no Thorp and Covich’s freshwater invertebrates, 4th edition (5 volumes) [1-5/1, 4 ed.] 9780123850263

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Thorp and Covich’s Freshwater Invertebrates

A Global Series of Books on the Identification, Ecology, and General Biology of Inland Water Invertebrates by Experts from Around the World Fourth Edition Edited by James H. Thorp Volume I : Ecology and General Biology Edited by James H. Thorp and D. Christopher Rogers Published 2015 Volume II: Keys to Nearctic Fauna Edited by James H. Thorp and D. Christopher Rogers Expected Publication Date: 2015 Volume III: Keys to Palaearctic Fauna Edited by D. Christopher Rogers and James H. Thorp Expected Publication Date: 2016 Planned Future Volumes of the Fourth Edition Keys to Palaearctic Insects and Aquatic Collembola Keys to Oriental Fauna Keys to Australasian and Oceana Fauna Keys to Neotropical and Antarctica Fauna Keys to Afrotropical Fauna

Related Publications in This Series Ecology and Classification of North American Freshwater Invertebrates Edited by J.H. Thorp and A.P. Covich First (1991), Second (2001), and Third (2010) Editions

Ecology and General Biology Thorp and Covich’s Freshwater Invertebrates - Volume I

Fourth Edition Edited by

James H. Thorp D. Christopher Rogers

AMSTERDAM • BOSTON • HEIDELBERG • LONDON • NEW YORK • OXFORD • PARIS SAN DIEGO • SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

Academic Press is an imprint of Elsevier 32 Jamestown Road, London NW1 7BY, UK 525 B Street, Suite 1800, San Diego, CA 92101-4495, USA 225 Wyman Street, Waltham, MA 02451, USA The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, UK Copyright © 2015 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangement with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library Library of Congress Cataloging in Publication Data A catalog record for this book is available from the Library of Congress For information on all Academic Press publications visit our website at http://store.elsevier.com/ ISBN: 978-0-12-385026-3 Printed and bound in China



Dedications from the Editors

“To the many dedicated authors and both old and new friends who over the years have contributed their efforts and hard-won knowledge to these books and research on invertebrate ecology and taxonomy.” James H. Thorp “To my family and anyone else who has wondered what was going on under the water’s surface.” D. Christopher Rogers

Contributors to Volume I

Michael T. Bogan [Chapter 41] Department of Environmental Science, Policy, and Management, University of California Berkeley, Berkeley, California, USA, email: [email protected] Matthew G. Bolek [Chapter 15]  Department of Zoology, Oklahoma State University, 415 Life Sciences West, Stillwater, Oklahoma 74078 USA, email: bolek@ okstate.edu John E. Brittain [Chapter 34]  Natural History Museum, University of Oslo, P.O. Box 1172 Blindern, NO-0318 Oslo, Norway, email: [email protected]

Peter S. Cranston [Chapter 40]  Evolution, Ecology, and Genetics, Research School of Biology, The Australian National University, Canberra ACT 0200 Australia; Also an Emeritus Professor at Entomology Department, University of California Davis, Davis, California 95616, USA, email: pscranston@ucdavis Neil Cumberlidge [Chapter 32]  Department of Biology, 2009 New Science Facility, Northern Michigan University, 1401 Presque Isle Avenue, Marquette, Michigan 49855 USA, email: [email protected]

Kenneth M. Brown [Chapter 18] Department of Biological Sciences, Louisiana State University, A343 Life Sciences Annex, Baton Rouge, Louisiana 70803, USA, email: [email protected]

Kevin S. Cummings [Chapter 19]  Illinois Natural History Survey, Division of Biodiversity and Ecological Entomology, University of Illinois, 607 E. Peabody Drive, MC-652, Champaign, Illinois 61820 USA, email: [email protected]

Francisco Brusa [Chapter 10] División Zoología Invertebrados, Museo de La Plata, FCNyM-UNLP, 1900 La Plata, Argentina, email: [email protected]

Cristina Damborenea [Chapter 10]  División Zoología Invertebrados, Museo de La Plata, FCNyM-UNLP, 1900 La Plata, Argentina, email: [email protected]

Carla E. Cáceres [Chapter 28]  School of Integrative Biology, Program in Ecology, Evolution and Conservation Biology, Morrill Hall, University of Illinois at Urbana-Champaign, 505 South Goodwin, Urbana, Illinois 61801, USA, email: [email protected]

L. Cristina De Villalobos [Chapter 15]  Museo de Ciencias Naturales, Paseo del Bosque FCNyM-UNLP, 1900 La Plata, Argentina, email: [email protected]

David R. Cook [Chapter 25]  7725 North Foothill Drive South, Paradise Valley, Arizona, USA Rickey D. Cothran [Chapter 31] Department of Biological Sciences, University of Pittsburgh, 4249 Fifth Avenue, Pittsburgh, Pennsylvania 15260 USA, email: [email protected] Gregory W. Courtney [Chapter 40]  Department of Entomology, Iowa State University, 432 Science II, Ames, Iowa 50011 USA, email: [email protected] Matthew R. Cover [Chapter 41]  Department of Biological Sciences, California State University Stanislaus, One University Circle, Turlock, California 95382 USA, email: [email protected] Alan P. Covich [Chapter 2, 27]  Odum School of Ecology, Ecology Building, University of Georgia, Athens, Georgia 30602-2202 USA, email: [email protected]

R. Edward DeWalt [Chapter 36]  Prairie Research Institute, Illinois Natural History Survey, University of Illinois, 607 East Peabody Drive, Champaign, Illinois 61820 USA, email: [email protected] Klaas-Dowe B. Dijkstra [Chapter 35]  Nederlands Centrum voor Biodiversiteit Naturalis, Leiden, Darwinweg 2, 2333 CR Leiden, The Netherlands, email: [email protected] Walter W. Dimmick [Chapter 1]  2612 Harper Street, Lawrence, Kansas 66046 USA, email: dimmick@ sunflower.com Genoveva Esteban [Chapter 7]  Conservation Ecology and Environmental Sciences Group, School of Applied Sciences, Bournemouth University, Talbot Campus Poole, Dorset BH12 5BB, UK, email: gesteban@­ bournemouth.ac.uk Bland J. Finlay [Chapter 7]  School of Biological and Chemical Sciences, Queen Mary University of London, The River Laboratory, Wareham, Dorset, BH20 6BB, England, email: [email protected] xix

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  Contributors to Volume I

Nadine Folino-Rorem [Chapter 9] Department of Biology, Wheaton College, IL, 501 College Avenue, Wheaton, Illinois 60187 USA, email: [email protected]

Tobias Kånneby [Chapter 12]  Department of Zoology, Swedish Museum of Natural History, PO Box 50007, SE-104 05 Stockholm, Sweden, email: tobias. [email protected]

Stuart R. Gelder [Chapter 22]  Department of Science & Mathematics, ME, University of Maine at Presque Isle, Presque Isle, Maine 04769 USA, email: stuart.gelder@ umpi.edu

Siegfried Kehl [Chapter 39]  Department of Animal Ecology II, Universität Bayreuth, Universitätsstr. 30, 95440 Bayreuth, Germany, email: siegfried.kehl@ uni-bayreuth.de

Jean-Jacques Geoffroy [Chapter 26] Département Ecologie et Gestion de la Biodiversité, Muséum National d’Histoire Naturelle, UMR 7204 CESCO CNRS-MNHN-UPMC, 4 avenue du petit Château 91800 Brunoy, France, email: [email protected]

Boris C. Kondratieff [Chapter 36]  C. P. Gillette Museum of Arthropod Diversity, Department of Bioagricultural Sciences and Pest Management, Colorado State University, 012A Laurel Hall, Fort Collins, Colorado 80523, USA, email: [email protected]

Stanislav Gorb [Chapter 35] Spezielle Zoologie, Universität Kiel, Am Botanischen Garten 1-9, 24098 Kiel, Germany, email: [email protected]

David M. Lodge [Chapter 32]  Department of Biological Sciences, University of Notre Dame, Notre Dame, Indiana 46556, USA, email: [email protected]

Frederic R. Govedich [Chapter 23]  Department of Biological Sciences, Southern Utah University, Cedar City, Utah 84720, USA, email: [email protected]

David A. Lytle [Chapter 37]  Department of Integrative Biology, Cordley Hall, Oregon State University, Corvallis, Oregon 97331, USA, email: [email protected]

Daniel L. Graf [Chapter 19]  Department of Biology, University of Wisconsin, Stevens Point, 2100 Main St., Stevens Point, Wisconsin 54481, USA, email: dgraf@ uwsp.edu

Renata Manconi [Chapter 8]  Dipartimento di Scienze della Natura e del Territorio (DIPNET), Università di Sassari, Via Muroni 25, I-07100 Sassari, Italy, email: [email protected]

Roberto Guidetti [Chapter 17]  Department of Life Sciences, University of Modena and Reggio Emilia, via Campi 213/D, 41125, Modena, Italy, email: roberto. [email protected]

Koen Martens [Chapter 30]  Freshwater Biology, Royal Belgian Institute of Natural Sciences, Brussels, Belgium; and Department of Biology, University of Ghent, Ghent, Vautierstraat 29, 1000 Brussels, Belgium, email: martens @naturalsciences.be

Ben Hanelt [Chapter 15]  Department of Biology, University of New Mexico, Albuquerque, 163 Castetter Hall, New Mexico 87131, USA, email: [email protected] Horton H. Hobbs [Chapter 32]  Department of Biology, Wittenberg University, Springfield, Ohio, USA, email: [email protected] Rick Hochberg [Chapter 12]  Department of Biological Sciences, University of Massachusetts-Lowell, One University Avenue, Lowell, Massachusetts 01854, USA, email: [email protected] Ralph W. Holzenthal [Chapter 38]  Department of Entomology, 219 Hodson Hall, University of Minnesota, 1980 Folwell Ave, St. Paul, Minnesota 55108, USA, email: [email protected]

Patrick J. Martin [Chapter 21]  Biologie des Eaux, Institut royal des Sciences naturelles de Belgique douces 29, rue Vautier, B-1000, Bruxelles, Belgium, email: [email protected] William E. Moser [Chapter 23]  Department of Invertebrate Zoology, National Museum of Natural History, Washington, DC, Smithsonian Institution, Museum Support Center, MRC 534, 4210 Silver Hill Road, Suitland, Maryland 20746, USA, email: [email protected] Diane R. Nelson [Chapter 17]  Department of Biological Sciences, East Tennessee State University, Johnson City, Tennessee 37614, USA, email: [email protected]

David J. Horne [Chapter 30]  School of Geography, Queen Mary University of London, Mile End Road, London E1 4NS, UK, email: d.j.horne@ qmul.ac.uk

Carolina Noreña [Chapter 10]  Departamento Biodiversidad y Biología Evolutiva, Museo Nacional de Ciencias Naturales (CSIC), Madrid, España, c/o Jose Gutierrez Abascal 2, 28006 Madrid, España, email: [email protected]

Vincent J. Kalkman [Chapter 35]  Nederlands Centrum voor Biodiversiteit Naturalis Leiden, Darwinweg 2, 2333 CR Leiden, The Netherlands, email: [email protected]

Brian J. O’Neill [Chapter 33]  Department of Biological Sciences, University of Wisconsin – Whitewater, 269 Wyman Mall, Whitewater, Wisconsin 53190 , USA, email: [email protected]

  Contributors to Volume I

George O. Poinar Jr. [Chapter 14] Department of Zoology, Oregon State University, Corvallis, Oregon 97331, USA, email: poinarg@science. oregonstate.edu Heather C. Proctor [Chapter 25] Department of Biological Sciences, CW405 Biological Sciences Building, University of Alberta, Edmonton, T6G 2E9, Canada, email: [email protected] Roberto Pronzanto [Chapter 8]  Dipartimento di Scienze della Terra, dell’Ambiente e della Vita (DI.S.T.A.V.), Università di Genova, Area Scientifico-Disciplinare 05 (Scienze biologiche), Settore BIO/05, Genova, Italy, email: [email protected] Mark Pyron [Chapter 18]  Department of Biology, Ball State University, Cooper Life Sciences Building, CL 121, Muncie, Indiana 47306, USA, email:mpyron @bsu.edu Lorena Rebecchi [Chapter 17]  Department of Life Sciences, University of Modena and Reggio Emilia, via Campi 213/D, 41125, Modena, Italy, email: lorena. [email protected] Vincent H. Resh [Chapter 6] Department of Environmental Science, Policy, and Management, University of California, 305 Wellman Hall, Berkeley, California 94720, USA, email: [email protected] Anthony Ricciardi [Chapter 5]  Redpath Museum, McGill University, Montreal, Quebec H3A 2K6, Canada, email: [email protected] Blanca Ríos-Touma [Chapter 38]  Centro de Investigación de la Biodiversidad y el Cambio Climático, Museo de Zoología, Universidad Tecnológica Indoamérica, Quito, Ecuador, email: [email protected]. [Also contact at: Department of Landscape Architecture & Environmental Planning, 300 Wurster Hall, University of California, Berkeley, California 94553, USA.] D. Christopher Rogers [Chapters 1, 3, 24, 27-28]  Kansas Biological Survey and Biodiversity Institute, Higuchi Hall, University of Kansas, 2101 Constant Avenue, Lawrence, Kansas 66047-3759, USA, email: [email protected]

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John B. Sandberg [Chapter 36]  California Department of Fish and Wildlife, CSUC Research Foundation, California State University, Holt Hall, 1205 W 7th St., Chico, California 95929, USA, email: jsandberg@ csuchico.edu Michel Sartori [Chapter 34]  Museum of Zoology, Palais de Rumine, Place Riponne 6, CH-1014 Lausanne, Switzerland, email: [email protected] Andreas Schmidt-Rhaesa [Chapter 15] Zoological Museum, University Hamburg, Martin Luther-King. Platz 3, 20146 Hamburg, Germany, email: andreas. [email protected] Isa Schön [Chapter 30]  Freshwater Biology, Royal Belgian Institute of Natural Sciences, Vautierstraat, 29, 1000 Brussels, Belgium, email: Schoen @naturalsciences.be Alison J. Smith [Chapter 30]  Department of Geology, Kent State University, Kent, Ohio 44242, USA, email: [email protected] Bruce P. Smith [Chapter 25]  Department of Biology, Ithaca College, Ithaca, New York 14850, USA, email: [email protected] Hilary A. Smith [Chapter 13]  Department of Biological Sciences, University of Notre Dame, Notre Dame, Indiana 46556, USA, email: [email protected] Ian M. Smith [Chapter 25]  Canadian National Collection of Insects and Arachnids, Agriculture and Agri-Food Canada, K.W. Neatby Building, 960 Carling Ave., Ottawa, Ontario K1A 0C6, Canada, email: [email protected] Terry W. Snell [Chapter 13]  School of Biology, Environmental Science and Technology, Georgia Institute of Technology, Atlanta, Athens, Georgia 30332, USA, email: [email protected] Malin Strand [Chapter 11] Coordinator, Marine Invertebrates, Swedish Species Information Centre, Swedish University of Agricultural Sciences, Box 7007, SE 75007 Uppsala, Sweden, email: [email protected] Eduardo Suárez-Morales [Chapter 29]  El Colegio de la Frontera Sur (ECOSUR), Unidad Chetumal, P.O. Box 424. Chetumal, Quintana Roo 77014, Mexico, email: [email protected]

David M. Rosenberg [Chapter 6]  Emeritus Scientist Freshwater Institute, Fisheries and Oceans Canada, 501 University Crescent, Winnipeg, Manitoba, R3T 2N6, Canada, email: [email protected]

Frank Suhling [Chapter 35]  Institut für Geoökologie, Technische Universität Braunschweig, Langer Kamp 19c, Raum 303, 38106 Braunschweig, Germany, email: [email protected]

Göran Sahlén [Chapter 35]  Ecology and Environmental Sciences, Halmstad University, Högskolan i Halmstad, Box 823, Kristian IV: s väg 3, 30118 Halmstad, Sweden, email: [email protected]

Per Sundberg [Chapter 11]  Department of Biological and Environmental Sciences, University of Gothenburg, P.O. Box 463, SE-405 30 Gothenburg, Sweden, email: [email protected]

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Robin E. Thomson [Chapter 38] Department of Entomology, University of Minnesota, Hodson Hall, 1980 Folwell Avenue, St. Paul, Minnesota 55108, USA, email: [email protected] James H. Thorp [Chapters 1–4, 24, 27, 33]  Kansas Biological Survey and Department of Ecology and Evolutionary Biology, Higuchi Hall, University of Kansas, 2101 Constant Avenue, Lawrence, Kansas 66047, USA, email: [email protected] Tarmo Timm [Chapter 21]  Institute of Agricultural and Environmental Sciences, Centre for Limnology, Estonian University of Life Sciences, Rannu, Tartumaa 61117, Estonia, email: [email protected] Jan van Tol [Chapter 35]  Nederlands Centrum voor Biodiversiteit Naturalis, Leiden, Darwinweg 2, 2333 CR Leiden, The Netherlands, email: jan.vantol@ ncbnaturalis.nl Piet F.M. Verdonschot [Chapter 20] Freshwater Ecology, Alterra, Wageningen UR, P.O. Box 47, 6700 AA Wageningen, The Netherlands, email: [email protected] Robert L. Wallace [Chapter 13]  Department of Biology, Ripon College, Ripon, 300 Seward Street, Ripon, Wisconsin 54791, USA, email: [email protected]

  Contributors to Volume I

Alan Warren [Chapter 7]  Department of Life Sciences, Natural History Museum, Cromwell Road, London SW7 5BD, UK, email: [email protected] Gary A. Wellborn [Chapter 31]  Department of Biology, University of Oklahoma, 730 Van Vleet Oval, Norman Oklahoma 73019, email: [email protected] Bronwyn W. Williams [Chapter 22]  Department of Zoology, Southern Illinois University, 1125 Lincoln Drive, MC 6501, Carbondale, Illinois 62901, USA, email: [email protected] Jonathan D.S. Witt [Chapter 31]  Department of Biology, University of Waterloo, 200 University Avenue West, Waterloo, Ontario, N2L 3G1Canada, email: [email protected] Timothy S. Wood [Chapter 16]  Department of Biological Sciences, Wright State University, 3640 Colonel Glen Highway, Dayton, Ohio 45435, USA, email: tim. [email protected] Donald A. Yee [Chapter 39]  Department of Biological Sciences, University of Southern Mississippi, 118 College Drive #5018, Hattiesburg, Mississippi 394060001, USA, email: [email protected]

Acknowledgments for Volume I

Many people contributed to this volume in addition to the chapter authors and those acknowledged in individual chapters. We greatly appreciate all our colleagues who have contributed information, figures, or reviews to Volume I and also thank those who provided similar services for the earlier editions, upon which the present book partially relies. In particular, we would like to thank Vince Resh, who suggested a number of possible chapter authors in addition to writing his own chapter.

Finally, we are again grateful to the highly competent people at Academic Press/Elsevier who helped in many aspects of the book’s production from the original concept to the final marketing. In particular, we appreciate our association with Sean Coombs and Candice Janco in the U.S. offices of Elsevier, as well as past editors who assisted us in producing T&C I–III. James H. Thorp D. Christopher Rogers

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About the Editors

using descriptive, experimental, and modeling approaches in the field and laboratory. While his research emphasizes aquatic invertebrates, he also studies fish ecology especially as related to food webs. He has published more than one hundred refereed journal articles, books, and chapters, including three single-volume editions of Ecology and Classification of North American Freshwater Invertebrates (edited by J.H. Thorp and A.P. Covich). Prof. Thorp is now embarked on a major project to expand from the previous North American emphasis on freshwater invertebrates to the fourth edition’s global coverage of this subject in perhaps nine volumes of Thorp and Covich’s Freshwater Invertebrates. Dr James H. Thorp has been a Professor in the Department of Ecology and Evolutionary Biology at the University of Kansas (Lawrence, KS, USA) and a Senior Scientist in the Kansas Biological Survey since 2001. Prior to returning to his alma mater, Prof. Thorp was a Distinguished Professor and Dean at Clarkson University, Department Chair and Professor at the University of Louisville, Associate Professor and Director of the Calder Ecology Center of Fordham University, Visiting Associate Professor at Cornell University, and Research Ecologist at the University of Georgia’s Savannah River Ecology Laboratory. He received his Baccalaureate degree from the University of Kansas and both Masters and PhD degrees from North Carolina State University. Those degrees focused on zoology, ecology, and marine biology, with an emphasis on the ecology of freshwater and marine invertebrates. Dr Thorp is currently on the editorial board of two journals (River Research and Applications and River Systems) and is a former President of the International Society for River Science (ISRS). He teaches three courses at the University of Kansas (Principles of Ecology, River and Lake Ecology, and Marine Biology) and has both Masters and doctoral graduate students working on various aspects of the ecology of organisms, communities, and ecosystems in rivers, reservoirs, and wetlands. Prof. Thorp’s research interests and background are highly diverse and span the gamut from organismal biology to the ecology of communities, ecosystems, and macrosystems. He works on both fundamental and applied research topics

D. Christopher Rogers is a research zoologist at the University of Kansas with the Kansas Biological Survey and is affiliated with the Biodiversity Institute. Christopher specializes in freshwater crustaceans (particularly the Branchiopoda and the Decapoda) and the invertebrate fauna of seasonally astatic wetlands on a global scale. He has numerous peer reviewed publications in crustacean taxonomy and invertebrate ecology, as well as published popular and scientific field guides and identification manuals to freshwater invertebrates. Christopher is an Associate Editor for the Journal of Crustacean Biology and a founding member of the Southwest Association of Freshwater Invertebrate Taxonomists. He has been involved in aquatic invertebrate conservation efforts all over the world.

