylindraceum, Salmo trutta, Salvelinus alpinus, Europe; Akhmerov (1954): unsightly lesions in O. kisutch, should be disposed of safely; D. Anderson (pers. comm., Col¬ orado, 1989): C. aitedi, plankton centrifuge head sample, western USA; Arai and Mudry (1983): Cottus cognatus, Prosopium williamsoni, Rhinichthys cataractae, Richardsonius balteatus, Thymallus arcticus, B.C.; Arthur et al. (1976): Y.T.; Awakura and Kimura (1977): description of milky condition in imported smoked coho salmon, caused by a myxosporidan, Japan; N. P. Boyce et al., 1985: distribution, detection, biology of H. salmincola (= zschokkei) in Pacific salmon; Boyd and Tomlinson (1965): milky condition of stored O. kisutch, B.C.; Bucke (1972): flesh soft and mushy in North American smoked salmon exported to England; N. O. Christensen and Elliman (1976): H. salmincola differentiated from H. zchokkei, from imported Pacific salmon, Denmark; Dogiel et al. (1958): ulcerative disease, former Soviet Union; Fish (1939a): tapioca disease caused by H. salmin¬ cola, WA; P. Janeke (pers. comm., USFWS, 1976): P. williamsoni, MT; Moles (1982): Oncorhynchusgorbuscha, O. keta, O. kisutch, O. nerka, O. tshawytscha, AK; Nepszy (1988): C. artedi, Lake Huron, Ont.; Newell and Canaris (1969): P. coulteri, P. williamsoni, MT; R. E. Olson (1978): muscle, O. kisutch but not O. tshawytscha, OR; Shaw (1947): in muscle of O. tshawytscha, OR; C. Smith (pers. comm., USFWS, 1973): cysts in muscle of whitefish, Yellowstone River, MT; W. Taylor (pers. comm., USFWS, 1977): gills, O. mykiss, OR. The lesions in salmon flesh are sometimes unsightly and the fishes are often discarded. Although the para¬ site is widespread and not a health hazard to fishes or to humans, heavily infected salmon should not be sold

66

Subkingdom Protozoa

for human food or placed where they might expose uninfected stocks. W. Taylor (pers. comm., Olympia, WA, (1975) stated that the plankton centrifuge method of O'Grodnick (1975) was excellent for recovering spores.

■ M. anisocapsularis Schulman, 1962. Ki and Ha (1971): Cyprinus carpio, northern Vietnam.

■ Henneguya sp. H. S. Davis (1923, p. 432): gill cartilage, Pomoxis annularis.

■ M. arcticus Pugachev and Khokhlov, 1979: in brain, Oncorhynchus kisutch, O. nerka, Salvelinus tnalma, S. neva, former Soviet Union. These authors clarify a long¬ standing confusion between M. arcticus and M. neurobius. M. arcticus is larger (14.3-16.5 pm long) than M. neurobius and is pyriform, whereas M. neurobius is oval. Unfortunately, most of the records for M. neurobius probably concern M. arcticus instead.

■ Henneguya sp. Delisle (1972): probably cause of death of Esox niger, Brome Lake, OR. ■ Henneguya spp.: see Margolis and Arthur, 1979, p. 20, for Canadian records.

Genus Myxobolus Biitschli, 1882 (Figs. 78-90, 96, Color Figs. 7-10)

Note: Lorn and Noble (1984) have synonymized Myxosoma with Myxobolus, so that all previous Myxosoma spp. appear here as Myxobolus spp. Shulman (1984) is not in agreement with this classification. Myxobolus spp. contain an iodinophilous vacuole and Myxosoma spp. do not; Lorn and Noble (1984), however, have not always found this reliable. Lentospora Plehn (1904) is a syn¬ onym of the former Myxosoma. Spores ellipsoid, ovoid, or subspherical in valvular view; biconvex in sutural view; shell valves smooth. Two, polar capsules, mostly pyriform; exceptionally, one capsule apparently missing and the remaining one not situated axially. Posteriorly, sutural ridge possibly extending into a crescentic ledge. Binucleate sporoplasm present, often with an iodinophilous vacuole, particularly before preservation. Trophozoites generally large, polysporous, with pansporoblast formation. Histozoic in freshwater fishes and in a few marine fishes. Some exotic species included here because their hosts have been imported to North America. ■ Myxobolus acutus (Fujita, 1912) Lorn and Noble, 1984: in kidney, Carassius auratus, Japan, former Soviet Union. ■ M. agolus Landsberg, 1985: in spleen, kidney, Tilapia, Israel. ■ M. akhmerovi Shulman, 1966. Ki and Ha (1971): Cyprinus carpio, northern Vietnam; Shulman (1984, p. 204). ■ M. albovae Krasliniikova (in Lorn et al., 1976): Cyprinus carpio, Europe. ■ M. amurensis Akhmerov, 1960 (in Lorn et al., 1976): Cyprinus carpio, East Asia; Alvarez-Pellitero et al. (1979): in viscera, C. carpio, Spain. ■ M. anguillae (Fujita, 1929) (Kudo, 1933) Lorn and Noble, 1984 (Syn. Myxosoma a.): in subcutaneuous tis¬ sue, Anguilla japonica, Japan. ■ M. angustus (Kudo, 1934): in gills, Pimephales vigilax, IL; Desser and Paterson (1978): probably M. angustus, ultrastructure and cytochemical observations, Notropis cornutus, Ont.; G. L. Hoffman (unpub., USFWS, 1983, 1984): Notemigonus crysoleucas, P. promelas, NC.

■ M. anurus (Cohn, 1895): in gills, fins, Esox Indus, Europe.

■ M. argenteus S. D. Lewis, 1968 (Color Fig. 10): cysts in subdermal connective tissue, Notemigonus crysoleucas, IL; G. L. Hoffman (unpub. 1979-1984): glycogen vacuole absent, no albuminoid capsule seen in India ink prepa¬ ration, spore slightly pyriform, AR; Li and Desser (1985b): in many organs of N. crysoleucas, Ont. ■ M. artus Akhmerov, 1960b: in intestinal wall, kidney, Cyprinus carpio, small cysts, small spores; Li and Desser (1985b): in gills, kidney, etc., Notemigonus crysoleucas, Ont; Lom et al. (1976): C. carpio, East Asia. ■ M. aureatus H. B. Ward, 1919: golden cysts in fins, Notropis anogenus, Lake Erie; Arai and Mudry (1983): in gills, Lota lota, Mylocheilus caurinus, Ptychocheilus oregonensis, Richardsonius balteatus, B.C. (because of the unusual organ location, this is questionable); Kudo (1934): in fins, Pimephales notatus, IL; Li and Desser (1985b): brain, gut, swim bladder, Perea flavescens, Ont.; Lom and Dykova (1992): blackspot cyst in fins, Pimephales promelas, IL. ■ M. bellus Kudo, 1934: in skin, Cyprinus carpio, IL. ■ M. bibullatus (Kudo, 1934) Landsberg and Lom, 1991: in skin, Catostomus commersoni, IL; Dechtiar (1972b): C. commersoni, Ont.; Hanek and Molnar (1974): C. com¬ mersoni, Ont.; Nepszy (1988): C. catostomus, C. commer¬ soni, Lake Huron, Lake Ontario; Nigrelli (1943): in gills, C. commersoni, NY. ■ M. bilineatum Bond, 1938a: in brain, Fundulus heteroclitus, MD. ■ M. bondi (Bond, 1939) Landsberg and Lom, 1991 (Syn. Myxosoma tnuelleri Bond, 1939). ■ M. brachyspora (J. R. Baker, 1963) (Syn. Myxosoma b.): spleen, Tilapia esculenta, T. variabilis, East Africa. ■ M. branchialis (Markewitsch, 1932) (Syn. Myxosoma b., M. circulus): gills, muscle, Cyprinus carpio and others, former Soviet Union; Lom et al. (1976): C. carpio, Europe. ■ M. bubalis Otto and Jahn, 1943: gall bladder, Ictiobus bubalus, IA, cysts attached by stalks. ■ M. capsulatus H. S. Davis, 1917: visceral connective tis¬ sue, Cyprinodon variegatus, NC. ■ M. carassii Klokacewa, 1914: in viscera, Carassius carassius, former Soviet Union; Lom et al. (1976): Cypri¬ nus carpio, Asia, Europe.

Family Myxobolidae

67

■ M. carpiodes (Meglitsch, 1937) Landsberg and Lom, 1991 (Syn. M. rotundus Meglitsch, 1937).

■ M. cuneatus (Bond, 1939) Lom and Noble, 1984 (Syn. Myxosoma c.): in gill arch, Esox masquinongy, NY.

■ M. cartilaginis (Hoffman, Putz, and Dunbar, 1965) (Syn. Myxosoma c.): in cartilage of head and spines, Lepomis macrochirus, L. cyanellus, Micropterus salmoides, WV, MD; Lom and Hoffman (1971): morphology of spores compared with M. cerebralis, PA.

■ M. cyprini Doflein, 1898: in viscera, muscle, Cyprinus carpio and other cyprinids, Europe; BykhovskayaPavlovskaya (1962): identification; Korting and Her¬ manns (1984): not considered pathogenic, Germany.

■ M. catostomi no. 1 (Kudo, 1923, in Kudo, 1926) (Syn. Myxosoma c.): in muscle, connective tissue, Catostomus commersoni, MI; Dechtiar (1972b): C. commersoni, Ont. ■ M. catostomi no. 2 Fantham, Porter, and Richardson, 1939: in mouth subepithelium, muscle, Catostomus com¬ mersoni, Que.; Fantham and Porter (1947): intestine, gall bladder, liver, kidney, C. catostomus, Que.; Mpoame (1982): C. insignis, AZ; Pugachev (1980): C. catostomus, Former Soviet Union. ■ M. cerebralis (Hofer, 1903) (Plehn, 1905) (Syn. Myxo¬ soma c., according to Lom and Noble, 1984). (Color Figs. 7, 8). G. L. Hoffman (1990): review, in cartilage of the following species, with the most susceptible first: Oncorhynchus mykiss, O. nerka, Salvelinus fontinalis, O. tshawytscha, Salmo salar, S. trutta, O. kisutsch; S. namaycush and lake and brook trout cross are refractory; Euro¬ pean salmonids have also been found infected. Other reviews and pertinent papers include El-Matbouli and Hoffmann (1989): experimental transmission of two species developing bisporogony via tubificid worms; Ghittino and Vigliani (1978); Halliday (1973, 1976); G. L. Hoffman (1970, 1976b); Lom and Hoffman (1971): morphology of spores; Lorz et al. (1989): comparison of digest and plankton centrifuge concentration methods; Lunger et al. (1975): electron microscopy; Markiw and Wolf (1974): concentration; O'Grodnick (1975): con¬ centration; Prihoda (1983): experimental infection; Uspenskaya (1978): experimental infection; Wolf (1986): review; Wolf and Markiw (1985): review. ■ M. circulus Akhmerov, 1960b: syn. of M. branchialis. ■ M. commersoni (Fantham et al., 1939) Lom and Noble, 1984 (Syn. Myxosoma c.): in skin, Catostomus commersoni, Que. ■ M. compressus Kudo, 1934: in skin, Notropis blennius, IL. ■ M. congesticus Kudo, 1934: in fins, Moxostoma anisurum, IL. ■ M. conspicuus Kudo, 1929: in skin, Moxostoma breviceps, IL; Dechtiar (1972b): Chrosomus neogaeus, M. anisurum, Ont.; Fantham et al. (1939): M. aureolum, Que.; Nepszy (1988): Phoxinus neogaeus, Lake Huron; Nigrelli (1943): M. aureolum, NY.

■ M. cyprini (Myxosoma cyprini) Spall, 1974 (is probably Myxobolus angustus or M. spalli): in gills, Notropis lutrensis, Notemigonus crysoleucas, OK. ■ M. cyprinicola Reuss, 1906. Bykhovskaya-Pavlovskaya (1962): in gills, Cyprinus carpio and Amur cyprinids, Rus¬ sia; Lom et al. (1976): review. ■ M. dechtiari Cone and Anderson, 1977a: gills, Lepomis gibbosus, Ont. ■ M. dentium (Fantham, Porter, and Richardson, 1939): sore mouth, Esox masquinongy, Que.; George et al. (1977): incidence, NY; Nepszy (1988): E. masquinongy, Lake Ontario, Ont. ■ M. dermatobius Ishii, 1916. Bykhovskaya-Pavlovskaya et al. (1962): skin, Anguilla japonicus, Oncorhynchus keta, O. kisutch, producing ulcers; Copland (1982): wild Anguilla, England; Hoshina (1953): skin, Cyprinus carpio, Japan; Lom et al. (1976): review. Author: it is not known whether this is one or two species. ■ M. diaphanus (Lantham, Porter, and Richardson, 1940) Lom and Noble, 1984 (Syn. Myxosoma d.): in testes, Fundulus diaphanus, N.S. ■ M. discrepans Kudo, 1920: in gills, Carpiodes diffonnis, IL; H. S. Davis (1923): in Ictiobus sp., but sometimes penetrating cartilage of gills; Mpoame (1982): gills, Catostomus insignis, AZ; Rice and Jahn (1943): gills, Pomoxis sparoides, IA. ■ M. dispar Thelohan, 1895. Gratzek (pers. comm., Univ. of Georgia, 1983): kidney, goldfish, GA; Halliday (pers. comm., 1973): cross section, black cyst, Scotland; Ivasik et al. (1967): erosion of Cyprinus carpio gills; Lom et al. (1976): C. carpio, Czech Republic and Slovakia. ■ M. divergens Ki and Ha, 1971: Aristichthys nobilis, northern Vietnam. ■ M. dogieli Bykhovsky and Bykhovskaya, 1940: in heart, Cyprinus carpio and grass carps, former Soviet Union; Lom et al. (1976): review, Europe. ■ M. drjagini Akhmerov, 1954: in skin, Hypophthalmichthys molitrix; Baohna et al. (1979): whirling dis¬ ease, in central nervous system, H. molitrix, China; Hsieh (1987): in brain and semicircular canals (twist disease), H. molitrix, China.

■ M. corneus Cone, Horner, and Hoffman, 1990: in cornea of eye, Lepotnis macrochirus, IL.

■ M. dujardini (Thelohan, 1899) Lom and Noble, 1984 (Syn. Myxosoma d.)\ in gills, Cyprinus carpio and others, Europe; Dogiel et al. (1958, p. 212); Lom et al. (1976): review; L. G. Mitchell (pers. comm., 1976): Ptychocheilus regonensis, MT.

■ M. couesii Fantham, Porter, and Richardson, 1939: in eye (posterodorsal corner of anterior chamber, on iris), Couesius plumbeus, Que.

■ M. ellipsoides Thelohan 1892. Bykhovskaya-Pavlovskaya (1962): in many organs of many fishes including Cypri¬ nus carpio and grass carps, Europe, Asia.

68

Subkingdom Protozoa

■ M. ellipticoicies (Fantham, Porter, and Richardson, 1939): behind operculum, Catostomus commersoni, Que.

(pers. comm., Ohio State Univ., 1971): bursts body wall of E. buccata. See M. fanthami.

■ M. encephalicus (Mulsow, 1911) (Syn. Lentospora e. Mulsow, 1911). Dykova et al. (1986): in brain, causing whirling disease of Cyprinus carpio fingerlings, Czech Republic and Slovakia; Plehn (1924): in brain blood vessels, C. carpio swim in circle or "stand on head."

■ M. gravidas Kudo, 1934: in skin, Moxostoma anisurum, IL.

■ M. endovasus (Davis, 1947) Lorn and Noble, 1984 (Syn. Myxosoma e.)\ in gills, Ictiobus bubalus, IA.

■ M. heterolepis Li and Desser, 1985a: brain and eye, Notropis heterolepis, Ont.

■ M. equatorialis (Landsberg, 1985) Lorn and Noble, 1984 (Syn. Myxosoma e.): spleen, Tilapia aureus x T. niloticus, Israel.

■ M. heterospora (Baker, 1963) Lom and Noble, 1984 (Syn. Myxosoma h.) liver, spleen, kidney, Tilapia, East Africa.

■ M. eucalii (Guilford, 1965) Lorn and Noble, 1984 (Syn. Myxosoma e.): cartilage, Culaea inconstans, Lake Michigan.

■ M. hoffmani (Meglitsch, 1963) Lom and Noble, 1984 (Syn. Myxosoma h.); in cartilaginous sclera of eye, Pimephales promelas, ND; Beckwith (pers. comm., 1973): P. promelas in farm ponds, Bangor, PA; Crutcher (pers. comm., USFWS, 1975): as above, MO; G. L. Hoffman and Putz (1965): additional data, same collection.

■ M. exiguus Thelohan, 1895. Shulman (1966): in gills of many fishes including Cyprinus carpio, Europe. ■ M. exsulatus Pugachev, 1980: Catostomus catostomus, Kolyma River, Russia. ■ M. fanthami (Fantham et al., 1939) Landsberg and Lorn, 1991 (Syn. M. grandis Fantham et al., 1939). ■ M. farionis Gonzalez-Lanza and Alvarez-Pellitero, 1984: in central nervous system, Salmo trutta, Spain. ■ M. filamentus (Rice and Jahn, 1943) Grinham and Cone, 1990 (Syn. Myxosoma okobojiensis Rice and Jahn, 1943; M. janricei (Rice and Jahn, 1943) Landsberg and Lorn, 1991). ■ M. funduli (Hahn, 1915) Kudo, 1920 (Syn. Myxo¬ soma f. Kudo, 1920): gills and muscle, Fundulus diaphanus, F. heteroclitus, F. majalis, Woods Hole, MA. Author: this species was apparently originally described as Myxobolus f. by Hahn and as Myxosoma f. by Kudo; it is probably one species. Bond (1938a): experimental infection, direct; Current et al. (1979) electron microg¬ raphy, F. kansae; Hahn (1915): experimental infection of Cyprinodon variegatus and F. diaphanus; Janovy and Hardin (1987): F. zebrinus, NE; Janovy and Hardin (1988): diversity of parasite assemblage; Knight et al. (1980): annual prevalence and geographic distribu¬ tion, NE; Wiles (1975b): F. diaphanus, N. S. ■ M. galilaeus Landsberg, 1985: spleen, Tilapia aureus x T. niloticus, Israel. ■ M. gibbosus Herrick, 1941: in connective tissue cov¬ ering gill arches, Lepomis gibbosus, Lake Erie. Note: M. gib¬ bosus Li and Desser, 1985 is probably M. osbumi. ■ M. globosus Gurley, 1894 (from Kudo, 1920, p. 139): gills, Erimyzon oblongus, ND, SC. ■ M. grandis (Kudo, 1934) Lorn and Noble, 1984 (Syn. Myxosoma g. Kudo, 1934; M. grandis Fantham et al., 1939): large abdominal cyst, Ericymba buccata, Notropis cornutus, N. hudsonius, Rhinichthys atronasus, IL, Que.; Dechtiar (1972b): N. hudsonius, Lake of the Woods, Ont.; Nepszy (1988): N. hudsonius, Lake Huron, Ont.; Ross

■ M. hendricksoni L. G. Mitchell, Seymour, and Gamble, 1985: brain, Pimephales promelas, description including electron microscopy, IA.

■ M. homeospora (Baker, 1963) Lom and Noble, 1984 (Syn. Myxosoma //.): in many organs, Tilapia, East Africa; Paperna (1973): Tilapia and Barbus, Africa. ■ M. hudsonius (Bond, 1938) Lom and Noble, 1984 (Syn. Myxosoma h.): in skin, Fundulus heteroclitus, NY. ■ M. hyborhynchi Fantham, Porter, and Richardson, 1939: in bone at end of mandible, Hyborhymchus notatus, Que.; Molnar et al. (1974): kidney, Pimephales notatus, P. promelas, Que. ■ M. ibericus Gonzales-Lanza and Alvarez-Pellitero, 1984: viscera, gills, Salmo trutta, Spain. ■ M. inaequus Kent and Hoffman, 1984: brain, knifefish (Eigematinia viriscens), South America (CA experimental fish). ■ M. inoniahts Fish, 1939b: in flesh, Microptems salmoides, MT; G. L. Hoffman (unpub., USFWS, 1979): M. salmoides, TX; Horner (pers. comm., State Fisheries, 1978): bases of fins, M. salmoides, IL; Sullivan (pers. comm., Auburn Univ., 1979): bases of fins, M. salmoides, AR. ■ M. insidiosus Wyatt and Pratt, 1963: muscle, Onchorhynchus tshawytschi, OR; Amandi et al. (1985): new host record, O. mykiss, live box infections, immunodiagnosis, differentiation from M. cerebralis, M. squamalis, Ceratomyxa shasta, Henneguya sp.; Wyatt (1978a): in muscle, O. kisutch, WA; Wyatt (1979): muscle, Oncorhynchus clarki, OR. ■ M. intestinalis Kudo, 1929: intestinal wall, Pomoxis nigro-maculahis, IL; Dechtiar (1972b): same, Ont.; Meg¬ litsch (1937): same, IL. ■ M. iowensis Otto and Jahn, 1943: gills, Pomoxis nigromaculatus. ■ M. israelensis Landsberg, 1985: spleen, Tilapia aureus x T. niloticus, Israel. ■ M. kisutchi Yasutuke and Wood, 1957: spinal col¬ umn, Oncorhychus kisutch, WA; G. L. Hoffman (unpub.,

Family Myxobolidae

1982, 1987): O. tschawytscha, S. Leek and W. Yasutake, WA; Horsch (1987): O. mykiss, CA (spores significantly larger than those from WA; perhaps n. sp.); Wyatt (1978a): cysts in brain, O. tshawytscha, OR. ■ M. koi Kudo, 1920: gills, Cyprinus carpio, Japan; Cren¬ shaw and Sweeting (1986): in koi carp imported from Japan, England; Hensley and Nahhas (1975): same, CA; Hoshina (1952): same, Japan; Hsieh (1987): 80% mor¬ tality in carp fry, China; Ishizaki (1964): Acheilognathus lanceolata, Japan; Ki and Ha (1971): C. carpio, northern Vietnam; Nakai (1928): described as n. sp. from carp fry, Japan; Yunchis (1978): killed carp brooders, former Soviet Union. ■ M. kosloffi Wyatt, 1979: kidney, Catostomus luxatus, OR. ■ M. kostiri Herrick, 1936: subcutaneous, Micropterus dolomieui, Lake Erie; Crider (pers. comm, and thesis, Western Kentucky Univ., 1969): Lepomis macrochirus; G. L. Hoffman (unpub., USFWS, 1966): L. macrochirus, WV. ■ M. krokhini Konovelov and Shulman, 1966 (in Shulman, 1966, p. 277): in viscera, Oncorhynchus nerka, Salvelinus alpinus, former Soviet Union; Moles (1982) O. nerka, AK. ■ M. lamellus Grinham and Cone, 1990: in secondary lamellae, Castotomus commersoni, NS. ■ M. lepomicus Li and Desser, 1985b: viscera and muscle, Lepomis gibbosus, Ont. ■ M. lintoni Gurley, 1893: subcutaneous, Cyprinodon variegatus, Woods Hole, MA; Bursey (1987): large lesions, death apparent, VA; Nigrelli and Smith (1938): tissue response, C. variegatus, NY. ■ M. macrocapsularis Reuss, 1906. Lorn et al. (1976): Cyprinus carpio, Europe, Asia. ■ M. magnaspherus Cone and Anderson, 1977a: parietal peritoneum, Lepomis gibbosus, Ont. ■ M. medius (Fantham et al., 1939) Lorn and Noble, 1984 (Syn. Myxosoma m.)\ large abdominal cyst, Notropis cornutus, Que. ■ M. meglitschi (Meglitsch, 1939) Grinham and Cone, 1990: see M. rotundus. ■ M. mesentericus Kudo, 1920: viscera, Lepomis cyanellus, IL; Chen and Hsieh (1960): Ophiocephalus, China. ■ M. microcystis Price and Mellen, 1980: gills, Micropterus salmoides, IL; Hoffman (unpub., USFWS, 1985): same, AR. ■ M. microlatus Lin and Nie, 1973. ■ M. microthecum (Meglitsch, 1942) Lorn and Noble, 1984 (Syn. Myxosoma m.): mesenteries, Minytrema melanops, IL; Arai and Mudry (1983): gall bladder, kid¬ ney, intestine, Rhinichthys balteatus, R. cataractae, B.C. ■ M. morone (Paperna, 1973) Lorn and Noble, 1984: head cartilage, Morone saxatilis, VA; Paperna and Zwerner (1976): same.