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Preface to the Fourth Edition

When Alan P. Covich and I initially conceived the first edition of Ecology and Classification of North American Freshwater Invertebrates in the mid-1980s (subsequently published in 1991 by Academic Press), the possibility of a second edition (2001) was only vaguely considered, much less a third (2010). Between the second and third edition, I tried to convince some European scientists to develop independently a similar approach for their fauna, but a lack of interest at that time doomed this proposed Elsevier book project. Less than a year after the third edition, an acquisition editor at Elsevier (Candace Janco) inquired about a fourth edition for 2015 at an annual meeting of the North American Benthological Society (now Society for Freshwater Sciences). Other project commitments prevented Alan from continuing as a coeditor, and I felt that 2015 was too early for a fourth edition without major changes in the scope of the project or detail of the keys. At that point a kernel of what was possibly a creative but certainly audacious idea began to sprout. This soon grew to a proposal I submitted with D. Christopher Rogers to develop an approximately 9-volume series covering inland water invertebrates of the world. (Christopher had coauthored two chapters in the third edition and has now coauthored A Field Guide to Freshwater Invertebrates of North America with me.) Although the prospectus ultimately submitted to Elsevier included books covering all major zoogeographic regions, the initial contract was limited to three volumes encompassing the world’s major book markets (Europe and the USA). While I am the sole editor of the book series at this time, Christopher has been a major partner in developing ideas for the fourth edition and is an editor on the other volumes (senior editor on the third). As we made significant progress on the first three volumes, we began contacting some potential coeditors and authors to develop volumes for other zoogeographic regions and negotiations with some of those volumes are now underway. Based on my feelings and recommendations of Academic Press, I have named our book series Thorp and Covich’s Freshwater Invertebrates in order to: (a) associate present with past editions, unite current volumes in the fourth edition, and link to possible future editions; (b) establish a connection between the ecological and general biology

coverage in volume I with the taxonomic keys in the remaining volumes; and (c) give credit to Alan Covich for his work on the first three editions. For the sake of brevity, I refer to the current edition as T&C IV. Whether T&C “V” will ever appear is certainly problematic, but who knows! Our concept for T&C IV included producing one book (volume I) with 6 chapters on general environmental issues applicable to many invertebrates, followed by 35 chapters devoted to individual taxa at various levels (order to phylum, or even multiple phyla in the case of the protozoa). Volume I was designed both as an independent book on ecology and general biology of various invertebrate taxa and as a companion volume for users of the keys in the regional taxonomic volumes, thereby reducing the amount of duplicate information needed for the taxonomic volumes. The perhaps eight taxonomic volumes will contain both keys for identifying invertebrates in specific zoogeographic regions and descriptions of detailed anatomical features needed to employ those keys. The multilevel keys are formatted to enable users to work easily at the level of their project/course need and their scientific experience. For that reason, we separated keys by major taxonomic divisions. For example, a student in a college course might work through one or more of the initial crustacean keys to determine the family to which a freshwater shrimp belongs. In contrast, someone working on an environmental monitoring project might need to identify a crayfish to genus or even species, and thus would use the relevant, detailed keys that require more background experience. We have also designed the keys, where possible, to proceed from a general to a specific character within a couplet. These changes to the key are one of Christopher’s major contributions to T&C IV. While the vast majority of authors in T&C I–III were from the USA or Canada, we attempted in T&C IV to attract authors from many additional countries in six continents. Although we largely succeeded in this goal, we expect future editions of T&C to continue increasing the proportion of authors from outside North America as our books become better known internationally. Our goals for T&C IV are to improve the state of taxonomic and ecological knowledge of inland water

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invertebrates, to help protect our aquatic biodiversity, and to encourage more students to devote their careers to working with these fascinating organisms. These goals are especially important because the verified and probable losses of species in wetlands, ponds, lakes, creeks, and

Preface to the Fourth Edition

rivers around the globe exceed those in most terrestrial habitats. James H. Thorp Editor-in-Chief, T&C IV

Preface to Volume I

Readers familiar with previous three editions of the Thorp and Covich (T&C) volumes will notice a dramatic change in T&C IV, as we have expanded from a North American concentration to worldwide coverage. This volume consists of 6 general chapters on inland water habitats and invertebrates, and 35 chapters on the ecology and general biology of specific groups. The greatest difference between this volume of T&C IV and the chapters in T&C III is that all taxonomic keys have been shifted to separate volumes. This has allowed an increase in the ecological coverage in volume I and more detailed keys in subsequent volumes. This enabled us to expand from the North American-centric coverage in previous editions to the global ecological perspective characterizing T&C IV, which is one of the reasons we increased the international representation of authors. In the process we added some groups not occurring in the Nearctic (e.g., aquatic millipedes) or whose numbers were not large in the Nearctic (e.g., freshwater crabs and shrimp, which dominate tropical freshwater habitats). Readers of the earlier editions will note that we have expanded coverage of the ecology and general biology of insects from a single chapter in T&C III to 9 chapters in volume I. However, the taxonomic keys in Volume II for the Nearctic Region remain limited to family level to avoid duplication with the excellent existing text on North American aquatic insects by Merritt, Cummins, and Berg (2008). The literature-cited sections in volume I represent a compromise between T&C II and III, in that we have returned to inclusion of author names and dates in the text (rather than just numbered citations) and have expanded the number of allowed citations at the end of each chapter, as in T&C II; however, the average number of references per chapter in this printed volume is often less than in the first two volumes in order to save space. T&C IV continues our policy from the previous

edition of being in color for volume I; indeed, we have considerably augmented the number of color figures. We believe this helps students, in particular, appreciate better that bright colorful invertebrates are not entirely limited to marine and terrestrial habitats. This edition is strongly focused on species found in fresh through saline inland waters, with a nonexclusive emphasis on surface waters. Again, most estuarine and parasitic species are not covered in this book, but we do discuss species whose life cycles include a free-living stage (e.g., Nematomorpha) and species that live in hard freshwaters through to brackish waters even though they may be normally associated with estuarine or marine habitats in some parts of their life cycles (e.g., some shrimp and crabs). It is our hope that scientists and students from around the world will enjoy this volume, but it is also important that we interest the younger generation (especially the kindergarten through middle school group of children) in freshwater invertebrates. One way that this could happen is to present them with interesting books to read on these organisms and freshwaters in general. Few have been written, but one of the more interesting ones includes biology, ecology, and even rudimentary taxonomy! That book is entitled The Secret Life of Streams and was authored by Lynell Marie Garfield, with contributions from Daniel Devine, Jason Barnes, and Sandra L. Silva (see www.clearmountainstream.com). This charming children’s book was published in 2013 by Lucy Bat Books (www.luckybatbooks.com) and has an ISBN number of 1-9390-5133-9). We strongly recommend you consider getting this book for your younger children. Editors, Volume I James H. Thorp D. Christopher Rogers

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Chapter 1

Introduction to Invertebrates of Inland Waters James H. Thorp Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, Lawrence, KS, USA

D. Christopher Rogers Kansas Biological Survey and Biodiversity Institute, University of Kansas, Lawrence, KS, USA

Walter W. Dimmick Lawrence, KS, USA

Chapter Outline Introduction3 Species and Phylogenies 4 Species and Concepts of Species 4 Biological Species Concept 5 Phylogenetic Species Concept 6 Evolutionary Species Concept 6 Cohesion Species Concept 6 The Role of Phylogenies in Studies of Ecology and Behavior 7 Phylogenetic Trees 8 Using Systematics and Taxonomy to Classify Organisms 9

INTRODUCTION At some point in almost every biologist’s life, an invertebrate stimulated their curiosity. It might have started when they peered through a microscope at pond water and saw a protozoan, hydra, rotifer, or some other creature. Or perhaps they watched a butterfly or bee pollinating a flower, a pill bug curled in a ball within their palm, ants marching steadily onward with heavy loads, anemones waving their tentacles in a tide pool, or a crayfish scurrying between refuges under stream rocks. It is true that many inland water invertebrates may initially seem drab and uninspiring next to their more colorful and often larger marine relatives. Yet, once the curious observer penetrates beyond superficial appearances and thoroughly examines their diverse structural, physiological, behavioral, and general ecological adaptations, few fail to

Taxonomic Keys to Invertebrates of Inland Waters 10 Key to Freshwater Invertebrates 11 Mollusca Classes 11 Annelid Groups 12 Arthropod Subphyla 12 Tetraconata: Crustacean Classes and Malacostracan Orders12 Tetraconata: Hexapoda Classes 13 Tetraconata: Hexapoda: Insecta Orders 13 References21

be impressed by these fascinating freshwater creatures. This childhood fascination led progressively, if not inevitably, to collegiate studies of invertebrates and then to a life-long career in science for many authors contributing to this and other volumes in our fourth edition. Aquatic scientists focusing on invertebrates conduct research in a wide diversity of fields. Some are fascinated by the diversity of invertebrates and the relationships among species over evolutionary history, or they may be involved in conducting environmental assessments in which knowledge of invertebrate identification is typically essential. Others seek to mitigate human diseases and parasitism by understanding and controlling those where freshwater invertebrates play a role. Many more study the population biology or culturing of invertebrates involved in food webs,

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00001-2 Copyright © 2015 Elsevier Inc. All rights reserved.

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which contribute directly (freshwater crayfish) or indirectly (such as zooplankton prey of lake fish) to human diets. Many more investigate the ecology of invertebrates either from an environmental monitoring perspective or just to satisfy their scientific curiosity. In this chapter, we discuss invertebrate taxonomy and classification. We also include a taxonomic key to help the reader begin the process of identifying inland water invertebrates at higher taxonomic levels. This dichotomous key will lead to the appropriate chapter in volume I for information on the ecology and general biology of various groups and to more detailed identification keys in the other volumes of this series. Additional information on freshwater invertebrates can be found in various textbooks on invertebrate zoology and aquatic ecology. For amateur invertebrate zoologists and students needing initial guidance to species in the field and laboratory, we recommend the use of more introductory texts, such the Field Guide to Freshwater Invertebrates of North America (Thorp and Rogers, 2011).

SPECIES AND PHYLOGENIES The identification and definition of natural biological units is one of the major guiding principles for biologists that endeavor to study, discover, and document the world’s biodiversity. Unless you properly identify the organism you are studying, you will be hindered in drawing any justifiable conclusions. Genera and species are examples of natural biological units as they result from evolution. The formal name of a natural biological unit is a taxon. In contrast, a population is an example of a qualitative unit that is often not a natural unit but rather a convenient partition of geographic variation. The principal intellectual tools used to define natural taxa have emerged from the scientific disciplines of taxonomy, systematic biology, and population genetics. Species concepts and phylogenetic analyses are central to systematic biology. Population genetics is a method of direct study of connectivity between populations. Sometimes, analyses of molecular variation enable the discovery of cryptic species, which are difficult to distinguish by morphological characters alone. This has resulted in the recognition of multiple species that were formerly thought to be a single wide-ranging species. There are several different species concepts, and no one species concept is necessarily better than another. There are various groups of organisms wherein one concept works better than another, depending on the evolution of that group. Similarly, most species can be defined using two or more species concepts. These species concepts are not competing concepts, but rather different tools to be used sometimes in concert, sometimes individually, to describe and define biodiversity. In fact, the appropriateness of a particular species concept can vary with the evolutionary

SECTION | I Introduction

patterns of that group, and most taxa can be defined using two or more species concepts. Although this situation may seem inconvenient for a new student or a researcher who is primarily interested in ecological or behavioral studies, an informed viewpoint about species concepts and phylogenetics can only help a serious scholar of invertebrates. Cataloging biodiversity is paramount in our modern world, given the alarming amount of local extirpation and global extinction of species (particularly because of anthropogenic factors). Although it may appear that taxonomy changes capriciously, the real issue is that taxonomic concepts are hypotheses to be tested. A species may move from one genus to another or move from species to subspecies and back as new data becomes available. The goal is to have taxonomic and systematic classification and hierarchy converge with biological reality.

Species and Concepts of Species Evolution is a phenomenon of lineages, and the termini of these lineages are called species. Species are the result of historical natural processes (Darwin, 1859). Anagenesis is the modification of lineages through mutation, gene flow, natural selection, and random genetic drift. Cladogenesis is the creation of a new lineage by splitting a preexisting lineage or the result of hybridization between two preexisting lineages. Consequently, the first logical step to understanding current and past biodiversity is the identification of species and other natural units that result from evolution (Mayden and Wood, 1995). Scientists often need to identify biodiversity units below the genus level, but ultimately, unit definitions must be based on appropriate species concepts. Species concepts play a critical role in the interpretation of variation because species provide an upper boundary for population comparisons. Whether a particular population or group of populations is distinctive enough to be recognized as a species continues to be a key issue in many taxonomic and conservation disputes. Taxonomists identify and describe species, and classify them according to a hierarchy. This is accomplished by following the scientific method, as is any branch of science. A species is a name for a discrete group of organisms (see species concepts below). The definition of that name is, in essence, a theory, constructed by the taxonomist based on available morphological, molecular, behavioral, and/or ecological data. Although these “theories” are meant to be tested, as are all scientific theories, it should be remembered that species are unstable through time unlike most scientific subjects. They evolved from other species, they are still under selective pressures, and they may evolve away from the original described theory or may become extinct or both.

Chapter | 1  Introduction to Invertebrates of Inland Waters

The scientific literature about the nature of species and the best method of identifying species is enormous and was previously contentious. Species concepts almost always fall into one of two categories: operational species concepts or ontological species concepts. Operational species concepts provide specific criteria to determine whether or not two different populations belong to the same species. The biological species concept of Mayr (1942) is a classic example of an operational species concept. The operation used by Mayr is the discovery of barriers to sexual reproduction that are assumed to define species boundaries. Ontological species concepts provide a theoretical species definition, but do not specify a method for their identification. Wiley’s (1981) evolutionary species concept is an example of an ontological species concept. Species are defined as lineages expected to persist through time, but no method (operation) is prescribed for the discovery of these lineages. In short, ontological concepts can be understood as theoretical species definitions. Operational species concepts provide only a method of how to discover species and are, by definition, limited by their particular methodology. Below are brief definitions of the biological, phylogenetic, cohesion, and evolutionary species concepts along with comments on their strengths and limitations. These influential concepts provide a good basis for understanding many issues regarding species concepts. For a more thorough discussion and evaluation of species concepts, we recommend an article by Mayden and Wood (1995). Suffice to say that not all species concepts will work with all groups, some groups can be defined by only one concept, some groups have different species depending on which concept is used, and still other groups have species that are defined the same way regardless of which concepts is used. This is a reflection of the diversity of life and the diversity of ways that species can evolve. A good intellectual benchmark to start from, or return to, when considering any aspect of species or speciation is Darwin’s basic proposition that new species originate by the splitting of preexisting species. Under this premise, we accept the idea that species are new lineages, but we are not constrained in any way on how to identify them. This leads us to an ontological species concept rather than an operational species concept. Because biological diversity is enormous, it seems highly unlikely that any single operational concept could be devised to encompass all species.

Biological Species Concept Species are groups of actually or potentially interbreeding populations that are reproductively isolated from other such groups (Mayr, 1942). This biological species concept has been prevalent in the evolutionary literature for the last several decades and is emphasized in many college level biology courses. It

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is probably the species concept most familiar to organismal biologists in diverse fields, such as conservation biology, forestry, fisheries, and wildlife management. Species defined by the biological species concept have also been championed as units of conservation (O’Brien and Mayr, 1991). Theodosius Dobzhansky, a prominent evolutionary geneticist and an important contributor to the modern evolutionary synthesis, characterized the concept of a biological species as a system of populations (Dobzhansky, 1935, 1970). The gene exchange between these systems (species) is limited or prevented by reproductive isolating mechanisms, such as species-specific breeding behaviors, hybrid sterility, and gametic incompatibility. Thus, under the biological species concept, species are simultaneously a reproductive community, a gene pool, and a genetic system. The study of reproductive isolating mechanisms is central to the biological species concept because these mechanisms provide gene flow barriers that define the boundaries of the reproductive community and gene pool thereby preserving the integrity of the species’ genetic system. In practice, however, isolating mechanisms are rarely studied and species are usually diagnosed by differences in morphological features. A fundamental drawback to the biological species is the concept is that it is exclusively defined in terms of sexual reproduction. Asexual and cyclically parthenogenetic taxa are obviously excluded from this concept, but it is also true that many species capable of sexual reproduction cannot be easily accommodated within the framework of the biological species concept. Species capable of self-fertilization (e.g., parasitic tapeworms and some plants) and those with mandatory sibling mating are more similar to asexual than to sexually outcrossing species (Templeton, 1989) from the viewpoint of population genetics. Species that freely hybridize (open mating systems) with one or more other species yet maintain their evolutionary identity as species also provide a serious challenge to the validity of the biological species concept. Freely hybridizing species are known from plants, insects, and vertebrates (Templeton, 1989). Another important limitation of the biological species concept concerns speciation. The most widely accepted model of speciation is the allopatric model. Generally speaking, the allopatric model entails the isolation of a subpopulation from the main population followed by the differentiation of the isolated subpopulations into new species. Historically, the notion of a correlation between geographic subdivision of populations and speciation grew out of the observation that the closest relatives tend to occupy separate but contiguous geographic areas. However, isolation mechanisms can be geographical, temporal, or behavioral. In each case, it is a barrier to reproduction and gene flow. Hence, the evolutionary forces responsible for allopatric speciation

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may not be influenced by the isolating mechanisms that are a fundamental aspect of the biological species concept. Because it is impossible to study gene flow and reproductive behavior of species known only from fossil remains, the biological species concept cannot be applied to the thousands of species known only from their fossils. In summary, the major limitations of the biological species concept are that it is inapplicable to: (1) fossil species; (2) organisms reproducing asexually or with extensive selffertilization; and (3) sexual organisms with open mating systems (species that freely hybridize).

Phylogenetic Species Concept A species is an irreducible cluster of organisms diagnostically distinct from other such clusters and containing a parental pattern of ancestry and descent (Cracraft, 1989). Phylogenetic methods focus primarily on evolutionary relationships and ancestry. Several species concepts have been proposed that are based on the methods and philosophy of phylogenetic systematics (Cracraft, 1989; Nixon and Wheeler, 1990; Nelson and Platnick, 1991). The definition presented above was proposed by Cracraft (1989), and it illustrates the historical viewpoint of phylogenetic species concepts. The identification of phylogenetic species is based on the discovery of characters unique to a population or group of populations. Morphological, molecular (e.g., allozymes, mtDNA haplotypes, nucleic acid sequences, and amino acid sequences), behavioral, and ecological characters are used to diagnose different species. Individual characters or groups of characters that are determined to be primitive or shared as a common character between groups are called plesiomorphies. Characters that are determined to be recently derived and unique to a lineage are called apomorphies. These characters are the result of the unique evolutionary history of the population(s) (e.g., species) and serve to distinguish them from other species. Phylogenetic species concepts are attractive to many biologists, in part, because they use a popular methodology (cladistics). Furthermore, the methodology for discovering phylogenetic species is clearly articulated in several of the proposed phylogenetic species concepts. In contrast to the biological species concept, phylogenetic species c­ oncepts can accommodate both asexual and sexual species, as well as species that may undergo occasional or extensive ­hybridization. Even though phylogenetic species concepts have several positive features, several drawbacks are inherent to these concepts (Mayden and Wood, 1995). One serious disadvantage is that all species are considered to be evolutionary units that are equivalent in comparative biological studies (Cracraft, 1983). However, not all species or any taxonomic level are equal, in that each has different ecological roles,

SECTION | I Introduction

functions, and evolutionary histories. Another disadvantage is that these concepts are prone to either underestimate or seriously overestimate the number of species present. Finally, it is frequently difficult to tell if a character is apomorphic or plesiomorphic, or even if that character evolved more than once. In these situations, a misunderstanding of the characters may result in obscured relationships or present false relationships.

Evolutionary Species Concept A species is a single lineage of ancestor–descendant populations that maintains its identity from other such lineages and has its own evolutionary tendencies and historical fate (Wiley, 1981). The evolutionary species concept was refined and popularized in Wiley’s book (1981) on the principles of phylogenetic systematics. The evolutionary and phylogenetic concepts are related but different. The evolutionary species concept is ontological and simply states that individual species exist as lineages of ancestor–descendant populations which maintain their own characteristic identity with its peculiar evolutionary tendencies and historical fate. It does not specify a method for identifying species, in contrast to the phylogenetic species concept, which is operational rather than ontological. The evolutionary concept is advantageous because it is applicable to living, extinct, sexual, and asexual groups. It also emphasizes that species can be maintained through gene flow and both developmental and ecological constraints. Interestingly enough, this is the concept that most biologists actually use whether or not they realize it. Decisions regarding species status are usually based on patterns of phenotypic cohesion within a group compared to phenotypic discontinuity with other groups. Although scientists rarely articulate which species concept they used when describing new species, the vast majority of species descriptions have been based on morphological comparisons. Even taxonomists that are ardent supporters of the biological species concept are essentially using the evolutionary species concept because their decisions are founded on phenotypic continuity or discontinuity and not on comparisons of gene flow among populations. The phylogenetic species concept does not contribute anything to the evolutionary concept, but the phylogenetic species concept has the drawback that there is an operational aspect to the definition. That is, it is limited to diagnosable species—an action predicated on human abilities.

Cohesion Species Concept A species is the most inclusive group of organisms having the potential for demographic and genetic exchangeability (Templeton, 1989). (Genetic exchangeability is the

Chapter | 1  Introduction to Invertebrates of Inland Waters

exchange of genes between individuals through sexual reproduction.) The cohesion species concept was proposed and described in detail by Templeton (1989). His concept emphasizes the evolutionary processes that hold evolutionary lineages (species) together through time. Because biological species are defined by gene flow, the evolutionary processes (e.g., isolating mechanisms) that curtail or promote gene flow define the boundaries of sexually reproducing species. Templeton pointed out, however, that gene flow is only one of the microevolutionary forces acting on populations and that random genetic drift, demography, and natural selection also have a role in defining evolutionary lineages because they can act on both sexual and asexual populations. Because genetic exchangeability can counter the subdividing effects of natural selection and genetic drift for sexually reproducing populations, it is crucial to know what kinds of mechanisms prevent or promote subdivision of asexual populations. Another explanation is required for the cohesion of asexual taxa, and this explanation is termed “demographic exchangeability.” The fundamental niche is defined by the genetic tolerances of individuals to some set of ecological conditions. If two individual members of an asexual population occupy the same fundamental niche, then as individuals they are demographically exchangeable. Thus, complete demographic exchangeability occurs when all individuals in a population display exactly the same range of tolerances for all relevant ecological variables. Another way of thinking about this is in the historical perspective of ancestor–descendant relationships. Consider that in a hypothetical sexual population with complete “genetic” exchangeability, any single individual organism could become the ancestor to all members of the population at some point in the future. In an asexual population with complete “demographic” exchangeability, the same result could occur. Thus, demographic exchangeability is an important mechanism for maintaining cohesion among asexual taxa. To better understand how demographic exchangeability works, we can compare it with genetic exchangeability in sexual taxa. Suppose a mutation arises in an individual of a sexual population which decreases the individual’s ability to mate with some subset of the members of its population. This means it would no longer have complete genetic exchangeability. This is obviously a weakening of the cohesion to the population for this individual and any of its descendants. Now further suppose that an asexual individual has a mutation that changes its tolerances to the ecological factors that define its fundamental niche. It will then pass on these changes to its descendants. Given that the parameters of their fundamental niche no longer exhibit complete demographic exchangeability, their cohesion to the overall population has consequently been diminished.

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Templeton’s (1989) concept of demographic exchangeability readily allows the incorporation of microevolutionary forces other than gene flow. Random genetic drift and natural selection are forces that operate as readily on asexual as sexual reproductive systems, and they act as cohesive forces for asexual species. The cohesion, evolutionary, and phylogenetic concepts each have in common their recognition of the importance of cohesion and ancestor–descendant relationships. Templeton, the author of the cohesion concept, criticized the evolutionary concept as dealing with the manifestation of cohesion and not its mechanics. Templeton greatly illuminated the mechanisms of cohesion, but his cohesion concept is really contained within the framework of the evolutionary concept. Arguably, the evolutionary species concept is the best available concept that can be used to identify species level units of biodiversity because this concept is consistent with what is known about naturally occurring units that are the result of evolutionary history (Mayden and Wood, 1995). Nevertheless, the cohesion concept articulated by ­Templeton (1989) complements the evolutionary species concept by identifying the mechanisms that are responsible for the speciation process. Templeton’s cohesion mechanisms deserve careful consideration because they play a critical role in the process of speciation and generation of biodiversity. Students are encouraged to consider Templeton’s 1989 article because it is especially relevant to the diverse reproductive strategies of aquatic species of invertebrates.

The Role of Phylogenies in Studies of Ecology and Behavior The arrangement of the present volume is based on the current classification of invertebrate phyla, and that classification rests upon an understanding of each group’s phylogenetic relationships. Phylogenies play a critical role because they provide an intellectual framework for comparative studies of morphology, behavior, and ecology at various levels. Three of the most eminent invertebrate zoologists in the modern history of biology—Charles Darwin, Willi Hennig, and Herbert H. Ross—consistently stressed the importance of integrating studies of phylogeny, ecology, and behavior (Brooks and McLennan, 1991). Although some of the reasons that a phylogenetic basis for comparative studies is needed are summarized below, the landmark book by Brooks and McLennan (1991) should be consulted for comparative studies of species. Phylogenetic analyses are perhaps the best way to discover biologically meaningful groups of species that are statistically comparable. (Many ecological questions can, of course, be studied at community and greater organizational levels using genera or even higher taxonomic

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levels.) Although statistical techniques routinely assume that samples are independently drawn from a model distribution of samples, species are not independent and are instead connected through history. Because some species are more closely related than others or arose at different times, species cannot be blindly treated as independent samples because of their historical, phylogenetic connections. The pattern of phylogeny must be accounted for, and that is why comparative studies of morphology, behavior, and organismal and population ecology are best examined in the context of a phylogenetic framework. Moreover, both the pattern and timing of speciation and character evolution must be considered because all species are a mosaic of characters that have evolved over time. A character that evolved two million years ago (mya) cannot necessarily be evaluated in the context of current ecological conditions. A phylogeny is also needed to ensure that a feature is evaluated in the correct historical context. The chief difficulty of integrating a phylogenetic approach to comparative studies is that the demand for accurate phylogenies greatly exceeds the supply!