69

■ M. moxostomi Nigrelli, 1948: snout (prickle cell hyper¬ plasia), Moxostoma aureolum. ■ M. muelleri (Bond, 1939b) Lorn and Noble, 1984 (Syn. Myxosoma m.)\ gills, Esox masquinongy, NY. See M. bondi. ■ M. mulleri Biitschli, 1882. Shulman (1966, p. 248): in many fish species; Li and Desser (1985a): gills, ureters, Semotilus atromaculatus, Algonquin Park, Ont. ■ M. multiplicatus (Reuss, 1906) (Rice and Jahn, 1943) Lorn and Noble, 1984 (Syn. Myxosoma m.)\ gills, Ictiobus bubalus, IA; Shulman (1966): the Iowa record is prob¬ ably in error. ■ M. musculi Keyserlitz, 1908: in many fishes, includ¬ ing Carassius auratus, Tinea tinea, former Soviet Union; Hahn (1913): skin, Fundulus, USA; Li and Desser (1985a): many organs, Notemigonus crysoleucas, Ont.; Lorn et al. (1976): Cyprinus carpio, Uzbekistan. ■ M. musseliusae Jakovtchuk, 1979: gills, Cyprinus car¬ pio, Krasnodar, Russia. ■ M. mutabilis Kudo, 1934: skin, Pimephales notatus, IL; Chen and Hsieh (1960): Ophiocephalus, China. ■ M. nemachili Weiser, 1949: connective tissue of head, Nemachilius barbalulus, Czech Republic and Slovakia; Li and Desser (1985b): many organs, Notropis heterolepis, N. cornutus, Algonquin Park, Ont. ■ M. neurobius Schuberg and Schroder, 1905: brain and spinal cord, Salmo trutta, Europe. Unfortunately, there has been a mix-up of names: M. neurobius is oval and 13.4-14 pm long (original description of 10-12 pm may have been from preserved material). Most of the fol¬ lowing references probably apply to M. arcticus, rather than M. neurobius. Arthur et al. (1976): central nervous system, Coregonus clupeaformis, Prosopium cylindraceum, Salvelinus namaycush, Thymallus arcticus, Y.T.; Awakura (pers. comm., 1983): Japan; R. E. Bailey and Margolis (1987): Oncorhynchus nerka, B.C., WA; D. Dana (1982): experimental transmission per os, but no pathogenicity; Kennedy (1974): S. trutta, S. salar, England; Kudo (1920, p. 144); D. Locke (pers. comm., State Fisheries, 1973): ME; Plehn (1924): spinal cord but not brain, Oncorhynchus mykiss, Thymallus thymallus, Europe: Pugachev and Khokhlov (1979): Salmo trutta, Thymallus arcticus, T. thy¬ mallus, former Soviet Union; Reichenback-Klinke (1954a, p. 597): Germany; W. Taylor (pers. comm., USFWS): WA. Author: this nonpathogenic species is common in the central nervous system of salmonids of circumpolar distribution. It has been confused with M. cerebralis but is larger. ■ M. neurophilus (Guilford, 1963) Lorn and Noble, 1984 (Syn. Myxosoma n.): midbrain, Etheostoma nigrum, Perea flavescens, Lake Michigan, WI. ■ M. nodosus Kudo, 1934: skin, Pimephales notatus, IL. ■ M. nodularis Southwell and Prashad, 1918: India; Mpoame (1982): Catostomus insignis, AZ. Author: This may be in error, since it is unlikely that the Indian host

70

Subkingdom Protozoa

would be transferred to Arizona, and Catostomus does not occur in India.

I A; Wagh (1961): transplanted to Notemigonus crysoleu¬ cas, IL (may be in error).

■ M. notatus Mavor, 1916: intermuscular in body, Pimephales notatus, Canada.

■ M. ovatus Kudo, 1934: skin, Ictiobus bubalus, IL.

■ M. notemigoni Lewis and Summerfelt, 1964 (Color Fig. 9): scale pockets, milk-scale disease, Notemigonus crysoleucas, AR; F. Meyer (1967): during acute phase, conspicuous hemorrhagic areas present at base of scales with ameboid stages, containing two to four spores. Author: many cases in Arkansas, 1974-1985; sometimes entire sporoplasm stains with iodine, and there is no albuminoid halo with India ink preparation. ■ M. notropis no. 1 Fantham, Porter, and Richardson, 1939: cysts in skin, Notropis comutus, N. heterolepis, Que.; Li and Desser (1985b): many organs, Notemigonus crysoleu¬ cas, Algonquin Park, Ont. ■ M. notropis no. 2 (Fantham et al., 1939) Lorn and Noble, 1984 (Syn. Myxosoma n.)\ large cysts in abdom¬ inal cavity and liver, Notropis comutus, Que.; Desser and Paterson (1978): morphology of spores and mucous envelope, Ont. ■ M. nuevoleonis Segovia Salinas, 1989a, 1989b: bone (cartilage?) at base of fins, Poecilia mexicana, P. reticulata, Nuevo Leon, Mexico. ■ M. obliquus Kudo, 1934: body muscle fibers, Carpiodes velifer, IL. ■ M. oblongus Gurley, 1893: beneath skin, Erimyzon sucetta, Fox River, NC. ■ M. okobojietisis no. 1 Otto and Jahn, 1943: intestine, Pomoxis nigro-maculatus, IA. ■ M. okobojietisis no. 2 (Rice and Jahn, 1943) Lorn and Noble, 1984 (Syn. Myxosoma o.): gills, Ictiobus bubalus, IA. ■ M. orbiculatus Kudo, 1920: muscle, Notropis gilberti, IL. ■ M. orbitalis (Fantham, Porter, and Richardson, 1939) (Syn. Myxosoma o. Fantham, Porter, and Richardson, 1939). ■ M. osburni Herrick, 1936: mesenteries and peri¬ toneum, Lepomis gibbosus, Micropterus dolomiuei, Lake Erie; Cone and Anderson (1977a, 1977b): pancreatic tis¬ sue, L. gibbosus, Ont.; Dechtiar (1972b): mesenteries, M. dolomieui, Ont.; IA; G. L. Hoffman (unpub., USFWS, 1980): L. macrochirus, ME; Ingram and Mitchell (1982): pancreas, L. gibbosus, includes histopathology, none of 341 L. macrochirus were infected; Li and Desser (1985a): viscera, L. gibbosus, Algonquin Park, Ont. (Author: this was published as M. gibbosus, so it may be M. gibbosus, no. 1, without glycogen vacuole); Otto and Jahn (1943): gall bladder, L. macrochirus, P. nigro-maculatus. ■ M. ovalis (Davis, 1923) (Kudo, 1933) Lorn and Noble, 1984 (Syn. Myxosoma o.): gills, Ictiobus bubalus, I. cyprinella, IA; G. L. Hoffman (unpub., USFWS, 1983): in heavy infections, in skin and gills, I. bubalus, I. cyprinella, AR; C. C. Long and Meglitsch (1970): Carpiodes velifer,

■ M. ovoidalis Fantham, 1930: skin, Salvelinus fontinalis, Que.; Fantham et al. (1939): same. ■ M. paralintoni Li and Desser, 1985a: heart, Lepomis gib¬ bosus, Algonquin Park, Ont. ■ M. parellipticoides (Fantham, Porter, and Richardson, 1939) (Syn. Myxosoma p. Fantham, Porter, and Richard¬ son, 1939): large abdominal cyst, Pfrille neogaeus, Que. ■ M. pavlovskii Akhmerov, 1954: gills, Hypophthalmichthys molitrix, former Soviet Union; Jakovtchuk (1987): biology and life cycle, including experimental infection, former Soviet Union; Lucky (1978): gills, H. molitrix, Aristichthys nobilis, Czech Republic and Slo¬ vakia, but probably imported from Hungary; Molnar (1979d) fresh and "aged" spores not infective, histopathology. ■ M. pellucides Li and Desser, 1985a: gills, Semotilus atrornaculatus, Algonquin Park, Ont. ■ M. pendula (Guilford, 1967) Lorn and Noble, 1984 (Syn. Myxosoma p.): gills, Semotilus atromaculahis, WI; Dechtiar (1972a): same, Lake Erie, Ont.; Nepszy (1988): same, cyst attached pendulously to gill arches. ■ M. percae Fantham, Porter, and Richardson, 1939: small cysts at base of pectoral fins, Perea flavescens, Que.; Dechtiar (1972b): cysts at fin base and eye, P. flavescens, Ont. ■ M. pfrille (Fantham, Porter, and Richardson, 1939) Lorn and Noble, 1984 (Syn. Myxosoma p.): large abdominal cyst, Phoxinus neogaeus, Que. ■ M. pharyngeus (J. D. Parker et al., 1971) Lorn and Noble, 1984 (Syn. Myxosoma p.)\ pharyngeal cavity of Gambusia affinis, pathogenic, OK. ■ M. poecilichthidis Fantham, Porter, and Richardson, 1939: creamy cysts in connective tissue of gut, Etheostoma exile, Que.; Li and Desser (1985b): gills, Lepomis gibbosus, Algonquin Park, Ont. ■ M. procerus (Kudo, 1934) Lorn and Noble, 1984 (Syn. Myxosoma p.)\ skin, Percopsisguttatus, IL; Dechtiar (1972a): Percopsis omiscomaycus, Ont.; Guilford (1965): skin, P. omiscomaycus, Lake Michigan; G. L. Hoffman (unpub., USFWS, 1980, 1981): P. omiscomaycus, Lake Erie, Pimephales promelas, Culaea inconstans, Saskatoon, Sask.; Nepszy (1988): P. omiscomaycus, Lake Huron, Ont. ■ M. pseudodispar Gorbunova, 1936. Lorn et al. (1976): many fishes, including Cyprinus carpio; Shulman (1966): same. ■ M. pseudokoi Li and Desser, 1985a: gills, skin, Notro¬ pis comutus, Algonquin Park, Ont. ■ M. rhinichthidis Fantham, Porter, and Richardson, 1939: cysts beneath skin, Rhinichtliys atratulus, Que.; Cone (pers. comm., St. Mary's Univ., MD, 1993): same.

Family Myxobolidae

■ M. robustus (Kudo, 1934) Lom and Noble, 1984 (Syn. Myxosoma r.)\ connective tissue, Notropis comutus, IL. ■ M. rotundahis Dogiel and Akhmerov, 1960. Lom et al. (1976): Cyprinus carpio, East Asia; Shulman (1966): intes¬ tine and fins of many fishes, including C. carpio. ■ M. rotimdus (Meglitsch, 1937) Lom and Noble, 1984 (Syn. Myxosoma r.): gills, Carpiodes cyprinus, IL; Dechtiar (1972a): same, Ont.; Landsberg and Lom (1991): trans¬ ferred to M. carpioides; Nepszy (1988): C. cyprinus, Lake Erie, Lake Huron, Ont. ■ M. sarigi Landsberg, 1985: Tilapia aureus x T. niloticus, Israel. ■ M. scardinii Reuss, 1906 (is M. bramae Reuss, 1906, according to Shulman, 1984). Li and Desser (1985b): gut, kidney, Notropis heterolepis, Algonquin Park, Ont. ■ M. schuberti Li and Desser, 1985b: brain, kidney, muscle, nares, spleen, Notropis comutus, Algonquin Park, Ont. ■ M. scleropercae (Guilford, 1963) Lom and Noble, 1984 (Syn. Myxosoma s.): cartilage of sclera of eye, Perea flavescens, Percina caprodes, Lake Michigan, WI; Dechtiar (1965a): Lake Erie; Dechtiar (1972a): same; Nepszy (1988): Percina caprodes, Lake Huron, and Perea flavescens, Lake Erie; J. Schachte (pers. comm., State Fisheries, 1982): P. flavescens, NY. ■ M. spalli (Spall, 1974) Landsberg and Lom, 1991 (Syn. Myxosoma cyprini Spall, 1974). ■ M. sparoidis Otto and Jahn, 1943: gall bladder, intes¬ tine, Pomoxis nigro-maculatus, IA. ■ M. squamae Keysselitz, 1908. Lom et al. (1976): Cypri¬ nus carpio, Asia; Shaw (1947): skin, Oncorhynchus kisutch, OR; see Shulman (1966, p. 250). ■ M. squamalis (Iversen, 1954) Lom and Noble, 1984 (Syn. Myxosoma s.): scales, Oncorhynchus keta, O. kisutch, O. mykiss; Hoskins et al. (1976): O. kisutch, O. keta, B.C.; C. Jensen (pers. comm., State Fisheries, 1961): causes red sore-like areas, handling may cause bleeding, fisher¬ men reject affected fish, OR; Olson (1978): O. kisutch, O. tshawytscha, Pacific Ocean, OR. Author: spore is similar to M. cerebralis but has two distinctive ridges, one on either side of suture. ■ M. squamosus Kudo, 1934: subepithelium, Hybopis kentuckiensis. ■ M. subcircularis Fantha, Porter, and Richardson, 1939: muscle of pelvic fin, Catostomus commersoni, Que.; Cone (1983): abdominal muscle, 2 of 14 juvenile C. commer¬ soni, N.B., includes photos.

71

■ M. teres Kudo, 1934: body muscle, Notropis whipplii, IL. ■ M. toyamai Kudo, 1915: gills, Cyprinus carpio, Japan. Single polar capsule; see Thelohanellus toyamai. ■ M. transovalis (Gurley, 1894) Lom and Noble, 1984 (Syn. Myxosoma t.): under scales, Clinostomum funduloides, VA; Rice and Jahn (1943): gills, Ictiobus bubalus, IA. ■ M. transversalis (Gurley, 1893) Fantham, Porter, and Richardson, 1939: muscle, Notropis comutus, Que.; Molnar et al. (1974): kidney, Hybognathus hankinsoni, N. heterolepis, Laurel Creek, Ont. ■ M. uvidiferensis Cone and Anderson, 1977a: in the cyst of Uvulifer ambloplitis (blackspot metacercaria), Lepomis gibbosus, Ont.; some spores with posterior and lateral extensions of shell valves. ■ M. vastus Kudo, 1934: corium, above scales, Moxostoma aureolum, IL. ■ M. wellerae Li and Desser, 1985b: muscle, Notropis comutus, Algonquin Park, Ont. The following list of inadequately described Myxobolus and formerly Myxosoma species is included here to help in the identification of undescribed species. ■ Myxobolus sp. Arai and Mudry (1983): in 14 fish species, B.C. ■ Myxobolus sp. Arthur, Margolis, and Arai (1976): Catostomus catostomus, C. cognatus, Arshihik Lake, Canada. ■ Myxobolus sp. Bardach (1951): cause of ulcerations and mortality, Perea flavescens, Lake Mendota, WI. ■ Myxobolus sp. Brienholt and Heckmann (1980): skin, Catostomus discoides, C. latipinnis, UT. ■ Myxobolus sp. Chung and Kou (1977): "boil" type in Cyprinus carpio, Taiwan. ■ Myxobolus sp. Cone and Wiles (1985b): trophs and spores in gills, Notemigonus crysoleucas (possibly M. angustus); a different form in gill arch of juvenile Catostomus commersoni, N.S. ■ Myxobolus spp. Dechtiar (1972a): intestinal wall, Ambloplites mpestris; gills, Cottus bairdi; muscle, Etheostoma exile, E. nigrum; gills, Ictalums nebulosus; connective tis¬ sue, Lepomis gibbosus; gills, Lota lota; gills, Moxostoma anisurum, M. erythrurum; gills, Noturus gyrinus; gills, Percina caprodes, Pungitius pungitius, Rhuiichthys cataractae, Lota lota, Ont. ■ Myxobolus sp. Fantham (1926): bile and duodenum, Carassius auratus, South Africa.

■ M. subtecalis (Bond, 1939a) (Syn. Myxosoma s. Bond, 1939a): viscera, fat of cranial cavity and kidney, Fundulus heteroclitus, Chesapeake Bay, Hudson River; Bond (1939a): spores induced infection in F. heteroclitus.

■ Myxobolus sp. Hanek and Molnar (1974): intestine, gall bladder, Anguilla rostrata, Que.

■ M. symmetricus Rice and Jahn, 1943: gills, Ictiobus bubalus, IA.

■ Myxobolus sp. Heckmann, Kimbal, and Short (1987): stomach wall and other muscle, Cottus bairdi, LIT.

■ Myxobolus sp. Heckmann and Farley (1973): Hesperoleucas symmetricus, CA.

72

Subkingdom Protozoa

■ Myxobolus sp. G. L. Hoffman (Chan, pers. comm., 1979) : in large intracapsular appendix, Pimephales prome¬ las, Ont. ■ Myxobolus sp. G. L. Hoffman (unpub., USFWS, 1965): cartilage, Cottus bairdi, Catostomus sp., PA. ■ Myxobolus sp. Horner (pers., comm., State Fisheries, 1980) : small cysts in meninges, connective tissue, near spinal column, hind kidney, Pomoxis nigro-maculatus, IL. ■ Myxobolus sp. T. L. Joyce (pers. comm., Ctr. Piscicole Natl., Cameroun, 1975): killed Tilapia nilotica fingerlings, West Africa. ■ Myxobolus sp. Kozel and Whittaker (1982): gills, Etheostoma caeruleum, KY. ■ Myxobolus sp. Mitchum (pers. comm., State Fisheries, 1966, WY): Catostomus ardens, C. catostomi, WY. ■ Myxobolus spp. Molnar and Fernando (1974): gills, Ambloplites rupestris, Etheostoma exile, Hybognathnus hankinsoni, Nocomis biguttattus, Notropis comutus, Pimephales promelas, Semotilus atromaculatus; gall bladder, intes¬ tine, H. hankinsoni, N. biguttattus, N. comutus, Pimephales notatus, S. atromaculatus, Ont. ■ Myxobolus sp. Muzzal, Whelan, and Peebles (1987): gills, Lota lota, MI. ■ Myxobolus sp. J. O'Grodnick (pers. comm., State Fish¬ eries, 1970, PA): cartilage, Semotilus atromaculatus, PA. ■ Myxobolus sp. R. L. Price and Jilek (1980): intestinal wall, Dorosoma cepedianum, IL. ■ Myxobolus sp. Sutherland and Holloway (pers. comm., Univ. of North Dakota, 1979): gills, Pimephales promelas, ND.

of carp fry caused dark fin pigmentation (blackspot?), Hungary; Desser et al. (1983): electron microscopy of sporogenesis, large tail lesions, C. carpio, Hungary; Jeney (1979): gills, fins, mesenteries, liver, C. auratus, C. carpio, Hungary (probably imported); Lom et al. (1976): C. car¬ pio, East Asia. ■ T. fuhrmani (Auerbach, 1909) (Syn. T. acuminatus, T. saurogobi): in gills, muscles, kidneys, liver, connective tissue, nine fish species, including Cyprinus carpio, Europe; Ki and Ha (1971): gills, C. carpio, northern Vietnam. ■ T. kitauei Egusa and Masumura, 1981: intestinal giant cystic disease, Cyprinus carpio, Japan; this species has a thin-walled, balloonlike sac around the spore. ■ T. notahis (Mavor, 1916) Kudo, 1929: subdermal white cysts, Pimephales notatus, IL; Fantham and Porter (1947): base of tail, Catostomus commersoni, Notropis comutus, Que.; G. L. Hoffman (unpub., USFWS, 1976, 1979): black cysts, Semotilus atromacidahis, MI, and N. hudsonius, OH; Kudo (1934): subdermal cysts, N. blennius, N. cornutus, P. notatus, P. vigilax, IL; Li and Desser (1985a): Notemigonus, comutus, N. crysoleucas, N. heterolepis, Semotilus atromaculatus, Algonquin Park, Ont.; Nepszy (1988; pers. comm., Fisheries, Ont., 1983): sections, head of P. promelas, MN; R. Walker (pers. comm., Rensselaer Inst., NY, 1983): N. hudsonius, Troy, NY. Author: some of these records include black-spot cysts and others do not for two possible reasons: species variant differ¬ ences and the age of the cyst. The same variation apparently occurs with European species: R. Hoffmann, Munich Univ., has blackspot Thelohanellus slides.

■ Myxobolus sp. Szalai and Dick (1987a): Carpiodes cyprinus, Man.

■ T. oculi-leucisci (Trojan, 1909): vitrous humor of eye, several fishes, including Carassius auratus, Cyprinus car¬ pio, Europe; Lom et al. (1987): brain, Gobiogobio, Czech Republic and Slovakia.

■ Myxobolus sp. Witala (1970): as pathogen of Cyprinus carpio, histopathology, Poland.

■ T. oviformis Li and Desser, 1985a: eye, muscle, Notemigonus crysoleucas, Algonquin Park, Ont.

Genus Thelohanellus Kudo, 1933 (Fig. 97) Spores pyriform, tear-shaped, or ellipsoid in valvular view; tear-shaped or pyriform in sutural view. Single pyriform, tear-shaped, or subspherical polar capsule present; binucleate sporoplasm usually with glycogen vacuole. Histozoic in freshwater fishes.

■ T. pyriformis (Thelohan, 1892) Kudo, 1933. Dogiel et al. (1958): cysts in muscle, kills coregonids; Dykova and Lom (1987): development in gill filament arteries with contact hypertrophy, Tinea tinea, Czech Republic and Slo¬ vakia; Ghittino (1962): in T. tinea from fish market, Italy; Kudo (1918): spleen, Perea flavescens, MA; Shul¬ man (1966): several European fish hosts, including T. tinea.

— Ref. Lom and Noble (1984); Shulman (1966, p. 227).

■ T. toyamai Kudo, 1915: gills, Cyprinus carpio, Japan; Ki and Ha (1971): C. carpio, northern Vietnam; Lom et al. (1976): C. carpio, Asia, Europe.

■ Thelohanellus callisporis Ki and Ha, 1971: skin, gills, Cyprinus carpio, northern Vietnam.

■ Thelohanellus sp. Dechtiar (1972a): Notropis hudso¬ nius, Lake Erie, Ont. (may be T. notatus).

■ T. dogieli Akhmerov, 1955 (Syn. T. amurensis, T. carassi, T. cyprini, T. hovorkae, T. nikolskii): gills, fins, skin, mesen¬ tery, liver, Carassius auratus, Cyprinus carpio, Asia; Cirkovic (1987): pathogenicity inconclusive, former Yugoslavia; Circovic and Jovanovic (1987): experimental infection

■ Thelohanellus sp. R. Hoffmann (pers. comm., Munich Univ., 1983): black-spot cysts, Scardinius erythrophthalmus, Germany. ■ Thelohanellus sp. R. Horner (pers. comm., Fisheries, 1991, IL): Notropis hudsonius, Lake Michigan, IL.

Family Chloromyxidae

■ Thelohanellus sp. A. J. Mitchell (pers. comm., USFWS, 1969): Carassius auratus, Stuttgart, AR.

73

Genus Ceratomyxa Thelohan, 1892 (Fig. 99)

Genus Unicauda Davis, 1944 (Fig. 98) Spore differing from Henneguya in that the single caudal appendage is not a continuation of the shell valves, but a structure made from different material and adher¬ ing to the shell valves along a distinct boundary. Histozoic in freshwater fishes. — Ref. H. S. Davis (1944a); Kudo (1934, 1971); Lorn and Noble (1984).

■ Unicauda brachyura (Kudo, 1934) (Syn. Henneguya b. Ward, 1919): fin ray, Notropis anogenus, Lake Erie. ■ U. clavicauda (Kudo, 1934) (Syn. Henneguya c. Kudo, 1934): subdermal, Notropis blennius, IL; G. L. Hoffman and J. Peterson (unpub., 1993): under scales, Notropis hudsonius, MT. ■ U. crassicauda (Kudo, 1934) (Syn. Henneguya c. Kudo, 1934): skin, fins, Capostoma anomalum, IL; Arai and Mudry (1983): gall bladder, kidney Ptychocheilus oregonensis, B.C. ■ U. fontinalis (Fantham, Porter, and Richardson, 1939) (Syn. Henneguya f. Fantham, Porter, and Richardson, 1939): skin, Salvelinus fontinalis, Que. ■ U. macrura (Gurley, 1894) H. S. Davis, 1944a: con¬ nective tissue of head, Hybognathus nuchalis, TX. ■ U. magna Minchew, 1981: fins, Piniephales promelas, fish hatchery, very long "tail," total length 110 (75-170) pm. ■ U. monura (Gurley, 1893) H. S. Davis, 1944a: sub¬ cutaneous intermuscular tissue, Phredoderus sayanus, NJ. ■ U. percae Fantham, Porter, and Richardson, 1939 (Syn. Henneguya p.): gills, Perea flavescens, Que.; Fan¬ tham and Porter (1947), same. ■ U. plasmodia (Davis, 1922) (Syn. Henneguya p. H. S. Davis, 1922): gills, Ictalurus punctatus. Unicauda sp. Joy, Tartar, and Franklin (1978): Campostoma anomalum, WV.

Transverse elongations of spore exceeding its axial diameter. ■ Ceratomyxa anguillae Tuzet and Ormieres, 1957: gall bladder, Anguilla anguilla, France. ■ C. shasta Noble, 1950: viscera, fingerling Oncorhynchus mykiss, CA; Bartholomew et al. (1989a, 1989b): very serious pathogen in fingerlings of Oncorhynchus clarki, O. gorbuscha, O. keta, O. mykiss, O. nerka, S. salar, S. trutta, O. tshawytscha, Salvelinus fontinalis, northwestern USA, western Canada. Author: current evidence indicates that, except for C. shasta, all Ceratomyxa species parasitic in fishes are in marine fishes; almost all are coelozoic. C. shasta is the "misplaced" member; not only is it in freshwater fishes, but it is histozoic, often producing fatal lesions. Diagnosticians are urged to identify the spores as well as search the intestinal scrapings for ameboid movement of the trophozoite (Yasutake, 1987). The O'Grodnick plankton centrifuge method (1975) can be used to concentrate the spores (W. G. Taylor, pers. comm., USFWS, OR, 1979). ■ Ceratomyxa sp. Arai and Mudry (1983): intestine, Catostomus macrocheilus, B.C.

Family Chloromyxidae Thelohan, 1892 Spores spherical, subspherical, or elongated; bisected by a straight meridional suture; possibly bearing caudal appendages. Four polar capsules present at apex of spore; either one pair in the level of the suture line and the sec¬ ond pair perpendicular to the suture, or both pairs diag¬ onally beyond the level of the suture. Trophozoite probably monosporous to polysporous. Coelozoic in freshwater and marine fishes; exceptionally in amphib¬ ians; rarely histozoic.

Suborder Variisporina

Genus Agarella

Lorn and Noble, 1984

Dunkerly, 1915

(Syn. Eurysporea Kudo, 1920)

(Fig. 100)

Family Ceratomyxidae Doflein, 1899 Spores with valves elongated enormously perpendicular to straight central suture; two shell valves sometimes asymmetrical. Polar capsules spherical or subspherical, close to suture line in plane perpendicular to it. Tropho¬ zoites monosporous to polysporous, usually disporous. Coelozoic in marine fishes (one in freshwater); rarely histozoic or in amphibians.

Spores elongate to ovoid; very slightly flattened paral¬ lel to the suture line; each valve extending in a caudal projection. Four pyriform polar capsules present, two larger and two smaller; one large and one small capsule in each shell valve. Histozoic in freshwater fishes. — Ref. Kudo (1971, p. 791).

■ Agarella gracilis Dunkerly, 1915: testis, South Ameri¬ can lungfish; Walliker (1969): testis, kidney, Lepidosiren paradoxa (lungfish), Brazil.

74

Subkingdom Protozoa

Genus Caudomyxum Bauer, 1948 (No fig.)

Essentially like Chloromyxum, but with one or two stout, tapering caudal projections.

■ C. fluviatilis Thelohan, 1892. Shulman (1966): in sev¬ eral fishes, including Carassius auratus, former Soviet Union, France. ■ C. fLmduli Hahn, 1915: muscle lesions, Fundulus, MA.

— Ref. Shulman (1966, p. 205).

■ C. gibbosum Herrick, 1941: gall bladder, Lepomis gibbosus, Fake Erie; Dechtiar (1972b): F. gibbosus, Ont.; G. F. Hoffman (1959c): same, WV.

■ Caudomyxum nanum O. N. Bauer, 1948: kidney, Lota lota, former Soviet Union.

■ C. giganteum Fujita, 1923: gall bladder, Oncorhynchus gorbuscha, Japan, large spore, 14-16 pm.

Genus Chloromyxum Mingazzini, 1 890 (Fig. 101)

Spore valves smooth or with ridges; rarely with caudal filamentous projections; the two pairs of polar capsules sometimes of unequal sizes. Other characters with those of the family. — Ref. Kudo (1971); Schaperclaus (1954, p. 351).

■ Chloromyxum barbi Dogiel, 1934. Sedlaczek (1987): gall bladder, Hypophthalmichthys molitrix, Germany. ■ C. carassii Akhmerov, 1960b: gall bladder, Carassius auratus, former Soviet Union. ■ C. catostomi Kudo, 1920: gall bladder, Catostomus commersoni; Dechtiar (1972b): same, Ont.; Listebarger and Mitchell (1980): same, IA, ultrastructure; L. G. Mitchell (1978a): Campostoma anomalum, Notropis dorsalis, Pimephales notatus, P. promelas, Semotilus atromaculatus, IA. ■ C. chitoseuse Fujita, 1923: gall bladder, Oncorhynchus keta, Japan. ■ C. coregoni O. N. Bauer, 1948. Shulman (1966): in sev¬ eral fishes, including 13 species of Coregomis, Oncorhynchus nerka, Salmo salar, S. trutta, Salvelinus alpinus, AK, Scandinavia, former Soviet Union; Bailey and Margolis (1987): O. nerka, B.C., WA; Moles (1982): O. nerka, AK. ■ C. cyprinus Fujita, 1927. Shulman (1966): in several fishes, including Ctenopharyingodon idella, Cyprinus carpio, Hypophthalmichthys molitrix, Amur River, Russia, Japan; Fom and Dykova (1981): liver, gall bladder, grass carps, causing massive necrosis, Czech Republic and Slovakia; Molnar (1979a): in grass carps, Hungary. ■ C. dubium Auerbach, 1908. Shulman (1966): gall bladder, Lota lota, former Soviet Union, Switzerland; Arthur et al. (1976): L. lota, B.C. ■ C. esocinum Dogiel, 1934. Shulman (1966): gall blad¬ der, Esox Indus, former Soviet Union; Arthur et al. (1976): E. Indus, B.C.; G. F. Hoffman (1978b): E. Indus, NE; Fi and Desser (1985b): gall bladder, Notemigonus crysoleucas, Ont.; Sedlaczek (1987): E. Indus, Germany. ■ C. externum H. S. Davis, 1947: gills, Semotilus margarita, Rhinichthys atratulus, WV. This is the only known exter¬ nal myxosporidean.

■ C. granulosum Davis, 1917. Arai and Mudry (1983): gall bladder, Catostomus macrocheilus, B.C. ■ C. hypophthalmichthys Hsieh 1987: gall bladder, silver carp fingerlings; swollen gall bladder; body cavity and body surface becoming yellow, China. ■ C. koi Fujita, 1913: gall bladder, Cyprinus carpio, Japan; Shulman (1966); same, former Soviet Union. ■ C. kovaljovae Evlanov, 1981: gall bladder, Lota lota, for¬ mer Soviet Union. ■ C. legeri Touraine, 1931. Molnar (1979a): gall bladder, C. carpio, Scardinius erythro, Hungary; Shulman (1966): gall bladder of several fishes, including Cyprinus carpio, former Soviet Union. ■ C. tnajori Yasutake and Wood, 1957: in glomeruli, Oncorhynchus mykiss, O. tshawytscha, WA. ■ C. montschadskii Shulman, 1962: former Soviet Union; Arai and Mudry (1983): gall bladder, Ptychocheilus oregonensis, B.C. ■ C. nanum Akhmerov, 1960b: kidney(?), Ctenopharyngodon idella, Amur River, Russia, very small spore, 3.5-5 pm; J. Brock (pers. comm., Dep. band Nat. Res., 1988): in sections of gall bladder, Ctenopharyingodon idella, HI. ■ C. opladeli Meglitsch, 1942b: gall bladder, Pylodictis olivaris (Opladelus o.), IF. ■ C. pseudomucronatum Kashkowsky, 1966: gall bladder, Lota lota, Ural River, Kazakhstan (see Shulman, 1966, p. 524). ■ C. quadrifonne Fujita, 1923: gall bladder, Oncorhynchus gorbuscha, O. keta, Japan. ■ C. renalis Meglitsch, 1947: kidney, Fundulus majalis, IF. ■ C. tanakai Fujita, 1936: gall bladder, Oncorhynchus keta, Hokkaido, Japan. ■ C. thompsoni Meglitsch, 1942b: gall bladder, Ictiobus bubalus, IF. ■ C. thymalli Febzelter, 1912. Shulman (1966): in gall bladder, Thymallus arcticus, T. thymallus, former Soviet Union. ■ C. trijugum Kudo, 1920: gall bladder, Pomoxis sparoides, Xenotis megalotis; Jameson (1931): CA; Janeke (pers. comm., 1983): gall bladder, Lepoinis macrochirus, CO; Fi and Desser (1985b): L. gibbosus, Algonquin Park, Ont.; Fistebarger and Mitchell (1980): scanning electron

Family Myxidiidae

microscopy; L. G. Mitchell (1978a): L. gibbosus, redescription, MT; L. G. Mitchell (1978b): Lepomis cyanellus, L. macrochirus, Pomoxis annularis, P. nigromaculatus, MT; L. G. Mitchell (1980): L. cyanellus, L. macrochirus, IA, epizootiology and histopathology; Otto and Jahn (1943): IA. ■ C. truttae Leger, 1906: gall bladder, Salmo trutta fario, France; Alvarez-Pellitero et al. (1982): S. trutta, Spain, redescription; H. S. Davis (1947): Salvelinus fontinalis, VT; Dogiel et al. (1958): Oncorhynchus mykiss, hypertrophy and hyperemia of bladder, sometimes no mortality but sometimes mass mortality, former Soviet Union; Ghittino et al. (1978): S. trutta, causes jaundice, Italy; Hanek and Molnar (1974): S. fontinalis, Osmerus mordax, Que.; E. R. Noble (1950): S. fontinalis, CA; Shulman (1966, p. 509): pathogenicity; Vismanis et al. (1976): serious dis¬ ease of cultured Baltic Salmo salar, pathogenicity and con¬ trol, former Soviet Union. ■ C. hiberadatum Konovalov, 1966. Shulman (1966): gall bladder, Thymallus arcticus, former Soviet Union. ■ C. wardi Kudo, 1920. L. G. Mitchell (1978a): gall bladder, Oncorhynchus clarki, O. mykiss, O. nerka, Salveli¬ nus fontinalis, S. malma, redescription; Moles (1982): O. nerka, AK; Uzmann (pers. comm., 1961): gall bladder, intestine of salmonids, WA. ■ Chloromyxum spp. Akhmerov and Martianova (1958): methods for describing species of Chloromyxum, former Soviet Union. ■ Chloromyxum spp. Arthur, Margolis, and Arai (1976): Coregonus clupeaformis, T. arcticus, B.C. ■ Chloromyxum spp. Fischthal (1947a): Catostomus catostomus, Nocomis biguttatus, Notropis cornutus, Pimephales notatus, WI.

75

smooth or with ridges; suture line bisecting the spore. Two usually pyriform capsules situated at each end of the spore; capsular foramina lying in suture plane, at or near end of spore and open mostly in opposite directions. One binucleate sporoplasm usually located between the capsules. Typically coelozoic; small or large tropho¬ zoites; monosporous, disporous, or polysporous; polysporous with pansporoblast formation; also histozoic; intracellular stages known. In marine and freshwater fishes; some species in amphibians and reptiles. Since the latter species might cause confusion with species in fishes, they are included here. — Ref. Jayasri and Hoffman (1982): worldwide review, species listed in order of increasing size; L. G. Mitchell (1967): review of North American species, key to species, illustrated.

■ Myxidium acinum Hine, 1975b: gills, Anguilla aus¬ tralis, A. dieffenbachii, scanning EM, New Zealand. Author: I am including Myxidium of eels worldwide because of the importance of eels in world trade. ■ M. americanum Kudo, 1920: kidney tubules of turtle, Trionyx apinifera, IL. ■ M. anguillae Ishii, 1915: syn. M. giardi. ■ M. aplodinoti Kudo, 1934: gall bladder, Aplodinotus gnumiens, IL. ■ M. bellum Meglitsch, 1937: gall bladder, Ictalurus punctahis, IL; G. L. Hoffman, (unpub. res., 1975): I. punctatus, AR; Janeke (pers. comm., 1975): same, CO. ■ M. chelonarum C. A. Johnson, 1969: gall bladder of turtle, Pseudemys scripta, redescription, North America. ■ M. ctenopharyngodonis Akhmerov, 1960b: kidney, Ctenopharyngodon idella, Amur River, Russia.

■ Chloromyxum sp. G. L. Hoffman (unpub., USFWS, 1966): Carson National Fish Hatchery, Carson, WA.

■ M. cuneiforme Fujita, 1924: gall bladder, Carassius carassius, Cyprinus carpio, Japan, China.

■ Chloromyxum sp. G. L. Hoffman and Putz (1963): Salvelinus fontinalis, PA.

■ M. enchelypterygii Hoshina, 1952: fins, Anguilla japonica, Japan (may be syn. of M. giardi).

Family Myxidiidae Thelohan, 1892 Spores spindle-shaped, sigmoid, or crescentic; some¬ times almost semicircular in valvular view; ellipsoidal; with two (one in Coccomyxa) polar capsules located in opposite ends, with terminal or slightly lateral capsular foramina. Longitudinal suture line straight, curved, or sigmoid. Mostly coelozoic, rarely histozoic, in marine and freshwater fishes.

Genus Myxidium Butschli, 1 882 (Figs. 91, 102)

Spores usually fusiform, straight, slightly crescentic, or sigmoid, with more or less pointed ends. Shell valves

■ M. folium Bond, 1938a: hepatic ducts, gall bladder, Fundulus heteroclitus, MD. ■ M. fusiforme Fujita, 1927: kidney, Anguilla japonica, Japan. ■ M. gasterostei Noble, 1943: gall bladder, Gasterosteus aculeatus, CA; Lester (1974): gall bladder, G. aculeahts, B.C. ■ M. giardi Cepede, 1906. Jayasri and Hoffman (1982): Anguilla vulgaris, Europe, review; Hine (1980): review of Myxidium of eels—the following are synonyms: M. anguil¬ lae, M. enchelypterygii, M. illinoisense, M. serum, M. zealandicum; Copland (1981): distribution in wild and cultured European eels in England; Copland (1983): histopathology of A. anguilla kidney, viscera, marked granulomatous changes in spleen and peritoneal fat, Scotland; Ghittino (1974): skin, A. rostrata, GA; Hanek and Molnar (1974): Que.; Hine (1975b): gills, A. australis, A. dieffenbachi as M. zealandicum, scanning EM of spores;

76

Subkingdom Protozoa

Hine (1978): variation in spores as M. zealandicum, New Zealand; Hulbert et al. (1977): reported as M. zealandicum from A. rostrata, ultrastructure, Canada; Komourdjian et al. (1977): large cyst in kidney, reported as M. zealandicum, first report from A. rostrata, Canada; Landsberg (1983): primarily in gills, hind kidney, and intestine, A. anguilla elvers, Israel; Molnar (1979a): A. anguilla, Hungary; New¬ man (1977): histopathology of cutaneous infection, A. rostrata, MD. Author: this species, or complex of sub¬ species, is apparently a serious eel pathogen worldwide. ■ M. illinoisense Meglitsch, 1937: kidney, Anguilla bostoniensis, IL; P. Hine (pers. comm., New Zealand, 1980): syn. of M. giardi; Nepszy (1988): A. rostrata, Lake Ontario, Canada.

■ M. oviforme Parisi, 1912: gall bladder, Salmo salar, Norway; Jameson (1931): Oncorhynchus mykiss, Phanerodon furcatus, CA; Jayasri and Hoffman (1982): O. gor¬ buscha, O. keta, O. mykiss, O. tshawytscha, S. salar, S. trutta, Europe, USA, former Soviet Union. ■ M. percae Fantham, Porter, and Richardson, 1939: subdermal, Perea flavescens, Canada. ■ M. pemiciosum Dogiel and Bogolepova, 1957. Shulman (1966): in many fishes, including Coregonus autumnalis, former Soviet Union. ■ M. pfeifferi Auerbach, 1908. Jayasri and Hoffman (1982): in many fishes, including Cyprinus carpio, Tinea tinea, Europe, former Soviet Union.

■ M. incurvation Thelohan, 1892. H. S. Davis (1917): gall bladder, Gambusia affinis, Fundulus majalis, NC.

■ M. phyllium H. S. Davis, 1917: gall bladder, Gambusia affinis, NC.

■ M. kudoi Meglitsch, 1937: gall bladder, Ictalurus furcatus, IL.

■ M. rhodei Leger, 1905. Jayasri and Hoffman (1982): in many fishes, including Cyprinus carpio, Europe; Dykova et al. (1987): morphology and pathology in Rutilus rutilus, Czech Republic and Slovakia.

■ M. lentiforme Fujita, 1927: kidney, A. japonica, Japan. ■ M. lieberkuhni Biitschli, 1882: urinary bladder, Esox niger, E. Lucius; Arthur et al. (1976): E. Indus, western Canada; Guilford (1965): E. Indus, Lake Michigan; Noble (1943): Lota lota, Canada, WI, Europe; Mavor and Strasser (1918): E. Indus, WI; Reichenbach-Klinke (pers. comm., Munich, 1975): muscle, E. Indus, Germany; Shulman (1966): pathogenic. ■ M. macrocapsulare Auerbach, 1910: gall bladder, Scardinius; Otto andJahn (1943): in Aplodinotis grunniens, IA; Li and Desser (1985b): kidney, Notropis heterolepis, Ont.; L. G. Mitchell (1978b): Catostomus commersoin, Ictalurus punctatus, Lepomis macrochirus, Notropis dorsalis, IA. ■ M. matsuii Fujita, 1929: gall bladder, intestine, Catosto¬ mus macrocheilus, B.C. ■ M. macrocheili L. G. Mitchell, 1967: gall bladder, bile and pancreatic ducts, Catostomus macrocheilus, MT; Arai and Mudry (1983): in gall bladder, intestine, C. macro¬ cheilus, B.C. ■ M. melum Otto and Jahn, 1943: gall bladder, Ictalurus melas, Pomoxis sparoides, IA. ■ M. minteri Yasutake and Wood, 1957: renal tubules of Chinook and silver salmon, rainbow, steelhead, and brook trout, WA; R. E. Olson (1978): Oncorhynchus kisutch, O. tshawytscha, OR; J. W. Wood (1979): in kidneys, O. kisutch, WA; Sanders and Fryer (1970): gall blad¬ der, O. clarki, O. kisutch, O. mykiss, O. tshawytscha, Prosopium williamsoni. ■ M. moxostomatis Kudo, 1921: gall bladder, Moxostoma sp., NY. ■ M. myxocephali Fantham, Porter, and Richardson, 1940: gall bladder, Myxocephalus octodecemspinosus, Canada. ■ M. obscurum Konvalov and Shulman, 1966: gut, uri¬ nary bladder, Oncorhynchus gorbuscha, O. keta, O. nerka, former Soviet Union.

■ M. rhomboidium Shurmans-Stekhoven, 1920: kidney, Gasterosteus pungitius, Netherlands. ■ M. salvelini Mavor and Strasser, 1918: urinary bladder, Salvelinus fontinalis, WI; Bailey and Margolis (1987): Oncorhynchus nerka, B.C., WA. ■ M. serotinum Kudo and Sprague, 1940: gall bladder, Rana pipiens, USA; J. G. Clark and Shoemaker (1973): in salamander, Eurycea bislineata, WV; Kudo (1943): gall bladder, Bufo terrestris, R. clamitans, R. pipiens, R. sphenocepliala, USA. ■ M. sennn Hine, 1975b: alimentary muscles, mesentery, Anguilla dieffenbachi, New Zealand; Hine (pers. comm., New Zealand, 1980): syn. of M. giardi. ■ M. shulmani Konovalov and Shulman, 1976: syn. of M. salvelini. ■ M. truttae Leger, 1931: gall bladder, Salmo trutta, France; Ghittino et al. (1978): S. trutta, cause of jaundice, Italy. ■ M. uchiyamae Fujita, 1927: kidney, Anguilla japonica, Japan. ■ M. umblae Leger, 1931: gall bladder, Salvelinus alpinus, France. ■ M. umbri Guilford, 1965: renal tubules, Lake Michi¬ gan. ■ M. ventricosum Shulman, 1962: renal tubules, Thymallus arcticus, Amur River, Russia. ■ M. wupeliensis Chen, 1984: kidney, Carassius auratus, China. ■ M. zealandicum Hine, 1975: syn. of M. giardi, accord¬ ing to Hine, 1980; Komourdjian et al. (1977): ultra¬ structure of sporogony, Canada. ■ Myxidium sp. Arai and Mudry (1983): kidney, Onchorhynchus tshawytscha, Prosopium williamsoni.

Family Sphaerosporidae

■ Myxidium sp. Brienholt and Heckmann (1980): gall bladder, Catostomus latipennis, UT. ■ Myxidium sp. H. S. Davis (1947): gall bladder, kidney tubules, trout. ■ Myxidium sp. Fantham and Porter (1947): gall blad¬ der, Micropterus dolomieui. ■ Myxidium sp. Gauthier (1926). ■ Myxidium sp. Guilford (1965): gall bladder, Umbra litni. Myxidium sp. Heckmann et al. (1987): gall bladder, Cottus bairdi, UT. ■ Myxidium sp. G. L. Hoffman (unpub. res., USFWS, 1980): in gall bladder, Gila robusta, OR; possibly n. sp. ■ Myxidium sp. Li and Desser (1985b): gall bladder, Perea flavescens, Algonquin Park, Ont. ■ Myxidium sp. Oka (1973): in eel, Anguilla, ingested by Ichthyophthirius. ■ Myxidium sp. Rice and Jahn (1943): gall bladder, Ictiobus bubalus, IA; Yasutake and Wood (1957): kidney, trout, WA.

Genus Zschokkella Auerbach, 1910 (Fig. 103) Spores ellipsoidal in sutural view and slightly bent; or semicircular in valvular view, with rounded or bluntly pointed ends; nearly spherical capsules opening slightly subterminally and both to one side. Single binucleate sporoplasm. Trophozoites disporous to polysporous, with pansporoblast formation. Coelozoic in marine and freshwater fishes; a few species in amphibians and reptiles. ■ Zschokkella chungshanensis Chen, 1984: intestine, Anguilla japonica, China. ■ Z. cyprini Qadri, 1962b: gall bladder, Cyprinus carpio, England; Kennedy (1974). ■ Z. linghuensis Chen, 1984: gall bladder, Cyprinus car¬ pio, China. ■ Z. nova Klokacewa, 1914. Shulman (1966): gall blad¬ der of several fishes, including Carassius auratus, Ctenopharyngodon idella, Salmo trutta, Tinea tinea, former Soviet Union; Molnar (1979a): gall bladder, liver, C. carassius, Gobio gobio, Hungary. ■ Z. orientalis Konovalov and Shulman, 1966 (in Shul¬ man, 1966): gall bladder, Oncorhynchus gorbuscha, O. keta, O. mykiss, O.tshawytscha, Salvelinus alpinus, for¬ mer Soviet Union. ■ Z. salvelini Fantham, Porter, and Richardson, 1939: kid¬ ney capsule, Salvelinus fontinalis, Canada; Hanek and Mol¬ nar (1974): intestine, S. fontinalis, Que. ■ Z. tilapiae Chen, 1984: intestine, Tilapia mossambica, China.

77

Family Ortholineidae Lorn and Noble, 1984 Genus Neomyxobolus Chen and Hsieh, 1950 (Fig. 104)

Spores ovoid in valvular view; anterior end slightly con¬ vex; posterior semicircular, flattened parallel to sutural plane. Trophozoites disporous to polysporous; coelo¬ zoic in urinary tract of freshwater fishes. ■ Neomyxobolus ophiocephalus Chen and Hsieh, 1960: kidney, Ophiocephalus spp., China; Arai and Mudry (1983): kidney, Catostomus catostomus, C. commersoni, C. macrochirus, B.C.

Family Parvicapsulidae Shulman, 1953 Asymmetrical, thin-walled spores elongated roughly in the sutural plane, with unequal valves meeting in a curved suture and two to four very small polar capsules in the apex. Trophozoites disporous to tetrasporous. Coelozoic in urinary system; histozoic in marine and anadromous fishes.

Genus Parvicapsula Shulman, 1953 (Fig. 105)

With characteristics of the family. — Ref. Shulman (1966, p. 215).

■ Parvicapsula sp. From 1979 to 1984 in Puget Sound, Washington, and Oregon, a Parvicapsula sp. was a seri¬ ous pathogen of marine pen-raised coho salmon, Oncorhynchus kisutch, causing kidney hypertrophy and tubular degeneration. In the same area, Gadus macrocephalus (Pacific cod) were found infected with fewer par¬ asites and no apparent pathogenicity, so this species was assumed to be the natural host. For further infor¬ mation, see G. L. Hoffman (1981): preliminary report; Johnstone (1984): pathogenesis and life cycle, including experimental infection and cod host, provisional name of P. kabatai; Kent et al. (1987): pathogenicity; P. Wag¬ ner (1980): report from the affected netpen farm.