Phylogenetic Trees From the time of Darwin until the 1950s, phylogenies were, for the most part, the result of personal opinion based on expertise and familiarity with a particular group. No repeatable methods were widely used to formulate phylogenetic hypotheses. The development of numerical taxonomy in 1957 in separate articles by Charles Michener and Peter Sneath and later extensive elaboration by Robert Sokal and Peter Sneath catapulted the enterprise of formulating phylogenetic hypotheses from the era of informed opinion into the age of empirical statistical analysis. Their basic premise was that to escape the subjectivity of the previous approach, the analysis of a large number of characters and a measure of the overall degree of similarity between pairs of species would provide a more reliable phylogenetic tree and, thus, a more stable scheme of classification. Their approach became known as “phenetics.” The fundamental problem with this approach is that the choice of characters and the method of weighing character values were based entirely upon the type of data selected for analyses. Although the method of phenogram development may have been objective, the data were still subjective. The idea of basing phylogenetic hypotheses on overall similarity was soon challenged by the emerging discipline of cladistics. Over a decade of often rancorous debate ensued between the disciples of phenetics and cladistics following the 1966 English translation of Hennig’s Phylogenetic ­Systematics. Mark Twain once wrote “A scientist will never show a kindness for a theory which he did not start himself.” Perhaps at no other time have these words rung truer than during the contest between cladists and pheneticists.

SECTION | I Introduction

In his book, Willi Hennig asserted that only one of three types of similarity could be used to formulate phylogenetic hypotheses. A character or attribute of two or more species may be similar because of convergent or parallel evolution. This type of similarity is not only devoid of phylogenetic information, it is also likely to be positively misleading because a character or an attribute of two or more species may be similar because of common descent. This type of character is termed a homologous character and exists in two forms: shared derived and shared primitive. The phenetic approach includes all three of these types of similarity. In contrast, Hennig’s cladistic methodology is based on the premise that only shared derived characters can be used to support phylogenetic relationships. Shared derived characters are also called synapomorphies and must meet the criterion of homology. Shared primitive characters (symplesiorphies) also must meet the criterion of homology, but they cannot be used to establish phylogenetic relationships. Whether a character is synapomorphic or symplesiomorphic is contingent upon its distribution on a phylogenetic tree in reference to the group being analyzed. For example, the character state “presence of hair” is a synapomorphy supporting the monophyly of mammals. In contrast, the character state “presence of hair” is a symplesiomorphy for the family Hominidae because it evolved before the group Hominidae and, thus, cannot inform us about relationships among members of the Hominidae. The widespread use of allozyme markers and mtDNA haplotypes stimulated the development of sophisticated computer algorithms to analyze these data. During the late 1980s, the development of polymerase chain reaction technology revolutionized molecular biology and enabled systematists to gather DNA sequence data for a relatively large number of species. The general approach used by many computer programs that are capable of analyzing large data sets can be modestly described as a two-step process. In the first step, all possible tree topologies are generated. In the second step, the characters are optimized along the tree branch and a criterion, such as maximum likelihood or maximum parsimony, is used to choose the best tree. An alternative approach is to generate a measure of genetic distance from the sequence data and fit the pairwise distances between all taxa to a branching diagram (tree). Because the number of trees scales up dramatically as the number of taxa under study is increased, it is rarely possible to optimize the characters for every possible tree topology. The currently available computer packages for conducting phylogenetic studies of morphological and molecular data present the user with a very large range of analytical choices. Each choice is burdened with a particular assumption about the data or how to analyze the data. The choices made can have a dramatic impact on the resulting phylogeny

Chapter | 1  Introduction to Invertebrates of Inland Waters

and the consumers of these phylogenies bear the burden of deciding whether or not the published phylogeny meets their standard of acceptability. As stated previously, it is often difficult to determine if a given character is apomorphic, plesiomorphic, or convergent. In these situations, a misunderstanding of the characters may result in obscured or even false relationships. One must use multiple data sets to determine which data provide the strongest signal and the least noise. For example, the choice of whether to analyze morphological data simultaneously with molecular data or to analyze the data separately and present the resulting phylogenies as a consensus tree remains a disputed point, but the two different approaches may produce dramatically different phylogenies or may converge. A long recognized but unresolved problem concerns the difficulty in determining the robustness of a phylogenetic tree or a subsection of a tree. A comprehensive review of the methods and controversies regarding the inference of phylogenetic trees can be found in the study by Felsenstein (2004). Our best general advice for prospective users of phylogenetic trees who are unfamiliar with the methods and controversies is caveat emptor.

USING SYSTEMATICS AND TAXONOMY TO CLASSIFY ORGANISMS Even though phylogenies and species are the result of evolution, the rules of taxonomy and hierarchical classification of groups of species are arbitrary human endeavors. Although it is vitally important for our classifications and named species to reflect biological reality, it is unrealistic to expect a perfect coherence between our classifications and the material results of more than three billion years of evolution. Thus, no phylum is exempt from an occasional reshuffling of species, renaming of constituent taxa, and recalculation of total species richness (number of species). This dynamic feature of the discipline is the expected response to the acquisition of new data and differences in data analysis and weighing. You will often encounter references to current taxonomic debates in this book and other volumes of our series. The final classification of taxa has been left to the authors of individual chapters. The editors of the fourth edition have allowed the chapter authors a great amount of leeway in this area, requesting only that they: (1) inform the reader about the principal areas of dispute; (2) only use species names for taxa previously described in the international refereed literature; and (3) follow the International Code of Zoological Nomenclature (ICZN). Species are named using a system of binominal nomenclature (names composed of two names) based initially on the eighteenth century proposal of the brilliant Swedish naturalist Carl von Linné (or Carolus Linnaeus, the Latinized form typically used in the scientific literature of his day). The conventions and rules governing

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TABLE 1.1  Principal Zoological Ranks and Suffixes Where Designated Kingdom Phylum Subphylum Superclass Class Subclass Cohort Superorder Order Superfamily Family Subfamily Tribe Subtribes Genus Subgenus Species Subspecies

the way animals are named are under the auspices of the ICZN. Table 1.1 lists the major categories for naming and classifying organisms. One can tell the hierarchical level of the higher categories below the “order” level by their suffix: superfamilies end in “-oidea,” tribes end in “-ini,” and subtribes end in “-ina.” A genus name in zoology is always a word that can be treated as a singular Latin nominative noun. The species name is a binomial (literally “two names”) composed of the genus name and a specific epithet. For example, the name for the red swamp crayfish, Procambarus clarkii (Girard, 1852), is composed of the genus (Procambarus) and the specific epithet (clarkii). The name clarkii by itself is not the species name. A specific epithet in zoology can be in any language and can be a noun or an adjective. The scientific name ends with the name of the original describer (Girard in the example above) and the year in which he described the animal. If the person’s name is enclosed in parentheses (as it is in the example above), it indicates that the species had been originally assigned to a different genus or species by that describer and was later transferred to the present taxon. This aids researchers in tracking the history of the name, and in finding the original description of that taxon. Traditionally, the species name is always shown in italics, or it is underlined if hand written. A subgenus name must be in parentheses when used in a trinomial (three names), but these are not used when it is used alone. For example, it is proper to discuss the fairy shrimp Branchinellites in a text sentence by itself, but when the full species name— Branchinella (Branchinellites) hardingi (Qadri and Baqai, 1956)—is used, the subgenus name must be in parentheses. This avoids some confusion with trinomial names involving a subspecies where no names are in parentheses and only the genus is capitalized. A subspecific epithet is used to denote a subspecies and is not separated

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by parentheses. For example, the scientific name of the diving water beetle, Thermonectus nigrofasciatus ornaticollis Aube, 1838 consists of the genus, specific epithet, and subspecific epithet. Scientific names in zoology are either in Latin or Greek, or they are Latinized from another language. An example of the latter is the cladoceran Bosmina (Sinobosmina) freyi, which was formed by Latinizing the last name of Professor David G. Frey to honor this now deceased coauthor of branchiopod chapters in previous editions of our series. Honoring colleagues by using their name to designate a new species is not uncommon among taxonomists; however, it is frowned upon by some taxonomists because it runs counter to original rules set by Linnaeus (even though he occasionally violated this rule) who recommended using a name referencing a distinguishing feature of that species to aid in identification. The final arbiter for naming taxa according to this system is the International Commission on Zoological Nomenclature, a body mandated by the International Union of Biological Sciences (IUBS) to evaluate taxonomic names proposed by scientists. The members are elected by the zoologists who attend the IUBS or other international zoological congresses. The guiding component of these rules is the principle of priority. Priority is important because it promotes stability by recognizing as valid the oldest name used for that organism; this is described in ICZN Article 23.2. In some situations, the oldest name is obscure whereas a later name is in far more widespread use. In these instances, the ICZN may overrule the principle of priority in order to maintain stability of the name. Stability is absolutely vital for communication and understanding among colleagues worldwide in the past, present, and future. The spelling of a species name remains constant among disparate languages. The International Commission on Zoological Nomenclature uses a set of agreed-upon articles (mandatory) and recommendations (non-mandatory) termed the ICZN. The commission bases its decisions on rules covering the priority of names and designation of types. As the name implies, “priority” specifies that the first name assigned in a publication is generally the official name of that taxon. The commission has plenary powers to set aside the “rightful” name of a species if by doing so the cause of science would be further advanced. For example, alteration of a familiar, long-standing name could cause a major confusion in the scientific literature. Occasionally, two distinct taxa are inadvertently assigned the same binominal name by different scientists. In that case, the species described last loses its name as a junior synonym and must be renamed. In the event that two species, or two genera, or other category are inadvertently given the same name, the later name becomes a junior homonym of the older, and that taxon must be renamed. To avoid confusion, one individual of

SECTION | I Introduction

each species is designated as the “type specimen” of that taxon and the species description is based primarily upon that specimen. Such type specimens are deposited in one of the many internationally recognized museums and made available for study. Type specimens can only be replaced if the original is inadvertently lost or destroyed, even if the type specimen turns out not to be the best representative of that species’ form. The ICZN passes judgment on names of genera and species but does not strictly control higher classifications (e.g., names of phyla and classes). Information on systematic zoology is published in various journals, such as the Bulletin of Zoological Nomenclature, Zeitschrift fur Wissenschaftliche Zoologie (Abteilung B), Zootaxa, and Bulletin Signaletique, as well as in numerous society publications and journals dealing with a broad range of subjects including classification (e.g., the Journal of Crustacean Biology). Readers interested in knowing where best to search for articles on a given taxon should consult the section on references found at the end of each chapter in our book, or use the Zoological Record, which is an online searchable data base available through many institutional libraries. The importance of correct identification and consistent approaches to classification soon becomes apparent to all people interested in learning about general biotic relationships. Vast amounts of information are cataloged in research libraries and museums that use the Linnean system of binominal classification. Once a specimen is correctly identified, a large number of pertinent references in specialized journals can be accessed, not only in systematics but also in ecology, evolution, animal behavior, agriculture, and medical sciences. Furthermore, development of the worldwide web and the Internet has opened a tremendous resource for taxonomists and other biologists. One can rapidly track down biological and taxonomic research on an incredible number of species by using computer-based information retrieval systems present in most libraries or via the worldwide web. Exercise caution when accessing web sites because the age and reliability of the information may vary greatly. In general, information that has been published in recent, refereed journals is much more reliable. Websites sponsored by scientific societies or large museums are usually good sources of information and are more easily accessed by most students.

TAXONOMIC KEYS TO INVERTEBRATES OF INLAND WATERS The following dichotomous, taxonomic keys are designed to lead the reader to appropriate chapters within volume I, and they may also be useful as an initial key for the subsequent taxonomic volumes in this series. It can be used for the larval and adult stages of aquatic insects, but it generally applies only to the adults of other taxa.

Chapter | 1  Introduction to Invertebrates of Inland Waters

11

Key to Freshwater Invertebrates 1 Multicellular, heterotrophic organisms as individuals or colonies (sometimes with symbiotic autotrophs)�������������������������������������������������������������2. 1’  Unicellular (or acellular) organisms present as individuals or colonies with nuclei irregularly arranged; heterotrophic and/or autotrophic; multiple phyla within the autotrophic protozoa phyla������������������������������������������������������������������������������������������������� Kingdom Protista. (Figure 1.1; Chapter 7.) 2(1) 2’

 adially symmetric or radially asymmetric organisms living individually or in colonies��������������������������������������������������������������������������������������3. R Individuals bilaterally symmetric�������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������4.

3(2)

 urface not porus; oral tentacles always present around a closeable mouth; colonial or single, mostly single polyp forms (primarily hydra) or rarely S medusoid form (freshwater jellyfish; adults with a single central body cavity opening to the exterior and surrounded by cellular endoderm, acellular mesoglea, and cellular ectoderm�������������������������������������������������������������������������������������������������� phylum Cnidaria). (Figure 1.2(a) and (b); Chapter 9.) Surface porus; colonial; tentacles absent; no closable orifices; without discrete organs; cellular-level (or incipient tissue-level) construction; variable, non-distinct colony shapes, including encrusting, rounded, or digitiform growth forms; skeleton of individual siliceous spicules and a collagen matrix; internal water canal system; may contain symbiotic algae; the sponges��������������������������������������������phylum Porifera. (Figure 1.3; Chapter 8.)

3’

4(2) 4

 ral region with numerous tentacles or cilia distributed around the mouth; organism never with eversible jaws and never vermiform as O adult��������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������5. Oral region with two or no tentacles, or tentacles behind the mouth������������������������������������������������������������������������������������������������������������������������������7.

5(4) 5’

 ral region with tentacle-like structures, organisms in gelatinoids or branching colonies�������������������������������������������������������������������������������������6. O Oral region ringed with cilia, muscular pharynx (mastax) with complex set of jaws; single free swimming, or semi-sessile living singly or in small colonies; wheel animals, or rotifers��������������������������������������������������������������������������������������������������������������� phylum Rotifera. (Figure 1.4; Chapter 13.)

6(5)

 ral tentacles (the lophophore) in a “U” or “horseshoe” shape around mouth; anus opens outside of lophophore; colonial animals, often in massive O colonies attached to hard surfaces; true bryozoans������������������������������������������������������������� phylum Ectoprocta (Bryozoa). (Figure 1.5(a); Chapter 16.) Both mouth and anus open within lophophore; individual (non-colonial) animals with a calyx containing a single whorl of 8–16 ciliated tentacles������������������������������������������������������������������������������������������������������������������������������������������������� phylum Entoprocta. (Figure 1.5(b); Chapter 16.)

6’

7(4) 7’

Not with the combination of characteristics described below���������������������������������������������������������������������������������������������������������������������������������8. Small (50–800  μm), spindle- or tenpin-shaped, ventrally flattened with a more or less distinct head bearing sensory cilia; cuticle usually ornamented with spines or scales of various shapes; posterior of body often formed into a furca with distal adhesive tubes; gastrotrichs (pseudocoelomates) ������������������������������������������������������������������������������������������������������������������������������������������������������������������phylum Gastrotricha. (Figure 1.6; Chapter 12.)

8(7) 8’

Anterior mouth and posterior anus present��������������������������������������������������������������������������������������������������������������������������������������������������������������9. Flattened or cylindrical, acoelomate worms with only one, ventral digestive tract opening; sometimes with evident head; turbellarian flatworms (commonly called planaria, a nonspecific, and usually incorrect name)����������������������������������������� phylum Platyhelminthes. (Figure 1.7; Chapter 10.)

9(8) 9’

 ermiform or not, eversible oral proboscis not present, although eversible jaws or other mouthparts may occur�����������������������������������������������10. V Long, flattened, unsegmented worms with an eversible proboscis; ribbon worms�����������������������������������phylum Nemertea. (Figure 1.8; Chapter 11.)

10(9) 10’

 ody not enclosed in a single, spiraled shell or in a hinged, bivalved shell; or if a bivalved shell is present, then animal has jointed legs��������11. B Soft-bodied coelomates whose viscera is covered (in freshwater species) by a single or dual (hinged), hard calcareous shell; with a ventral muscular foot; fleshy mantle covers internal organs; snails, clams, and mussels������ ­phylum Mollusca. (Figure 1.9(a) and (b); see key to classes of Mollusca; Chapters 18-19.)

11(10) S  egmented legs absent in all life stages; if jaws are present, then body with at least 20 segments�����������������������������������������������������������������������12. 11’ Adults and most larval stages with legs; if larvae without legs or prolegs (some insects), then cephalic region with paired mandibles, or eversible head, always with less than 15 body segments��������������������������������������������������������������������������������������������������������������������������������������������������������������14. 12(11) Organism vermiform, not segmented���������������������������������������������������������������������������������������������������������������������������������������������������������������������13. 12’ Organism vermiform or not, body segmented������������������������������phylum Annelida. (Figure 1.10(a–c); Chapters 20–23; see key to annelid groups.) 13(12) B  ody cylindrical, usually tapering at both ends; cuticle without cilia, often with striations, punctuations, minute bristles, etc.; 1 cm long (except family Mermithidae, 1 year) are sometimes necessary. At middle latitudes, univoltine, bivoltine, and multivoltine patterns are present in insects and other invertebrates.

Tropical Zone The tropics are traditionally defined as those latitudes where the sun is directly overhead at its zenith at least once per year, with the northernmost area being the Tropic of Cancer (23° 26″ 14.675″ north) and the southernmost being the Tropic of Capricorn (23° 26″ 14.440″ south). The implications for aquatic organisms is that seasonality is less associated with solar radiation and water temperatures and is

SECTION | I Introduction

26

instead linked to precipitation patterns. Although present below the major 30° N and S band of deserts, this area still contains some significant desert regions, such as portions of the Atacama Desert in Chile, the Saharan Desert of northern Africa, the Namib of southern Africa, and most of the Arabian Peninsula. Contrasting with these dry environments are patterns of high precipitation in much of the tropics which provides seasonality for terrestrial and aquatic organisms. Precipitation averages 200–400 cm/year with a minimum monthly average of 10 cm in tropical rainforests; most rainforests are found between 10° N and S latitudes. Tropical dry forests (mostly at 10–25° N and S latitudes) average 50–200 cm rain annually but have a pronounced dry season lasting several months with 100 years), history (1–100 years), and pulse (245,000 km2), Lake Victoria (69,485 km2), Lake Tanganyika (32,893 km2), and the Great Bear Lake (31,500 km2), or the large land-locked saline Caspian Sea (∼371,000 km2) (Downing and Duarte, 2010)—the majority of surface waters in lentic systems is present in lakes and ponds no larger than 1 km2 (Downing et al., 2006).

Geomorphology and Physicochemical Zonation of Lakes Although rivers and streams are important biologically, more than 99% of the inland surface waters are in fresh and saline lakes. These lentic ecosystems occur on every continent but are concentrated in the formerly glaciated

44

SECTION | I Introduction

FIGURE 2.19  (a) Lunzer Obersee, a karstic lake in Austria (photograph courtesy of Robert Wallace); (b) El Lago de Atitlán in Guatemala, which is Central America’s deepest lake at 320 m; this crystal clear lake is increasingly threatened by algal blooms from pollution; shown is part of the caldera and two of the surrounding volcanoes; (c) Castle Lake, a subalpine cirque (tan) lake in the Siskyou Mountains of northern California, USA, which features the longest-running (since 1958) ecological data set in North and South America; and (d) Crater Lake in Oregon, USA, whose depth of light penetration is greater than the thermocline. Photographs (b)–(d) courtesy of Sudeep Chandra.

regions of the northern hemisphere. Except for ∼20 lakes with depths greater than 400 m, most natural and humanconstructed lentic systems have average depths of less than 20 m (Wetzel, 2001; Scheffer, 2004; Moss et al., 2008). Most are geologically young, dating from the last glacial period in the case of natural lakes or from the last century for manmade lakes. Natural lakes can be formed in many ways; indeed, Hutchinson (1957) listed 76 distinct processes for their origin. Many result from catastrophic phenomena (landslide, glacial, tectonic, and volcanic activities), but some form less violently from the action of rivers (channel cutoffs, or oxbows), waves, and rock solution. Before the advent of extensive, commercial fur trapping in North America, ponds built by beavers were a pervasive feature of the headwaters of many North American streams (Naiman et al., 1986). After nearly being extirpated from the lower 48 states, and population densities of this species have risen dramatically over the last 50 years.

In fact, the increase in number of beaver ponds over the last half century has been responsible, according to some models (Naiman et al., 1991) for a 1% rise in atmospheric methane! Lentic ecosystems can be divided into several abiotic zones, primarily based on distance from shore, light penetration, and temperature change (Figure 2.20). The photic zone extends downward to the depth of 1% light penetration; all primary producers and most heterotrophic animals live within this zone. Areas below these depths are variously called the aphotic or profundal zone (the latter often in reference to benthos). The shallow, nearshore region of the photic zone where rooted macrophytes can exist is termed the littoral zone. The entire mass of open water located away from both the shore and the littoral zone is known as the limnetic or pelagic zone. The benthic zone can be further divided into somewhat arbitrary categories based on other ecosystem

Chapter | 2  Overview of Inland Water Habitats

45

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FIGURE 2.20  Biotic and abiotic zones within a lake along with a list of some representative freshwater invertebrates found within these zones.

characteristics, such as substrate type, wave action, and vegetation type. The importance of substrate type for lentic environments is similar to that described earlier for lotic ecosystems. On a seasonal basis, most lakes in temperate zones become stratified with a layer of lighter water, the epilimnion, floating over the denser hypolimnion (Figure 2.20). During at least the summer, the epilimnion is considerably warmer than the hypolimnion. These two zones are separated by a layer of rapid temperature change called the metalimnion. The boundary between the epilimnion and hypolimnion where temperature changes occur most rapidly with depth is defined as the thermocline (it is typically within the metalimnion). Because water exchange is minimal between upper and lower zones during stratification, the hypolimnion is frequently lower in oxygen, higher in nutrients, and different in pH and other chemical concentrations. When lake stratification breaks down for a short period during one or more seasons and the surface wind velocity is high enough, much or all of the water mass re-circulates in a process referred to as lake turnover. If a lake mixes completely twice a year, it is considered dimictic; if it turns over only once a year, it is called monomictic. In other lentic systems, complete mixing either rarely takes place (oligomictic) or occurs more than twice a year (polymictic). In addition to thermal zones, some lakes are chemically stratified, commonly with a heavier layer of saline water in the hypolimnion. Some of these lakes never mix completely and are termed meromictic; they circulate only in the upper zones. As a consequence, they are typically devoid

of oxygen in deeper zones, where nutrients can accumulate over time. Meromictic conditions can be natural or induced by certain pollutants.

Biotic Zonation of Lakes Lake invertebrate assemblages can be subdivided primarily into “zooplankton” (mostly protozoa, rotifers, cladocera, and copepods) within the water column and “benthos” that live in, on, or just above the bottom (often in association with macrophytes). A third, smaller group encompasses the “neuston” which lives at the air–water interface. These first two categories remain quite distinct except in three general cases. A few species, such as larvae of the phantom midge Chaoborus, migrate upward at dusk into the plankton and return to the benthos near dawn; adults are winged terrestrial individuals. Meroplankton, in contrast to the much more diverse holoplankton, spend only a portion of their life cycle in the water column. For example, chironomid midges are benthic for most of the larval period (the short-lived adults are aerial) but swim within the water column during the earliest stages of their larval existence. The Ponto-Caspian dreissenid mussels, which have invaded much of Europe and North America (e.g., Gallardo et al., 2013) have a veliger stage that is planktonic for up to 3 weeks. Although they are not true members of the plankton, many near-shore species, such as some littoral-zone water fleas (cladocerans such as Bosmina), alternate periods of resting or foraging on the bottom with intervals of swimming and foraging in the water column.