Family Sphaerosporidae Davis, 1917 With two polar capsules opening at anterior tip and sit¬ uated in plane perpendicular to straight suture line (suture passing "between" polar capsules). Spores spher¬ ical, rounded, pyramidal, with tapering anterior end;

78

Subkingdom Protozoa

or elongated, often with appendages. Trophozoite mono- to polysporous; mostly coelozoic in marine and freshwater fishes; sometimes histozoic. — Ref. Lorn and Noble (1984).

Genus Acouda

N n

(Fig. 106) This was formerly Mitraspora elongata Kudo, 1920. Mitraspom was declared a synonym of Hoferellus (Lorn, 1986), but this species has no caudal filaments or "tails" and does not share characteristics of the other genera of Sphaerosporidae. Spores elongated, spherical in cross section; with two polar capsules, shell finely striated without posterior filaments; sutural line passing between polar capsules. In kidneys of centrarchids; one species. ■ Acauda elongata (Kudo, 1920) (Syn. Mitraspora elongata Kudo, 1920): in kidney, Lepomis cyanellus, IL; G. L. Hoff¬ man (1959c): L. cyanellus, L. macrochirus, Micropterus salmoides, WV; G. L. Hoffman (unpub., USFWS, 1985): L. cyanellus, AR.

Genus Hoferellus Berg, 1898 (Syn. Mitraspora Fujita, 1912) (Figs. 92-95, 108, 109, 115, 478) Spores pyramidal or ovoid, with pointed anterior end and two posterior processes from the lateral faces of the valves; between these processes many rigid, short fila¬ ments extending from the posterior surface. Suture line passing between the polar capsules (there has been some confusion about this). ■ Hoferellus carassii Akhmerov, 1960. (Syn. Mitraspora cyprini Fujita, 1912). This species was erroneously described as Mitraspora cyprini by Fujita (1912). It was described and correctly named Hoferellus carassii by Akhmerov (1960b) but was known in the literature as Mitraspora cyprini until corrected by Lorn (1986). Ahmed (1973, 1974): description of kidney enlargement dis¬ ease, morphology and life cycle (as M. cyprini), Japan; Alvarez-Pellitero et al. (1979): in cloaca, C. carassius, Spain; Gratzek (pers. comm., Univ. of Georgia, 1983): saw spores, C. auratus, CA; G. L. Hoffman (1984): description of parasite and polycystic kidney enlargement (kidney bloater), strangely sometimes unilateral, AR; Lorn (1986): correction of name and differentiation from H. cyprini of carp, Czech Republic and Slovakia; Molnar et al. (1989): only some goldfish developed polycystic kidney enlargement disease, histopathology and sites of devel¬ oping parasites, Hungary; Schlumberger (1950): descrip¬ tion of polycycstic kidney enlargement of goldfish (probably caused by H. carassii), AR; Shulman (1966): in uriniferous tubules, Carassius auratus, Amur River, Rus¬ sia; Takahashi and Kawana (1974a): on the first infection

site; Takahashi and Kawana (1974b): decline of occur¬ rence, Japan. ■ H. cyprini (Doflein, 1898) Berg, 1898 (Syn. Hoferia c. Doflein). Shulman (1966): epithelial cells and lumen of uriniferous tubules, Cyprinus carpio, former Soviet Union, Europe, spores more stubby than H. carassii; Korting (1986): in one-year-old carp, Germany; Lorn (1986): differentiation from H. carassii; Molnar et al. (1986): in renal tubules of carp, discussion of identity between Hoferellus cyprini and Mitraspora cyprini (=H. carassius), Germany. ■ H. gilsoni (Debaisieux, 1925) Lorn, Molnar, and Dykova, 1986 (Syn. Sinuloinea g. Debaisieux, 1925): attachment, urinary bladder, Anguilla anguilla, Hungar¬ ian fish farms.

Genus Myxobilatus Davis, 1944 (Fig. 107) Spores elongated, anteriorly pointed; shell valves, often with fine ridges, extending posteriorly in two caudal appendages, superficially resembling Henneguya; polar capsules pyriform. Binucleate sporoplasm possibly con¬ taining iodinophilous vacuole. Trophozoites small to large; disporous to polysporous. In the urinary system, rarely histozoic, in freshwater and marine fishes. — Ref. H. S. Davis (1944a); Lorn and Noble (1984).

■ M. asymmetricus H. S. Davis, 1944a: urinary bladder, Stizostedion vitreum, IA. ■ M. caudalis H. S. Davis, 1944a: urinary bladder, Aplodinotus grunniens, IA. ■ M. cotti Guilford, 1965: urinary bladder, Cottus bairdi, Lake Michigan, WI; Li and Desser (1985a): kidney, ureters, Lepomis gibbosus, Algonquin Park, Ont. ■ M. gasterostei (Parisi, 1912) (Syn. Henneguya g. Parisi, 1912). Shulman (1966): urinary bladder, ureter, Gasterosteus aculeatus, Pungitius pungitius, former Soviet Union, Italy; Arthur and Margolis (1975b): G. aculeatus, B.C.; Hoskins et al. (1976): G. aculeatus, B.C.; Lester (1974): G. aculeatus, B.C. ■ M. legeri (Cepede, 1905) (Syn. Henneguya l. Cepede, 1905). Shulman (1966): urinary bladder, ureters of sev¬ eral fishes, former Soviet Union, Lrance; Li and Desser (1985a): kidney, ureters, Notemigonus crysoleucas, Algon¬ quin Park, Ont.; Molnar (1988): development in cyprinids, Hungary. ■ M. medius (Thelohan, 1892) (Syn. Henneguya in. Thelohan, 1892). Shulman (1966): urinary tubules, Gasterosteus aculeatus, Pungitius pungitius, former Soviet Union, France. ■ M. mictosporus H. S. Davis, 1944a (Syn. Henneguya in. Kudo, 1920): urinary bladder, Lepomis spp., Micropterus salmoides; Booker and Current (1981): M. salmoides, ultrastructure, AL.

Family Sphaerosporidae

■ M. nostalgicus Lom, 1986: renal tubules, Tinea tinea, Czech Republic and Slovakia. ■ M. noturi Guilford, 1965: urinary bladder, ureters, Noturus gyrinus, Lake Michigan, WI. ■ M. ohioensis Davis, 1944a (Syn. Henneguya o. Herrick, 1941): urinary bladder, Lepomis gibbosus, Lake Erie; Cone and Anderson (1977a): urinary bladder, ureters, L. gib¬ bosus, Ont. ■ M. rupestris Davis, 1944a (Syn. Henneguya r. Herrick, 1941): urinary bladder, Ambloplites rupestris. ■ M. semotili Li and Desser, 1985a: ureters, Semotilus atromaculatus, Algonquin Park, Ont. ■ M. wisconsinensis Davis, 1944a (Syn. Henneguya w. Mavor and Strasser, 1916): urinary bladder, Perea flavescens, WI. ■ M. yukonensis Arthur and Margolis, 1975b: renal tubules, urinary bladder, Cottus cognatus, YT, and Gasterostus aculeatus, B.C.; Arthur et al. (1976): C. cognatus, Y.T.

Genus Sphaerospora Thelohan, 1892 (Fig. 110) Spherical or subspherical spores with valvular diameter not significantly exceeding sutural diameter; valves smooth or with ridges, often with lateral protuber¬ ances; sutural ridge often prominent; polar capsules subspherical to pyriform; two uninucleate sporoplasms present. Mono- or disporous trophozoites; coelozoic in the urinary system of freshwater and marine fishes; some histozoic. Often with intracellular stages and presporogonic development cycle in various body sys¬ tems. Recent findings of the vegetative stages causing disease in various organs, including the blood, make this an important genus in fish pathology. — Ref. Lom and Noble (1984); Lom Paloskova, and Dykova (1985): studies on blood-inhabiting forms; Shulman (1966, p. 176).

■ Sphaerospora amurensis Akhmerov, 1960. Shulman (1966): in kidney tubules, Hypophthalmichthys molitrix, Amur River, Russia. ■ S. angulata Fujita, 1912. Shulman (1966): kidney tubules, ureters, urinary bladder, Carassius auratus x Cypritms carpio, former Soviet Union; Csaba et al. (1984): possible etiology of carp disease, Hungary; Korting (1982): in swim bladder disease, Germany; Kovacs-Gayer et al. (1982): correlation between appearance of swim bladder inflammation and S. angulata; Kovacs-Gayer (1983): cutaneous sphaerosporosis in small C. carpio, Hungary. ■ S. carassii Kudo, 1919. Author: This is a European species recently found in goldfish and grass carp in USA.

79

■ S. cristata Shulman, 1962. Arthur et al. (1976): urinary bladder, ureters, Lota lota, B.C.; Shulman (1966) same, former Soviet Union. ■ S. cyprini (Fujita, 1912) (Syn. Mitraspora c. Fujita, 1912). Shulman (1966): kidney tubules, ureters, urinary bladder, Carassius auratus, Cyprinus carpio, Japan, Korea, Amur River, Russia. ■ S. diminuta Li and Desser, 1985a: kidney, ureters, Lep¬ omis gibbosus, Algonquin Park, Ont.; Lom, Desser, and Dykova (1989): description, Ont. ■ S. elegans Thelohan, 1892. Hanek and Molnar (1974): Apeltes quadroons, G. aculeatus, P. pungitius, Que.; Lester (1974): G. aculeatus, B.C.: Shulman (1966): kidney tubules, urinary bladder, and occasionally connective tis¬ sue of ovaries of several fishes, including Gasterosteus aculeatus, Lota lota, Pungitius pungitius, Europe, former Soviet Union. ■ S. galinae Evlanov, 1981: Tinea tinea, former Soviet Union; Lom, Korting, and Dykova (1985): renal tubules, T. tinea, light and electron microscopy, Czech Republic and Slovakia. ■ S. gasterostei Schurmans-Stekhoven, 1920: kidney, Pungitius pungitius, Netherlands. ■ S. hankai Lom, Desser, and Dykova, 1989: renal tubules, Ictalurus nebulosus, Ont. ■ S. ictaluri Hedrick, McDowell, and Groff, 1990: blood, kidney, Ictalurus punctatus, CA. ■ S. molnari Lom, Dykova, Pavlaskova, and Grupcheva, 1983: gills, skin, blood, Cyprinus carpio, Europe. ■ S. notropis Fantham, Porter, and Richardson, 1939: cysts under mouth epithelium of N. cornutus and in muscle of Catostomus commersoni, Que. ■ S. paulini Lom, Desser, and Dykova, 1989: renal tubules, Semotilus atromaculatus, Ont. ■ S. renalis Bond, 1938a: renal tubules, Fundulus heteroclitus, Chesapeake Bay; Bond (1939a): spores induced infection in F. heteroclitus. ■ S. renicola Dykova and Lom, 1982: kidney, Cyprinus carpio, pathogenicity, possible relationshiop of Csaba's blood forms, Czech Republic and Slovakia; Csaba (1976) and Csaba et al. (1984): possible relationship of blood and swim bladder forms to Sphaerospora; Korting and Her¬ manns (1984): high incidence in 5000 carp, patho¬ genicity, Germany; Korting (1986): present in carp with swim bladder disease; Landsberg (1986): first record in Israel; Lom et al. (1982): ultrastructure; Molnar (1988): presporogonic development; Molnar and Kovacs-Gayer (1986b): experimental evidence that blood form is S. reni¬ cola, Hungary; Odening et al. (1988): seasonal dynam¬ ics of S. renicola in carp, Germany. ■ S. tincae Plehn, 1932 (Syn. S. pernicialis Leger, 1930): head kidney, causing kidney enlargement and death of young tench, Tinea tinea, Europe; Hermanns and Kort¬ ing (1985): epizootiology and histopathology, T. tinea,

80

Subkingdom Protozoa

Germany; Lom, Korting, and Dykova (1985): redescrip¬ tion with light and electron microscopy, Germany. ■ 5. truttae Fisher-Scherl et al., 1986: kidney, Salmo tnitta, description of spores, developmental stages, Germany.

ent. Tissue parasites of freshwater fishes; two species. ■ Wardia lucii Kudo, 1921: in glomeruli, renal tubules, connective tissue of kidney, Esox niger, NY. ■ W. ovinocua Kudo, 1920: in ovary, Lepomis humilis, IL.

■ Sphaerospora sp. Baska and Molnar (1988): blood stages in cyprinids, Hungary. ■ Sphaerospora sp. Bennet and Wolke Sphaerospora-like, in kidney, S. salar, N.B.

(1986):

■ Sphaerospora sp. Fantham and Porter (1947): gall blad¬ der, Fundulus heteroclitus, Que. ■ Sphaerospora sp. Groff, McDowell, and Hedrick (1989): kidney, blood, Ictalurus punctatus, CA; may be related to proliferative branchitis of catfish. ■ Sphaerospora sp. R. C. Hamilton (1980): kidney, Carassius auratus, ultrastructure, Australia. ■ Sphaerospora sp. Hedrick, Kent, and Smith (1986): kidney of Gila bicolor and Gasterosteus aculeahis in waters containing trout infected with PKD (proliferative kidney disease). ■ Sphaerospora sp. Lom and Dykova (1981): one species in the gill filaments of cyprinids; a different species in the kidney, Cyprinus carpio, Czech Republic and Slovakia. ■ Sphaerospora sp. Molnar, Hanek, and Fernando (1974): ureter, Notropis cornutus, Laurel Creek, Ont. Proliferative Kidney Disease of Salmonids (Color Fig. 12)

In proliferative kidney disease (PKD) of salmonids one usually finds an unclassified protozoan parasite (PKX) whose primary cells often contain secondary and tertiary daughter cells (Kent and Hedrick, 1985) and "haplosporosomes" (membrane-bound osmophilic bodies, 0.1 pm) (Ferguson and Needham, 1978). The latter are usually found in haplosporidean species. PKD causes the kidneys to become greatly enlarged and sometimes causes high mortality of salmonids, usually trout, in Europe and the western United States. Sporogonic stages have been found in the lumen of kidney tubules in fishes in California (Kent and Hedrick, 1987), and spores of an unidentified Sphaerospora have been found in PKDinfected trout in Germany (Odening et al., 1988). It is possible that PKX is not a normal parasite in salmonids— not sporulating properly—but nonetheless sporulates normally in other fishes of enzootic waters, e.g., Gas¬ terosteus aculeatus and Gila bicolor (Feist, 1988; Hedrick, Kent, and Smith, 1986). For reviews of North American work on PKD, see Hedrick, Kent, and Smith (1986) and Hedrick, Kent, and Toth (1986).

Genus Wardia Kudo, 1920 (Fig. Ill) Spores in the shape of an isoceles triangle, with two con¬ vex sides; oval in profile; two large polar capsules pres¬

Order Multivalvulida Shulman, 1959 Shell of radially symmetrical spores composed of three to seven valves meeting in three to seven sutures; polar capsules, one to each valve, grouped together at the apex of the spore.

Family Kudoidae Meglitsch, 1960 Spores with four shell valves, each containing one polar capsule.

Genus Kudoo Meglitsch, 1947 (Fig. 112)

Spores stellate, quadrate, or rounded quadrate in apical view, with suture lines often indistinct; polar capsules pyriform. Two uninucleate sporoplasms present, one enveloping the other. Trophozoites small, producing one to seven spores, and also large, polysporous. No pan¬ sporoblast formation observed. Histozoic; mostly intra¬ cellular in muscles; exceptionally coelozoic or otherwise in marine fishes. ■ Kudoa cerebralis Paperna and Zwerner, 1974: in con¬ nective tissue of nervous system, Morotie saxatilis, Chesa¬ peake Bay; Paperna and Zwerner (1976): same. ■ K. funduli (Hahn, 1915) Meglitsch, 1947 (Syn. Chloromyxum f. Hahn, 1915): in muscles, fins, Fundulus hetero¬ clitus, MA, NJ. ■ K. histolytica (Perard, 1928) Meglitsch, 1947 (Syn. Chloromyxum h. Perard, 1928): muscle, Scomber scomber, causing muscle liquefaction, France; Prudhomme and Pantaleon (1959): muscle liquefaction, Salmo salar, France. ■ K. tliyrsitis (Gilchrist, 1924) Meglitsch, 1947 (Syn. Chloromyxum t. Gilchrist, 1924): body muscle liquefac¬ tion, Thyrsites atum; Harrell and Scott (1985): pseudocysts and muscle liquefaction, Salmo salar in pen culture, WA. ■ Kudoa sp. E. Mateo (pers. comm., Lima, Peru, 1972): causing serious muscle liquefaction in marine food fishes in Peru. Author: this species is strictly a marine par¬ asite but is included here because anadromous fishes may be affected. Infected fishes should not be shipped because of the unsightly muscle liquefaction.

Phylum Microspora

Family Pentacapsulidae Naidenova and Zaika, 1970 Five shell valves, each containing one polar capsule.

Genus Pentacapsula Naidenova and Zaika, 1970 (Fig. 113)

Spores stellate in a pentaradiate form in apical view. Large polysporous trophozoites histozoic in muscles of marine fishes. ■ Pentacapsula muscularis Cheung et al., 1983: causing soft and mushy muscle in marine aquarium butterfly fish ('Chaetodon collare), N.Y. Aquarium.

Class Actinosporea

81

■ Triactinomyxon dubium Granata, 1924. Marques (1984): Tubifex tubifex, sporoplasm containing 32 sporo¬ zoites, France, Poland. Author: this species is included because Hamilton and Canning (1987b) found it in their Myxobolus cerebralis experimental host, Tubifex. They believed it not to be a part of the M. cerebralis life cycle yet the same Triactinomyxon species that Wolf and Markiw (1984) found in their infection experi¬ ments with M. cerebralis. Because Triactinomyxon is a part of the M. cerebralis life cycle, the name T. gyrosalmo is a synonym of T. dubium (Corliss, 1985). If Triactino¬ myxon is in fact part of the cycle of Myxobolus cotti, as claimed by El-Matbouli and Hoffmann (1989), and if it is an undescribed species, then the name T. myxoboli cotti should stand.

Phylum Microspora Sprague, 1977

Noble, 1980 Included because of reported connection to the life cycles of Myxobolus cerebralis, M. cotti, and M. arcticus (El-Matbouli and Hoffmann, 1989; Kent, pers. comm., 1992; Wolf and Markiw, 1984). Parasites in tissues of aquatic invertebrates, especially Annelida. Spores with three polar capsules, each containing a coiled filament; spore membranes with three valves. Several to many sporozoites in sporoplasm. Four families: two with no tail-like processes; one with more than three tail-like processes; and the family Triactinomyxidae, described below, with three processes on each spore. Spores single or connected in nets. — Ref. Janiszewska (1955, 1957): new systematics, new genera and species, sexual cycle, morphology, ecology and history of investigations; Lee et al. (1985): brief review; MacKinnon and Adam (1924): life history of Tri¬ actinomyxon; Marques (1984): ultrastructure, life cycle, systematics.

Order Actinomyxida Lom, 1980

Family Triactinomyxidae Janiszewska, 1957 Long tubular processes protruding from each of the three valves; spores separate or connected by ends of processes ("tails"). Four genera.

Genus Triactinomyxon Stoic, 1899 (Fig. 114)

Spore anchorlike in shape; sporoplasm (main body of spore) containing 8-100 sporozoites and with elon¬ gated conelike shape. Nine species known.

Microsporidia are unicellular, intracellular parasites that do not have visible polar capsules as do the myxosporidia; instead, they have a minute tube, the polar filament, that lies coiled in the intact spore. On stim¬ ulation, the filament (tube) is everted through the spore wall with force, enabling the tip to penetrate a host cell membrane. The infective agent, known as the sporo¬ plasm, passes from the spore through the tube and enters the cytoplasm directly. The spores are small, 2-6 pm, rarely 20 x 6 pm, vary from spherical to cylindri¬ cal (those of fish are cylindrical), and are highly refractile. In fresh mounts, a clear area (vacuole) may often be seen near one end, sometimes at both ends. No other structures are visible with light microscopy. When spores enter the digestive tract of a susceptible host, the polar tubes are extruded, and the sporoplasms pass through as amoebulae to enter the gut epithelium. They then pass into the blood stream or body cavity and reach the target site of infection. There they grow and undergo repeated binary or multiple divisions, filling the host cells with continuously increasing numbers of meronts (round, oval, or elongated), which eventually transform into sporonts. Sporoblasts develop inside the sporonts and usually become sporophorous vesicles (SPOV). The number of spores in each sporont varies with genus and species. In some genera, e.g., Pleistophora. the SPOV has a durable membrane, so that a charac¬ teristic number of spores are enclosed and visible in wet mounts. In other genera, e.g., Glugea and Loma, the SPOV membrane is thin and fragile and breaks in wet mounts, thereby releasing the spores. Ultramicroscopic and taxonomic aspects are covered in Sprague (1977) and Sprague et al. (1992). — Ref. Canning et al. (1986): classification, methods; Dykova and Lom (1980): histopathology; G. L. Hoffman (1980): blackspot cyst (see Microsporidiuni sp.); Hussey (1971): cercariae as hosts, in trematodes of fishes (see Nosetna stri-

82

Subkingdom Protozoa

geoideae); Larsson (1983): identification key methods; Leida et al. (1980): concentration, detection; Loubes (1979): meiosis, ultrastructure; Mathews and Mathews (1980): methods; Modin (1981): rodlet cell as host; Muzzioli (1975): ultrasonic methods used to liberate spores; Niederkorn et al. (1980): antigenicity of several genera; R. E. Olson (1976): experimental infection; Parker and Warner (1970): measurement of spores, including shrinkage effects of fixation and dehydration; Porte and Chilmonczyk (1974): transovarial unnamed species; Sprague (1977): classification; Summerfelt and Warner (1970): variation in spore size can be significant; Undeen and Avery (1983): concentration, detection; Vavra and Maddox (1976): methods; Vavra and Sprague (1976): glossary; Weidner et al. (1984): sporoplasm discharge through polar tube.

Class Microsporea Delphy, 1963 (Fig. 116) The revised classification by Sprague et al., 1992, is not included in this section. The reader is referred to the orig¬ inal article listed earlier. This main group of microsporidia includes species with elongated, oval, to tubular spores, each with a long, coiled, polar tube and one uninucleate or binucleate sporoplasm. The pansporoblast membrane is thin and may be persistent or resorbed. The number of nuclei within the spore, as identified with the Feulgen reaction or staining with Giemsa after HC1 hydrolysis (Weiser, 1976), divides the class into two orders: Pleistophoridida Stemple, 1906, with uninucleate spores, and Nosematidida Labbe, 1899, with binucleate spores. The fish microsporidia are in order Pleistophoridida; there are 108 species in fishes and 2 in myxosporidans. In some cases, electron microscopy is needed to determine genera.

Order Pleistophoridida

Key to the Genera of Microsporidians in Fishes (Edited byJ. Lom, Czech Republic and C. Morrison, Canada)

1. Xenoma (hypertrophied host cell) produced; sporophorous vesicle (pansporoblast membrane) fragile, usually not persisting in wet mounts, thus freeing the spores.2 1. No xenoma produced.3 2. (1) Large xenoma (up to 10 pm) with most vegetative stages at periphery; many spores in sporophorous vesicle (see also Heterosporis) .(Figs. 1 17-121, 129-132, 135, 137, 138) Glugea 2. (1) Usually smaller xenoma, with vegetative stages distributed throughout; spores fewer, usually eight in sporophorous vesicle.(Figs. 123, 133, 138c) Loma 3. (1) Very small spores, 1 x 2 pm, visible intracellularly. ... 4 3. (1) Spores larger, 2-4 x 5-8 pm, not recognizably intracellular.5 4. (3) Found in hematopoietic cells of salmonids .(no fig.) Enterocytozoon salmonis 4. (3) Found in rodlet cells of salmonids .(Figs. 124, 125, 136) Microsporidium rhabdophilia 5. (3) Sporophorous vesicle membrane durable; spores usually in packets of variable numbers; packets usually persisting in wet mounts; spores within each packet in same developmental stage, but packets at different developmental stages .(Figs. 126-128, 134) Pleistophora 5. (3) Similar to Pleistophora, but forming xenoma; found once in Germany.(Fig. 122) Heterosporis * Unable to classify (see also no. 4) .(Figs. 124, 125, 136) Collective group Microsporidium

Stempell, 1906

Family Pleistophoridae Stempell, 1909 Spores uninucleate, thin-walled; with direct peroral infectivity. In sporogony, nuclei dividing irregularly and producing variable numbers of sporoblasts and spores.

Genus Enterocytozoon Desportes, Le Charpentier, Galian, Bernard, Cochand-Priollet, Lavergne, Ravisse, and Modigliani, 1985

Diplokaryotic meronts lying in direct contact with host cell cytoplasm; sporogony polysporoblastic—probably octosporoblastic—by multiple fission of spherical sporo-

gonial plasmodia with isolated nuclei. Development of spore organelles, including polar tube, well-advanced before fission of sporogonial plasmodium into sporob¬ lasts. Spores uninucleate, with endospore layer of the wall poorly developed. Found in the enterocytes of a man suf¬ fering with AIDS; also found in hematopoietic cells of Oncorhynchus spp. ■ Enterocytozoon salmonis Chilmonczyk, Cox, and Hedrick, 1991: ovoid spores (1x2 pm) in nuclei of hematopoietic cells, Oncorhynchus tshawytscha, causing anemia, lymphoblastosis, leukemia, CA, WA; Elston et al. (1987): first account; Hedrick et al. (1980): causing leukemia; Kent and Dawe (1990): experimental trans¬ put continues on page 86)

polar cap basal portion of filament polaroplast, laminated area sporoplasm nucleus polar filament coils spore membrane system saccate termination of polar filament polaroplast, granular area 116. Microsporidian

- Microsporea FIG. 116. Schematic of a microsporidian

spore (from Putz and McLaughlin, 1970, by permission of the American Fisheries Society). FIG. 117. Glugea cepedianae, development: a, schizont showing scopula-like end, b, binucleate meronts; c, young sporont; d, binucleate sporont; e, quadrinucleate sporont; f, multinucleate sporont; g, sporont with young sporoblasts; h, sporont; i, section of a cyst (from Putz et al., 1965). FIG. 118. Glugea anomala, spore (from Canning et al., 1986, by permission of Academic Press). FIG. 119. Glugea hertwigi, spore (from Canning et al., 1986, by permission of Academic Press). FIG. 120. Glugea cepedianae, spore. FIG. 121. G. hertwigi, spore (from Schrader, in Kudo, 1954). FIG. 122. Heterosporis finki, binucleate initial stages, probably meronts, within hypertropohic connective tissue cells (from Canning et al., 1986, by permission of Academic Press). FIG. 123. Loma salmonae, spore (from Putz et al., 1965). FIG. 124. Microsporidium rhabdophilia spores (from Canning et al., 1986, by permission of Academic Press). FIG. 125. Microsporidium spores: a, M. valamugili; b, M. sauridae, marine species given as examples (from Canning et al, 1986, by permission of Academic Press). FIG. 126. Pleistopliora ovariae, spores (from Canning et al., 1986, by permission of Academic Press). FIG. 127. P. ovariae, spore (from Putz et al., 1965). FIG. 128. Pleistophora hyphessobryconis, spores (from Canning et al., 1986, by permission of Academic Press).

117. Glugea

121. Glugea

118. Glugea

122. Heterosporis

1 |jm

123. Loma

124. Microsporidium

- Microsporea FIG. 129. Glugea pimephales, infection of Pimephales promelas; note distended, whitish abdomens of top two fishes (from Morrison et al., 1985). FIG. 130. G. pimephales, sagittal section of infected P. promelas; visceral mass (arrow) consists mostly of spores. FIG. 131. G. pimephales, spores, squash preparation of formalin-fixed xenoma, phase contrast (from Morrison et al., 1985). FIG. 132. Glugea cepeclianae, infection of Dorosoma cepedianum; large xenoma filled with spores, centimeter scale (from Putz et al., 1965). FIG. 133. Loma salmonae, xenoma in gill of Oncorhynchus mykiss. FIG. 134. Pleistophora hyphessobryconis, from massive visceral and muscular lesions of Capoeta tetrazona; note that many spores are still contained in the sporoblastic vesicle. FIG. 135. Glugea anomala (skin cysts) and Schistoceplialus (visceral tapeworm) in Gasterosteus aculeatus (courtesy of E. Elkan, Northwood, England). FIG.136. Microsporidium rhabdophilia, spores in disintegrating rodlet cell in intestinal scaping of Oncorhynchus tshawytscha (from Modin, 1981, by permission of Blackwell Scientific Publishers).

- Life cycle of Glugea attomala FIG. 137. Diagrammatic representation of part of a xenoma from a stickleback: A = layer of apposed connective tissue, product of the host response; B = retractile xenoma wall, composed of layers of the cell coat; C = cell membrane of the xenoma; D = peripheral layer of the xenoma, with increased pinocytotic activity; E = host cell cytoplasm; F = uninucleate, initial meronts; G = division of meronts; H = cylindrical meronts, some with nuclei in the process of dividing, and one with the sporophorous vesicle (SPOV) being formed around it (arrow); I = host cell nucleus; J = rounded meronts; K = elongate sporogonial plasmodium within an SPOV starting to segment into sporoblast mother cells; L = radial segmentation of sporognial plasmodium into sporoblast mother cells; M = SPOV with sporoblast mother cells; N, O = sporoblast mother cells dividing; P = SPOV with sporoblasts; Q = SPOV with spores; R = SPOV preserved even after breakdown of many of the vesicles, with release of mature spores into the central space; S, T = agglomeration of free spores in the space occupying the center of the xenoma (from Canning et al., 1986, by permission of Academic Press).

86

Subkingdom Protozoa

- Microsporea FIG. 138a,b. Glugea pimephales: a, EM of spore, showing polar filament with 15-16 turns, nucleus (N), and posterior vacuole (PV); b, early sporoblast, nucelus (N) in anterior end and membranes (M) of sporophorous vesicle present (from Morrison et al., 1985). FIG. 138c. Loma salmonae, EM of spore (courtesy of C. Morrison, Halifax, N.S.).

mission; J. K. Morrison et al. (1990): causing lympho¬ blastosis in freshwater fishes. Note: Microsporidiwn rhabdophilia may belong to this group.

Genus Glugea Thelohan, 1891 (Figs. 1 1 7-121, 129-1 32, 1 35, 1 37, 1 38, Color Figs. 13, 14)

Nuclei isolated throughout development; meronts cylin¬ drical, ribbonlike, with electron-dense coat on plasma membrane; surrounded by cisterna of host endoplasmic reticulum. Polysporoblastic sporogony in sporophorous vesicle: multinucleate sporogonial plasmodia dividing by multiple fission to give multiple sporoblast mother cells, these cells in turn dividing by binary fission into sporoblasts. Sporophorous vesicle wall fragile, not persisting in wet mounts, thsu releasing spores. Large and variable number of uninucleate spores produced within sporophorous vesicle. Inducing usually large xenomas encapsulated in sloughing layers of host cell coat, in which the host cell nucleus is highly branched in periph¬ eral cytoplasm. Developmental stages of the parasite lying peripherally and spores lying centrally in the xenoma. Electron microscopy needed for some details. Hosts usually fishes. Glugea anomala (typical of genus) producing xenomas in "cysts," sometimes 10 mm in diameter, in the stickleback. Spores usually oval. — Ref. Canning et al. (1986): review, including ultrastruc¬ ture; Dykova and Lorn (1980): histopathology, patho¬

genicity, Czech Republic and Slovakia; Kudo (1971, p. 813); Lee et al. (1985, p. 377); Sprague (1977): taxo¬ nomic review; Weidner (1972, 1976a, 1976b): ultra¬ structure; Weidner et al. (1984): spore discharge and transfer of polaroplast membrane.

■ Glugea anomala (Moniez, 1887) Gurley, 1893 (Syn. Nosema a. Moniez, 1887): formation of large xenomas in Gasterosteus aculeatus, Pungitius pungitius, Europe, North America; Arme (1972): G. aculeatus, Ireland; Can¬ ning et al. (1986): review; Canning et al. (1982): redescrip¬ tion; Dykova and Lorn (1978a): histopathology; G. L. Hoffman (unpub. res., USFWS, 1976): G. aculeatus, Que.; Kennedy (1974): same, England; Landry (1976): same, Que.; Lester (1974): same, B.C.; Lorn and Laird (1976): same, Nfdl.; Nepszy (1988): Culaea inconstans, P. pungi¬ tius, Lake Huron, Ont.; Voronin (1974): description of sporophorous vacuole, former Soviet Union. ■ G. cepedianae (Putz, Hoffman, and Dunbar, 1965) Canning, Lorn, and Dykova, 1986 (Syn. Pleistophora c. Putz et al., 1965): large xenomas present in viscera, Dorosoma cepedianum, OH, probably causing mortality; Dechtiar (1972a): D. cepedianum Lake Erie, Ont.; Dykova and Lorn (1980): placed in Glugea; Lopinot (pers. comm., State Fisheries, 1968): Lake Decatur, IL; R. L. Price (1982, 1983): D. cepedianum, including ultrastructure, southern IL; W. Rogers (pers. comm., Auburn Univ., 1972): P. cepedianae (G. cepedianae) common in southeast USA. ■ G. gasterostei Voronin, 1974: connective tissue, G. aculeatus, former Soviet Union; Canning et al. (1986): possibly syn. of G. anomala.

Order Pleistophoridida

■ G. hertwigi Weissenberg, 1911: large xenomas in intes¬ tine and other organs, Osmerus eperlamis, nearly 100% prevalence, northern Europe; Bogdanova (1957): O. eperlanus, former Soviet Union; Chen and Power (1972): loss of fecundity in infected female smelt, but in males, infection mostly in intestinal wall, Ont.; Dechtiar (1965b, 1972a): in smelt, Lake Erie; Delisle (1972): many infected smelt, xenomas mostly in intestinal wall but ubiquitous, Que.; Fantham et al. (1941): several infected smelt col¬ lected, many xenomas, including description, Que.; Haley (1954a, 1954b): many organs, Osmerus eperlanus, O. mordax, marine and freshwater, NH; G. L. Hoffman (unpub. res., USFWS, 1982): tentative identification, in muscle of Perea flavescens, Lake Erie, Ont.; Legault and Delisle (1967): acute disease in "0" age rainbow smelt, Que.; Locke (pers. comm., State Fisheries, 1965): in smelt in freshwater lakes, ME; Nepszy (1988): Lake Ontario, Canada; Nepszy and Dechtiar (1972): mortal¬ ity of adult smelt, Ont.; Nepszy et al. (1978): mortality of "0" age smelt, Ont.; Niederkom et al. (1980): immunofluorescent cross reaction with other genera; ScarboroughBull and Weidner (1985): discharge of sporoplasm through polar filament, ultrastructure; Sherburne and Bean (1979): incidence of G. hertwigi and piscine ery¬ throcyte necrosis, MA to Canada; Weiser (1949): O. eper¬ lanus, Czech Republic and Slovakia; Wellings et al. (1969): histopathology in English sole, WA. ■ G. pimephales (Fantham, Porter, and Richardson, 1949) Morrison, Hoffman, and Sprague, 1985 (Syn. Nosema p. Fantham et al., 1941) (Color Figs. 13, 14): large xenomas in body cavity, Pimephales promelas, Que.; Dechtiar (1972a): in viscera, muscle, Pimephales notatus, muscle, P. promelas, Lake of the Woods, Ont.; G. L. Hoff¬ man (unpub. res., USFWS, 1965, 1966): P. notatus, WI; Nepszy (1988): P. notatus, P. promelas, Lake Ontario, Ont.; Sprague (1977): transfer to Microsporidium; R. Walker (pers. comm., Rensselaer Inst., 1978): P. prome¬ las, NY. Author: this species was transferred from Nosema to Glugea because the xenoma has the characteristics of Glugea. ■ G. takedae Awakura, 1974: trunk muscle, Oncorhynchus masore, O. mykiss, Japan; Miki and Awakura (1977): experimental infection and ultrastructure, Japan; Takahashi (1978): water temperature greatly influenced rate of schizogony and sporogony, O. mykiss apparently acquired a strong immunity to infection; Vialova and Voronin (1987): O. gorbuscha infected 70-100%, O. mason 9-46% in Sakhalin Rivers, Russia. ■ G. truttae Loubes, Maurand and Walzer, 1981: in yolk sac of three-week-old Salmo trutta, trout farm, Switzer¬ land, no xenoma formed; development, ultrastructure. ■ G. weisetibergi Sprague and Vernick, 1968 (possible syn. of G. anomala; cf. Canning and Lorn, 1986): large subperitoneal and subcutaneous xenomas, Apeltes quadracus, Patuxent River, Solomons, MD, description including ultrastructure; Erickson et al. (1968): EM of everted polar

87

filament; Vernick, Sprague, and Lloyd (1969): ultra¬ structure; Vernick, Tousimis, and Sprague (1969): surface ultrastructure. ■ Glugea sp. Bond (1938a): small xenomas in mucosa of stomach and bile duct, Fundulus heteroclihis, Chesa¬ peake Bay, MD. ■ Glugea sp. Chacko (1982): xenomas in feral Ptychocheilus oregonensis, Salvelinus alpinus, ID. ■ Glugea sp. Crandall and Bowser (1981): large xenomas in viscera and subdermal connective tissue, of Gambusia affinis, southern CA. ■ Glugea sp. Molnar, Hanek, and Fernando (1974): in intestine, gall bladder, Notropis cornutus, Pimephales promelas, Ont. ■ Glugea sp. Voronin (1974): small xenomas in skin, Lota lota, near St. Petersburg, Russia.

Genus Heterosporis Schubert, 1969 (Fig. 122)

Similar to Pleistophora, but forming a xenoma. — Ref. Canning et al. (1986): review; Schubert (1969a, b): description.

■ Heterosporis finki Schubert, 1969a: in connective tis¬ sue around esophagus, Pterophyllum scalare, freshwater pet fishes, Germany; ultrastructure; Michel et al. (1990): large muscle lesions, P. scalare, histopathology, ultra¬ structure, France; Schubert (1969b): ultrastructure. ■ H. schuberti Lorn, Dykova, Korting, and Klinger, 1989: in myocytes, freshwater pet fishes Ancistrus cirrhosus, Pseudocrenilabrus multicolor, macro- and microspores, latter with large posterior vacuole, Europe.

Genus Loma Morrison and Sprague, 1981 (Figs. 123, 1 33, 138c)

Nuclei unpaired throughout life cycle; uninucleate meronts with simple plasma membrane lying in direct contact with host cell cytoplasm and developing into multinucleate plasmodia. Sporogony polysporoblastic within a sporophorous vesicle (SPOV); sporoblasts num¬ bering up to 8 (occasionally up to 16) transforming into spores within the vesicle. Infected cells hyper¬ trophic, forming xenomas up to about one mm in diameter, with single, centrally located, hypertrophic nucleus. Xenomas all a simple host cell membrane coated with fibrillar layer. Developmental stages of par¬ asite intermingled with spores throughout the xenomas. Fish are hosts. — Ref. Canning et al. (1986): review; Morrison and Sprague (1981a, b, c): generic description, ultrastructure.

88

Subkingdom Protozoa

■ Loma fontinalis Morrison and Sprague, 1981a: gill lamellae, Salvelinus fontinalis, xenomas up to 0.5 mm, N. S.; Morrison and Sprague (1983): same, comparison with other species. ■ L. salmonae (Putz, Hoffman, and Dunbar, 1965) Mor¬ rison and Sprague, 1981a (Syn. Pleistophora s. Putz, Hoff¬ man, and Dunbar, 1965): Pleistophora sp. Wales and Wolf, 1955: gill lamellae, Cottas sp. Oncorhynchus mykiss, O. nerka; Awakura (1974): gills, Oncorhynchus mason, Hokkaido, Japan; Behkti (1984): variation in sporogony, France; Dykova and Lorn (1980): histopathology; Hanek and Molnar (1974): Salvelinus fontinalis, Que.; G. L. Hoffman (unpub., USFWS, 1965, 1968): S. fontinalis, PA, Oncorhynchus mykiss, AZ; Horner (pers. comm., State Fisheries, 1986, 1987): Oncorhynchus kisutch, O. mykiss, IL; Janeke (pers, comm., USFWS, 1972): O. mykiss, AZ; Poynton (pers. comm., Univ. of Southampton, 1986): O. mykiss, Salmo trutta, fish farms, England; J. W. Wood (1979): possibly L. salmonae in Cottus sp., O. kisutch, O. mykiss, O. nerka, WA. ■ Loma sp. Behkti (1984): gills, Tilapia melanopleura, Benin, Nigeria. ■ Loma sp. Hauck 1984: arteries, gills, kidney, other ves¬ sels, Oncorhynchus tshawytscha, AK. ■ Loma sp. Kent, Elston, and Harrell (1987): gills, O. kisutch, marine cage, very similar to L. salmonis, WA.

Genus Pleistophora Gurley, 1893

(Figs. 126-128, 134) Life cycle and development: nuclei are isolated through¬ out development. Meronts are rounded plasmodia with a thick amorphous wall external to the plasma mem¬ brane. At the onset of sporogony, the wall separates from the surface and becomes the thick wall of the sporophorous vesicle (SPOV). Sporogony is polysporoblastic: the sporogonial plasmodium divides by repeated segmentation, producing a variable and large number of spores within the SPOVs, which often persist in wet mounts. Spores are uninucleate, and some species produce macrospores. Infections are diffused in tissues without xenoma formation. Hosts are usually fishes or arthropods, but at least one species (P. myotrophica) occurs in amphibia. ■ Pleistophora anguillarum Hoshina, 1951: skeletal muscle, small whitish spots beneath skin, sometimes in many organs, may cause mushy muscle, Anguilla japonica, Japan; Canning, Lorn, and Dykova (1986, p. 95): review, descriptions, therapy; Egusa et al. (1974): review; Sprague (pers. comm., Univ. of Maryland, 1980): eel farmers in Formosa being ruined by P. anguillarum. P. dallii Zhukov, 1964. Canning et al. (1986, p. 97): subcuta¬ neous tissue near base of pectoral and caudal fins, Dallia

pectoralis, Chukotka, Russia, might be Glugea sp.; Shulman (1962): description. ■ P. hyphessobryconis Schaperclaus, 1941. Canning et al. (1986, p. 101): a very serious parasite, usually in skele¬ tal muscle but found anywhere in heavy infections, from 17 pet fish species, including Carassius auratus; invaded muscles appear grayish-white to white, some¬ times with emaciation, scoliosis, kyphosis, and raised scales; Dykova and Lorn (1980): histopathology; G. L. Hoffman (unpub., 1980, 1982): in Metynis sp. Brazil, Capoeta tertrazona, FL; Lorn and Corliss (1967): ultra¬ structure, Hyphessobrycon sp. ■ P. laclogensis Voronin, 1978. Canning et al. (1986, p. 107): muscle of Lota lota, Osmerus eperlanus, near St. Petersburg, Russia. ■ P. myotrophica Canning, Elkan,and Trigg, 1964: caused mortality of Bufo and Xetiopus (amphibia), experimen¬ tal infection and life cycle. Included here to avoid con¬ fusion with fish infections. ■ P. oolytica Weiser, 1949. Canning et al. (1986, p. 15): in oocyte, Esox Indus, Hucho hucho, Rutilus rutilus, Czech Republic and Slovakia, Austria. ■ P. ovariae Summerfelt, 1964. Canning et al. (1986, p. 118): oocytes, Notemigonus crysoleucas, 13 midwest and southern states and CA; Modin (pers. comm., State Fish¬ eries, 1978): second record in Pimephales promelas, CA; Nagel and Hoffman (1977): oocytes, P. promelas, AR; Summerfelt and Warner (1970): review. Author: the off¬ color (opaque-white) infected eggs are easily detected, and the intensity of infection may be severe in cultured golden shiners. Since the intensity is usually less severe in younger brooders, however, these may be used suc¬ cessfully for stocking new ponds. There is no evidence of stock depletion among feral shiners. ■ P. tahoensis Summerfelt and Ebert, 1969: body wall, Cottus beldingii, Lake Tahoe, CA. ■ Pleistophora sp. Arai and Mudry (1983): pyloric cae¬ cum, intestine, Oncorhynchus mykiss, Richardsonius balteatus, B.C. ■ Pleistophora sp. Bond (1937b): muscle, outer layer of spinal cord, Fundulus heteroclitus, MD. ■ Pleistophora sp. Heckmann, Kimball, and Short (1987): muscle, Cottus bairdi, UT. ■ Pleistophora sp. Herman and Putz (1970): liver, Ictalurus punctatus, WV. ■ Pleistophora sp. G. L. Hoffman (unpub., USFWS, 1988): muscle, Perea flavescens, large spores (9 pm), Lake Ashtab¬ ula, ND. ■ Pleistophora sp. Hoshina (1951): epicardium, Oncorhyn¬ chus mykiss, Japan. ■ Pleistophora sp. Wellborn (1966) (in Putz and McLaughlin, 1970): Dorosoma petense, USA (may be Glugea cepedianae).

Sporozoa of Uncertain Classification

Genus Thelohania Henneguy, 1892 (No fig.) Nuclei diplokaryotic or isolated, according to stage of development. Merogony inadequately known. Meronts sometimes diplokaryotic; early sporont diplokaryotic, giv¬ ing rise to eight uninucleate sporoplasts in sporophorous vacuole. Spores uninucleate. Hosts usually arthropods, but two fish hosts known. ■ Thelohania baueri Voronin, 1974: Finland, near St. Petersburg, Russia; Canning et al. (1986): review. ■ T. ovicola (Auerbach, 1910) Kudo, 1924: oocytes, Coregonus exiguus, Switzerland.

Collective Group (Genus?)

Microsporidium Balbiani, 1884 (Figs. 124, 125, 136) Catchall "genus" for microsporidians that cannot be more properly classified at present. ■ Microsporidium pseudotumefaciens (Pflugfelder, 1952) Canning et al. 1986 (Syn. Glugea p. Pflugfelder, 1952): in ovarian follicles and other viscera, also subcutaneous, Stuttgart, Germany. ■ M. rhabdophilia Modin, 1981: in rodlet cells [Rhabdospora thelohani (?)] of skin, gills, intestine, Onto rhynchus kisutch, O. mykiss (steelhead and regular), O. tshawytscha, clusters of 16 spores measuring 2.6-3.5 x 0.8-1.2 pm, freshwater, CA. Note: this species may belong in Enterocytozoon. ■ Microsporidium sp. Buttz and Holloway (1987): Pungitius pungitius, ND. ■ Microsporidium sp. de Kinkelin (1980): spinal cord, zebra danio, Brachydanio rerio, pet fish, Germany. ■ Microsporidium sp. Elston, Kent, and Harrell (1987): intranuclear in hemoblasts, Oncorhyjichus tshawytscha, associated with anemia, vegetative stages only, WA. ■ Microsporidium sp. Fischthal (1947a, 1950c): dorsal muscle, Notropis comutus, WI. ■ Microsporidium sp. Herman and Putz (1970): heart muscle, intestinal submucosa, Ictalurus punctatus, MD.

89

in trophozoites of Myxidium sp., in gall bladder of Notro¬ pis atherinoides, Troy, NY.

Sporozoa of Uncertain Classification Genus Dermocystidium Perez, 1907 (Figs. 139-142, Color Fig. 17)

This parasite has been considered a fungus by some and a haplosporidean protozoan by others. To the author's knowledge, no one has successfully classified those Dermocystidium species that parasitize lower ver¬ tebrates. The vegetative stage, as shown by Elkan (1962), resembles that of myxosporidean protozoa; the spores, however, have no characteristics of Myxosporidea or Haplosporidea, and further classification remains uncertain. Dermocystidium marinum, which parasitizes oysters and has been studied extensively (see Mackin, 1961), does possess some fungal characteristics—notably the spore wall, which reacts with iodide, and tubular outgrowths of the spore. However, only protoplasmic masses and plasmodia have been seen in fishes. Immature stages. Cysts with hyaline walls (probably of parasitic origin) of varying thickness, filled with amorphous protoplasmic mass in which chromatin granules are more or less evenly distributed. Granules coa¬ lesce to form nuclei, splitting up protoplasmic mass to form plasmodia and spherical portions 8 to 10 pm in diameter, which produce spores 3 to 12 pm in diameter. Spore stage. Spores enclosed in very rigid elongate, oval, or round cyst; hyaline cyst wall probably of para¬ sitic origin. Spores round, 3-12 pm in diameter, con¬ taining nucleus and large, conspicuous peripheral vacuole. Parasites of fishes. The Dermocystidium species asso¬ ciated with oysters may be unrelated but does produce identical spores. — Ref. O. N. Bauer (1981): review of those in cultured fishes, former Soviet Union; Cervinka et al. (1974): review and a new species in carp; Reichenbach-Klinke (1982): review of species in tropical pet fishes; Scheer (1957): key to species, Germany.

■ Microsporidium sp. G. L. Hoffman (unpub., USFWS, 1980): blackspot cysts in skin, Metynnis argenteus, (silver dollar) aquarium fish, spores 3.5 x 2.1 pm, Brazil.