46

Reservoirs: Artificial Lakes Artificial lentic systems are extremely abundant worldwide and cover at least 335,400 km2, or about 0.2% of the total land surface area (compared to 2.8% by natural lakes; Downing and Duarte, 2010). Farm ponds between 0.001 and 0.01  km2 (∼77  million worldwide) represent about 23% of that impounded area. Indeed, humans have built so many small and large reservoirs that there has been an actual measurable, albeit minute change in the earth’s rotation from the additional heavy weight of water retained in the northern hemisphere (Chao, 1995)! Humans have increased sediment transfer through rivers to the ocean because of soil erosion (2.3  ±  0.6  ×  109 metric tons/year), but reservoirs have reduced that by 1.4  ±  0.3  ×  109 metric tons/year (Syvitski et al., 2005). Scientists estimate that over 50  ×  109 metric tons of sediment and 1–3  ×  109 metric tons of carbon are now sequestered in reservoirs (Syvitski et al., 2005), with most of this accumulated since the 1960s when the drive to build more large reservoirs started. Faced with possible increased droughts in the future and continued energy shortages, many governments are accelerating construction of storage and electricity-generating reservoirs in the false belief that they constitute totally green energy and without factoring in the cost to aquatic life. Farm ponds are very common worldwide and function much like natural, shallow lentic systems. In contrast, large reservoirs (especially those generating electricity, and thus having significant discharge are a hybrid between a lake and a river. Large reservoirs often retain something of a channel with minimal flow rates, which reduces but does not eliminate stratification. Reservoirs often have a central area and then many deep-to-shallow bays with heavy vegetation, whereas rivers have a main channel and sometimes multiple lateral channels. The quasi-lentic nature of reservoirs has led to increased planktonic components, altered benthic invertebrate dominance, and replacement of much of the normal lotic fish fauna with introduced lentic species.

Wetlands In its broader definition, the term freshwater wetlands refers to non-tidal ecosystems whose soils are saturated with water on a permanent or seasonal basis (Mitsch and ­Gosselink, 2007; Brinson et al., 2008; Mitsch et al., 2009; Keddy, 2010; van der Valk, 2012). Emergent aquatic vegetation is prominent and may alternate with annual terrestrial plants in these marshy, or palustrine, habitats. The extensive biomass of trees, herbaceous vegetation, grasses, and other plants in many wetlands is responsible for their recently recognized abilities to help filter pollutants from the environment. ­Wetlands vary from shallow, seasonally ephemeral pools (e.g., Carolina bays, prairie playas, vernal pools, pocosins, and gnammas, tinajas, or rock pools; Taylor et al.,

SECTION | I Introduction

1999; Sharitz, 2003; O’Neill and Thorp, in press) (Figure 2.21(a)–(d)) to relatively permanent vegetation-choked marshes to semi-lotic alluvial swamps and hardwood depressions (Figure 2.22) (e.g., Batzer et al., 2005; Batzer and Baldwin, 2012), some of which are protected parks and wildlife refuges such the Everglades and Okefenokee. The world’s largest wetlands are located in the South American tropics (Amazon rainforest and wetlands and the Pantanal farther south) and the north temperate zone of Russia (West Siberian Plain) (Fraser and Keddy, 2005). The origin of wetlands varies tremendously, including, for example, wind-formed depressions (playas), tectonic basins, and alluvial formations (e.g., Rains et al., 2008). Because wetlands are generally more ephemeral than other lentic ecosystems, their biotic communities are strongly influenced by hydroperiod length (Jocqué et al., 2007; Batzer and Ruhi, 2013; O’Neill and Thorp, in press). These invertebrates are affected by ecosystem permanency (e.g., length of individual hydroperiods and time period since last exposure), predictability of drying, and season of drying (if at all), as well as nutrient and light limitations (Williams, 1987; Caceres et al., 2008). The nature of the biotic community also reflects volume and depth of open water, current velocity (for alluvial wetlands connected to the river), and types of predators. For example, the presence of fish completely changes the crustacean fauna of wetlands (e.g., Colburn, 2004). Invertebrates of seasonal wetlands, such as the Carolina Bays of the southeastern U.S., have evolved various characteristics that have directly or indirectly adapted them for ecosystems that alternate between aquatic and terrestrial states (Sharitz, 2003). Wiggins et al. (1980) divided animals of temporary pools into four groups based on their adaptations for tolerating or avoiding drought and their period of recruitment. “Group 1 are year-round residents incapable of active dispersal, which avoid desiccation either as resistant stages or by burrowing into pool sediments. Group 2 are spring recruits that must oviposit on water but subsequently aestivate and overwinter in the dry basin in various stages. Group 3 are summer recruits ovipositing in the dry basin and overwintering as eggs or larvae. Group 4 are nonwintering migrants leaving the pool before the dry phase, which is spent in permanent water; they return in spring to breed.” When a temporary pool refills after a drought, the first colonizers are usually detritivores that exploit the abundant biomass of recently dead and decaying terrestrial vegetation; these species provide the animal tissue that supports the later arrival of predaceous invertebrates (Wiggins et al., 1980). Alluvial swamps are at an opposite continuum from temporary wetland pools. Contrasting with the typical picture of a swamp as a stagnant marsh, alluvial (or riverine) swamps contain many areas of slowly to rapidly moving waters; portions of these wetlands somewhat resemble a highly braided stream. Although it is true that

Chapter | 2  Overview of Inland Water Habitats

47

FIGURE 2.21  Various wetlands. (a) playa wetland in the western Great Plains, USA. Photograph courtesy of Christopher Rogers; (b) an ephemeral wetland pool (a “gnamma”) formed in a rock pool in western Australia. Photograph courtesy of Christopher Rogers; (c) small portion of the large Cheyenne Bottoms wetland in southcentral Kansas, USA. Photograph courtesy of Kelly Kindscher; (d) a Spagnum bog, Mud Pond in New Hampshire, USA. Photograph courtesy of Robert Wallace.

FIGURE 2.22  Alluvial swamp adjacent to the Savannah River in South Carolina, USA. Photograph by Thorp, J.H.

alluvial swamps in the southeastern United States such as those bordering the Savannah River between South Carolina and Georgia (Figure 2.22) are ideal habitats for alligators and poisonous snakes, they are beautiful

ecosystems with very few aerial insect pests once one has moved inward from the surrounding brush and terrestrial forest. Alluvial swamps are extremely heterogeneous, organically rich environments; oxygen does not appear to be limiting, at least in the shaded, flowing water regions. The many soft- and hard-sediment habitats of swamps (including cypress and tupelo trees in the southeastern United States) are home to a great diversity and density of aquatic invertebrates. In a study of an alluvial swamp in South Carolina, Thorp et al. (1985) showed that invertebrates rapidly colonized wood snags, reaching a rough steady state in numbers within the first week of an 8-week study. Peak densities were equivalent to nearly 18,000 animals/m2! Filter-feeding taxa were numerically dominant early but soon were subordinate to gatherer and scraper functional feeding groups. Current velocity, suspended organic matter, dispersal capacity and competition for space may be important factors affecting community structure and colonization patterns in these aquatic ecosystems.

48

Hypersaline Lakes and Pools Natural saline lakes and soda springs/pools (Figure 2.23(a) and (b)) are common worldwide but generally have neither the average size nor abundance of freshwater lakes (Eugster and Hardie, 1978; Hammer, 1984; Jellison et al., 2008). They are frequently encountered in temperate steppe and other semi-arid regions of most continents, especially Australia (Timms et al., 2009) and western North America (Hammer, 1984). For example, Australia has more salinized soils than any other continent (Brinkman, 1980) and, as a result, has more halophilic, anostracan crustaceans than any other continent (Timms et al., 2009). This radiation of species should be expected because hypo- and hypersaline ionic strengths are known to accelerate molecular evolution rates in halophilic crustaceans (Hebert et al., 2002). Saline lakes usually occur in more arid regions where closed basins promote high concentrations of salts. These lentic ecosystems can include the following: (1) saltern lakes, which are high in sodium chloride; (2) acidic lakes with large concentrations of sulfates and borates; and (3) soda lakes characterized by abundant sodium carbonates and bicarbonates. Salinities in the Great Salt Lake (Utah, USA) have been recorded as high as 200 g/l (versus an average of 35 g/l in the world’s oceans). These salinities result in high production of brine shrimp and a unique food web (Belovsky et al., 2011; Wurtsbaugh, 2014). In addition to high salinities, hypersaline lakes differ in pH, metal concentrations, and alkalinity from more typical inland freshwater lakes. The abiotic characteristics of saline lakes make them extremely rigorous environments. Cyanobacteria colonize saline lakes as do some sedges, but the submerged macrophyte flora is relatively sparse in comparison to that in freshwater lakes. Because invertebrate density and diversity are enhanced by abundant littoral plants, the depauperate flora

SECTION | I Introduction

of saline lakes and the lack of certain other microhabitats undoubtedly depress the invertebrate fauna of briny pools. Few invertebrates of inland waters can regulate their osmotic and ionic concentrations in a hypertonic environment. As a result, hypersaline environments contain very few species, although their densities may be high. For example, the only permanent metazoa in Mono Lake (California, USA) are the brine shrimp Artemia monica Verrill, 1869, and the brine fly Ephydra gracilia Packard, 1871 (Herbst, 1999). High densities of brine shrimp in Mono Lake are crucial to survival of migratory eared grebes, Podiceps nigricollis Brehm 1831, which molt flight feathers while at Mono Lake; these birds depend on the crustaceans as their sole food source while in the lake (Cooper et al., 1984). Until courts in 1994 ordered a halt to the massive removal of inflowing freshwater from the mountains for transport to large coastal cities, this simple food chain was seriously threatened by rising salinities and the impending demise of the brine shrimp. When water diversions were stopped for several years and the salinity declined, the zooplankton community expanded to include rotifers— demonstrating the resiliency of the ecosystem and the importance of water-management policies (Jellison et al., 2001). Other saline lakes with variable but more persistent lower salinities typically support more species diversity, including the rotifer Brachionus plicatilis (Müller, 1786), numerous fairy shrimp, clam shrimp, copepods, ostracods, and several families of flies, beetles, and true bugs. As saline lakes become less salty, the diversity and density of their flora and fauna are enhanced. For example, the meromictic Waldsea Lake of Saskatchewan, which is one of the well-studied saline (≥3 ppt total dissolved solids) inland lakes in Canada, has a much higher species diversity than the much saltier Mono Lake. It contains two macrophyte species, some filamentous algae, a sparse phytoplankton population, and a dense but rather uniform population of zooplankton,

FIGURE 2.23  (a) The saline Mono Lake in arid central California; calcium carbonate deposits with the lake (tufa, as shown in the foreground) reflect the lakes chemical composition and hypersalinity. Photograph by Thorp, J.H; and (b) CaSO4 soda spring near Cuatro Cienegas, Mexico. Photograph courtesy of Robert Wallace.

Chapter | 2  Overview of Inland Water Habitats

dominated by the calanoid copepod Diaptomus connexus Light 1938, the rotifers Hexarthra fennica (Levander, 1892) and B. plicatilis, and the water flea Daphnia similis Claus 1876 (Hammer, 1984). The littoral zone supports high numbers of relatively few species, including include several species of true bugs, nine genera of beetles, midges (mostly Cricotopus), a few caddisflies, the damselfly Enallagma clausum Morse, 1895, one snail species (Lymnaea stagnalis (L., 1758)), and several genera of crustaceans. Although Waldsea Lake is certainly diverse compared to the hypersaline Mono Lake, its community is depauperate in comparison to most permanent, freshwater lentic environments. Some salt lakes that are present at least moderately close to the sea coast have viable land-locked populations of “marine” barnacles, especially Amphibalanus improvisus Darwin, 1854 (see Chapter 29). An example is the Salton Sea (­California, USA), which is below sea level and was created by an 18-month flood of the Colorado River beginning in 1905. It has a thriving population of Balanus amphitrite saltonensis, Rogers, 1949 in its saline waters, which currently average 44 g/l (compared to the average ocean value of 35 g/l). There is a highly variable community of zooplankton as a result of fluctuating salinities and high sulfate concentrations (Tiffany et al., 2002, 2007). The benthos includes dense populations of polychaetes that seasonally recycle phosphorus from the sediments, leading to high primary production and widespread anoxia in the deep-water sediments (Swan et al., 2007).

Phytotelmata Phytotelmata (singular “phytotelma”) are water bodies enclosed by a terrestrial plant; they occur on all continents except Antarctica (e.g., Derraik, 2009). They include living plant reservoirs (e.g., bromeliad tanks, pitcher plants (Figure 2.24) and other carnivorous plants holding water,

FIGURE 2.24  Some freshwater invertebrates colonize phytotelmata, such as these pitcher plants, Sararcinia purpurea, which are found in boggy areas. Photograph courtesy of Robert Wallace.

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water-filled tree hollows), water collected at bamboo internodes and axial water (bases of leaves, petals, and bracts) (Kitching, 2009), and, in some classifications, water collecting in dead plant material separate from the living plant (Greeney, 2001). Water-filled, artificial containers (e.g., automobile tires, cans, bird baths) can mimic the conditions of some natural phytotelmata. Eukaryotic biodiversity in phytotelmata is mostly composed of very small non-flying invertebrates, such as protozoa, rotifers, gastrotrichs, and nematodes (e.g., Maguire, 1971; Dunthorn et al., 2012), mosquitoes and other flying insects with a short life history, and tadpoles of tree frogs and other small frogs (e.g., Ryan and Barry, 2011). Some examples of complex life histories of juvenile Jamaican crab species can be demonstrated by using small containers to replace natural plant and shell phytotelmata (Diesel and Horst, 1995; Diesel, 1997; Diesel and Schubart, 2007). Some mosquito species are found exclusively in one species of phytotelmata, whereas others occur in multiple species but usually of the same general form (Albicócco et al., 2011). Those from tree holes are also well-known dwellers of artificial containers and ground water habitats, such as Ochlerotatus sierrensis. Research on phytotelmata communities has accelerated because of concerns about diseases transmitted by mosquitoes.

General Human Impacts on Lentic Communities Unlike the situation described earlier for streams and rivers, pollution in lakes and wetlands is often easily visible through effects on water quality (e.g., taste and odor problems in municipal drinking supplies), primary productivity (e.g., development of eutrophic conditions, especially lake algal scum), and fish kills. Those conditions can easily result in immediate demands by the public to rectify the deteriorating conditions. Determining the health of the system without obvious conditions such as algal scum, however, can be a more difficult task. For example, salt accumulation in the bottom of a lake from factory effluents can change a dimictic lake into one that only partially turns over (meromictic), and this may not be immediately obvious. On the other hand, the lack of strict environmental laws, an apathetic or resigned public, and/or an unresponsive government can quickly cause environmental deterioration of a closed system like a lake or wetland. Although lakes are valued by most people, for many years the public has ignored or labeled as undesirable the vast acreage of wetlands, ephemeral ponds, and swamps in North America, considering them breeding grounds for mosquitoes and snakes. Many have been drained, filled, and bulldozed for housing and commercial development without regard to their intrinsic value to wildlife and the environment. In the United States, about half of the original

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wetlands (∼220 million in the 1600s) have been drained and converted to other uses. “The increase in flood damages, drought damages, and the declining bird populations are, in part, the result of wetlands degradation and destruction” (http://water.epa.gov/type/wetlands/vital_status.cfm; accessed April 2014). Following public recognition of the importance of losing large numbers of wetlands, strict government environmental laws to protect wetlands have been passed in some countries; however, these are still being debated and variously interpreted nationally and internationally (e.g., Brinson et al., 2008). Unfortunately, most laws are not designed to protect ephemeral wetlands, although some environmental organizations (e.g., Playa Lakes Joint Venture, in the USA) and waterfowl hunting organizations (e.g., Ducks Unlimited in the USA) are working to extend greater protection to these wetlands. Reservoirs, although they are not natural lentic systems, are critical to the survival of modern civilization in terms of water storage, flood control, and sometimes electric power generation. In many countries, poor land management practices (e.g., removal of riparian cover and poor agricultural practices) have led to surprisingly rapid filling of reservoirs with sediments, thereby reducing the water storage capacity and useful life of the reservoirs (McCully, 2001; Hargrove, 2008). This has become a serious problem around the world, especially in some soil types and in areas with extensive row-crop agriculture where runoff into streams is extensive, such as the U.S. Great Plains.

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SECTION | I Introduction

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Chapter | 2  Overview of Inland Water Habitats

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SECTION | I Introduction

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Chapter | 2  Overview of Inland Water Habitats

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SECTION | I Introduction

Wurtsbaugh, W.A., 2014. The great salt Lake ecosystem (Utah, USA): long term data and a structural equation approach: comment. Ecosphere 5. art36. http://dx.doi.org/10.1890/ES13-00335.1. Yoshida, K., Hoshikawa, K., Wada, T., Yusa, Y., 2013. Patterns of density dependence in growth, reproduction and survival in the invasive freshwater snail Pomacea canaliculata in Japanese rice fields. Freshw. Biol. 58, 2065–2073.

Chapter 3

Collecting, Preserving, and Culturing Invertebrates D. Christopher Rogers Kansas Biological Survey and Biodiversity Institute, University of Kansas, Lawrence, KS, USA

James H. Thorp Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, Lawrence, KS, USA

Chapter Outline Introduction57 Collecting Legally 57 Collecting Safely 58 Collecting Conservatively 58 Collecting and Sampling 58 Obtaining Benthic Invertebrates 59 Obtaining Planktonic Invertebrates 60

INTRODUCTION1 Aquatic invertebrates are collected for various reasons, including fishing bait, human use (such as crayfish for etouffee, or mussels for pearls and buttons), scientific studies, and educational purposes. In past decades there have been an increasing number of programs sampling aquatic invertebrates as part of environmental baseline and impact analyses, particularly as relates to water quality. These programs have been implemented mostly in North America, Europe, and Australia, with new programs emerging in other parts of the world. Typically these programs (biological monitoring, or bioassessment) have specific field survey and sample collecting protocols that are designed to collect data to find answers to specific ecological questions.

Collecting Legally Before collecting any organism, make certain that you have legal access to the selected sites; trespassing is punishable 1 Some of the material in this chapter was extracted from: Thorp, J.H., Rogers, D.C., 2011. Field Guide to Freshwater Invertebrates of North America. Elsevier, Boston, MA, 274 p.

Record-Keeping61 Preserving and Fixing Specimens 61 Culturing Invertebrates 62

by law. Access may be limited by law to some types of public lands, especially watersheds of municipal drinking water supplies, which are generally protected for public health. It is generally illegal to collect in state and province parks, national parks and monuments, and wildlife refuges, without proper permits. Most countries require collecting permits for foreigners and occasionally residents. Similarly, most countries require permits to import or export organisms across their borders. International law governs the movement of specific taxa across borders, and one should consult the most recent Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES) before collecting in foreign countries (http://www.cites. org/). In many nations and states, a fishing license is required to collect most decapod species, and bag limits may apply. Most collecting limitations were instituted to protect local natural resources and to control the spread of potentially harmful and invasive species. It is always illegal to collect threatened and endangered species without the required permits, and copies of those permits are generally required if the specimens are to be moved across international borders. Many molluscs and crustaceans are legally protected in many nations.

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00003-6 Copyright © 2015 Elsevier Inc. All rights reserved.

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Prosecution may result even from inadvertently disturbing a protected species or its habitat. Be informed about the legal conditions of any area where you want to explore. Different counties, states, provinces, and nations have different laws, often for different reasons; do not assume the laws of one area apply to another. The more you know, the better protected you will be from inadvertent, unlawful activities and potential fines or imprisonment. Please be considerate of certain natural features, such as caves, waterfalls, rocks, or valleys, as some may have local cultural significance. Similarly, if you turn over a rock, return it to its original position before leaving, to maintain habitat conditions for the resident biota.

Collecting Safely Exercise normal, rational caution around aquatic habitats, as any activity near or in water can be potentially hazardous. When standing in water, maintain a stance with your feet well separated for more stability; felt-bottomed boots or waders help sustain a grip on slippery substrates. Always wear a life vest when using a boat, canoe, or kayak or when wearing chest waders. Such personal flotation devices are also good for working in or near swift, cold water. Swift currents can draw even the strongest swimmer under water. Flowing waters may undercut banks, making them liable to collapse. Flash floods occur during monsoons or where distant rainfall is funneled into channelized streams, rapidly changing stream depth from a few centimeters to a few meters in a matter of minutes. Muddy shorelines and lake bottoms can be more easily traversed in “coot boots” or other stabilizing footwear, which will help prevent you from becoming mired. Quicksand may also trap the unwary; if immersed in quicksand, the best strategy for escape is to lie out as flat as possible and swim to a firm substrate. Caves can have dramatically different temperatures inside than out, and one should only enter caves bringing adequate lights and protective gear. It is particularly dangerous to sample in caves where water flows in rather than out of the cavern, as you will have less warning of rising waters that could trap you below ground. Learn in advance and avoid the potential biological hazards associated with aquatic habitats in your collecting area. For example, in much of the American tropics where snails of the genus Biomphalaria occur, avoid touching water with bare skin. These molluscs are intermediate hosts of human liver fluke. Other parts of the world have similar hazards. Various invertebrate species that may present human health concerns include schistosomes (i.e., blood flukes, swimmer’s itch, duck itch, liver flukes), leeches, mites, ticks, and chiggers, as well as various biting flies (e.g., Simuliidae, Culicidae, Ceratopogonidae, Phlebotomidae, Glossinidae, Tabanidae). Biting flies may transmit a variety of diseases including various forms of viral encephalitis (such as West

SECTION | I Introduction

Nile Virus), sleeping sickness (trypanosomiasis), river blindness (onchocerciasis), yellow fever, dengue fever, and malaria. Other diseases and disease causing organisms in inland waters include cholera, amebiasis, Escherichia coli, Giardia, Legionella, and hepatitis. Some vertebrate organisms that occur in or near inland water bodies can be dangerous, such as sting rays, sharks (near coastal areas and some deep rivers), snakes, monitor lizards, snapping turtles, crocodilians, emu, cassowaries, water buffalo, pigs, hippos, elephants, large cats, bears, and humans. These lists of potential hazards are by no means exhaustive. The cardinal rule, therefore, is to be careful and cautious and not take risks.

Collecting Conservatively As a responsible scientist or student of nature, it is important to collect only the minimum number of specimens needed for your particular project. For purposes of environmental impact analyses, the number of samples and specimens are determined statistically. In studies of systematics, it is vital to collect enough invertebrates to identify new taxa by establishing patterns of morphology and genetics; but at the same time, one should avoid over-collecting living specimens, as one might do with inanimate stamps. To illustrate this problem with a marine example, invertebrate scientists visiting the Duke University Marine Lab in Beaufort, North Carolina in the 1970s collected so many of the very unusual, intertidal parchment worms (Chaetopterus variopedatus) to add to their personal collections that they essentially wiped out the species locally (James H. Thorp, personal observation). Finally, for educational purposes, some preserved samples will be required; but in other cases, only a few specimens are necessary, and these can sometimes be returned alive to their original habitats. In the latter case, be careful to avoid collecting rare species in the first place.