■ Dermocystidium anguillae Spangenberg, 1975: cysts in gills, Anguilla anguilla, cysts about 1.0 x 3.8 mm, spores 4.8 x 8.5 pm, Germany; Arlati and Bresolin (1986): in elvers, description of pathology, Italy; Hatai et al. (1979): A. anguilla, Japan.

■ Microsporidium sp. Plehn (1924): small "dots" in ovary, Sahno salar, S. trutta. Norway.

■ D. branchialis Leger, 1914: small round cysts, 0.2-0.6 mm, in gills, Sahno trutta, Europe.

■ Microsporidium sp. Porte and Chilmonczyk (1974): skin lesions, Oncorhynchus mykiss, Sahno salar, S. trutta, resembles UDN and is transovarial, France.

■ D. cuticulare Scheer, 1956: elongated 3 mm cysts in skin, Gasterosteus aculeatus, Germany.

■ Microsporidium sp. Walker (pers. comm., Troy, NY, 1987): description incomplete, spores about 8x2 pm,

■ D. cyprini Cervinka, Vitovek, Lorn, Hoska, and Kubu, 1974: oval cysts 1-2 mm in gills, Cyprinus carpio, patho¬ genic, Czech Republic and Slovakia.

90

Subkingdom Protozoa

-Dermocystidium spp.FIG. 139. Dermocystidium sp., cysts in skin of young Osmerus mordax, Ont. (courtesy of the late A. Dechtiarenko, Maple, Ont.). FIG. 140. Dermocystidium sp., elongated cysts in skin of young Lepomis macrochirus (courtesy of D. DeMont, Raleigh, NC). FIG. 141. Dermocystidium sp., spores from skin cysts on Salmo trutta, MD. FIG. 142. Dermocystidium sp.: a, cyst; b, spore.

■ D. erschovi Garvaki, Denisow, and Afanasjew, 1980:

■ D. kobicevi Allamuratov, 1965: oval cysts 0.2 mm

cysts up to 16 mm long, Cyprinus carpio, former Soviet Union, may be D. koi.

long, in skin, Cyprinus carpio, former Soviet Union.

■ D. gasterostei Elkan, 1962: cysts 2-3 mm long in skin, Gasterosteus aculeatus, England; Kennedy (1974), same. ■ D. kamalovi Allamuratov, 1965: oval cysts 0.6-1.3

mm long, Cyprinus carpio, former Soviet Union.

■ D. koi Hoshina and Sahara, 1950: elongated cysts,

0.04-0.3 mm in skin and muscle, Cyprinus carpio, Japan; G. L. Hoffman (unpub., USFWS, 1954): mas¬ sive lesions, Korea; G. L. Hoffman (unpub., 1980): in koi carp, USA.

Sporozoa of Uncertain Classification

■ D. macrophagi Moer, Manier, Bouix, et al., 1987: intra¬ cellular, Oncorhynchus mykiss, associated with PKD, France. ■ D. pusula Perez, 1913: spherical cysts, 1 mm, in gills, Salmo trutta, Ireland. ■ D. ranae Guyenot and Naville, 1922: elongated cysts 0.9-2 mm long, in frogs, Switzerland; Lorn and Vavra (1963a): spore has a mucus envelope. Included here in case of contamination. ■ D. salmonis H. S. Davis, 1947: spherical cysts, 1 mm, in gills, Oncorhynchus tshawytscha, CA; R. L. Allen and Meekin (1968): histopathology; Barney (pers. comm., USFWS, 1961): WA; Hauck (pers. comm., State Fish¬ eries, 1980): probably D. salmonis, in gills, arteries, cap¬ illaries, kidney tubules, O. tshawytscha smolts in water reuse system, AK; Leitritz (1988): CA; R. E. Olson et al. (1991): waterborne transmission, description of flagel¬ lated stage, OR; Pauley (1967): in gills, histopathology, O. tshawytscha, WA; J. W. Wood (1979): WA. ■ D. vejdovskyi Jirovec, 1939: small oval cysts, 0.1-1.2 mm, in gills, Esox Indus, Europe; Reichenbach-Klinke (1954a): same. ■ Dermocystidium sp. Arai and Mudry (1983): in gills, Coregonus clupeaformis, Lota lota, Prosopium williamsoni, Thymallus arcticus, B.C. ■ Dermocystidium sp. Bell and Margolis (1976): mas¬ sive mortality of prespawning, feral Oncorhytichus nerka, Vancouver Island, B.C. Author: this may have been D. salmonis. ■ Dermocystidium sp. Dechtiar (pers. comm., Ontario Fisheries, 1973, 1978): massive infection and mortality, young Esox lucius, Osmerus mordax, elongated cysts up

91

to 1.5 mm long, spores 5-7 pm and 7-9 pm, Lake Erie, Ont. ■ Dermocystidium sp. Millen Aquarium, Millen Natl. Fish Hatchery, GA 1975 (USFWS, Fish Hatchery Biol. Rep. No. 2, 1975, p. 7): in goldfish fins. ■ Dermocystidium sp. Heckmann and Ching (1987): reported as Ichthyophonus, on skin of Oncorhynchus clarki, Yellowstone Lake, MT. ■ Dermocystidium sp. Hoskins (pers. comm., Pacific Biol. Sta., 1989): repeatedly caused high mortality of early run Oncorhynchus nerka B.C. ■ Dermocystidium sp. Kennedy (1974): Gasterosteus aculeatus, Perea fluviatilis, Salmo trutta, England. ■ Dermocystidium sp. McVicar and Wootten (1980): in visceral fat and skin, Salmo salar, organism cultured in test tubes, Scotland. ■ Dermocystidium sp. Walker (pers. comm., Rensselaer Inst., 1982): in newt, Troy, NY. Included here to avoid confusion with species in fishes. ■ Dennocystidium sp. Wootten and McVicar (1982): in cultured Anguilla anguilla gills, apparently not D. anguillae, Scotland. During 1958-1985, the author made cursory exam¬ inations of Dennocystidium spp. from Carassius auratus (NC), Lepomis macrochirus (NC, WV), Microptenis dolotnieui (WV), Salmo salar (VT), Salmo trutta (ME), and Salvelinus fontinalis (ME, NH). All of these were in elongated worm¬ like cysts of varying sizes. Most were located in the fins. The spores ranged from 6 to 9 pm. It is not known whether these represent one species with variations or several species. D. DeMont, of Raleigh, NC, plans to publish on this subject soon.

Phylum

Coelenterata (Cnidaria) (Figs. 143-147, Color Fig. 18)

---Life cycle of Polypodiuin hydriformeFIG. 143. Schematic of life cycle, roman numerals representing months: a = mature stolon with internal tentacles from egg before spawning, b = stolon with external tentacles emerging from egg at time of spawning, c = free stolon, d-i = stolon becoming polyp, which may divide; route back to fish unknown, j-k = early embryonic stages in egg, 1-n = juvenile stolons, o = stolon with internal tentacles (from G. L. Hoffman et al., 1974, J. Parasitol.). 92

Phylum Coelenterata

93

- Polypodinmn hydriforme FIGS. 144, 145: Polypodium hydriforme from sturgeon eggs, MI. FIG. 144. Enlarged gray eggs on the right containing mature stolons; normal black eggs on the left. FIG. 145. Infected eggs; egg at lower left is intact and egg at upper right has been torn to release fragments of the stolon. FIGS. 146, 147: P. hydriforme from Acipenser ruthenus, Russia. FIG. 146. Infected eggs appear larger, and lighter-colored. FIG. 147. Mature stolon. (Figs. 144-147 from G. L. Hoffman et al., 1974, J. Parasitol.).

Multicellular animals. Body consists of two layers of cells: ectoderm (external) and endoderm (lining internal cav¬ ity of body). The internal cavity communicates with the external environment through the oral aperture. A characteristic feature of coelenterates is the presence of stinging cells (nematocysts) in their tissues; these cells are used for attachment to substrate, capture of prey, and protection against enemies. Most species are radi¬ ally symmetrical, free-living, sometimes colonial, usu¬ ally marine. Parasitism is extremely rare. Acipenserids (sturgeons) in North America and the former Soviet Union may be infected by one species, Polypodium hydri¬ forme. In the United States, paddlefish, Polyodon spafhula, may also be infected. This aberrant species of coelenterate does not seem to be closely related to any other form. Members of the genus Hydra may attack fish larvae. ■ Polypodium hydriforme Ussov, 1895 (Color Fig. 18). With only one species, the generic description is the same as the species. Raikova (1984): general account; Bykhovskaya-Pavlovskaya (1962): general account; Gusev (1985): in eggs, Acipenser guldenstaedti, A. nudiventris, A. ruthenus, A. stellatus, Huso dauricus, H. huso, former Soviet Union; G. L. Hoffman, et al. (1974): A. fidvescens,

MI; Mokhayer (1976): A. stellatus, Iran; Raikova (1988): embryonic stages; Raikova et al. (1979): histological evi¬ dence that the U.S. material is P. hydriforme in Polyodon spathula, MO; Suppes and Meyer (1975): first report from P. spathula, MO. Until 1974, this very unusual parasite had been found only in large rivers throughout the former Soviet Union, including the Amur River in the extreme east. State Fisheries employees of Michigan found it in A. fidvescens in 1974, and Missouri Fisheries employees found it in feral Polyodon spathula being used for spawning pur¬ poses. In 1976 it was reported from A. stellatus in Iran. Since then it has been found in California (Kent, pers. comm., Univ. of California, Davis, 1984), New Brunswick, Canada (Appy, pers. comm., 1979) and North Dakota (Holloway, pers. comm., Univ. of North Dakota, 1989). Thus it appears that P. hydriforme has a wide distribution in the northern hemisphere. The very early stages of infection in sturgeon eggs were not known until 1988, when Raikova reported on the unicellular and early cleavage stages of the parasite. The planula stage can be recognized in immature fish eggs,

94

Phylum Coelenterata

but detection usually occurs when the larger stolon stage with buds and tentacles is apparent and convoluted in the enlarged egg. The egg assumes the form of a large, striped sphere with white, convoluting lines against a dark background of yolk. Throughout its devel¬ opment in the egg, the Polypodium is characterized by the inversion of its germinal layers—the endoderm lying externally, in contact with the yolk, and the ectoderm lying in the cavity of the stolon. Immediately before spawning, the parasite turns inside out, leaving the ectoderm now on the outside. The parasite is ejected with the eggs, becomes free-living, and soon becomes a polyp with 12 tentacles, 6 on each side. The mouth of the polyp is directed superiorly. The organisms at this stage mul¬ tiply by longitudinal binary fission. Male and female gen¬ ital glands arise in midsummer. The mode of reinfection of fish eggs with the unicellular stage is not known.

Hydra Linneaus

Recently hatched larvae of fishes are killed by the nematocysts of Hydra if the coelenterates are present in large numbers. Reports include Beardsley (1904): destruction of trout fry held in anterior portions of hatchery troughs; Clady and Ulrikson (1968): mortal¬ ity of bluegill fry; Cordero (1941): damage to the mouth region of a teleost; Eisler and Simon (1961): mortality of recently hatched larvae of Pacific salmon; Gudger (1927, 1934): mortality to small fishes; Lerch (1980): control methods; Shuberg (1905): nematocysts found in the epithelium of recently dead trout; E. Williams (pers. comm., Univ. of Puerto Rico, 1994): carried on dried Artemia eggs, killing freshwater shrimp in hatch¬ eries worldwide.

Phylum

Platyhelminthes Class

Monogenea Carus, 1 863

Methods.96 Maintaining Gyrodactylus in the Laboratory .97 ■ Key to the Suborders of Monogenea .97 Subclass Monopisthocotylea . . .97 Family Gyrodactylidae.97 ■ Key to the Families of Monopisthocotylea .98 Genus Fundulotrema .98 ■ Key to the Genera of Gyrodactylidae.98 Genus Cyrodactyloides ... .100 Genus Gyrodactylus.100 Genus Laminiscus.104 Genus Polyclithrum .104 Genus Swingleus .104 Order Dactylogyrida .105 ■ Key to the Families of Dactylogyrida .105 Family Ancyrocephalidae ...105 Genus Actinocleidus.105 ■ Key to the Genera of Ancyrocephalidae.106 Genus Aethycteron.112 Genus Anchoradiscoides . . .112 Genus Anchoradiscus.112 Genus Cichlidogyrus.113 Genus Clavunculus.113 Genus Cleidodiscoides.113 Genus Cleidodiscus.113

Genus Genus Genus Genus Genus Genus Genus Genus Genus Genus Genus Genus Genus Genus

Enterogyrus.115 Haplocleidus .115 Leptodeidus.116 Ligictaluridus.116 Lyrodiscus.117 Macrohaptor.117 Onchocleidus.117 Pterocleidus.118 Sahuginis .119 Syndeithrum.119 Tetradeidus.120 Trianchoratus.120 Urodeidoides.120 Urodeidus.120

Addendum to Ancyrocephalidae . .121 Family Capsalidae.121 Genus Nitzschia.122 Family Dactylogyridae.122 ■ Key to the Genera of Dactylogyridae.122 Genus Acolpenteron.122 Genus Aplodiscus.124 Genus Dadylogyrus.124 Genus Pelluddhaptor.128 Genus Pseudacolpenteron . .129 Genus Pseudodadylogyrus .129 Family Diplectanidae.129 Genus Dipledanum .129 Family Pseudomurraytrematidae.130 ■ Key to the Genera of Pseudomurraytrematidae ... .130

Monogenea are small (microscopic) to medium-sized (5 mm) flatworms that complete their life cycle on one host. The immature worms are usually morphologically similar to the mature forms. The chief organ of attach¬ ment and best identification aid is the haptor, which is posterior (lateral in one freshwater species). There are 12 nearly always to 16 hooks (marginal hooks), and in most freshwater genera 2 to 4 larger anchors (hamuli)

Genus Anonchohaptor ... .1 30 Genus Icelanonchohaptor . .130 Genus Myzotrema.130 Genus Pseudomurraytrema .132 Family Tetraonchidae.132 Genus Tetraonchus.132 Suborder Polyopisthocotylea .133 ■ Key to the Genera of Polyopisthocotylea ... .133 Superfamily Diplozooidea ... .1 36 Family Diplozooidae.136 Genus Diplozoon.136 Superfamily Diclybothrioidea . 136 Family Diclybothriidae.136 Genus Didybothrium .136 Genus Paradidybothrium . .136 Superfamily Mazocraeoidea . . .136 Family Discocotylidae.136 Genus Discocotyle .136 Genus Odomacrum.137 Genus Neodiscocotyle.137 Family Heteroaxinidae.137 Genus Lintaxine.137 Family Mazocraeoidae.137 Genus Mazocraeoides.137 Genus Pseudanthocotyloides 1 38 Genus Pseudomazocraeoides 138 Family Microcotylidae .138 Genus Microcotyle.138 Genus Pauciconfibula.138

are centrally located in the haptor. In the Polyopistho: cotylida, however, the larval hooklets disappear and 6 to many cuticular adhesive units (clamps or suckers or both) develop. In some groups the copulatory organ or cirrus (penis) and its accessory piece, which are both com¬ posed of very resistant, hardened protein, are useful in species identification; they have varying shapes and sizes. Anterior adhesive organs (head organs), suckers, or 95

96

Phylum Platyhelminthes: Class Monogenea

pseudosuckers may be present. The eggs are compara¬ tively few and large, often with polar prolongations. Vitellaria are nearly always minutely follicular, coextensive with intestinal caeca. Some species live on the gills only, some on the body and fins only, and some on both. Some species are found in nasal fossae and lateral line pits. Enterogyrus is parasitic in the anterior gut of African Tilapia and a marine butterfly fish. One genus is found in the urinary system. Most are capable of moving around on the host, but some Dactylogyrus species apparently remain in one spot on the gills. Monogenea feed on mucus, epithelium, and sometimes blood. The feeding organ (pharynx) of at least one species emits a proteolytic substance that erodes the epidermis (Kearn, 1963). Some Monogenea can be serious pests in fish culture. Certain species of Gyrodactylus may become so numer¬ ous on cultured trout, goldfish, and bluegills that the fishes suffer great distress and must be treated to survive. Some Dactylogyrus species cause great damage to the gill filaments of carp and goldfish in hatcheries. Other Monogenea probably are potential threats to fish culture but have not been adequately studied. In fish populations that have become crowded in nature, similar results have been observed. — Ref. The most comprehensive references are BeverleyBurton (1984): key to Canadian species; BykhovskayaPavlovskaya (1962); Bykhovski (1957); Ergens and Lorn (1970); Gusev (1978, 1985); Hargis and Thoney (1983); Lebedev (1988); Paperna (1979); C. E. Price and McMahon (1967); Roman-Chiriac (1960); Skarlato (1977); Sproston (1946): synopsis; and Yamaguti (1963a): synopsis. Other pertinent references are O. N. Bauer et al. (1977): pathogenicity; Berry and Mizelle (1956): com¬ position of hooks; Chubb (1977): review of seasonal¬ ity; Cone (1979b): description of onchomiracidium of Urocleidus adspectus; Cone and Burt (1982b): experi¬ mental host specificity methods; Cone and Odense (1984): pathogenicity; Dranichak (1973): demonstra¬ tion of copulatory canal; Gelnar (1987b): effect of tem¬ perature on Gyrodactylus; Gusev (1970, 1985): variability due to age of host; Gusev (1973/1974): treatise on Monogenea of India; Gusev (1978): classification; Kearn (1966, 1968): parasite attachment; Kearn (1980, 1986): eggs and onchomiracidia; Kearn (1987): Method of locomotion; Koratha (1960): Monogenea on amphibia, review; Kritsky et al. (1986): neotropical Monogenea; Kuperman and Shulman (1978): effect of environment on population; Lambert (1979): onchomiracidia and phylogeny; Lamothe-Argumedo and Jaimes Cruz (1982): list of Mexican species; Lester and Adams (1974a): determination of partial immunity; Lyons (1966): chemical nature of hard parts; Musselius (1977): path¬ ogenicity; 1. L. Owen (1970): description of larvae; Paperna (1979): Monogenea of freshwater fishes of Africa; Pomales and Williams (1980): effect of tem¬ perature; C. E. Price and Arai (1967): anatomical terms; M. E. Scott and Robinson (1984): immunity; Vladimirov (1971): immunity.

Methods

It is preferable that fishes be examined immediately after killing, which can be accomplished by severing the spinal cord just behind the head with a bone cutter, by pithing, or by narcotizing; cutting with a sharp knife causes excessive bleeding. Anesthetics such as Sandoz MS 222 (ethyl m-aminobenzoate) can be used to immo¬ bilize the fishes that must be kept alive. Small pieces of infected gills may be removed with little harm to the fishes. Preserved fishes may be used but are far less sat¬ isfactory than freshly killed ones. If the fish is small, it should be immersed in chlorinefree water and examined at lOx with a dissection micro¬ scope. Monogenea are usually very active and thus can more easily be detected. The outer body, the fins, the inside of the mouth (including the olfactory and lateral line pits), and particularly the gills should be observed. After examining the intact fish, the opercula should be carefully cut off to expose the gills, which can then be excised as needed for thorough examination. Acolpenteron species should be sought in the urinary system. If the fish is too large to examine as described, excised gills and fins, mucus scrapings, and the inside of the mouth should be examined individually. Flukes may be removed from the fish mechanically, but the attendant mucus often makes this difficult. Gyrodactylus may be removed dead, but fresh enough for making permanent preparations, by immersing the host in formalin (1:4000) for 15 to 45 minutes. The flukes leave the fish and seldom become entangled in mucus. Other flukes may be removed likewise but do not become free from the gills as readily. Flukes so removed may be studied fresh or fixed for permanent preparations. Freez¬ ing the gills for 6 to 24 hours will also free flukes from the gills (Mizelle, 1936). Chloretone (Hargis, 1953b) and Nembutal (Ikezaki and Hoffman, 1957) have also been used. Monogenea are best studied when fresh, or better yet, alive. Such material may be studied under a cover slip with enough water to prevent flattening. Later the fluid may be removed with pieces of paper towel or filter paper to flatten the specimen and best observe the haptoral armature and copulatory complex. Permanent preparations may be made after fixing wonns removed by the foregoing methods, providing the worms are fixed promptly after they are immobilized. Freezing sometimes destroys the architecture of the soft parts, but the hard parts, which are usually the most important, will be satisfactory. Mizelle (1936) describes a method for affixing Monogenea to a slide: The specimens were transferred to slides coated with Mayer's albumin. They were then placed in a finger bowl on inverted Stender dishes. Alcohol was poured in the bottom of the container to a level below the top of the Stender dishes, the top of the bowl was covered and the apparatus set in

Family Gyrodactylidae

an incubator. It was found that exposure for one to five minutes at 55°C was sufficient to coagulate the albumin and securely attach the specimens to the slide. After attachment of the specimens to the slides they were fixed in Gilson's fluid and treated with iodine for removal of mercuric salts. Hematoxylin and carmine stains are usually employed for staining Monogenea. Unstained semipermanent mounts may be made by mounting in a very small amount of glycerine jelly, polyvinyl alcohol, Turtox CMC, or CMC-S (Turtox Bio¬ logical Supply House) and sealing with lacquer or per¬ manent mounting medium. The "two-cover-glass" method may also be used: Place the specimen in a small amount of water on a 22-mm square cover glass; add a small amount of mounting agent (as above); place a smaller cover glass on the specimen; place four drops of mounting medium on a slide; invert the cover glass preparation and place it on the mounting medium, which will seal the mount. Glycerine jelly may be used for fixed or preserved specimens (Mizelle and Seamster, 1939). The usefulness of such mounts will depend on how well the anchors are separated in the flattening process. — Ref. Berry (1966): sectioning; Glaser (1974): osmic acid fixative, clear in glycerine; Gusev (1983): methods book¬ let; G. L. Hoffman and Moore (unpub., 1982): acetoorcein in wet mounts stained the Gyrodactylus shield; Kayton, Kritsky, and Tobias (1982): histochemistry of hard parts; Kritsky, Leiby, and Kayton (1978): rapid stain for haptoral bars; Lester and Adams (1974a): neu¬ tral red caecal stain; Lynn et al. (1981): silver stain for chaetotaxy of onchomiracidia; Malmberg (1956a): ammonium picrate/glycerine fixative mountant, and excretory system; Mizelle and Klucka (1953): measuring methods; Parker and Haley (1960): method for count¬ ing Gyrodactylus.

Maintaining Gyrodactylus in the Laboratory Lifecycle. Gyrodactylus bullatarrudis Turnbull, 1956, of the guppy, completes its life cycle from larva to larvaproducing adult in about 60 hours at 25-27°C (Turnbull, 1956). It probably takes two weeks for a population of one Gyrodactylus to build up to 100. Maintaining on trout. Usually small fingerlings (2-5 cm) are most heavily infected. As fish become older, the population of Gyrodactylus gradually dimin¬ ishes. Yearlings (23-30 cm) usually are still infected, but with a very small number of parasites. Minimal water supply and crowding are usually con¬ ducive to increasing the Gyrodactylus population. Gyrodactylus can be transferred to uninfected fishes by placing unifected fishes in the same tank with infected fishes. They can also be transferred by skin scrapings (G. L. Hoffman and Putz, 1964). Maintaining on bluegills. Gyrodactylus macrochiri (Hoffman and Putz, 1964) can be maintained at 12°C as described above, they disappear in aquaria at room temperature.

97

Maintaining other species of Gyrodactylus. Goldfish Gyrodactylus populations follow the same pattern as above but can be maintained at room temperature. Turnbull (1956a) maintained the guppy Gyrodactylus at room temperature with no difficulties, but Malmberg (1956) was unable to maintain Gyrodactylus spp. To ensure large populations of Gyrodactylus, it is best to have young or small uninfested fishes available to infest new lots from time to time, possibly monthly, while maintaining the proper temperature.

Key to the Suborders of Monogenea Haptor (posterior attachment organ) as a single unit; larval haptor retained, more or less unchanged, in adult; anterior end with gland organs; mouth not surrounded by oral sucker; paired suckers within mouth absent; genitointestinal canal absent; haptor usually with 1 or 2 pairs of anchors (hamuli) and 12 to 16 hooks .(p. 97) Monopisthocotylea Functional haptor of adult developed anterior to larval haptor with lateral rows of suckers or clamps always supported by cuticular sclerites, usually acting as clamps; larval haptor vestigial; anterior end without adhesive glands.(p. 133) Polyopisthocotylea

Subclass Monopisthocotylea Odhner, 1912 Haptor a single unit derived from larval haptor; arma¬ ture usually consisting of large anchors (hamuli) and smaller hooks, sometimes marginal. Anterior end fre¬ quently with adhesive organs in form of "head organs"; genitointestinal canal absent. Most species highly hostspecific. — Ref. See Monogenea references.