COLLECTING AND SAMPLING There are two basic ways to gather aquatic invertebrates: qualitative collecting and quantitative sampling. Collecting consists of qualitatively gathering specimens of one or more taxa. Specimens may be collected for general faunal inventories, for student collections, or for material to use in the laboratory. Conversely, quantitative sampling is the gathering of one or more invertebrate taxa according to a statistically replicable method. Quantitative sampling is designed to gather specific, repeatable data in order to answer one or more questions posed as part of a scientific study. Any of the following methods can be used either qualitatively or quantitatively. There are myriad ways of quantitatively sampling, and new methods are developed every day. The method

Chapter | 3  Collecting, Preserving, and Culturing Invertebrates

employed should be defined by the study question and the hypothesis being tested. It is not possible to describe all aquatic invertebrate sampling methods here, but a few basic methods are outlined, which can be adapted for different study scenarios. Many of these methods lend themselves to statistical replicability; however, any discussion of statistical field or laboratory methods is beyond the scope of this chapter.

Obtaining Benthic Invertebrates There are numerous ways to collect aquatic invertebrates, using traps, dredges, samplers, nets, and grabs. In this section general collecting methods and apparatus are discussed. The subsequent specific chapters of each taxonomic group will provide more detailed and specific collecting information. A great variety of invertebrates can typically be found on, among, and under rocks, under overhanging banks, among aquatic plants, on and in submerged or floating woody debris, and in log jams and leaf packs. Some invertebrates live in sand bars (many molluscs and odonates) or even in loose shifting sands in some stream bottoms (some ephemeropterans, some trichopterans). Exposed hard surfaces, such as dock pilings, boat bottoms, dam faces, and rocks, provide attachment sites for sponges and bryozoans as well as homes for trichopterans, dipterans, and many molluscs. Current velocities, temperatures, and oxygen conditions influence the distribution of many species (see Chapters 2 and 4). In flowing waters, the most oxygenated portions are the areas just below riffles, and in main channel vs side channel habitats. Because oxygen levels are high, more invertebrates congregate here than in other parts of a creek or river. Many predatory fish wait below riffles for invertebrates that are carried out by the current. For these and other reasons, invertebrate abundance peaks in side channels of multi-channeled rivers. In lakes and wetlands, invertebrate distribution can be affected by wave action, bottom composition, oxygen levels, light intensities, and presence of vertebrate predators. Many of the non-pelagic invertebrates congregate within the littoral zone among aquatic vegetation. The prevalence of easily collectable invertebrates varies with time of day. Many invertebrates, such as crayfish, shrimp, and some snails, are nocturnal. Many crayfish live in deep burrows or in the deepest portions of rivers during the day and only emerge into shallow areas at night. In many cases this seems related to predator avoidance. The basic collecting equipment for benthic habitats is a net and shallow bucket or pan. The pan or bucket should be partially filled with water from the collecting site. One can easily use a hand or a soft brush to remove seen and unseen invertebrates from the rocks, wood, or other debris from the habitat into the bucket or pan. Aquatic plants or handfuls of gravel or sand may be placed into the bucket or pan and the

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invertebrates will soon become visible as they move about. White plastic or enamel containers are usually best, as the invertebrates stand out against the bright surface. The best standard net is a dip net with a broad aperture (at least 45 cm wide and broad) and a long, sturdy, handle (1.5–2 m). The net bag should have a mesh size of 500 μm or less. Sweep the net through the lentic water column and across or through vegetation, woody debris, and rocks. The net can be inserted into the water and drawn toward shore if wading is not an option in shallow-margined ponds and lakes. The net is used to sweep under overhanging vegetation or banks, or can be used in the main current of lotic systems. Hold the net by the handle perpendicular to the substrate in the current, with the bag end on the substrate, facing upstream. The substrate upstream of the net is then disturbed by hand, and rocks, plants, and debris cleaned or shaken into the water; the current then carries material into the open net bag. Dump the contents of the bag into the pan or bucket and sort the invertebrates from the remaining material. A similar result can be achieved with a kick net, which is a 1 m square sheet of net mesh stretched between two 1.5 m long poles. Most decapods such as shrimp, crayfish, and crabs generally are captured with any good commercial crayfish trap using appropriate bait; however, most shrimp genera in the Atyidae and Xiphocaridae are periphyton feeders and will not come to bait. Mussels can be turned up by pulling a rake gently across the gravel bottom of a shallow stream or can be collected by hand, although sometimes snorkeling or scuba gear are required. An aquatic light trap is an effective trap for capturing positively phototaxic organisms, such as many insects and crustaceans, especially predatory beetles, mysids, and zooplankton. Commercial aquatic light traps are available, but also are easily constructed from a small bucket with a lid (bearing perforations large enough to admit the target organisms), a rope tether, a rock (heavy enough to keep the bucket from floating away), and a battery-operated glow stick. This trap should be deployed at dusk or just after dark. The activated glow stick should be hooked under the rock so that it will stay on the bottom of the bucket. Be sure to retrieve the trap before dawn, as specimens may swim out when they see the morning light. Littoral benthic substrate is typically sampled using a grab or a suction sampler. Among the former are Ekman, Kellen, Allen, Ponar, and Macan. These are basically an open, weighted box with spring-loaded jaws that close the box on an area of substrate. For deep water, the larger Ponar and Ekman’s grabs are dropped by a tether, and a weighted messenger slides down and trips the jaws shut. A suction sampler is similar but has a compressed rubber bulb that is released at the substrate surface, expands quickly, and sucks up material beneath it.

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You can also sample benthic invertebrates using artificial substrates, such as cages with rocks, sticks, or leaf pack, or a stack of uniformly separate plates (e.g., HesterDendy multiplate samplers). These are left submerged for colonization over a set period in quantitative sampling. One semi-quantitative method for small habitats, such as small springs or seeps, is a timed collection method, where aquatic invertebrates are collected for a specified time period to make one single sample. This can be repeated as many times as required for statistical replication. Many decapods leave the main water source as they age. Many crabs move at night from rivers and ponds to the surrounding uplands, or even climb trees to feed. Many crayfish and crabs burrow in banks and shorelines, sometimes with their burrow entrances above the water line, but with the bottom of the burrow below ground water. Burrowing species can be trapped, collected by hand, dug from their burrows, or suctioned out using a yabby pump. Emergence traps are large, tent-like canopies that one can build over a section of stream or shoreline. As adult insects emerge from the water, they are captured in the canopy. Black lights or mercury vapor lights with a bright white background sheet may be employed to attract adult aquatic insects that fly at night. Large numbers of ephemeropterans, plecopterans, trichopterans, hemipterans, and coleopterans may be collected using lights. A beating sheet is a square of light-colored canvas spread on an “x” shaped frame. The beating sheet is held by one corner, horizontally under the branches of vegetation along the shoreline. A stick or net handle is used to gently tap the branches, causing adult aquatic insects to drop off onto the beating sheet, where they can be collected. Malaise traps and flight intercept traps are similar to an emergence trap in that they capture flying insects. Both basically form a barrier that insects will fly into and be captured. The malaise trap is a tent like affair, with mesh walls (to allow penetration by air and light) and guides along the margins. An insect flies into the mesh and crawls along the surface toward the edges. Canvas guides along the edges direct the insect toward the top and middle of the barrier. At the top is a receptacle to capture the insects. Flight intercept traps are generally a large sheet of plexiglass held upright between two posts. The bottom edge of the plexiglass stands in a trough or gutter filled with soapy water. Flying insects encounter the plexiglass and drop into the soapy water, where they drown and are preserved temporarily until they can be removed.

Obtaining Planktonic Invertebrates The methods used to obtain planktonic invertebrates depend partially on their size, lifestyle, and time of day. Inland water invertebrates classified as planktonic include many “holoplankton” (those living their entire lives above the

SECTION | I Introduction

bottom and unattached to any substrate), a few “meroplankton” (usually larvae or nymphs that are benthic as adults), and some species that live on or near the bottom during the day but enter the plankton at night. Invertebrate holoplankton primarily consist of minute rotifers and larger crustaceans (mostly nauplii through adult copepods, and small (cladocera) and large (e.g., fairy shrimp) branchiopods). Meroplankton, which are abundant in the ocean, are relatively uncommon in freshwaters. However, they include a few insects (e.g., the phantom midge Chaoborus), larval mites (Arachnida), and the veligers of some bivalve molluscs (e.g., dreissenid mussels). Larval phantom midges live near the bottom of lakes during the day but migrate upwards into the epilimnion at night. Many harpacticoid copepods are also generally found on or near the bottom during the day but migrate upwards somewhat during the evening. Finally, rotifers and microcrustaceans living in the open water, pelagic zone of lakes are true plankton; whereas those inhabiting the vegetated littoral zone often are primarily benthic on upright plant stems and leaves. Some of the littoral forms are attached, but many others spend some time swimming among the vegetation, especially at night. Some evidence of migration from littoral zone to open water areas has been noted in littoral species, possibly because they are then less susceptible to visually hunting fish planktivores. The location of plankton and the equipment used to collect them depend in part on the type of aquatic ecosystem. Plankton in “permanent” lakes occur either in open water or littoral areas, with major species differences. In contrast, those in seasonal and moderately permanent wetlands show little migration and are usually widely distributed throughout the ecosystem. Copepods, cladocera, and rotifers in small streams are primarily benthic, even though they may be closely related to true planktonic forms. As the stream deepens and becomes less turbulent, more true planktonic forms appear; these are then termed potamoplankton (potamos is an anglicized form of the Greek word for river). Potamoplankton of large rivers are most abundant and diverse in the slower-moving, side channels. Rotifers are the dominant plankton in the main channels of rivers, but smaller copepods and cladocera also occur there. To collect true zooplankton, use either a plankton net (open or closeable), plankton trap (e.g., Schindler-Patalis plankton trap), water bottle sampler (vertical or horizontal; appropriate only for rotifers), or high-volume water pump and mesh. Specific volumes of water are collected at known depths using one of the three latter methods and then passed through a mesh (size related to group sought) to produce a quantitative sample. Plankton nets allow the investigators to obtain a species list of organism but are at most a semi-quantitative sampling method because the investigator rarely knows the exact amount of water sampled. The mesh size employed for all three methods will vary depending on the taxonomic group being investigated. They can be

Chapter | 3  Collecting, Preserving, and Culturing Invertebrates

obtained with mesh sizes ranging from 10 to 1000 μm (a few are standard size and thereby cheaper). Collect crustaceans using a mesh size ranging from 80 to 250 μm (smaller for copepod nauplii), and obtain rotifers by employing a final mesh size of at least 80 μm and preferably 20 μm. High-speed water pumps are an effective and relative inexpensive way of collecting zooplankton from known depths. The critical requirement is to have a pump velocity and tube diameter large enough to prevent copepods from fleeing the lower tube opening. Pump the water into a bucket of known volume and then gently pour this water through a mesh sieve (you can make one with PVC pipe and purchased mesh). If you pump directly through the sieve, some plankton will be damaged and even forced through the sieve. If the volume of water to be filtered is large or if the water is especially sediment-laden, you will need to periodically rinse the contents into a separate collecting bottle or plastic cup. Littoral zone plankton are more difficult to sample quantitatively. They can be collected qualitatively with a plankton net (which will miss some attached species), or you can examine some of the vegetation in the lab shortly after collecting it. In the latter case, add the plant material to a bucket with water while you are in the field. Once in the lab, place some of the vegetation in a dish with water and examine the material for live crustaceans and rotifers under a dissecting microscope. Once located, the organisms can be selected and then preserved before later detailed identification. Some rotifers alter their shape when placed in 75% ETOH, and this hinders later identification. Consequently, various investigators recommend killing them rapidly (to prevent distortion) by immersing the sieve and contents first in 95% EtOH or in hot water before preserving the sample in 75% EtOH. Unlike samples of benthic invertebrates, the EtOH in these samples does not need to be changed after 24 h. After returning the samples to the laboratory, you may need to split the sample into smaller volumes. This can be accomplished quantitatively with a Folsom or Motodo Plankton Splitter. Rotifers may need to be “settled” in special chambers to concentrate these small animals on a slide for later identification and counting. Identification of zooplankton requires either a dissecting microscope or a compound microscope, depending on the size of the organism and whether the investigator needs to dissect and identify appendages or other structures. Details on these processes can be found later in this book within the appropriate taxonomic chapter.

RECORD-KEEPING Specimen records are absolutely vital. Material lacking basic collecting data is typically worthless for scientific purposes. This standard information allows anyone to resample

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the same place. In this way, the data allow users to understand the organism’s distribution, habits, habitat, and activity periods. A log or field notes recording a day’s collecting should start with the date, a description of where the collecting took place, the time and weather, a habitat description, who was present, and a list of organisms observed and collected. The locality data should be in the following format: Country State, Province, or Territory County, Parish, or Regional District Locality description In this way, the data record goes from the larger to smaller area. This helps other researchers using the original data to find the collecting site easily and in a logical format (see example below). GPS or latitude/longitude coordinates can be added to this information to make the local site easier to find. These same data should be placed on collection labels for individual specimens, followed by the date of collection in the following format: day/month/year. Specimen labels should be concise and contain sufficient data so the sample can stand alone from the field notes. For example: USA: CA: Shasta County: Poison Lake at intersection of railroad tracks & Pittville Road, north of Highway 44. 2JUN2009. D.C. Rogers. This abbreviated format uses the least characters and thereby the least space. This is important when there is so little space on the specimen label for writing, and little space inside the specimen vial. Following the date, a catalog number correlating the specimen with other records or a collection catalog is used by some, and may be important to account for all specimens from a single site or study.

PRESERVING AND FIXING SPECIMENS The best preservative is ethyl alcohol (sometimes abbreviated as ethanol or EtOH), diluted no more than 70% with water. Isopropyl alcohol (isopropynol) will work, but ethyl alcohol is required for any later genetic studies. Specimens needed for genetic studies should be preserved in 90% or stronger ethyl alcohol. Glycerin can also be used (and may prevent desiccation if a container lid is loose), but it tends to make most arthropod specimens brittle. Formalin is a fixative, not a preservative, and in general it is not recommended for use due to its carcinogenic properties. Formalin also fixes proteins, making it impossible to get genetic data from your specimens. Some invertebrate groups have specialized preservation requirements. For example, ostracods and copepods are best kept in 5% formalin or glycerin, and nemerteans should be placed in Bouin’s formula. Lepidopteran larvae are best placed in KAAD in order to see the required characters

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needed for identification. Specific guidance for proper preservation, fixation, and preparation for each group is supplied in the appropriate chapters in this volume. When specimens are particularly soft bodied or if the specimen(s) volume is equal to or more than one-third the volume of alcohol, the preservative must be decanted and replaced after 24 h. If there are large amounts of vegetative material (e.g., leaves, algae, or woody debris), the preservative should be changed after 6 h and again after 24 h to prevent later decomposition (rot) of the specimens. The large amount of organic material can absorb the preservative, while simultaneously releasing water and diluting the preservative to a concentration that will not prevent decomposition. Museum-quality specimens, study specimens, and voucher specimens should be placed in a glass vial with alcohol filled nearly to the top. Always include a label (with the required data described above) inside the vial, written with a pencil or an alcohol-proof pen. Laser jet printers can also be used to make labels; however, once the labels are printed, they should be heated in a microwave oven for a few minutes. Laser jet printers place the characters on the paper using wax, which will dissolve in alcohol, thus causing all the characters to fall off the paper and lie in a jumble in the bottom of the vial after a few months.

CULTURING INVERTEBRATES There are many specialized methods for culturing aquatic inland invertebrates, and those methods are described in subsequent chapters in this volume pertaining to the specific organismal groups. Consequently, only general culture methods are described here. Most aquatic invertebrates can be kept in a tank set up as an aquarium (aquatic habitat) or paludarium (aquatic and terrestrial habitats), provided that the basic needs of dissolved oxygen, current, food, refuge, and waste removal are met. Paludaria are often important for amphibious species (many crabs and some insect and snails), especially for larval insects that need drier substrate in which to pupate, or for those insects and snails that deposit their eggs out of water. Large aquaria with an abundance of live plants are generally best for most inland aquatic invertebrates. The plants help oxygenate the water, absorb animal metabolic wastes, and keep the water clear. The best way to set up an aquarium for plants is to fill the bottom couple of centimeters of a dry, empty glass tank with a rich potting soil. Cover this soil with at least 4 cm of fine, clean sand and then add water by pouring it onto a rock or plate so that the sand is not disturbed. Add plants by pushing their roots through the sand into the soil. The sand keeps the soil from fouling the water. This approach is generally not recommended for strong burrowing taxa, such as decapods or larger mussels, as their activities will release the soil from beneath the sand. Larger crustaceans also tend to eat most plants.

SECTION | I Introduction

Floating (Lemna, Pistia, Salvinia, Limnobium, Riccia, Azolla, Anacharis) or epiphytic (Taxiphyllum, Vesicularia) plants will absorb large amounts of nitrate, nitrite, and ammonia from the water and help keep algae under control through direct competition. Rooted aquatic plants will filter the water to a lesser extent, but will also help oxygenate the water. Filtration is generally needed; but as most filtration systems rely on strong suction, sponge filters are probably safest for most swimming invertebrates. Benthic and sessile taxa often benefit from strong currents generated by power heads or other filtration pump devices. Filtration should be accomplished by an outside filter or a sponge filter. We strongly discourage use of “under gravel” filters because they promote strong root growth, while stems and leaves grow and mature little. It is generally best to exchange one-fourth of the aquarium water volume each week with rainwater, distilled or R/O (reverse osmosis) water, or water from the habitat where the specimens originated. The larger and better functioning the aquarium, the less frequently the water will need changing. Tap water should not be used unless it has been dechlorinated. The latter can be accomplished by vigorously bubbling air through the water for one or more days. Most aquatic invertebrates obtain oxygen that is dissolved in the surrounding water through gills or across the body surface. Many insects and some snails collect air bubbles from the meniscus. The basic rule is that the cooler the water, the more dissolved oxygen it contains. Unfortunately, it is difficult to keep aquarium temperatures cool enough to meet the dissolved oxygen requirements of most flowing-water invertebrates. You can increase oxygen levels by employing one or more air pumps and air stones. Providing the appropriate type and amount of food (but not too much) is often a major challenge. Predatory species need other invertebrates to feed upon, and their remains may foul the tank. Most grazing species scrape algae attached to rocks and other hard surfaces, or they browse on vascular plants. Nontoxic tree leaves (e.g., Populus, Alnus, Falcataria) that provide food for a variety of grazing and shredding invertebrates can be collected and stored in a refrigerator for many months. Add one or more leaves to an aquarium or paludarium as needed to provide food for crustaceans, insects, and snails. Refugia are equally important. Many snails and flatworms are nocturnal and need dark daytime hiding places. In a community-type aquarium, potential prey animals need to be able to hide from predators or else the aquarium becomes an abattoir. Some organisms, such as crayfish, require individual shelters to avoid damage from frequent fighting. Finally, many species will escape unless the tank has a tight-fitting lid. Some insects, crabs, and crayfish are adept at climbing air-line tubing or power cords. Similarly, snails can climb most surfaces, and most flying insects can fly directly from the water surface.

Chapter 4

Functional Relationships of Freshwater Invertebrates James H. Thorp Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, Lawrence, KS, USA

Chapter Outline Introduction65 Distribution in Space 65 Physical and Ecological Characteristics of Lotic Ecosystems66 Longitudinal Patterns in Habitat and Organisms 66 Hydrogeomorphic Patterns 67 Lateral Riverscape Patterns of Organisms in Medium to Large Rivers 68 Physical and Ecological Characteristics of Lentic Ecosystems68 Temporal Factors 68 Spatially Distributed Habitat Features 69 Aquatic–Terrestrial Ecotones 70 Acquiring Energy 71 Biodiversity Traits and Functional Feeding Groups 71 Traits in General 71

INTRODUCTION The task of summarizing the functional ecology of inland water invertebrates of the world is only a little less daunting than composing a whole book on the ecology of all invertebrates (terrestrial, freshwater, and marine) or a college text on general ecology—and I am limited to a single chapter! In tackling this task, one must limit the number of topics, cover only selected taxa and write at a somewhat cursory level. Chapter 2’s coverage of inland water habitats helps substantially in this overwhelming task, but the breadth of other topics deserving coverage is still enormous. My approach, therefore, is to focus here on “life constraints” of critical importance to species survival. The primary task of all organisms from a genome perspective is to reproduce successfully. All other tasks are subservient and are basically organismal characteristics that

Habitat Traits 71 Functional Feeding Groups 71 Aquatic Food Webs 72 Food Web Techniques 73 Lotic Food Webs 74 Lentic Food Webs 75 Regulating Populations and Communities 75 Density-Dependent Control: Top-Down and Bottom-Up 76 Competition (Bottom-Up Control) 76 Predation (Top-Down Control) 76 Density-Independent Control: Environmental Variability77 Acknowledgment79 References79

promote living long enough and acquiring sufficient energy to form gametes, acquire a mate (if necessary), and produce viable progeny. Of these, I focus in this chapter on habitat characteristics, food webs, species interactions, and environmental control. My coverage extends here from species to communities; ecosystem and macrosystem ecology are covered in other ecology texts.

DISTRIBUTION IN SPACE Invertebrates are distributed in aquatic space based on habitat characteristics (discussed primarily in Chapter 2), interactions with other organisms (competition, predation, or parasitism), type and abundance of food, historical access to the site, and chance events. The current distribution also reflects to some extent adaptations to past environmental

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00004-8 Copyright © 2015 Elsevier Inc. All rights reserved.

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conditions and ongoing (but difficult to perceive) transitions to new community organizations.

Physical and Ecological Characteristics of Lotic Ecosystems Longitudinal Patterns in Habitat and Organisms Downstream distribution patterns are affected by the hydrogeomorphic patch structure of the river, flow direction (especially toward or away from the poles), tributary inputs, and where the river terminates (dry basin, underground, lentic system, larger stream, or directly into an estuary/ocean). For brevity’s sake, I limit the options in this section to the main channel (but see the subsequent discussion of lateral distribution) of streams flowing toward the equator and the ocean, and I do not consider hydrogeomorphic patch composition (but see the following section). We can assume under these restricted conditions that the lotic system along this pathway will get more permanent (seasons to millennia), less temporally dynamic in discharge over a 24-h period, deeper, wider, faster, less turbulent, warmer, more turbid, less oxygenated, and characterized by finer bottom sediments. If the headwaters are in a forested region, the riparian zone may occlude light hitting the stream initially, but as the stream widens, more light reaches the bottom (supporting autotrophy) until it gets too deep. If instead the headwaters start either in grasslands, shrublands, or above the tree line, then the downstream pattern will generally be increasing canopy cover for a while, followed by the typical pattern of decreasing percent canopy cover downstream. Wood substrate will occupy more of the stream bed in forested headwaters and gradually decrease downstream in the main channel. The nature of the riparian zone—such as the general land cover and whether existing trees are deciduous versus evergreen—affects thermal patterns and allochthonous food abundance and digestibility. As these physical habitat conditions change in a downstream direction, the type and abundance of predators will shift accordingly. Predators within shallow headwaters consist mostly of invertebrates (e.g., some stoneflies, hellgrammites, odonates, planaria, true bugs, various crustaceans) and low densities of demersal fish (e.g., darters, sculpins), salamanders, snakes, and some mostly terrestrial vertebrates (e.g., raccoons in North America). As the stream widens and deepens, the diversity and size of potential fish predators increase on average and planktivorous fish appear (e.g., small emerald shiners, large paddlefish in the Mississippi River system). The change in predators introduces different biotic constraints on the invertebrate community. Also along this downstream flow path, food resources change in absolute and relative abundances. The historical and still predominant perspective has been that food resources in forested headwaters are primarily based on

  SECTION | II  General Ecology and Human Impacts

allochthonous leaf litter from the riparian zone because of insufficient light to support adequate instream (autochthonous) benthic algae, suspended algae, moss, and vascular macrophytes. Recent stable isotope research, however, has indicated that algae may be a primary food source in some canopied stream even though the algae are not readily visible to investigators (see the section on Lotic Food Webs) and that autochthonous carbon predominates as a food source farther downstream. Food webs are discussed in greater detail later in this chapter. The combination of downstream changes in habitat, food resources, and potential competitors and predators alters both total invertebrate biodiversity and dominant types of species in the main channel. In headwater springs, amphipods and isopods at the spring source quickly give way downstream to aquatic insects (e.g., Barquín and Death, 2006; Carroll and Thorp, 2014). Intermittent headwaters and larger, intermittent sections of rivers favor species that can either colonize and reproduce rapidly, seek refuge in the hyporheic zone during drying of surface waters, or flee downstream to more permanent waters (e.g., Datry et al., 2014). Species favoring hard substrates in headwaters give way downstream to those able to burrow in soft substrate, though many persist near shore on available rocks and wood snags (e.g., Greenwood and Thorp, 2001) where conditions are more similar to smaller streams. Hyporheic species flourish best where water flow is sufficient and continuous and life-sustaining oxygen percolates through the substrate; such environmental demands constrain some species in intermittent and low-order streams or in areas where substrates are too small to allow adequate hyporheic water exchange. Likewise, infauna, nonhyporheic species can be restricted by substrate size and oxygen availability. Holoplanktonic invertebrates—principally many rotifers and microcrustaceans (branchiopods and copepods)— occur from headwaters to the river terminus, but they are not classified as zooplankton until the assemblage lives primarily in the water column (planktonic) rather than on the bottom (epibenthic). Some species of all three groups are present in headwaters (Schram et al., 1990), but their density, diversity, and planktonic nature do not increase until turbulence drops and suspended algae is sufficient to support populations long enough for reproduction. Population densities tend to peak in lateral slackwater areas of the main channel (Casper and Thorp, 2007) (see also the section on lateral riverscape patterns) where lower velocities and turbulence and greater temperatures and light penetration support higher production of algal food resources (Wehr and Thorp, 1997; Reynolds, 2000). However, life in the main channel of rivers is challenging in comparison to lakes because of the downstream transport and the helical flow that regularly takes the zooplankton’s largely algal food in and out of the photic zone. Animal “potamoplankton” in the main channel also need to be able to reproduce rapidly

Chapter | 4  Functional Relationships of Freshwater Invertebrates 

(thereby favoring rotifers and smaller species of microcrustaceans) before they are transported out of the system. Consequently, maximum diversity, density, and productivity of animal potamoplankton occur in lateral slackwaters, including side channels and backwaters of rivers (Reckendorfer et al., 2001; Casper and Thorp, 2007), as discussed in the following sections.