See the Key to the Families of Monopisthocotylea on p. 98.

Family Gyrodactylidae Cobbold, 1864 Small, elongated Monogenea; anterior end bilobed; each lobe with head organ. Haptor well-developed, bearing one pair of large anchors (hamuli) with dorsal and ventral supporting bars and 16 marginal hooks. Intestine bifurcate, the two limbs not uniting posteri¬ orly. Eye spots absent. Cirrus armed with row of minute spines and usually with trianglular cuticular plaque. Ovary V-shaped or lobed, posterior or ventral to testis; vitellaria absent or united with ovary; vagina absent. Viviparous.

98

Phylum Platyhelminthes: Class Monogenea

Key to the Families of Monopisthocotylea 1. Viviparous; two anchors (hamuli) with supporting dorsal and ventral transverse bars; ventral bar with shield; 16 marginal hooks, eye spots absent; anchor hooks of embryo usually visible about midway in parent worm .(p. 97) Gyrodactylidae 1. Oviparous, usually with eye spots; usually 12 to 14 hooks, sometimes absent.2 2. (1) Body flat, large; haptor with well-developed muscles, utilizing suction for attachment to host skin; anterior body with pair of disclike adhesive organs; on marine fishes, including

Acipenser.

.(p. 121) Capsalidae 2. (1) Body usually cylindrical and small; haptor lacking suction disc, with anchors (hamuli) for primary attachment to gills of host; anterior end with head organs.3 3. (2) Haptor with one pair of anchors that articulate

— Ref. Kritsky and Thatcher (1977): Fundulotrema created to contain Gyrodactylus foxi, G. megacanthus, G. prolongis, G. stableri, G. trematoclithrus.

■ Fundulotrema foxi (Rawson, 1973) Kritsky and Thatcher, 1977 (Syn. Gyrodactylus f.)\ Fundulus heteroclitus, GA. ■ F. megacanthus (Wellborn and Rogers, 1967) Kritsky and Thatcher, 1977 (Syn. Gyrodactylus m.): Fundulus olivaceus, MS; Kozel and Whittaker (1985): F. notatus, KY. ■ F. prolongis (Hargis, 1955) Kritsky and Thatcher, 1977 (Syn. Gyrodactylus p.): Fundulus grandis, FL; Cone and Odense (1988): F. diaphanus, scanning EM, N.S.; Dick¬ inson and Threlfall (1975): F. heteroclitus, Nfdl.; Hanek and Fernando (1971b): on F. diaphanus, Ont.; A. R. Lawler (1982): cause of death of F. heteroclihts in aquaria. ■ F. stableri (Hathaway and Herlevich, 1973) Kritsky and Thatcher, 1977: F. kansae, CO; Cloutman (1974): same, KS; Janovy and Hardin (1987): F. zebrinus, NE; Janovy and Hardin (1988): same, diversity of parasite assemblage, NE.

with dorsal transverse bar; reduced ventral anchors and transverse bar present or not, usually on Cypriniformes.(p. 122) Dactylogyridae 3. (2) Haptor with two pairs of anchors (except for

Anonchohaptor and Icelanonchohaptor, with

Key to the Genera of Gyrodactylidae

one pair), which may or may not articulate each with a transverse bar.4 1. Marginal hooks evenly distributed on haptor; 4. (3) With two transverse bars that usually articulate each with one pair of the anchors; cirrus (penis) generally curved; usually on

Perciformes

.(p. 105) Ancyrocephalidae

each anchor with single superficial root; two transverse bars present, dorsal and ventral.2 1. Marginal hooks in three groups, two lateral, one posterior; each hamulus with two roots,

4. (3) Without two transverse bars supporting the usual two pairs of anchors; cirrus curved or U-shaped.5 5. (4) Ovary looping around right intestinal caecum; cirrus U-shaped with complex accessory piece comprising two or three arms; on catostomids

deep and superficial; one tranverse bar ventral; on marine and brackish water fishes.3 2. (1) Peduncle simple; on freshwater, brackish, or marine teleosts .(Figs. 148-150, Color Fig. 19)

Gyrodactylus

.(p. 130) Pseudomurraytrematidae 2. (1) Peduncle surrounded by anteriorly directed 5. (4) Ovary not looping around right intestinal caecum; cirrus curved with simple accessory piece; on gills of salmonids and esocids.(p. 132) Tetraonchidae

"flange" armed with sclerotized points; on Cyprinodontidae (Fundulus spp.) .(Figs. 151, 153)

Fundulotrema

3. (1) Peduncular transverse bar and sclerotized lateral haptoral accessory bars present .(Fig. 152)

Swingleus

Genus Fundulotrema

3. (1) Peduncular transverse bar absent.4

Kritsky and Thatcher, 1977

4. (3) Sclerotized bar or plate present between

(Figs. 151, 153)

Gyrodactylidae. Body elongated, with distinct peduncle, surrounded by anteriorly directed flange armed with sclerotized points; hooks distributed evenly on periph¬ ery of haptor. Anchors with elongated superficial root; ventral bar with pronounced anterolateral processes and elongate, rectangular shield; dorsal bar butterfly¬ shaped. On body surface of North American cyprinodontid fishes.

anchor roots.5 4. (3) Bar or plate between anchor roots absent, three pairs of accessory bars present.(no Fig.)

Polydithrum

5. (4) Sclerotized bar present between anchor roots; no lateral haptoral accessory bars .(no Fig.)

Gyrodactyloides

5. (4) Sclerotized plate present between anchor roots; eight hooks in posterior group.(no Fig.)

Laminiscus

head organ

pharynx cephalic gland cirrus intestine

embryo

ovary testis vitellaria

dorsal bar ventral bar ventral shield

haptor anchor cirrus

b

a

148. Gyrodactylus

150

151. Fundulotrema

152. Swingleus

153. Fundulotrema

-Gyrodactylid generaFIG. 148. Gyrodactylus spp.: a, Gyrodactylus eucaliae; b, G. macrochiri, haptor. FIG. 149. Terminology of haptoral armament of gyrodactylids. 1, anchor showing roots on base: a = general base area, b = deep root, c = superficial root, d = shaft, e = point; 2, anchor: a = base, b = knob, c = fold, d = superficial root, e = shaft, f = point, g = arc membrane; 3, hook: a and b = shank, c = heel, d = toe, e = shelf, f = shaft, g = point (recurved), h = point (open) (from Mizelle and Kritsky, 1967a). FIG. 150. Methods for measuring hard parts of haptor. 1, anchor: a = overall length, b = length of main body, c = length

of inner process, d = length of point; 2, dorsal connecting bar: a = length, b = width; 3, marginal hook: a = overall length, b = length of hook proper; 4, ventral connecting bar: a = length, b = width, c = length of “ears," d = length of ventral shield (from Ergens and Lorn, 1970, by permission of the Czechoslovak Academy of Sciences). FIG. 151. Fundulotrema, haptoral characters (from Beverley-Burton, 1984). FIG. 152. Swingleus polyclithroides, showing haptor and peduncular bar (from Rogers, 1969a, Tulane Studies in Zoology and Botany). FIG. 153. Fundulotrema megacanthus, showing haptoral and peduncular bar (from Wellborn and Rogers, 1967, J. Parasitol.).

100

Phylum Platyhelminthes: Class Monogenea

■ F. trematoclithrus (Rogers, 1967) Kritsky and Thatcher, 1977 (Syn. Gyrodactylus t.): Lucania goodei, AL.

Genus Gyrodactyloides Bykhovski, 1947 (No fig.)

Gyrodactylidae. With two median anchors, both with connecting bars and a supplementary chitinoid arma¬ ment that is typically well-developed. Supplementary armament consisting of plate or bar lying between inner processes (more powerfully developed than outer processes), with two filaments usually extending from it, surrounding inner processes of anchors and con¬ verging at center of posterior edge of connecting bar. Other characteristics as in the family. Parasites of marine and occasionally anadromous fishes. ■ Gyrodactyloides strelkowi Bykhovski and Polyanskii, 1953: gills, Oncorhynchusgorbuscha, former Soviet Union; (1982): O. nerka, AK.

Genus Gyrodactylus Nordmann, 1832 (Figs. 148-150, Color Fig. 19)

Viviparous; larva, including anchors, usually present in parent in utero. Very host-specific in nature, except possibly G. elegans. Many species. — Ref. The most comprehensive references are BeverleyBurton (1984); Ergens and Lorn (1970); Gusev (1978); Sproston (1946); Yamaguti (1963a, p. 10). Other pertinent references include Arcadi (1948): silver stain; F. Braun (1966): morphological variation from a clone; Cone and Wiles (1989): ultrastructure of attachment sites with necrosis; Ergens (1965): development of hard parts, vari¬ ability; Gelnar (1987b): effect of physical condition of host on the parasite, Czech Republic and Slovakia; G. L. Hoffman and Putz (1964): effect of temperature, also development; Kayton et al. (1982): histochemistry of hard parts; Kollman (1968): mechanism of hook; Lester (1972): penetration depth of anchors; Lester and Adams (1974b): simple population model; Lynn (1966): chemistry of hard parts; Lynn (1972a): sense organs; Lynn et al. (1981): silver impregnation for oncomiracidia chaetotaxy; Malmberg (1956a): Swedish Gyrodactylus spp.; Malmberg (1969): grouping and identification by excretory sys¬ tem; Malmberg and Malmberg (1970): comparison of excretory systems of Gyrodactylus and Dactylogyrus; F. Meyer (1970b): seasonality; Mizelle and Kritsky (1967a): key to species, host list, terminology of haptoral arma¬ ment; Mizelle and McDougal (1970): on amphibia; Putz (unpub., USFWS, 1962): methyl green wet mount to show cirrus spines (hooks); Yin and Sproston (1948): parasitized by Trichodina.

■ Gyrodactylus aculeati Malmberg, 1956a: Gasterosteus aculeatus, Sweden. ■ G. alabamensis Rogers, 1968: Nocomis leptocephalus, AL. ■ G. albeoli Rogers, 1968: Notropis albeolus, VA.

■ G. aldrichi Threlfall, 1974: gills, Couesius plumbeus, Lab. ■ G. alexanderi Mizelle and Kritsky, 1967b: skin, Gas¬ terosteus aculeatus, CA; Cone and Odense (1984): G. aculeatus, B.C.; Cone and Wiles (1985a): comparative study of five species on gasterostids, B.C. and Japan; Lester (1972): attachment to fish, B.C.; Lester (1974): skin, G. aculeatus, B.C.; Lester and Adams (1974a): reproduc¬ tion, morphology, effect on host; Lester and Adams (1974b): simple model of Gyrodactylus population, par¬ tial immunity. ■ G. ambystomae Mizelle, Kritsky, and McDougal, 1969: Ambystoma macrodactylum, CA. Amphibian species listed here to avoid confusion. ■ G. anguillae Ergens, 1960: Anguilla anguilla, Albania; Eversole (1981): A. rostrata, North America; Ogawa and Egusa (1978b, 1980): A. anguillae reared in Japan; Paperna and Lahav (1971): A. anguilla, Israel. ■ G. aphredoderi Rogers and Wellborn, 1965: Aphredodenis sayanus, AL. ■ G. aphyae Malmberg, 1956. Ergens (1970): Salmo trutta, Phoxinus phoxinus, Czech Republic and Slovakia; Ergens (1975): effect of water temperature on size of hard parts. ■ G. aquilinus Threlfall, 1974: fins, Catostomus catostomus, Lab. ■ G. arcuatus Bykhovski, 1933. Kennedy (1974): Gas¬ terosteus aculeatus, England. ■ G. asperus Rogers, 1967b: on Notropis baileyi, AL. ■ G. atratidi Putz and Hoffman, 1963: Rhinichthys atratulus, Semotilus margarita, WV; Hanek and Fernando (1971b): Rhinichthys atratulus, R. cataractae, Ont.; Kirby (1981): seasonality on Notropis spilopterus, most numer¬ ous at less than 15°C and when schooling; Threlfall (1974): sparse population on Catostomus commersoni, Lab. ■ G. aurorae Mizelle, Kritsky, and McDougal, 1969: on tadpoles of Rana aurora (frog), CA. Listed here to avoid confusion. ■ G. avalonia Hanek and Threlfall, 1969: Gasterosteus aculeatus, G. wheatlandi, Nfdl.; Cone et al. (1983): skin, Oncorhynchus mykiss in net pens, brackish, probably abnormal host; Cone and Wiles (1985a): Gasterosteus aculeatus, G. wheatlandi, Puntius puntius, N.B., N.S. (Syn. G. lairdi, G. memoriales, G. terranovae); Cusack and Cone (1985): microcolonies of bacteria on parasite; Hanek and Fernando (1971b): Fundulus diaphanus, G. aculeatus, Lepomis gibbosus, Rhinichthys atratidus, Ont.; Hanek and Molnar (1974): skin, G. aculeatus, Pungitius pungitius, Que.; Hanek and Threlfall (1969, 1970b, 1970c): on sticklebacks, Nfdl.; Threlfall and Hanek (1970b): Salvelinus fontinalis, Nfdl. ■ G. baeacanthus Wellborn and Rogers, 1967: Notropis venustus, GA; Kritsky and Mizelle (1968): N. atnoenus, VA.

Family Gyrodactylidae

■ G. bairdi Wood and Mizelle, 1957: Cottiis bairdi, IN; Arthur et al. (1976): Cottus cognatus, Aishihik Lake, Y.T.; Cone and Wiles (1983b): G. labradorius is syn.; Dechtiar (1972a, 1972b): C. bairdi, Ont.; Hanek and Threlfall (1970a): C. bairdi, Lab.; Lubinsky and Loch (1979): C. bairdi, Man.; Muzzal (1986): C. bairdi, ML ■ G. birmani Konovalov, 1967: Salvelinus alpinus, former Soviet Union. ■ G. bretinae Wellborn, 1967: Etheostoma stigmaeum, AR. ■ G. brevis Crane and Mizelle, 1967b: skin, Hesperoleucas navarroensis, Oncorhynchus rnykiss (steelhead), CA; Cone et al. (1983): probably accidental on O. mykiss. ■ G. bulbacanthus Mayes, 1977: Fundulus zebrinus, NE; Janovy and Hardin (1987, 1988): F. zebrinus, diversity of parasite assemblage, NE. ■ G. bullatarudis Turnbull, 1956: Poecilia reticulata, Canada, original source unknown; Rogers and Well¬ born (1965): same, AL; Cone and Odense (1984): patho¬ genicity minimal; G. L. Hoffman (unpub., USFWS, 1967): P. reticulata, CO; Kritsky and Fritts (1970): Poecilia sphenops, Costa Rica; Reichenbach-Klinke (pers. comm., Univ. of Munich, 1960): Germany. ■ G. bychowskyi Sproston, 1946: new name for Bykhovski's G. medius from Gasterosteus aculeatus, East Prussia. ■ G. callawayensis Mayes, 1977: Notropis lutrensis, NE. ■ G. cameroni Hanek and Threlfall, 1970: Apeltes quadracus, Nfdl.; Cone and Wiles (1985b): same, N.B., N.S. ■ G. campostomae Wellborn, 1967: Campostoma anomalum, AL; Cloutman (1976): same, AR; Kritsky and Mizelle (1968): same, VA. ■ G. canadensis Hanek and Threlfall, 1969: Gasteros¬ teus aculeatus, G. wheatlandi, Nfdl.; Cone and Wiles (1985a): same, Nfdl., Que.; Hanek and Molnar (1974): gills, fins, G. aculeatus, Pungitius pungitius, Que.; Hanek and Threlfall (1970a, 1970b, 1970c): Apeltes quadracius, Gasterosteus aculeatus, G. wheatlandi, Lab., Nfdl. ■ G. carassii Malmberg, 1956a: Carassius auratus, Swe¬ den; Ergens (1971a): same, possibly syn. with G. decorus and G. magnificus, Czech Republic and Slovakia. ■ G. carpio Kritsky and Mizelle, 1968: Cyprinus carpio, AR. ■ G. chologastris Mizelle, Whittaker, and McDougal, 1969: Chologaster agassizi (cave fish), C. comuta (swamp fish), NC, KY. ■ G. colemanensis Mizelle and Kritsky, 1967b: skin, Oncorhynchus mykiss, CA; Cone et al. (1983): skin, O. mykiss, Salvelinus fontinalis, S. namaycus, AR, Nfdl., N.S., Ont.; Cone and Cusack (1988): O. mykiss, Salmo salar, Salvelinus fontinalis, comparison with G. salmonis, N.S.; Cone and Wiles (1989): not pathogenic to fry of O. mykiss; Cusack (1986): ultrastructure of attachment; Cusack et al. (1988): bacterial colonies on parasite; Hath¬ away and Herlevich (1973): S. fontinalis, CO.

101

■ G. commersoni Threlfall, 1974: gills, Catostomus commersoni, Lab. ■ G. couesius R. A. Wood and Mizelle, 1957: Couesius plumbius dissimilis, B.C. ■ G. crysoleucas Mizelle and Kritsky, 1967b: Notemigonus crysoleucas, CA. ■ G. ctenopharyngodontis Lin M.-E., 1962. BykhovskayaPavlovskaya (1962): skin, Ctenopharyngodon idella, China; Edwards and Hine (1974): C. idella, New Zealand. ■ G. cylindriformis Mueller and Van Cleave, 1932: Umbra limi, NY. ■ G. cypiini Diarova, 1964: Cyprinus carpio, former Soviet Union; Glaser (1969): comparison of Gyrodactylus species of C. carpio, Germany; Rogers (1968): C. carpio, AL. ■ G. cyprini Kollman, 1968. Gusev (1985): syn. of G. katharineri. ■ G. cyprinodontis Mizelle and Kritsky, 1967b: Cyprinodon nevadensis, Death Valley, CA. ■ G. dakotensis Leiby, Kritsky, and Peterson, 1972: skin, Ictiobus cyprinellus, ND; Kritsky et al. (1972): Carpiodes car¬ pio, ND. ■ G. dechtiari Hanek and Fernando, 1971b: gills, Rhinichthys atratuli, R. cataractae, Ont. ■ G. dorosomae Rogers, 1975: Dorosoma cepedianum, AL. ■ G. egregius R. A. Wood and Mizelle, 1957: Richardsonius egregius, NV. ■ G. egusae Ogawa and Hioki, 1986: Anguilla japonica, Japan. ■ G. elegans von Nordmann, 1832: common species in Europe, occurrence in North America needs more study; Anthony (1969): effect of temperature increase; Brienholt and Heckmann (1980): Catostomus discobolis, C. latipinnis, UT; Cone and Wiles (1983c): comparison of G. elegans var. A with other gyros of Carassius caras¬ sius, N.S.; Curtin (1956): Cyprinus carpio, PA; Haderlie (1953): Salmo gairdneri (=Oncorhynchus mykiss), CA; Kennedy (1974): Gasterosteus aculeatus, England; Lewis and Lewis (1963): Notemigonus crysoleucas, IL; Malm¬ berg (1956a): G. elegans Nordmann, 1832, from Abramis bratna is true species, that from North American gold¬ fish is probably Wagner's species, and that reported from trout by Mueller (1936) is different species; Mueller (1936a): var. A on C. carassius, var. B on trout, NY; Nowlin et al. (1967): C. auratus, TX; Seamster (1938a, 1938b): Ictalurus melas. ■ G. elegans muelleri (G. elegans var. A Mueller, 1936) Yin and Sproston, 1948: Carassius auratus, North America. ■ G. elegans salmonis (G. elegans var. B Mueller, 1936) Yin and Sproston, 1948: trout, North America. ■ G. ensatus Mizelle, Kritsky, and Bury, 1968: larvae of salamander, Dicamptodon ensatus; review of Monogenea on animals other than fish. Included here to avoid confusion.

102

Phylum Platyhelminthes: Class Monogenea

■ G. eos Mayes, 1977: skin, Phoxinus eos, NE. ■ G. etheostomae Wellborn and Rogers, 1967: Etheostoma radiosum, AR; Hanek and Fernando (1971a): Etheostoma nigrum, Ont.; Kozel and Whittaker (1982): Etheostoma caeruleum, KY; Kritsky and Leiby (1971): Etheostoma asprigens, ND; Molnar et al. (1974): Etheostoma exile, Ont. ■ G. eucaliae Ikezaki and Hoffman, 1957 (Fig. 148): Culaea inconstans, ND; Cone and Wiles (1985a): review of Gyrodactylus on gasterosteids; Dechtiar (1972a): C. inconstans, Ont.; Hanek and Fernando (1971a): same, Ont.; Kritsky (1978): study of cephalic glands; Kritsky and Kruidenier (1976): ultrastructure; Kritsky and Mizelle (1968): C. inconstans, ND; Fubinsky and Foch (1979): same, Man. ■ G. fairporti Van Cleave, 1921b: Cyprinus carpio, Ictalurus melas IA; Kritsky and Mizelle (1968): redescription; Frost (1973): L nebulosus marmoratus, AF ■ G. foxi Rawson, 1973: is Fundulotrema foxi. ■ G. freemani Hanek and Fernando, 1971b: fins, Perea flavescens, Ont. ■ G. fryi Cone and Dechtiar, 1984: skin, Esox masquinongy, Ont.

■ G. khendensis Ergens, 1974: gills, fins, nasal fossae, skin, C. carpio, Mongolia; Fux (1987): C. carpio, Germany; Ogawa and Egusa (1978b): same, Japan. ■ G. kobayashii Hukuda, 1940. Cone and Wiles (1983c): comparison with other Gyrodactylus from Carassius auratus; Ergens and Ogawa (1978): redescription from C. auratus, G. elegans is syn., Japan. ■ G. labradorius Hanek and Threlfall, 1970a. Cone and Wiles (1983b): syn. of G. bairdi. ■ G. lacustricolae Rogers, 1967b: Erimyzon sucetta, AF. ■ G. lacusgrandis Hanek and Threlfall, 1970a: Cottus bairdi, Fab. ■ G. lacustris Mizelle and Kritsky, 1967a: Pimephales promelas, ND. ■ G. lairdi Hanek and Threlfall, 1969: Gasterosteus aadeatus, G. wheatlandi, Nfdl.; Cone and Wiles (1985a): pos¬ sibly syn. of G. avalonia; Hanek and Threlfall (1970, 1971): Nfdl., Fab. ■ G. lambanus Rogers, 1967b: Notropis longirostris, AF. ■ G. laruei Kritsky and Mizelle, 1968: Campostoma atiotnalum, VA.

■ G. funduli Hargis, 1955a: Fundulus similis, FF; Hutton (1964): same; Sogandares (pers. comm.): same, FA.

■ G. limi Wood and Mizelle, 1957: Umbra limi, IN; Hanek and Fernando (1971a): same, Ont.; Peckham and Dineen (1957): same, IN.

■ G. gambusiae Rogers and Wellborn, 1965: Gambusia affinis, FF.

■ G. lineadactylus Wellborn, 1967: Pomoxis nigromaculatus, MS.

■ G. gloriosi Rogers, 1968: Enneacanthus gloriosus, GA; Mayes (1973): same, NC. ■ G. goerani Hanek and Fernando, 1971a: Ambloplites rupestris, Ont. ■ G. gurleyi E. H. Price, 1937b: Carassius auratus, TX; Cone and Wiles (1983c): redescription and comparison with other Gyrodactylus from C. auratus. ■ G. hargisi E. W. Williams and Rogers, 1971: gills, Cyprinodon variegatus, syn. of Gyrodactylus sp. Hargis, 1955, AR. ■ G. heterodactylus Rogers and Wellborn, 1965: Elassoma zonatum, AF. ■ G. hoffmani Wellborn and Rogers, 1967: skin, Pimephales promelas, AF; Holloway (1986): same, ND; Mizelle and Kritsky (1967a): same, ND; Molnar et al. (1974): same, Ont.; G. ictaluri Rogers, 1967: Ictalurus punctatus, FF. ■ G. illigatus Rogers, 1968: Notropis shumardi, AF. ■ G. imperialis Mizelle and Kritsky, 1967b: skin, Gillichthys mirabilis, Salton Sea, CA. ■ G. joi Ogawa and Hioki, 1986: Anguilla japonica, Japan. ■ G. katharineri Malmberg, 1964: Cyprinus carpio, Swe¬ den; Cone and Dechtiar (1985): same, Fake Erie, Ont.; Gelnar (1987a, 1987c): effect of temperature, scaled C. carpio more heavily parasitized; Glaser (1969): C. car¬ pio, comparison of Gyrodactylus of carp.