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Given these physical and chemical differences, ecological characteristics vary among hydrogeomorphic patches (Poole, 2002; Thorp et al., 2006, 2008) (Figure 4.2). Patches of similar type but farther apart should be more similar ecologically than adjacent, but dissimilar patches (Poole, 2002). Algal and vascular plant productivity and levels of system metabolism (cf. Dodds et al., 2013) should increase with greater habitat complexity at this spatial scale,

Hydrogeomorphic Patterns The previous section assumed for the sake of simplicity that rivers are longitudinal clines, exhibiting nearly continuous and predictable changes in habitat characteristics from headwaters to the river terminus. In fact, the opposite is a more accurate portrayal. Rivers are usually hydrogeomorphically complex systems with only partially predictable, and nonclinal variation downstream in bed slope, bed composition, main channel width, and channel number and connectivity (Thorp et al., 2006, 2008; Poole, 2010). For example, one section of a river may be constricted, another meandering, a third braided, and a fourth with relatively permanent anastomosing side channels and true backwaters. As the geomorphic structure of the river changes downstream, the diversity and types of flow habitats are altered (Poole, 2010), each with different mean and variability of flow over short to long time scales (Thorp et al., 2008). These can vary from higher velocity habitats in the central main channel, to slower slackwater areas along the margins of the main channel, and to moderate-to-zero flow habitats in the lateral parts of the riverscape. They also vary in floodscape characteristics, such as the number, size, and connectivity of usually disconnected channels (backwaters and some anabranches), oxbow lakes (=billabongs of Australia; Figure 4.1), wetlands, and normally dry terrestrial floodplains (Thorp et al., 2008). Flow habitats can vary in substrate composition and size and physicochemical conditions (e.g., O2, temperature, pH).

FIGURE 4.1  Billabong (oxbow channel) of the Murray River of Australia shown in upper left. A small portion of the main channel appears in the lower right. Photograph by J.H. Thorp.

FIGURE 4.2  A conceptual riverine landscape is shown depicting various functional process zones (FPZs) and their possible arrangement in the longitudinal dimension. Not all FPZs and their possible spatial arrangements are shown. Note that FPZs are repeatable and only partially predictable in location. Information contained in the boxes next to each FPZ depicts the hypothesized hydrological and ecological conditions for that FPZ, with symbols explained in the information key at the bottom. Hydrological scales are flow regime, flow history, and flow regime, with the scale of greatest importance indicated for a given FPZ. The ecological measures (food chain length, nutrient spiraling, and species diversity) are scaled from long to short, with this translated as low to high for species diversity. The light bar within each box is the expected median, with the shading estimating the range of conditions. Size of each arrow reflects the magnitude of vertical, lateral, and longitudinal connectivity. Figure reproduced from a revision of Figure 1.1 in Thorp et al. (2008).

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whereas nutrient spiraling lengths should decrease. Carbon sequestration and denitrification should be maximized in complex channel patches, especially those with true backwaters characterized by periodically zero directional flow, heavy detrital buildup, and some anaerobic conditions on the bottom and in surface waters. Interactions with riparia and terrestrial environments are maximized in hydrogeomorphically complex patches. Likewise, species diversity, density, and productivity of fish and invertebrates should be greatest in these complex habitats.

Lateral Riverscape Patterns of Organisms in Medium to Large Rivers Stream ecologists working in headwaters have focused strongly on the stream’s main channels or have generally grouped empirically all relatively permanent and ephemeral channels into a single channel. In contrast, scientists working in medium to large rivers are more likely to explicitly distinguish different lateral sites and flow habitats from the main channel because of easily demonstrable differences in ecosystem processes and community structure of vertebrates, invertebrates, and autotrophic organisms. Perspectives on the nature of longitudinal patterns (headwaters to the mouth) in invertebrate (and fish) assemblages differ greatly when focusing on lateral riverscape communities, slackwater sites in the main channel, and the principal portion of the main channel where flows exceed 10 cm/s. In medium to large rivers, the latter often consists of infauna (e.g., bivalve molluscs, chironomid midges, oligochaete worms), epifauna able to maintain position in higher flows (more common on rocky bottoms), and zooplankton such as rotifers and relatively small copepods and cladocera. Lateral slackwater sites in the main channel are characterized by minimal water currents, deposited sediment sometimes mixed with cobble or larger rocks, some living tree roots/branches from riparian trees, wood snags, and some rooted, aquatic macrophytic vegetation. In these slackwater areas are found various infauna and a substantial variety of epifaunal insects, snails (Brown et al., 1998; Greenwood and Thorp, 2001), and other invertebrates, including crustaceans (e.g., shrimp, crabs, crayfish, peracarids, depending on the region). Communities in the side (=lateral) channels of the riverscape as a group are often highly diverse and vary in composition according to flow habitats and the affected type and complexity of organic (living and dead) and inorganic substrates. These can range from those more typical of slowly flowing river sites to those more typical of placid wetlands. Zooplankton in these areas are much more diverse and larger on average than those in the main channel (Casper and Thorp, 2007); they also more closely resemble invertebrates of true lentic habitats. However, all lateral riverscape flow habitats are subject to periodic flow and flood pulses, with the frequency and magnitude

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of these altering the benthic and planktonic invertebrate communities.

Physical and Ecological Characteristics of Lentic Ecosystems The distribution, density, and diversity of invertebrates in lentic habitats are related to temporal (daily, seasonal, yearly, and evolutionary) and spatially distributed habitat features.

Temporal Factors Temporal effects on lentic systems span the gamut from millennia to seasonal changes. About 20 lakes worldwide are classified as ancient (>1 million years old) and include Africa’s Lake Tanganyika (2–12 Mya, depending on the basin), Asia’s Lake Baikal (oldest at 25+ million years), Europe’s Lake Ohrid (>4 Mya), Japan’s Lake Biwa (∼4 Mya), and South America’s Lake Titicaca (∼3 Mya). This time span is long enough for evolutionary diversification of species, such as ∼250 species of cichlid fish in Lake Tanganyika (Takahashi and Koblmüller, 2011) and more than 265 amphipod species in Lake Baikal (MacDonald et al., 2005). At time scales of tens of thousands of years, lakes can establish diverse and relatively permanent communities via aquatic, aerial, and rarely terrestrial connection to other water sources (e.g., overland movement of crayfish). If the lake regularly maintains water throughout the year and does not freeze to the bottom, it is likely to support fish, many of which consume invertebrates. Fish affect the invertebrate assemblage either directly by predation or indirectly by altering interspecific competitive results or habitat conditions (e.g., resuspending sediments, changing macrophyte abundance). In lentic systems that lose all surface waters at least every few years and lack a surface water connection to a stream or another lake, “relatively permanent” wetland communities develop. These can contain abundant and diverse populations of aquatic invertebrates, but they usually lack fish (exceptions occur when the fish can estivate in formerly muddy bottoms). Finally, when the ecosystem dries regularly and fills for only weeks to a few months seasonally or over multiple years, specialist invertebrate communities develop in ephemeral wetlands (e.g., prairie playas, vernal pools, pocosins, gnammas, tinajas, rock pools). The ephemeral wetland fauna consist primarily of species able to hatch from estivating embryos (the egg bank) in the formerly dry bottom and species migrating from terrestrial or other nearby aquatic habitats (especially insects and some amphibians). The hydroperiod of wetlands and lakes substantially affects species type and diversity as well as food web relationships (e.g., Schriever and ­Williams, 2013; O’Neill and Thorp, 2014). Seasonal changes can affect the distribution of invertebrates even in many relatively permanent lakes, and these

Chapter | 4  Functional Relationships of Freshwater Invertebrates 

are often associated with ice or oxygen tension. The entire water column of some polar lakes freeze completely to the bottom, whereas in some extreme but still cold latitudes, ice may scour the bottom of the littoral zone and eliminate most invertebrates. Oxygen tension varies seasonally in most lentic systems (except very deep ones with no significant turnover). It can also fluctuate diurnally in shallow wetlands from either temperature changes (especially in arid and semiarid ecoregions) or oxygen uptake at night in pools with high autotrophic production.

Spatially Distributed Habitat Features Lakes exhibit strong patterns of vertical distribution of invertebrates, whereas water depth is a minor factor in most wetlands. The four primary, depth-related factors in lakes are thermal and chemical stratification, light penetration, and substrate characteristics. Lakes are typically stratified vertically into an upper epilimnion and a lower (and usually much larger) hypolimnion, with the metalimnion separating them and defined by the thermocline (=area of maximum change per meter) (Figure 4.3). The thermocline also represents a strong water density gradient, with liquid water being most dense at ∼4 °C. As decomposition occurs in the hypolimnion, the oxygen content drops and this layer may become anoxic. The chemical state of a number of elements (e.g., iron) are then shifted from the surface water condition. Lakes turnover and redistribute water, oxygen, and nutrients infrequently or once,

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twice, or multiple times per year (see Chapter 2), depending on the lake turnover pattern (affected by depth, latitude, and altitude). The main channel of an undammed river, by the way, lacks a thermocline, but ephemeral thermoclines may occur in nonflowing areas of the lateral riverscape. Lake vertical stratification in physicochemical conditions is an important factor causing vertical stratification in invertebrates, primarily because of often low oxygen conditions in many hypolimnetic areas and a lack of habitat above the sediment in which to hide. Indeed, the hypolimnion continuously accumulates inorganic sediments and decomposing organic matter, thereby maintaining a small benthic particle size. Few macroinvertebrates tolerate conditions in the profundal zone beneath the seasonal thermocline, but microand meiofauna can be abundant in deep water. Littoral Zone The depth of the littoral zone (Figure 4.3)—the bottom area defined as having ≥1% of the surface’s photosynthetically active radiation (PAR)—restricts the area and amount of primary productivity by algae and other autotrophs. In highly turbid lakes and reservoirs, this may be only a few centimeters, but in very clear lakes (e.g., Crater Lake, Oregon, United States), the 1% PAR level may be below the metalimnion. Areas of high autotrophic production typically begat higher secondary production and greater diversity of invertebrates. The autotrophic organisms, especially vascular macrophytes, also contribute physical structure to

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the lake, which is an important factor in increasing density and diversity of all animals because of greater habitat niche complexity and potential protection from some fish predators. Studies of vegetated and nonvegetated regions of the littoral zone demonstrate the great value of macrophytes in reducing predation rates on benthic macrofauna (Hershey, 1985). This refuge is especially crucial because benthic animals are generally poor swimmers and have difficulty escaping highly motile predators. The littoral zone is also characterized by greater wave action and often greater size diversity of benthic substrates (e.g., more rocks). Because of generally higher oxygen conditions, the littoral zone of some lakes may support large assemblages of meiofauna, including nematodes, gastrotrichs, and rotifers, which can move between sediment particles. Interstitial organisms living among sand grains are called psammon. Neuston The term neuston refers to the assemblage of organisms associated with the surface film of lakes, oceans, and slowmoving portions of streams. It generally includes species living just underneath the water surface (hyponeuston), individuals that are above but immersed in the water (epineuston), and taxa that travel over the surface on hydrophobic structures (superneuston or, more properly, a form of epineuston). This name is similar to, or a subset of the older name, pleuston (sometimes neuston is used in reference to the microscopic components of the more encompassing pleuston). The density of neustonic organisms decreases with increasing turbulence. Consequently, most neuston is confined to lentic habitats or some lateral components of the riverscape. The neustonic food web is primarily supported by a thin bacterial film on the upper surface of the water, a concentration of phytoplankton near the surface, and allochthonous inputs from trapped terrestrial and aquatic organisms. Protozoa are common in this assemblage, which also includes bacteria, algae, and floating macrophytes (Munster et al., 1998; Butler et al., 2007), but other typically planktonic taxa are rare (one exception is the cladoceran Scapholeberis). Neustonic organisms may be entirely aquatic or they may move over the water surface (Krieger, 1992). The latter include springtails (Collembola), some arachnids (mites and water spiders), and various families of true bugs (e.g., water striders, Gerridae). Larger neustonic species are especially vulnerable to predation from both aquatic and terrestrial predators, and all species must be adapted to the higher ultraviolet radiation present near the water surface. Zooplankton The distribution of plankton in lentic systems varies with the taxonomic group and characteristics of the ecosystem. Plankton consist of bacterioplankton, phytoplankton (often

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combining true algae with cyanobacteria), and zooplankton. The last consists of holoplankton (permanent members of the plankton) and meroplankton (species that enter the plankton only during certain life stages, such as veligers of zebra mussels). They interact extensively with each other and with “organisms” capable of destroying/consuming them such as viruses, freshwater jellyfish, some benthic invertebrates (e.g., hydra), and planktivorous adult and larval fish. Zooplankton include all animals in the water column that float, drift, or swim weakly—that is, they are at the mercy of currents. Zooplankton are considered by some scientists to restricted to open water, “pelagic” species, which in freshwater consist mostly of rotifers and two kinds of crustaceans: cladocera (Branchiopoda) and copepods (harpacticoids, cyclopoids, and calanoids). However, the same broad groups of rotifers and crustaceans occur near shore among vegetation; these littoral zooplankton may be almost entirely benthic or either periodic or frequent swimmers among the littoral zone flora. Wetlands contain a greater proportion of littoral zooplankton, whereas lakes are dominated numerically by true pelagic zooplankton. Zooplankton are well known to migrate among habitats on a daily basis. Most migration is vertical from colder and darker waters during the day to shallow waters at night (e.g., Loose and Dawidowicz, 1994), but some littoral zone species migrate into open waters at night (e.g., Burks et al., 2002). The two primary explanations for this movement are: (1) to escape visually feeding, fish planktivores in the photic zone; and/or (2) to reduce metabolic costs by feeding on algae near the surface but then returning to deeper water during the day. Migration away from the surface during the day also reduces potential damage from ultraviolet radiation, but this would not account for deeper migration because most ultraviolet radiation is absorbed in the first meter, especially if much dissolved organic matter (DOM) is present (Morris et al., 1995). Potamoplankton are also known to migrate vertically and laterally in rivers (Casper and Thorp, 2007), but probably less so than in lakes and mostly in association with lateral river channels and bays.

Aquatic–Terrestrial Ecotones Ecotones can be defined in various ways (Décamps and Naiman, 1990), but in essence they are dynamic boundaries and transitional areas between two very distinct ecological habitats containing different communities and physicochemical features. In terrestrial systems, these are typically located between habitat patches or ecosystems, such as forest to grassland, whereas in aquatic systems, a major ecotonal transition is between land and water (cf. Riser, 1990). The physicochemical change for animals in these aquatic–terrestrial ecotones are more dramatic than for animals in a terrestrial ecotone because the habitats vary

Chapter | 4  Functional Relationships of Freshwater Invertebrates 

more substantially in a suite of characteristics, such as temperature, oxygen content, chemical conditions (e.g., pH, salinity, osmotic/ionic state), physical support for the animal (buoyancy), and predators. The riparian zone is a major transitional zone for all types of aquatic systems (Clary and Medin, 1999). Aquatic–terrestrial ecotones (henceforth: aquatic ecotones) occur around all aquatic systems (Gratton and Vander Zanden, 2009), but they are most spatially and temporally unstable in lotic systems on short time scales because of the more pronounced rise and fall of water levels. This instability will vary with river size (greater in arid-zone rivers and headwaters other than springs), lateral river complexity (from constricted channels to anastomosing systems with large floodscapes; Pinay et al., 1990), and the presence of levees and both water supply and power generating reservoirs. The greatest regular temporal fluctuations occur in reservoirs designed to provide peak-demand electrical generation. Ephemeral wetlands typically contain the most transitory, unregulated aquatic ecotones. These ecotones can be evaluated at multiple spatiotemporal scales. For example, at larger spatial and temporal scales, the connectivity between the riverscape and both aquatic and terrestrial elements of the floodscape represent an ecotone (Ward et al., 1999). On a finer scale, the transition from terrestrial to aquatic habitat along a river or lake bank (Dreyer et al., 2012) is also an ecotone. The focus of ecological investigations will vary with these ecotonal scales; for example, the biology of individual invertebrate taxa is usually more appropriately measured at the smaller spatiotemporal scales. Connectivity within either ecotonal scale can be defined as the rate at which organisms, organic matter, nutrients, and energy traverse the ecotones between adjacent ecological units (after Ward et al., 1999). Muehlbauer et al. (2014) identified a 50% “stream signatures” where subsidy resources from stream to land are still half their maximum (in- or nearstream) level. Such calculations help identify the spatial extent of streams, which can be important for protecting streams from pollution and channel manipulation (Doyle and Bernhardt, 2010). Biodiversity can be very high in terrestrial-to-terrestrial ecotones, but are typically low in terrestrial-to-aquatic ecotones. Prominent quasi-resident vertebrates exploiting this area at smaller spatial scales are adult frogs and shore birds, both preying on terrestrial and aquatic invertebrates. Terrestrial hunting spiders and semiaquatic spiders are very prominent in this ecotone. Representatives of aquatic invertebrates are many surface dwelling bugs and some beetles that occupy ecotone close to the waters edge or temporarily on land, many springtails (Collembola), and some pulmonate snails. Moreover, the many taxa of emerging insects that temporarily occupy this habitat could be considered ephemeral ecotonal residents (Karaus et al., 2013).

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ACQUIRING ENERGY Biodiversity Traits and Functional Feeding Groups Traits in General An alternative to the typical taxonomic description of community composition (Angermeier and Winston, 1999) is an approach based on functional or life history attributes under selection pressure from environmental conditions (Poff, 1997), such as size, drift, feeding behavior, and life cycle attributes. These “biodiversity traits” can link biodiversity patterns to food webs and ecosystem functioning (e.g., Reiss et al., 2009; Heino et al., 2013). This approach is founded on the habitat template theory, predicting that species possess traits that evolved to fit particular aquatic habitats (Townsend and Hildrew, 1994). Traits have even been used as targets for ecological restoration (­Laughlin, 2014). At species or higher taxonomic levels, trait-based guilds exhibit redundancy in stream ecosystems (Bêche and Statzner, 2009) and thus are more effective in comparing biogeographically distinct ecosystems than are communities based on taxonomic composition alone (Heino et al., 2013). Useable traits should vary somewhat predictably from upstream to downstream, across environmental gradients, and among regions as a result of landscape filters (Tonn, 1990). Heino et al. (2013) identified the need for studies of trait and taxonomic composition of stream communities among regions and drainage basins. Trait-based approaches using macroinvertebrate communities have been used frequently in Europe (Charvet et al., 2000; Bonada et al., 2007; Dolédec et al., 2006; Statzner and Bêche, 2010; Dolédec et al., 2011), but are less common in North America, with most U.S. research focused on western streams (Finn and Poff, 2005; Bêche and Resh, 2007) or based on existing environmental databases established across large geographic regions (Bêche and Statzner, 2009; Poff et al., 2010).

Habitat Traits A common trait used for aquatic invertebrates is based on habitat. This can include burrowers (moving into substrates), sprawlers (sitting on the bottom or slightly covered by a dusting of sediment), clingers (holding on to vegetation above the bottom usually), and swimmers (moving freely in the water column or regularly swimming from one substrate to another) (e.g., Carroll and Thorp, 2014).

Functional Feeding Groups Probably the oldest and certainly the most widely used trait is functional feeding group (e.g., Carroll and Thorp, 2014). These functional traits can be subdivided by the type of

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food ingested, how it is obtained, and/or where it is collected. The specificity of trait definitions can vary with the particular investigation, but a common division for food sources is based on animal prey, autotrophic organisms, and dead organic matter (benthic or suspended detritus). Invertebrates can be classified based on observation of feeding behavior, description of “mouth” structures, or analysis of either gut contents or food assimilated (for example, through use of stable isotopes). Animal prey can be consumed live (“predation”) or recently dead (“scavenging”) from other causes, with scavenging being functionally similar to predation. Predators consist of species either that ingest whole or pieces of recently killed prey (“engulfers”) or that pierce the body wall and suck out liquefied tissue (“animal piercers”). Ecto(external) and endo- (internal) parasites also consume living or dead tissue, but the prey are rarely killed by the parasite. Ectoparasitism occurs in various taxa, such as some rotifers, mites, a few copepods, and many branchiobdellids, though the latter inhabitants of the external surfaces of crayfish may be functionally more commensals than parasites (Füreder et al., 2009). The most commonly consumed living autotrophic organisms are true algae, but some invertebrates eat tissue (liquid or solid) of vascular macrophytes or rarely mosses (McWilliam-Hughes et al., 2009), with the macrophytes more commonly consumed as microbially colonized detritus. Cyanobacteria (blue-green algae) are occasionally consumed, but many cyanobacteria are distasteful or even toxic, or they may be colonial and difficult to ingest. Algae can be eaten from benthic surfaces by various “scrapers” or from the water column by “filterers” (a group with some members that consume dead organic matter) and “planktivores” (a broad term encompassing consumption of both phytoplankton and zooplankton). Herbivorous invertebrates may consume vascular macrophytes by burrowing into the plant (“burrowers”) or by sucking out liquid food (“plant piercers”). The food group “dead organic matter” includes suspended organic matter (seston, which also includes a separate living component), benthic particulate detritus, and rarely pieces of dead wood (eaten by “gougers” or “wood burrowers”). Benthic and suspended, dead organic matter can be subdivided by size into coarse, fine, and ultrafine particulate organic matter. Fine particulate organic matter (FPOM) and ultrafine particulate organic matter may include both dead organic matter and some living components of algae and both autotrophic and heterotrophic bacteria. Invertebrates eating benthic detritus are known as “collector/gatherers” or “deposit feeders,” whereas those extracting detritus from the water column are called “filterers.” The nutritive value of detritus is derived partially from the animal or plant components and partially from associated bacteria and fungi.