■ G. lingulatus Rogers, 1968: Hypentelium etowanum, AF. ■ G. longoacuminatus Zitnan, 1964: Carassius auratus, Europe: Cone and Wiles (1983c): comparison with other Gyrodactylus of C. auratus; Ergens (1980b): redescrip¬ tion, Czech Republic and Slovakia; Ogawa (pers. comm., Japan, 1980): C. auratus, Japan. ■ G. lotae Gusev, 1953: Lota lota, former Soviet Union; Cone and Dechtiar (1985): skin, L. lota, Algonquin Park, Ont. ■ G. lucii V. G. Kulakovskaya, 1952: skin, Esox lucius, for¬ mer Soviet Union; Cone and Dechtiar (1985): E. lucius, Fake Huron, Ont.; Ergens (1971b): same, Mongolia. ■ G. lythruri Rogers, 1975: Notropis atrapiculis, N. bellus, AF. ■ G. macrochiri G. F. Hoffman and Putz, 1964 (Syn. G. elegans of Hargis, 1953): Lepomis cyanellus, L. macrochirus, and experimentally on Micropterus salmoides, WV; Dechtiar (1972b): Lepomis gibbosus, Ont.; Hanek and Fernando (1971a): Micropterus dolomieui, M. salmoides, Ont.; Fubinsky and Foch (1979): Man.; Molnar et al. (1974): M. salmoides, Ont.; Rawson and Rogers (1972): population peak in February; Rogers and Wellborn (1965): L. macrochirus, AF, Lepomis punctatus and Chaenobryttus gulosua, FF; J. R. Sullivan (1977): Micropterus sp., AF.

Family Gyrodactylidae

103

■ G. magniclypeus Rogers, 1968: Moxostoma cervinum, VA.

■ G. parvicirrus Rogers, 1975: Notropis atherinoides, AL.

■ G. magnus Konovalov, 1967: Thymallus arcticus, for¬ mer Soviet Union; Ergens (1971b): same, Mongolia.

■ G. percinae Rogers and Wellborn, 1965: Percina nigrofasciata, AL.

■ G. margaritae Putz and Hoffman, 1963: Semotilus margarita, WV.

■ G. planensis Mayes, 1977: skin, Notropis dorsalis, NE.

■ G. masu Ogawa, 1986: skin, cultured Oncorhynchus masou, O. mykiss, O. rhodurus, Japan.

■ G. prolongis Hargis, 1955a: Fundulus grandis, FL; Sogandares (pers. comm.): will transfer to Cyprinodon variega¬ tus in aquaria, LA.

■ G. medins Kathariner, 1894: Micropterus dolomieui (questioned by Price, 1937b); Cooper (1915): same, Canada; Ergens (1962, 1970, 1974): Cyprinus carpio, Czech Republic and Slovakia; Glaser (1969): comparison of Gyrodactylus from C. carpio, Germany; M. C. Meyer (1954): Salvelinus fontinalis, ME; Paperna (1964a): C. carpio and others, Israel; Reichenbach-Klinke (1956b): Poecilia reticulata, aquarium fishes, Germany. ■ G. mernorialis Hanek and Threlfall, 1969: Gasteroseus aculeatus, G. wheatlandi, Nfdl; Hanek and Threlfall (1970a); Cone and Wiles (1985a): syn. of G. avalonia. ■ G. micropogonus Wood and Mizelle, 1957: Micropogon undulatus, TX. ■ G. minytremae Wellborn, 1967: Minytrema melanops, AL. ■ G. mirabilis Mizelle and Kritsky, 1967b: Cottas asper, CA. ■ G. mizelli Kritsky and Leiby, 1971: Stizostedion canadense, S. vitreum, ND. ■ G. tnobilensis E. H. Williams and Rogers, 1971: gills, Cyprinodon variegatus, AL. ■ G. nannus Rogers, 1968: Menidia beryllina, AL. ■ G. nataliae Rogers, 1967b: Hybopsis amblops, AL. ■ G. navarroensis Crane and Mizelle, 1967b: Hesperoleucas navarroensis, CA. ■ G. nebraskensis Mayes, 1977: skin, Phoxinus nevgoeus, NE. ■ G. nebulosus Kritsky and Mizelle, 1968: fins, Ictalurus nebulosus, ND; Hanek and Fernando (1971a ): same, Ont.; Molnar et al. (1974): same. ■ G. nerkae Cone, Beverly-Burton, Wiles, and McDonald, 1983: skin, Oncorhynchus nerka, B.C.; R. E. Bailey and Margolis (1987): same. ■ G. nevadensis Mizelle and Kritsky, 1967b: skin, Cyprin¬ odon nevadensis, CA. ■ G. nigrum Rogers, 1975: Etheostoma nigrum, AL. ■ G. nipponensis Ogawa and Egusa, 1978b: gills, Anguilla japonicum, Japan; Ogawa and Egusa (1980): A. anguilla imported from France. ■ G. olsoni Mizelle and Kritsky, 1967b: skin, Gillichthys mirabilis, Salton Sea, CA. ■ G. osoblahensis Ergens, 1964. Ergens and Dulmaa (1969): Cyprinus carpio, Mongolia.

■ G. plumbeae Threlfall, 1974: Couesius plumbeus, Lab.

■ G. protuberus Rogers and Wellborn, 1965: Notropis uranoscopus, AL. ■ G. pungitii Malmberg, 1964. Kennedy (1974): Gasterosteus aculeatus, England. ■ G. rachelae Price and McMahon, 1967: gills, Notemigonus crysoleucas, TN. ■ G. rarus Wegener, 1909. Chappel (1969a, 1969b): fins, Gasterosteus aculeatus, England; Kennedy (1974): same. G. rhinichthius Wood and Mizelle, 1957: Rhinichthys osculus, NV; Lang and Edson (1976): same, WA. ■ G. richardsonius Wood and Mizelle, 1957: Richardsonius egregius, NV. ■ G. salaris Malmberg, 1956a: Salmo salar, Sweden; Cone (1981): on salmonids at a fish hatchery, B.C.; Johnsen and Jensen (1988): spread in Norway, mortal¬ ity of smolts and parr of S. salar in streams; Kennedy (1978a): S. trutta, England: Lucky (1963): O. mykiss, Czech Republic and Salovakia; Malmberg (1973): S. trutta, WV, former Soviet Union; Malmberg (1987b): intraspecific variation. Author: this species seems to be widespread and on occasion is devastating to young salmonids, feral as well as in culture. Importation of G. salaris should be avoided. ■ G. salmonis Yin and Sproston, 1948 (=G. elegans salmonis): trout, USA; Cone et al. (1983): emended description, on Oncorhynchus kisutch, B.C.; Oncorhynchus mykiss, AR, ID, MT, B.C., N.S.; Salmo aquabonita, CA; Salmo clarki {-Oncorhynchus clarki), B.C., Salmo salar, N.S.; Salmo trutta, NC; Salvelinus fontinalis, N.S.; Cone and Odense (1984): pathogenicity and penetration of hooks; Cone and Cusack (1988): Oncorhynchus mykiss, Salmo salar, Salvelinus fontinalis, N.S., widespread on body, hook penetration; Mitchum (pers. comm., State Fisheries, 1990): common in WY. ■ G. saratogensis Mizelle and Kritsky, 1967b: skin, Cyprin¬ odon nevadensis, Death Valley, CA. ■ G. schmidti Kritsky and Leiby, 1971: Stizostedion vit¬ reum, ND. ■ G. shulmani Ling, 1962 (=G. chinensis Ling, 1962). Ergens (1970): synonymy, gills, skin, mouth, Carassius auratus, Cyprinus carpio, Hypophthalmichthys molitrix, Czech Republic and Slovakia; Lux (1987): C. carpio,

Germany. ■ G. spathulatus Mueller, 1936a: Catostomus commersoni; Dechtiar (1972a, 1972b): C. commersoni, Moxostoma

104

Phylum Platyhelminthes: Class Monogenea

anisurum, Ont.; Hathaway and Herlevich (1973): C. commersoni, CO; Krueger (1954): same, OH; Lubinsky and Loch (1979): Man.; Threlfall (1974): C. catosto¬ nuis, Lab. ■ G. sprostonae Ling M.-E., 1962. Mattheis and Glaser (1970): Cyprinus carpio, biology and treatment, Ger¬ many; Molnar and Ghittino (1977): C. carpio, Po River, Italy. ■ G. stableri Hathaway and Herlevich, 1973: syn. of Fundulotrema s. ■ G. stankovici Ergens, 1970: Cyprinus carpio, Czech Republic and Slovakia; Ergens (1971): C. carpio, vari¬ ability, distribution in Europe; Molnar (1976b): C. car¬ pio, Hungary. ■ G. stegurus Mueller, 1937a: Fundulus diaphanus, NY. ■ G. Stephanas Mueller, 1937a: Fundulus heteroclitus, MD; Dickinson and Threlfall (1975): same, Nfdl.; Hanek and Threlfall (1976): Pungitius pungitius, Nfdl.; Dillon (1966): list of parasites of Fundulus; Hargis (1955a): Fun¬ dulus grandis, FL; A. R. Lawler (1982): secondary to G. prolongis in death of Fundulus heteroclitus in aquaria. ■ G. shinkardi Kritsky and Mizelle, 1968: Catostonuis occidentalis, CA; Hanek and Fernando (1971a): Etheostoma nigrum, Rhinichthys atratulus, R. cataractae, Ont.; Molnar et al. (1974): on fins, C. commersoni, Ont.; Threlfall (1974): Lab. ■ G. tennesseensis Rogers, 1968: Chrosomus eythrogaster, TN. ■ G. terranovae Hanek and Threlfall, 1969: Gasterosteus aculeatus, G. wheatlandi, Nfdl.; Cone and Wiles (1985a): syn. of G. avalonia. ■ G. trematoclithrus Rogers, 1967b: syn. of Fundulo¬ trema t. ■ G. truttae Glase, 1974. Ergens (1981): Salmo tnitta, vari¬ ability, Czech Republic and Slovakia; Malmberg (1987a): identification problems. ■ G. variabilis Mizelle and Kritsky, 1967b: Notemigonus crysoleucas, CA. ■ G. wardi Kritsky and Mizelle, 1968: Catostonuis occidentalis, CA; Hathaway and Herlevich (1973): Catostonuis catostonuis. ■ G. wagneri von Nordmann, 1832. Ergens (1979a, 1980b): is nomen nudum. ■ G. viriosus Wellborn and Rogers, 1967: Hypentelium etowanus, GA; Chien (1969): Hypentelium nigricans, GA. ■ G. wellborni Nowlin, 1968: skin, Notemigonus crysoleu¬ cas, TN. ■ Gyrodactylus spp. Because every freshwater fish or group of closely related fishes seems to have one or more species of Gyrodactylus, the complete list will not be recorded here; there are probably many undescribed species.

Genus Laminiscus Palsson and Beverley-Burton, 1983 (No fig.) Gyrodactylidae. Marginal hooks, each with relatively short, straight shaft, distributed on periphery of haptor in three groups: two anterolateral groups of four hooks each, and one posterior group of eight more widely dis¬ tributed hooks. Anchors with short, deep root and elon¬ gated superficial root. One connecting transverse bar present; thin, sclerotized plate extending from distal end of anterolateral roots to bar, supporting anchors and anterolateral groups of hooks. On gills of marine and brackish water teleosts. ■ Laminiscus strelkowi (Bykhovski and Polyanskii, 1953) Palsson and Beverley-Burton, 1983 (Syn. Gyrodactyloides s.). Margolis (1956): Oncorhynchus gorbuscha, O. nerka, B.C.

Genus Polyclithrum Rogers, 1967 (No fig.) Gyrodactylidae. Head bilobed, head organs present. Opisthohaptor with one pair of anchors, dorsal and ventral bars, and three pairs of accessory bars. Sixteen marginal hooks present; sclerotized processes support¬ ing haptor. Each anchor base with short, deep root and elongated superficial root; dorsal bar not attached to anchor; ventral bar with long posteriorly directed process; three pairs of accessory bars articulating with anchors. Marginal hooks located dorsally on haptor, divided into two main groups: four pairs located on anterolateral margin; remaining four pairs on posterior margin. Phar¬ ynx pyramidal; gut bifurcate, not confluent posteriorly. Cirrus pouch with one large spine; with or without small spinelets. Testis postovarian but partially encircling ovary. Uterus usually containing developing embryo. Par¬ asitic on fishes. ■ Polyclithrum nutgilini Rogers, 1967e: on fins, body, Mugil cephalus, Lake Seminole, GA.

Genus Swingleus Rogers, 1969 (Fig. 152) Gyrodactylidae. Head bilobed, head organs present. Pharynx pyramidal; gut bifurcate; not confluent poste¬ riorly. Cirrus with spine and spinelets. Peduncular bar present. Haptor with one pair of anchors, ventral bar, pair of lateral, winglike bars, and 16 hooks. Dorsal bar absent. Anchor base with cap of dense tissue. Marginal hooks dis¬ tributed in two groups: three pairs located on antero¬ lateral margin; remaining five pairs on posterior margin. Parasitic on fishes.

Family Ancyrocephalidae

■ Swingleus polycrithroides Rogers, 1969a: skin, Fundulus grandis, AL. ■ Swingleus sp. Billeter (1974): Fundulus heteroclitus, NY.

ent or absent; if present, one or two pairs, supported by one, two, or three transverse bars. Mouth lacking oral sucker and buccal suckers. Oviparous.

Order Dactylogyrida

Family Ancyrocephalidae

Bykhovski, 1937

(Bykhovski, 1937, as subfamily) Bykhovski and Nagibina, 1978

Fourteen or 16 marginal hooks, with most posterior pair, I, migrating anteriorly from peripheral to more central position; pair II between anchors. Anchors pres-

Key to the Families of Dactylogyrida (Modified from Beverley-Burton, 1984) 1. Body usually flat, large; haptor septate or nonseptate, with well-developed muscles, utilizing suction for attachment to skin or gills of fish; anterior body with pair of adhesive organs. On marine fishes, but included here because one genus on Acipenser .(p. 121) Capsalidae 1. Body slender, haptor with accessory adhesive plaques.(p. 129) Diplectanidae 1. Body usually cylindrical, small; haptor lacking "suction discs" musculature, with anchors of prime importance for attachment to gills of fish; anterior body with head organs. Primarily on freshwater fishes.2 2. (1) Haptor with 16 marginal hooks, pairs II to VIII peripheral, and pair I central; two pairs of anchors present, one dorsal, and one ventral; single transverse bar present. On gills of esocids and salmonids.(p. 132) Tetraonchidae 2. (1) Haptor with 14 marginal hooks, pairs II to VII peripheral and pair I central; one or two pairs of anchors and one, two, or three transverse bars present; anchors and bars sometimes secondarily reduced or lacking. Primarily on gills of freshwater fishes other than esocids and salmonids.3 3. (2) Ovary looping around right intestinal caecum; cirrus U-shaped, with complex accessory piece comprising two or three arms. On catostomids .(p. 130) Pseudomurrytrematidae 3. (2) Ovary not looping around right intestinal caecum; cirrus usually curved tube of varying size and shape, but not forming "U".4 4. (3) Haptor usually with one pair of anchors (dorsal) that articulate with dorsal transverse bar; reduced ventral bar and vestigial ventral anchor present or not. Typically, but not always, on Cypriniformes .(p. 122) Dactylogyridae 4. (3) Haptor with two pairs of anchors, each supported by transverse bar. Typically, but not always, on Perciformes.(p. 105) Ancyrocephalidae

105

Dactylogyrida. Body small, anterior with cephalic lobes; head organs present; cephalic glands in two groups in anterolateral region; two pairs of eyespots present. Haptor more or less distinct, with 14 marginal hooks, 12 peripheral and two central; two pairs of anchors present, usually one pair dorsal and one ventral, each sup¬ ported by a transverse bar. Mouth ventral, muscular pharynx present; intestinal caeca confluent. Testes intercaecal, postovarian; vas deferens looping dorsoventrally around left caecum, expanding to form variable acces¬ sory piece. Ovary pyriform or elongated, not looping around intestine; vaginal pore lateral, submarginal, or median. Freshwater or brackish water; typically, but not exclusively, on Perciformes. Author: There are 20 North American genera in this family; I have also included one genus that occurs on exotic Tilapia that have been established on this conti¬ nent. In the previous edition, I accepted considerable synonymization of genera, but in view of the work of others, particularly Beverley-Burton (1984, 1986, and more), I have reversed most of these to aid in the iden¬ tification of more species. I have, however, condition¬ ally retained Haplocleidus (anchors markedly dissimilar in size) and Petrocleidus ("winged" anchors). — Ref. Beverley-Burton (1984, 1986); Hargis and Thoney (1983); Mizelle and Hughes (1938): key to species; Mizelle and Price (1964b): key to genera; Mizelle et al. (1956): review; Sproston (1946); Yamaguti (1963a).

See the Key to the Genera of Ancyrocephalidae on p. 106.

Genus Actinocleidus Mueller, 1937 (Fig. 154)

Ancyrocephalidae. Two pairs of anchors present, both projecting ventrally; transverse bars articulating to form single supporting unit; marginal hooks of similar size and shape, except that hooks of pair II have smaller handle. Copulatory complex (Type I of Beverley-Burton, 1984) comprising sclerotized, tubular cirrus with inflated base and heavily sclerotized accessory piece with distal, fin¬ gerlike projection and bifid base. Vagina sclerotized or not, opening on left side of body, leading to seminal receptacle. Vitellaria coextensive with intestine, extend¬ ing laterally to body margin. On gills of centrarchids. — Ref. Beverley-Burton (1986): review and key; Mizelle et al. (1956): review.

Key to the Genera of Ancyrocephalidae Trianchoratus

1. Possessing three anchors. . (p. 120, Fig. 178)

1. Possessing four anchors.2 2. (1) In stomach or intestine.(p. 115, Figs. 163, 164)

Enterogyrus

10. (5) Vagina on left.14 11. (10) Anchors each with blunt, fingerlike projection on inner curve of blade; cirrus with loosely spiraling filaments supporting large sheath into which accessory piece is incorporated; on centrarchids. . . (p. 118, Fig. 174)

2. (1) On gills or in branchial chamber, sometimes mouth . . 3 3. (2) FHaptor with two transverse bars.4 3. (2) FHaptor with one transverse bar; anchors

Leptocleidus

with coiled cirrus. . . . (p. 116, Figs. 166, 167)

4. (3) Found on freshwater and brackish cyprinodonts; curved, tubular cirrus with inflated base, accessory piece variable and not articulated basally .(p. 119, Fig. 175)

Salsuginis

4. (3) Found on ictalurids; curved, tubular cirrus with inflated base, accessory piece basally attached .(p. 116, Figs. 168-170, Color Fig. 20)

Ligictaluridis

than body; anchors similar; transverse bars about same size; on centrarchids.(p. 120, Fig. 177)

Tetradeidus

12. (11) Cirrus with spirals; haptor not narrower than body; "keyhole" accessory piece not basally articulated with cirrus.13 13. (12) Anchors similar and about same size; transverse on centrarchids.(p. 117, Fig. 173)

Onchodeidus

13. (12) Dorsal anchor and transverse bar markedly larger than ventral; marginal hooks similar in shape and size;

4. (3) Found on fishes other than cyprinodonts and ictalurids.5

on centrarchids.(p. 115, Fig. 165)

Haplodeidus

14. (10) Haptor much wider than body; dissimilar anchors

5. (4) With two articulating (attached) transverse haptoral bars.6 5. (4) With two nonarticulating transverse haptoral bars. . 10 6. (5) Haptor distinct, umbrella-like, with scalloped margin; two pairs of anchors on small protuberance in center of ventral haptor surface; otherwise very

Actinodeidus.(p. 113, Fig. 160) Clavunculus

and transverse bars.(p. 117, Fig. 172)

Macrohaptor

14. (10) Haptor about same width as body; anchors and transverse bars usually similar.15 15. (14) Anchors large but lacking roots; anchor points extending beyond haptoral margin; on centrarchids .(p. 117, Fig. 171)

Lyrodiscus

15. (14) Anchors not extremely large and

6. (5) Flaptor distinct, but anchors not on central protuberance.7 7. (6) Haptoral bars dissimilar but relatively small; anchors moderately sized; cirrus with inflated base and articulated accessory piece with distal, fingerlike projection; on centrarchids;

Crinideidus.(p. 105, Fig. 154) Actinodeidus

7. (6) Haptoral bars dissimilar but relatively large.8

solid, shieldlike plate.(p. 119, Fig. 176)

not possessing roots.16 16. (15) Ventral bar consisting of three attached pieces (compound bar); genus is intermediate between

Actinodeidus and Ancyrocephalus; on cichlids, including Tilapia spp., Israel . . (p. 113, Figs. 158, 159) Cichlidogyrus 16. (15) Ventral bar not compound as above; not on cichlids.17 17. (16) Possessing coiled cirrus and accessory piece; on

8. (7) Anchor bases relatively small; dorsal bar a

Syndeithrum

8. (7) Anchor bases large to very large.9

Poecilia reticulata.(p. 120, Fig. 179b) Urocleidoides 17. (16) Cirrus not coiled.18 18. (17) Cirrus with spiraling filaments supporting sheath

9. (8) Haptor markedly larger than body;

to which accessory piece is attached; on etheostomids

anchor bases so large that anchors occupy all of haptor.(p. 112, Fig. 157)

Anchoradiscus

9. (8) Haptor about same width as body; anchors triangular in outline, with expanded proximal region; transverse bars articulating with each other to form massive central support.(p. 112, Fig. 156)

12. (11) Cirrus without spirals; haptor markedly narrower

bars about same size; marginal hooks varying in size;

and bearing hooklike projections distally

includes

11. (10) Anchors without fingerlike projection; with or without cirrus spirals.12

dissimilarly sized; in pharyngeal cavity of centrarchids;

similar to

Pterodeidus

Anchoradiscoides

10. (5) Vagina opening medially; anchors similar; transverse bars about same size; cirrus tubular, slightly curved; accessory piece not basally articulated with cirrus and with two distally fused rami; on percids and percopsids.(p. 120, Fig. 179a)

Urodeidus 10.

.(p. 112, Fig. 155)

Aethycteron

18. (17) Cirrus without spiraling filaments; tubular cirrus relatively short and robust; accessory piece a single unit with distal, fingerlike projection and bifurcated base; anchors relatively short; on centrarchids .(p. 113, Fig. 162) 18. (17) As with

Cleidodiscus

Cleidodiscus, but vagina not observed;

anchors with spur at base of anchor shaft and joining the anchor point distally; conspicuous double anchor wings present; cirrus with basally articulated accessory piece; on centrarchids . . (p. 113, Fig. 161)

Cleidodiscoides

10. (5) Vagina on right.11 (text continues on page 111)

head organs eyes pharynx

intestinal caecum accessory piece cirrus seminal vesicle vaginal opening vas deferens seminal receptacle ovary testis vitellaria

marginal hook anchor

transverse bars

154. Actinocleidus

157. Anchoradiscus

156. Attchoradiscoides

158. Cichlidogyrus

-Ancyroceplialid monogeneansFIG. 154. Actinocleidus, generalized schematic. FIG. 155. Aethycteron, generalized schematic. FIG. 156. Anchoradiscoides serpentinus. FIG. 157. Anchoradiscus triangularis (Figs. 154-157 from Beverley-Burton, 1984). FIG. 158. Cichlidogyrus tilapiae, drawn from living specimens (from Paperna, 1960). FIG. 159. C. tilapiae, haptoral armature (from Paperna, 1960). FIG. 160. Clavunculus okeechobeensis: 1 = haptor, 2 = hook, 3 = anterior anchor, 4 = posterior anchor, 5 = anterior bar, 6 = posterior bar, 7 = accessory piece, 8 = cirrus (from Mayes and Miller, 1973, Proc. Helm. Soc. Wash.).

HO

163

Enterogyrus

- Ancyrocephalid monogeneans FIG. 161. Cleidodiscoides sulcata (from Mayes and Miller, 1973, Proc. Helm. Soc. Wash.). FIG. 162. Cleidodiscus, generalized schematic. FIG. 163. Enterogyrus cichlidarum: AG = anchoral gland, EJ = cirrus, HG = haptoral gland, HO = head organ, O = ovary, PR = prostate, T = testis, U = uterus, VS = seminal vesicle (from Paperna, 1963b). FIG. 164. E. cichlidarum, haptoral armature; anterior is at the left, cirrus at the bottom. FIG. 165. Haplocleidus, generalized schematic. FIG. 166. Leptocleidus, generalized schematic. FIG. 167. Leptocleidus megalonchus: e = cirrus; f = accessory piece (Figs. 162-167 from Beverley-Burton, 1984).

168. Ligictaluridus

head organs eyes pharynx accessory piece cirrus seminal vesicle vaginal opening seminal receptacle

E E

vas deferens ovary vitellaria