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A few invertebrates, such as blackfly larvae (e.g., Hershey et al., 1996) are thought to consume bacteria directly and may even derive nutrition from DOM (Ciborowski et al., 1997). As useful as functional feeding groups can be, they also suffer from some severe limitations. First, constant or periodic omnivory is very common (except in specialist feeders like piercers), making it challenging to assign a single feeding trait (e.g., Mihuc, 1997). For example, a snail or mayfly scraping a rock surface for algae may also be deriving nutrition from associated detritus, microinvertebrates, and bacteria consumed simultaneously. Likewise, food choice at any one time may depend on availability. For instance, a net-spinning caddisfly may be considered a filter-­feeding detritivore if it subsists mostly on suspended detritus entrapped by its net, but it may discard detritus from the net if sufficient (and more nutritious) animal prey are caught on the drift (making it a predator; Benke and Wallace, 1980), and it may also eat algae growing on its retreat or nearby rock surface (making it an herbivore). Second, functional feeding groups can change with developmental stage in some organisms, especially aquatic insects. For example, a dragonfly nymph may be a collector-gatherer detritivore in early instars and then change to an engulfing predator as it matures. Third, ingestion and assimilation may not be equivalent from a food source perspective, thereby complicating trait assignment. This is especially true for collectors and detritivores that may consume a large amount of dead organic matter along with lesser volumes of bacteria, fungi, algae, and microinvertebrates, but then assimilate primarily the microflora and/or animal matter.

Aquatic Food Webs Food web ecology has long been a popular topic in aquatic ecology, with emphases on sources of energy (terrestrial “allochthonous” carbon or aquatic “autochthonous” carbon), factors controlling web complexity and food chain length, and regulation of web pathways and species abundance (see top-down and bottom-up control in a subsequent section). Because a thorough summary of these studies would require at least a full chapter in this book, the discussion is limited to a brief summary of the debates on organic sources for different food webs. The relative importance of autochthonous versus allochthonous carbon sources can be evaluated in terms of both secondary (heterotrophic) production and system metabolism (primary production plus respiration by all animals, plants, fungi, and bacteria). Decomposition of allochthonous carbon is thought to be the primary pathway for bacterial respiration and thus may account for most aquatic respiration (Wetzel, 2001). The primary source of carbon for invertebrate production is more controversial and probably varies among types of aquatic ecosystems and possibly

Chapter | 4  Functional Relationships of Freshwater Invertebrates 

even within a given ecosystem (e.g., main channel vs lateral areas and backwaters of a river and shallow bays vs. deep pelagic areas of a lake).

Food Web Techniques Depending on the species involved and the habitat characteristics, food webs can be analyzed using behavioral observations (and experiments), gut content analysis, and chemical techniques (stable isotopes and fatty acids). Given the difficulty of directly observing most stream organisms in situ, food web research initially used gut content analyses to determine energy sources for aquatic organisms (e.g., Benke and Wallace, 1997), but later shifted in part to chemical analyses (primarily stable isotopes and to a lesser extent lipids) (Delong and Thorp, 2006). The advantage of gut contents analysis is that you can often identify the specific taxa consumed, thereby making it much easier to construct food web models. The disadvantages are: (1) the process is very time-consuming; (2) you only know what is ingested (e.g., sometimes multiple prey items immersed within an inorganic and organic matrix) and may not be able to identify the specific target prey; (3) determining the absolute or sometimes even relative abundance of prey can be difficult if only small parts remain (e.g., crustacean appendages, worm setae); (4) only recent (e.g., last 24 h) feeding episodes are evident (which may not be a typical food source); and (5) the percent of an item ingested and identified from the gut content is not necessarily equivalent to the percent of an item assimilated into the consumer’s tissues. As an example, a shredder may ingest large quantities of terrestrial leaf particles but actually assimilate mostly the microbial flora (e.g., Chung and Suberkropp, 2009). An alternative approach involves chemical analyses. Although behavioral observations and gut content analysis can inform the scientist of what was consumed in the last few minutes to 24 h, chemical techniques integrate diets spanning longer periods (e.g., up to 3 months in the muscle tissue of fish preying on invertebrates). Analysis of lipid concentrations in consumer tissue can enable an investigator to distinguish between broad categories of autotrophs (Larson et al., 2013), such as diatoms versus green algae as the primary food source or even more easily algae versus leaves of a terrestrial plant (Kainz et al., 2004). The analytical process is laborious, more challenging in the field, and relatively expensive; consequently, the technique is not widely used by aquatic ecologists. A more widely used approach over the past few decades has involved stable isotope analysis (Fry, 2006). The most commonly used isotopes are carbon (rarer 13C and common 12C) and nitrogen (rarer 15N and common 14N), but sulfur (34S), hydrogen (2H), and oxygen (18O) are also occasionally used. Sulfur, for example, can be effectively used for tracking food webs that have a partial marine component

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(Peterson, 1999; Wissel and Fry, 2005). Researchers can examine relationships among different consumer species at the same and different trophic levels using biplots of δ15N (15N/14N) versus δ13C (13C/12C) and indices such as Layman’s metrics (Layman et al., 2007; Layman and Post, 2008). A multiple-source mixing model, such as IsoSource (Phillips and Gregg, 2003) or SIAR (Parnell et al., 2010), or Bayesian techniques (e.g., Moore and Semmens, 2008) can be used to determine the relative contribution of possible food sources to consumer diets, although selection of specific mathematical approaches can be controversial (Fry, 2013a, b; Semmens et al., 2013), especially when the number of potential food sources exceeds the number of tracers by more than one. Stable isotopes can be analyzed from bulk tissue (the most common approach) or using compound specific isotopes, such as those associated with amino acids (­Chikaraishi et al., 2009, 2014; Hannides et al., 2009). The bulk-tissue approach is relatively cheap but requires the investigator to link the higher trophic level consumer with either the original autotrophic source (e.g., algae) or an herbivore consuming that autotroph (e.g., a benthic feeding snail, a suspension feeding mussel) to determine the number of trophic transfers. Obtaining an accurate value for the basal 15N signature (the original autotroph or an herbivore eating it) can be challenging either because of spatiotemporal variability in autotrophic signatures or the frequent difficulty of locating longer lived herbivores in a given river, lake, or wetland. In a newer approach using stable isotopes of amino acids, one can calculate food chain length by subtracting the 15N level of phenylalanine (which changes minutely between trophic levels) from the 15N level of glutamic acid (which changes substantially and predictively between trophic levels). The major advantages of this method are that only the consumer signature is needed (halving the number of samples for analysis) and the variability of the data is significantly less, based on results of a comparative laboratory experiment (Bowes and Thorp, in review). However, the current cost of this technique (as of spring 2014) is 6 × higher for N alone and 9× higher for C and N analysis together because of more complex analytical techniques and the dearth of laboratories currently running these analyses. The newest approach to identifying carbon sources involves stable isotope fingerprinting. This links the signature for 13C (and presumably 15N) in tissue of the target animal with characteristic signatures of potential food sources (Larsen et al., 2009, 2013; Ellis et al., 2014). A significant problem with chemical techniques occurs when the number of potentially important food sources exceeds the number of tracers (e.g., δ13C, δ15N) by more than one. This makes it difficult to reliably determine the actual percentage use for each source. Various mathematical and graphical approaches have been proposed to partially overcome this problem (Fry, 2013a, b; Semmens

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et al., 2013). However, one nontracer and nonmathematical approach is to use ecological knowledge to reduce the number of some potential sources that need to be considered mathematically; these might include some limited amount of information from gut content analysis, behavioral observations, or other ecological approaches. Behavioral, gut content, and chemical techniques applied to individuals merely tell the investigator what a given consumer has eaten to some degree, thereby allowing construction of a topological food web. Experiments are necessary to determine whether predation/herbivory or food/energy abundance controls density and secondary production of a given organism. To determine whether a food web pathway is important to the community as a whole, it may be necessary to construct a bioenergetic food web model with taxa evaluated for their secondary production or energy value. Such models are rarely constructed nowadays because of the needed time and energy that must be devoted to measuring density and secondary production of the prey and predator (but see Carroll and Thorp, in review, for karst springbrooks in the U.S. Ozarks).

Lotic Food Webs Food web research across broad spatial scales has often focused on the relative contributions of allochthonous (terrestrial) and autochthonous (in-stream) organic matter to food webs. This research initially involved gut content analyses, but has focused on stable isotope techniques for the past two decades. Most of this research in both lotic and lentic ecosystems has involved topological models, C/N analyses (with values >12 often suggesting a terrestrial origin), and isotope C–N biplots. Relatively few studies have evaluated the contributions of food sources to secondary production. A major impetus for studying spatial patterns of food webs in lotic systems was publication of the River Continuum Concept (RCC; Vannote et al., 1980). The RCC stipulated that allochthonous leaf litter is the primary organic source in headwaters (which the model initially analyzed as forested) because algal production is severely depressed by riparian shading. This leads to abundant populations and diversity of leaf shredders in headwaters. In mid-order regions, the widening of the still shallow stream allows sufficient light to reach the water column and stream bottom to support abundant autochthonous productivity by algae and aquatic macrophytes. Consequently, scrapers and leaf miners increase in these areas, whereas shredders should decrease. As the river continues to widen and deepen, insufficient light reaches the bottom and algae are frequently carried below the photic zone by the helical flow in the river’s main channel. Therefore, the food webs of large rivers are hypothesized by the RCC to be based energetically on FPOM leaking from upstream food processors and

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minor amounts of phytoplankton. As a result, collectors are thought to dominate this portion of the river. Unfortunately, because the RCC was based on an overly simplified hydrogeomorphic model of rivers (Poole, 2002, 2010), it failed to include any lateral channel (riverscape) or floodplain (floodscape) areas of the river (Junk et al., 1989; Thorp and Delong, 2002), ignored the shallow lateral margins of rivers where invertebrate diversity is higher (Thorp, 1992), and may have underestimated the importance of autochthonous algal production in large rivers for invertebrate and fish production (Thorp and Delong, 1994, 2002; Delong and Thorp, 2006). Criticisms from proponents of the Flood Pulse Concept (Junk et al., 1989; Junk and Wantzen, 2004) led to revision of the RCC for large rivers (Sedell et al., 1989). However, that RCC revision still tended to seemingly underestimate contributions from autochthonous production in the main and side channels of rivers. The weight of isotopic evidence now seems to indicate that autochthonous carbon (mostly algae) is the primary source of energy supporting “animal production” in most but not all lotic systems considered as a whole (e.g., ­Hamilton et al., 1992; Bunn et al., 2003; Delong and Thorp, 2006; Hoeinghaus et al., 2007). A partial exception is the lower, tidally influenced Hudson River (a flooded valley with almost no tributary input), where terrestrial detritus contributes 40–60% of the carbon used by zebra mussels (>90% of the metazoan biomass), oligochaetes, midges, and Bosmina zooplankton (Cole and Solomon, 2012). However, other studies generally show increasing transition from terrestrial to algal carbon sources for many lotic food webs as watersheds increase above 10 km2 (Finlay, 2001). Although it still may be the case that allochthonous leaf litter is the dominant source of carbon-based energy supporting animal production in canopied headwater streams for much of the year, some studies have begun to show the importance of autochthonous production even in headwater streams (e.g., Finlay et al., 2002; Dudgeon et al., 2010; Carroll and Thorp, in review) and waterholes in rivers of arid regions (e.g., ­Jardine et al., 2013). The relative importance of autochthonous vs allochthonous carbon for the main channels of large, deep rivers, however, has not been satisfactorily resolved. The relative importance of autochthonous carbon in the entire riverscape of large rivers may depend on the hydrogeomorphic complexity of the local river valley (Thorp et al., 2006, 2008), flow characteristics (e.g., Bunn et al., 2003), time of the year (e.g., Delong and Thorp, 2006; Hoeinghaus et al., 2007), depth, and turbidity. Note, however, that these conclusions for the importance of autochthonous primary production pertain to its role in supporting “secondary production” of metazoa. It is more likely that allochthonous carbon is the driver of overall system metabolism in most lotic systems through bacterial decomposition of refractile carbon, with consequent production

Chapter | 4  Functional Relationships of Freshwater Invertebrates 

of CO2 and resulting overall heterotrophic metabolism (cf. Howarth et al., 1996; Thorp and Delong, 2002). Food webs in lotic systems are apparently well-integrated vertically in the main channels because they are mixed continually by the helical flow of water. However, there are significant differences in species composition, density, and food web complexity of aquatic species from the main channel to lateral channels and backwaters (e.g., Roach et al., 2009).

Lentic Food Webs Lentic food webs usually vary in complexity with hydrological permanency, size, depth, age (all direct relationships), salinity (inverse), and both temperature and pH (often greater at intermediate values). Webs are also influenced by watershed size, botanical features of both the riparian zone (presence/absence of trees) and surrounding watershed (e.g., grasslands vs forested basin), and whether streams enter and/or leave the lake. Lake food webs are somewhat partitioned by the vertical dimension into a shallow littoral zone web (especially where shallow bays are common), an epilimnetic plankton web (surface to moderately shallow depths), and a deep hypolimnetic web. Aspects of physical or physicochemical patchiness are present in all three habitats, and this can affect food webs, especially in the littoral zone where both littoral-pelagic and benthic habitats are available to promote more complex food webs than are generally found in the less diverse surface and deep pelagic zones. The food webs of relatively permanent lakes are based on both autochthonous (eukaryotic algae, autotrophic bacteria, vascular macrophytes, and occasionally mosses) and terrestrial carbon (e.g., Cole et al., 2011), with the majority coming from autochthonous sources in most cases. The relative amount of autochthonous and allochthonous carbon present in a lake varies with basin and lake characteristics and either carbon source may predominant in total abundance and effect on system metabolism (e.g., Wilkinson et al., 2013). However, algal production has been identified as the primary source of carbon supporting most lake food webs (Batt et al., 2012), though this can be from living and detrital components (Weidel et al., 2008). Lake autotrophy is more important in clear water lakes and decreases somewhat in more humic lakes with greater allochthonous inputs (Pace et al., 2007). Depth is a major factor indirectly controlling lentic food webs, especially as it relates to lake hydrologic permanency (Schriever and Williams, 2013). Food webs are starkly different in very deep, ancient lakes such as Baikal with its many indigenous species of amphipods (MacDonald et al., 2005) and other invertebrates compared to wetlands, most of which dry periodically and many of which are consistently ephemeral (e.g., playas of the U.S. Great Plains) and

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dry seasonally. Playas are often dominated numerically by crustaceans, especially cladocerans, copepods, and an assortment of tadpole, clam, and fairy shrimps, but those supporting abundant insect diversity are more complex trophically (O’Neill and Thorp, 2014). Trophic complexity in these ephemeral systems is dependent on the length of the hydroperiod and when the playas fill (e.g., spring when fewer aquatic insects are present or in the early fall when they are abundant). Food chain length and the relative abundance of predators changes predictably through the hydroperiod (O’Neill and Thorp, in review). Carbon sources for wetlands are primarily autochthonous and algal based (e.g., Hart and Lovvorn, 2003; Sobczak et al., 2005; Williams and Trexler, 2006), but macrophytes and other sources of carbon can contribute to food webs and system metabolism, including methane (e.g., Sanseverino et al., 2012).

REGULATING POPULATIONS AND COMMUNITIES Over the past century, aquatic scientists have periodically shifted major research emphases, though older focal areas never seem to disappear entirely. Taxonomy of aquatic species has survived through the past half-century in part because of concerns over worldwide loss of biodiversity and the need to develop and enforce environmental protection laws. New techniques, however, have partially shifted the emphasis in taxonomy from morphology to genetics as a way of classifying species and establishing phylogenetic relationships. Within aquatic ecology, the emphasis has tended to increase in scale from studies of individual species to communities, ecosystems, and macrosystems (including global change). Shifts in ecological research emphases have often followed landmark conceptual papers (e.g., on keystone predation, trophic cascades, river longitudinal patterns, and nutrient cycling), specific environmental concerns (e.g., pollution, invasive species), development of new research techniques (e.g., field cage, whole-lake experiments, stable isotope analysis, and advanced genetic techniques), advent of new equipment (e.g., respirometers, sondes for chemical analyses, global positioning systems), and development of mathematical analytical techniques (e.g., Bayesian statistics, geographic information systems). At other times, changes in ecological emphases seem stimulated, unfortunately, by nothing more than a desire to leave a now “passé topic” and seek new research horizons. The latter is further fostered by a subsequent change in government funding priorities. These changes in popularity of ecological foci rarely seem to follow complete resolution of an ecological question. Two broad ecological research topics related to invertebrate species, populations, and communities concern environmental variation and biotic control (via food availability and consumer interactions) as they influence species

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distribution and abundance. The following sections focus on especially important topics and questions that have never been satisfactorily resolved, in my opinion, despite more than half a century of research that continues today, albeit at a reduced level in most cases. These sections are also designed to highlight the progression of research in studies involving freshwater invertebrates.

Density-Dependent Control: Top-Down and Bottom-Up Competition (Bottom-Up Control) More than a century ago, ecologists began seeking to identify mechanisms allowing coexistence in habitats where species shared apparently limiting resources. The concept of competitive exclusion in response to potentially limiting resources was developed over time by scientists such as Joseph Grinnell (1904) and Gregory Gause (1932). Decades later, Hutchinson (1957) described a niche as an ­“n-dimensional hypervolume,” and Hardin (1960) captured the idea of competitive exclusion in his succinct phrase “complete competitors cannot coexist.” That is, species whose niches overlap in fundamental ways cannot thrive permanently in the same environment. Hutchinson (1959), while contemplating the diversity of corixid beetles in an Italian mountain pool, asked how so many species could coexist simultaneously in an environment of seemingly limited niches, and he later posed a similar paradox for lake plankton ­(Hutchinson, 1961). In these landmark papers, Hutchinson speculated that environmental change, predation, and minute but important niche differences could explain this coexistence in the face of fundamentally overlapping niches. These studies led directly or indirectly to decades of ecological research, with initial studies of competition and niche diversification, followed by observational and experimental research on the effects of predation, and lastly by investigations on the role of nonequilibrial ­environmental conditions (see the next two sections). Biological interactions leading either to coexistence or exclusion were then analyzed in terms of exploitative (resource) or interference competition (also known as territoriality or space competition). In the first case, species differ in how rapidly and efficiently they can harvest a resource, whereas in the second instance the difference is in ability to control space which is later used in obtaining food or a mate. Strictly speaking, actual resource limitation of the current population or its ability to produce healthy offspring must be present for resource competition to exist. Interference competition usually involves agonistic or aggressive interactions and does not necessarily require an immediate resource limitation. As an example of resource competition, the snail Elimia outcompeted the caddisfly Neophylax for algae in a stream (Hill, 1992). In contrast, the ability of the

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hydropsychid caddisfly larva Leucotrichia to drive away the mayfly nymph Baetis (a potential food competitor) from foraging space around its retreat (which also includes a net for filter-feeding) is an example of interference competition (Hart, 1985). Although agonistic space competition seems rarer in nature, the effects on microhabitat distribution of hydropsychid caddisflies and demonstrable access to highquality resources were also shown with in situ experiments in the small Mianus River of the northeastern United States (Georgian and Thorp, 1992). All these instances are examples of bottom-up control of species abundance in that the availability of food supposedly limits the density and possibly distribution (at least on a microhabitat scale) of freshwater invertebrate species. Competition is thought to involve density-dependent control of populations. That is, the intensity of the effect varies with the density of competitors. It is also implied that the competitive effect will extend to the next generation. In other words, reduction in the size of one generation (larvae or adult) is assumed to result in a lower population size in the next generation. However, this vital assumption is almost never tested and may, in fact, be wrong in many cases if other factors exert a dominant control on the population or if reproductive propagules easily replace lost individuals from the previous generation.

Predation (Top-Down Control) Publication of the keystone predation concept by Robert Paine (1966, 1969) led first to an expansion of field experiments and appreciation of the role of predators in reducing competition in marine systems and later to a similar focus over nearly two decades in freshwaters (e.g., Allan, 1982), with multiple books published on predation effects (e.g., Zaret, 1980). These studies focused, therefore, on top-down control, with a strong emphasis on predation (but minimal studies of parasitism). Initial freshwater experiments included manipulation of predators in experimental ponds (Hall et al., 1970) and field cages in reservoirs (Thorp and Bergey, 1981), with later experiments spreading to wetlands and streams. Stream studies often were inferential or involved artificial streams because cages were easily destroyed by the force of stream spates. Other field cage experiments, however, survived flow fluctuations and demonstrated significant effects of fish on benthic invertebrates in both shallow rivers (e.g., Power, 1990) and shoreline areas of large rivers (Thorp et al., 1998) as well as cascading effects on river zooplankton (“potamocorrals”; Figure 4.4) (e.g., Jack and Thorp, 2002; Thorp and Casper, 2003). Examples in nature where the keystone predation hypothesis was most applicable involved competition for space in a two-dimensional environment (e.g., rocky intertidal), where food webs were relatively simple and/or where equilibrium conditions prevailed. This requirement

Chapter | 4  Functional Relationships of Freshwater Invertebrates 

(a)

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(b)

FIGURE 4.4  (a) Shown here are six of 20 connected wood and aluminum frame modules (each 10′  ×  10′) of a potamoccoral rafter shown floating downstream in the Ohio River; (b) each module supports a 3500 L enclosure for manipulating river zooplankton (animal potamoplankton); shown here are two forms of the enclosures (use depending on the experiment): one impermeable (limnocorral next to A.P. Casper) and the other constructed of 80 μm nitex mesh (potamocorral, next to J.H. Thorp). Photographs courtesy of J.H. Thorp.

was critical to the early marine studies of Paine (1966, 1969). The intermediate disturbance hypothesis (Connell, 1978) included the role of predators in conditions of environmental equilibrium but also extended the theory to the role of fluctuating abiotic conditions (disequilibrium) in promoting higher species diversities at intermediate levels of environmental stress (see next section). The concept for “biotic” intermediate disturbance is that prey diversity will initially rise as predator densities increase (from reduced interspecific competition) and then decrease when the stress level reaches a critical point, producing a sigmoid or “bellshaped” curve. Explicit experimental tests of this predator effect on prey diversity and mathematical model are rare in freshwater (but see Thorp and Cothran, 1984). The vast majority of the studies published on the role of predation in freshwater have reported only effects on prey densities, not species diversity. The assumption—whether justified or not—was that reduction in densities would eventually lead to decreased species diversity before the environment changed or reproduction compensated for the loss by the next generation. An important way that predators could potentially regulate populations and communities other than by eliminating species (apparently rare) is by reducing population densities sufficiently so that interspecific competition for food or space no longer significantly controls species abundance. Equal consumption of competing prey species, however, may not lead to higher diversities in the prey assemblage. Another way that a predator could enhance prey species diversity is by switching its search image, behavior, or habitat in a way that favors capturing prey species B when densities of prey species A fall below some critical level. This promotes regrowth of the latter’s population and thereby overall community diversity. This density-dependent, type III functional feeding response has been shown in terrestrial

and some marine vertebrate predators, but has been rarely been shown in freshwaters. However, a laboratory experiment (Cothran and Thorp, 1985) showed that dragonfly nymphs changed foraging areas from the bottom of containers (where midges were abundant) to perches on artificial vegetation (where cladocera were prevalent) in responses to an experimental switch in the relative abundances of the two prey species. Aside from the ideas of keystone predation and the intermediate disturbance hypothesis, the most popular concept in predator-prey studies has probably been “trophic cascades” (Paine, 1980; Carpenter and Kitchell, 1996). This concept of top-down control in ecosystems stipulates that selective predation at the top of the food web cascades down the “food chain” in such a way that there is an alternating decrease and increase in abundance of lower trophic levels, thereby stabilizing the community at higher trophic levels (Kitchell and Carpenter, 1996). Trophic cascades have been reported in freshwater streams (e.g., Power, 1990) and lakes (e.g., Thorp and Casper, 2003). The effects of predators in producing trophic cascades vary among freshwater, marine, and terrestrial systems (Pace et al., 1998; Shurin et al., 2002), but they are generally more prominent in more productive but less complex food webs (which more closely resemble simple food chains). However, these environmental conditions do not always produce strong cascades (Borer et al., 2005).

Density-Independent Control: Environmental Variability Scientific support for either competition and predation being the dominant force in regulating aquatic communities began to fade in the mid- to late 1980s as ecologists began shifting from a long-standing equilibrium to

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a nonequilibrium (=stochastic) view of the world. From this perspective, competitive exclusion rarely occurs either because: (1) the mantle of competitive dominant switches among species with changes in abiotic conditions or (2) resource competition disappears when population densities of both species fall, thereby eliminating at least exploitative competition. Note: this still dominant, nonequilibrium perspective does not say that competitive type interactions disappear or that predators stop eating prey. Instead, it postulates either that: (1) environmental fluctuations eliminate resource scarcity and with it the forces that favored one species over another or (2) habitat variability in space and time favor one species in some places or times and other species in different conditions, thus providing an escape in space or time for the subordinate species and eliminating species extinction and loss of diversity. Stochastic processes do not necessarily eliminate stability but may, instead, constitute a mechanism underlying the apparent stability of ecological systems at different spatiotemporal scales (Wu, 1999). “Thus, equilibrium and nonequilibrium are not absolute and context-free, but relative and scale-dependent” (Wu, 1999). The environmental fluctuations that are most common and important for population and community regulation in aquatic systems vary between lotic and lentic habitats, and most of these have some direct or indirect relationship to hydrologic changes. Hydrological changes can directly or indirectly affect the force of water (hydraulics) on organisms, the displacement of substrate and species, oxygen tension, temperature (overheating or ice scour), and of course complete desiccation of the aquatic system. Global climate changes could affect invertebrates via both hydrologic changes (especially predictability and seasonal timing of flow events) and water temperatures (e.g., Woodward et al., 2010; Stoks et al., 2014). In streams, these stresses are principally related to the predictability in timing and degree of hydrologic events: short but intense spates, extensive floods, or long droughts. Flood events can devastate stream invertebrate populations. For example, a flood event in the ephemeral Sycamore Creek of the arid southeastern United States removed more than 90% of the benthic insect biomass and much of the attached algae (Grimm and Fisher, 1989). Cessation of flow is common in low-order headwaters (intermittent streams) and rivers of arid and semiarid regions, but it is not clear whether communities in these regions regularly reach the point where biotic interactions regulate community structure (and thus whether environmental fluctuations could disrupt this process). Two exceptions seem probable. First, rivers in regions such as much of the Australian continent, are often dry to isolated waterholes where both predation and food limitation could be important (e.g., Lake, 2003; Jardine et al., 2014). Second, flow in some mountain streams in semiarid regions (e.g.,

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southern California, United States) may mostly cease seasonally, trapping predaceous fish like trout in some pools but not others, thereby affecting selective consumption of invertebrates and possibly competitive outcomes (Hemphill and Cooper, 1984). Springs systems are notable in that environmental fluctuations are rare in comparison to other headwater streams, and thus one might expect to find greater importance of predation and competition in regulating invertebrate populations and communities (e.g., Carroll and Thorp, 2014). In comparison to stream ecology, environmental fluctuations are less important in relatively permanent lakes (except on an evolutionary time scale) but are approximately equally important in ephemeral wetlands. Although some wetlands are fairly consistent in the timing of their dry and wet periods (especially large tropical wetlands), others are at the mercy of local fluctuations in weather. For example, playa wetlands may fill multiple times during the year (with the fill season affecting community characteristics; O’Neill and Thorp, in review), once a year, or not for multiple years or even many decades (O’Neill and Thorp, 2014). Consistent colonists of ephemeral wetlands, such as tadpoles and fairy shrimps, take advantage of these vagaries of nature through their reproductive adaptations (resistant eggs) and may view playas as equilibrial environments, but many other nonspecialists in these systems (many insects) may experience the system ecology through a stochastic perspective. A major early thrust of studies of stochastic processes in streams started in the late 1980s and dealt with patch dynamics and the role of the resulting patch mosaics in affecting community assembly and disassembly (e.g., White and Pickett, 1985; Pringle et al., 1988; Wu and Loucks, 1995; Palmer et al., 2000; Wiens, 2002; Arrington et al., 2005; Thorp et al., 2008). As an example of this process, food competition between two grazing invertebrates in a stream could easily result in one species (A) being eliminated by exploitative competition from a second species (B). However, if species A is a faster at dispersing (reaching uncolonized habitats) and reproducing, then it may survive or even flourish in habitats where stream spates disturb the substrate sufficiently and displace most resident grazers. The effects of fluctuating conditions are sufficiently important in modern times, where most aquatic systems are at least somewhat impacted environmentally, that applied and fundamental environmental research has begun focusing increasingly on the characteristics of species, communities, and ecosystems to resist (resistance) or bounce back (resilience) from perturbations. Our abilities to predict and avoid conditions which will lead to “state or threshold changes” in communities will be valuable attributes for future aquatic ecologists and invertebrate zoologists to have.

Chapter | 4  Functional Relationships of Freshwater Invertebrates 

ACKNOWLEDGMENT My discussion of trait-based characteristics of aquatic organisms was taken in part from an NSF macrosystems grant proposal (April 2014; JHT as P.I.) comparing Mongolian and U.S. rivers; I especially appreciate contributions from Dr. Barbara Hayford for that proposal section.

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Chapter | 4  Functional Relationships of Freshwater Invertebrates 

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Chapter 5

Ecology of Invasive Alien Invertebrates Anthony Ricciardi Redpath Museum, McGill University, Montreal, QC, Canada

Chapter Outline Rates and Global Extent of Freshwater Invertebrate Invasions83 Human Vectors of Dispersal 84 Traits Conferring Invasion Success 86 Ecological Impacts 86

RATES AND GLOBAL EXTENT OF FRESHWATER INVERTEBRATE INVASIONS Under human influence, species are spreading faster and farther into new regions than at any other time in Earth’s history. This process of spread and establishment is termed biological invasion, and species introduced beyond their native (historical) range are referred to as alien species (or nonnative, nonindigenous, or exotic). Those that spread and establish rapidly, or that become pests, are often described as “invasive.” Freshwater ecosystems worldwide are highly vulnerable to being invaded and transformed by alien invertebrates. Large aquatic ecosystems are known to contain scores of alien species, especially crustaceans and molluscs (Table 5.1). The true numbers of alien species, even in these relatively well-studied systems, are probably grossly underestimated, resulting from (1) irregular monitoring, (2) incomplete faunal inventories, (3) poor knowledge of the historical biogeography of many of the species present, and (4) insufficient taxonomic expertise for recognizing alien invertebrates. Consequently, virtually all freshwater ecosystems contain cryptogenic species (taxa that cannot be classified with certainty as either alien or native) and pseudoindigenous species (alien species mistaken as natives) (Carlton, 2009). These are typically small-bodied taxa (e.g., bryozoans, annelids, nematodes, rotifers, gastrotrichs) that are often misidentified despite being common and widespread. The inverse correlation between an organism’s body size and its likelihood

Can the Impacts of Alien Invertebrates Be Predicted? 88 Management of Invasive Aquatic Invertebrates 88 References89

of being recognized as alien is termed the smalls rule (Carlton, 2003). This problem, along with a number of other factors (e.g., highly selective transport by humans, both intentionally and inadvertently), has contributed to an unbalanced representation of taxa among alien invertebrates in freshwater systems (Panov et al., 2004; Karatayev et al., 2009). Aquatic insects, for example, are rarely on alien species lists despite dominating benthic fauna in lakes and rivers (Table 5.1). Most winged insects are likely to have colonized their current ranges from glacial refugia much earlier (beyond the detection of biologists) than other taxa. The few unequivocal cases of alien aquatic insects are from invaded habitats very remote from their historic range and clearly beyond their natural dispersal abilities (e.g., Van de Meutter et al., 2010). Many species believed to have natural cosmopolitan distributions are likely alien in origin (Carlton, 2009). Several of these species have globally disjunct distributions, in which records of occurrence include isolated populations apparently far removed from a more extensive and contiguous historical range. Genetic analyses are beginning to reveal cryptic invasions in freshwater systems worldwide (e.g., Walther et al., 2006; Folino-Rorem et al., 2009). In contrast, some invasions have involved conspicuous, even spectacular, range extensions. The freshwater jellyfish Craspedacusta sowerbyi Lankester, 1880, native to the Yangtze River valley, is established on every continent except Antarctica, and it continues to be discovered in lakes and ponds across the world. Global introductions of the Asian clam Corbicula fluminea (Müller,

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00005-X Copyright © 2015 Elsevier Inc. All rights reserved.

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TABLE 5.1  Number of Alien Macroinvertebrate Taxa Recorded as Established in Large Freshwater Ecosystems System

Molluscs

Crustaceans

Annelids

Insects

Other

Reference

Laurentian Great Lakes

18

21

6

2

23

Lake Champlain

11

5

0

0

2

Marsden and Hauser 2009

Hudson River

19

6

1

0

2

Mills et al., 1997

Mississippi River

8

6

0

0

1

Rasmussen 1998; Grigorovich et al., 2008; Wood et al., 2011

Columbia River

3

4

7

0

3

Sytsma et al., 2004

10

23

6

1

6

Leuven et al., 2009

Thames River

8

15

3

2

13

Ebro River (Spain)

7

11

1

0

5

Oscoz et al., 2010

Lake Balaton (Hungary)

3

10

0

0

1

Muskó et al., 2007; Muskó et al. 2008; Gábor 2009

Rhine River

1774) and its congeners began more than 100 years ago and continue today. Within the past 25 years, more than a dozen invertebrate species from the Black and Caspian sea basins have invaded the North American Great Lakes (Ricciardi, 2006); these include the zebra mussel Dreissena polymorpha (Pallas, 1771) and the quagga mussel Dreissena rostriformis bugensis (Andrusov, 1897), which have since spread farther across the Rocky Mountains into the western United States (Stokestad, 2007). Numerous gastropods have become invasive globally, including Chinese mystery snails (Bellamya/Cipangopaludina spp.), Central American apple snails (Pomacea spp.), the New Zealand mud snail Potamopyrgus antipodarum (Grey, 1853), and a variety of pulmonate snails (e.g., Physella spp.). Intercontinental transfers of crustaceans are even more frequent. Alien crustaceans, including cladocerans, copepods, amphipods, mysids, and crayfish, are found in most large freshwater lakes and rivers worldwide (Table 5.1). Some crustaceans that were previously confined to coastal environments, such as the copepod Eurytemora affinis (Poppe, 1880), are now being discovered more frequently in freshwater lakes and rivers, suggesting a recent adaptation to low-salinity waters (Lee and Bell, 1999). Alien invertebrates are being discovered at increasing rates in large freshwater systems (Figure 5.1). Multiple factors may contribute to this apparent pattern, the most plausible being increased opportunities for introduction through human activities. In the Great Lakes, the composition of alien species discovered during the past century correlates with changes in shipping and other vectors. Thus, species known to be moved by particular vectors become more prevalent when that vector is most active (Ricciardi, 2006). Rates of invasion by freshwater invertebrates appear to be orders of magnitude

Mills et al., 1993; Ricciardi, 2006; Ricciardi, unpublished data

Jackson and Grey 2013

greater now than before human influence (Ricciardi, 2006). Molecular evidence has revealed that the modern rate of invasion of North American lakes by European cladocerans is ∼50,000 times greater than the prehistoric rate (Hebert and Cristescu, 2002).

HUMAN VECTORS OF DISPERSAL Invertebrates are being dispersed worldwide primarily by vectors associated with both shipping and live trade in aquatic organisms. Carlton (1993) identified more than 20 distinct human-mediated mechanisms of dispersal of the zebra mussel involving canals, shipping activities (ballast water, hull fouling, movement of navigational buoys, and boatyard equipment), and fishing activities, including the transport and release of live bait. On a global scale, molluscs and crustaceans are distributed through the commercial importation of aquatic organisms for the aquarium trade or for human consumption (Padilla and Williams, 2004; Keller and Lodge, 2007). For example, the red swamp crayfish Procambarus clarkii (Girard, 1852), native to the southeastern United States, has been introduced to two dozen countries in South America, Africa, Asia, and Europe, often deliberately for use in aquaculture and the aquarium trade, and as an agent to control pest snails (Gherardi, 2011). The importation of water plants for ornamental gardens likely contributed to the inadvertent introduction of the Asian bryozoan Lophopodella carteri (Hyatt, 1866) into North America during the 1930s (Ricciardi and Reiswig, 1994). Shipping is assumed to be responsible for hundreds of freshwater invertebrate invasions worldwide (e.g., Ricciardi, 2006; Leuven et al., 2009; Jackson and Grey, 2013) including, for example, the occurrence in the Panama Canal of sponges

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FIGURE 5.1  Accumulation of alien invertebrate species recorded as established in (a) the Laurentian Great Lakes, (b) the Rhine River, (c) the Thames River, and (d) the Ebro River, as a function of year of discovery. Sources: Ricciardi, 2006; Leuven et al., 2009; Jackson and Grey, 2013; Oscoz et al., 2010.

Eunapius carteri (Bowerbank, 1863) and Trochospongilla leidii (Bowerbank, 1863), the former previously unrecorded in the western hemisphere and the latter found previously only in the United States (Jones and Rützler, 1975; Poirrier, 1990). Resting eggs of cladocerans, copepods, rotifers, and bryozoans occur commonly in the bottom sediments of the ballast tanks of ships originating from freshwater and estuarine ports, in numbers that reach tens of millions of eggs per ship (Bailey et al., 2005; Duggan et al., 2005; Kipp et al., 2010). The number of freshwater bryozoan taxa found as statoblasts in the ballast sediments of 33 ships visiting the Great Lakes during a 2-year period accounted for more than 10% of all described species (Kipp et al., 2010). Indeed, the intercontinental movement of statoblasts by ships is assumed to be responsible for introducing an Asian bryozoan, Asajirella gelatinosa (Oka, 1891), into the Panama Canal region (Wood and Okamura, 1998).

The development of canals and navigational waterways can promote intracontinental invasions, as in the case of the spread of Ponto-Caspian animals across Europe (bij de Vaate et al., 2002). Several alien invertebrate species have dispersed through European waterways at mean rates of 40–130 km/yr (Leuven et al., 2009). Recreational boating is responsible for the overland dispersal of invertebrates attached to the boat hull or in bilge water (Johnson et al., 2001; Havel, 2012). The long-distance dispersal of dreissenid mussels has been aided by their attachment to barge traffic (Keevin et al., 1992) and to vessels trailered over land (Stokestad, 2007). For asexually reproducing organisms in particular, any form of transport of small amounts of water could potentially facilitate invasions over distances of thousands of kilometers. The interbasin movement of fishing gear [e.g., nets (Jacobs and MacIsaac, 2007)] almost certainly contributes to the spread of some

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invertebrate species. An interesting case involves a North American triclad flatworm (Phagocata woodworthi Hyman, 1937) that was introduced to Loch Ness, Scotland, at some point before 1977 (Reynoldson et al., 1981), conceivably on equipment transported by American monster hunters!

TRAITS CONFERRING INVASION SUCCESS Some hypothesized general attributes of successful aquatic invertebrate invaders are listed in Table 5.2. Although these generalizations have been widely cited, few have been tested rigorously. In a statistical comparison of European stream invertebrates, Statzner et al. (2008) found that genera with invasive species tended to reproduce more frequently and achieve greater abundances where they occurred, and also had significantly more ovoviviparity, which could enable colonization by a single individual that provides parental care and thus releases more viable offspring. Trait-based comparisons of alien and native gammarid amphipods in central Europe suggest that invasive aliens often have a combination of early maturity, greater fecundity (e.g., larger brood sizes), a greater number of generations per year, and a broad tolerance to habitat stressors (Grabowski et al., 2007); however, these traits may not be universal (e.g., Devin and Beisel, 2007). For bivalves, invasive taxa can be distinguished from natives by a set of biological traits that include shorter life spans, early maturity, and rapid growth (Morton, 1997; McMahon, 2002). Potentially confounding these patterns are traits that happen to promote uptake and transport by human vectors. For example, unionids are generally poor colonizers, and many have experienced range declines, although their larval stages are dispersed by fish.

One notable exception, however, is the Chinese pond mussel Sinanodonta (Anodonta) woodiana (Lea, 1834), which uses of broad range of fish hosts—including, most importantly, commercially cultivated species—that have caused it to be distributed to various continents inadvertently through aquaculture activities (Douda et al., 2012). Tolerance to aerial exposure may also aid in dispersal between water bodies, such as by overland transport on recreational boats (Johnson et al., 2001; Havel, 2012). Some studies have reported enhanced salinity tolerance among alien invertebrates, particularly crustaceans, compared with their native counterparts (Devin and Beisel, 2007; Grabowski et al., 2007). Similarly, introduced invertebrates discovered in the Great Lakes in recent decades consist predominantly of euryhaline species (Ricciardi, 2006). This pattern can be attributed to an enhanced capacity for euryhaline species to become abundant around estuarine ports and to tolerate fluctuating salinities within ballast tanks during long voyages. Many successful invaders also have resting (diapausing) stages that allow them to survive periods of unfavorable conditions, including hypoxia and temperature fluctuations, such as those typical of ballast tank environments (Panov et al., 2004). Invertebrate resting eggs may remain viable even during transport in mud attached to road vehicles and footwear (Waterkeyn et al., 2010).

ECOLOGICAL IMPACTS Although the impacts of invertebrate invasions are often inconspicuous and rarely studied, documented impacts are broad in scope and include native population declines, altered nutrient cycling, changes to contaminant bioaccumulation

TABLE 5.2  Hypothesized Attributes of Highly Invasive Freshwater Invertebrates Attribute

Reference

Abundant and widely distributed in original range

Miller et al., 2007

Broad environmental tolerance

Morton 1997; Miller et al., 2007; Alonso and Castro-Díez 2008

Short generation time (early sexual maturity, short life span)

Morton 1997; McMahon 2002; Grabowski et al., 2007

High, rapid growth rate

Morton 1997; McMahon 2002; Morrison and Hay 2011

High reproductive capacity

Morton 1997; Keller et al., 2007; Grabowski et al., 2007; Alonso and Castro-Díez 2008; Statzner et al., 2008

High rate of resource consumption

Morrison and Hay 2011; Dick et al., 2013

Aggressiveness (especially among crustaceans)

Gamradt et al., 1997; Dick 2008

Possession of natural mechanisms of active and passive dispersal

Alonso and Castro-Díez 2008

Strong potential to exploit human transportation systems

Morton 1997; Alonso and Castro-Díez 2008;

Asexual/hermaphroditic modes of reproduction

Panov et al., 2004

Adaptive capacity to colonize new environments (e.g., high plasticity)

Hänfling and Kollmann 2002

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pathways, indirect food web effects on plants and algae (trophic cascades), and physical habitat alteration (Ricciardi and MacIsaac, 2011). Some species have farreaching effects through their ecosystem engineering activities. These are best exemplified by small, prolific suspension-feeding organisms, such as the dreissenid mussels (Dreissena spp.), the golden mussel Limnoperna fortunei (Dunker, 1856), Asian clams (Corbicula spp.), and the mud amphipod Chelicorophium curvispinum (Sars, 1895). Chelicorophium inhabits networks of mud tubes that it builds on firm surfaces, thereby altering the physical habitat of other benthic invertebrates. During the 1980s, C. curvispinum invaded the Rhine River and, within a few years, it established densities of hundreds of thousands per square meter and altered the rocky substrate drastically with its muddy encrustations. The habitat engineering and suspension-feeding activities of this dense population apparently caused the near extirpation of the zebra mussel and a native hydropsychid caddisfly from the lower Rhine (van den Brink et al., 1993). Dense populations of Corbicula and Dreissena achieve filtration rates that exceed those of native bivalves by up to three orders of magnitude (Strayer et al., 1999). They can remove enormous quantities of suspended particles from the water column, causing a doubling or tripling of water clarity and promoting the expansion of weed beds (Phelps, 1994; Strayer et al., 1999; Vanderploeg et al., 2002). The influence of C. fluminea in the Potomac River was extensive enough to cause the appearance of submerged aquatic vegetation for the first time in 50 years, with concomitant increases in the abundances of littoral fish and waterfowl. These dramatic ecosystem changes were rapidly reversed after the clam population exceeded its carrying capacity and crashed (Phelps, 1994). Similarly, zebra mussel activities enhanced the water clarity drastically and thus stimulated luxuriant macrophyte growth in Lake St. Clair and the western basin of Lake Erie, which provoked a major shift in the fish community by displacing species sensitive to light (walleye, Sander vitreus (Mitchell, 1818)) and by favoring species adapted to foraging in weed beds (Vanderploeg et al., 2002). In littoral areas, byssally attached mussels (Dreissena and Limnoperna) generally increase benthic community richness and biomass through provision of microhabitat (interstices of clumped shells), grazing surfaces (shells), and nourishment in the form of fecal/pseudofecal deposits (Ricciardi et al., 1997; Ward and Ricciardi, 2007; Burlakova et al., 2012). However, these invasive bivalves also exclude some species of invertebrates through resource competition and interference (Ricciardi et al., 1997; Lauer and McComish, 2001). Intense fouling of the exposed shells of native molluscs by dreissenids (Figure 5.2) causes negative impacts (Ricciardi et al., 1998; Van Appledorn et al., 2007; zu Ermgassen and Aldridge, 2010; Sousa et al., 2011), including severe population declines (Ricciardi et al., 1998).

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FIGURE 5.2  Dreissenid mussels fouling a unionid bivalve (Lampsilis radiata) from the St. Lawrence River.

FIGURE 5.3  The killer shrimp Dikerogammarus villosus. Photo courtesy of J.T.A. Dick.

Introduced predatory invertebrates have caused conspicuous biodiversity loss in benthic and pelagic communities. The amphipod Dikerogammarus villosus, known as the “killer shrimp” (Figure 5.3), is a voracious predator of benthic invertebrates. It is rapidly eliminating native and alien gammarid populations from parts of western Europe (Dick et al., 2002), and these species displacements may have consequences for ecosystem functioning (MacNeil et al., 2011). The introduction of the Eurasian spiny water flea Bythotrephes longimanus Leydig, 1860, into Canadian Shield lakes has apparently resulted in rapid and persistent declines in native zooplankton richness (Boudreau and Yan, 2003). A similar predator, the fishhook water flea Cercopagis pengoi (Ostroumov, 1891), has invaded the Baltic Sea and the lower Great Lakes. Cercopagis feeds readily on small-body cladocerans; its population expansion during a 3-year period in Lake Ontario coincided with sharp declines in the abundance of dominant members of the zooplankton community, including two daphniid species that have been observed to be preyed upon by C. pengoi in lab experiments. One of these

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species declined despite exhibiting high fecundity during all 3 years (Laxson et al., 2003). After the appearance of C. pengoi in the Baltic Sea, densities of small-body cladocerans and copepods declined, particularly above the thermocline (Kotta et al., 2006). Where it occurs in dense patches, the C. pengoi population is estimated to consume tens of thousands of copepods per cubic meter per day, which implicates it as the cause of a drastic decline in copepod abundances in the Gulf of Finland (Lehtiniemi and Gorokhova, 2008). Another group of crustacean predators, mysid shrimp were stocked as a supplementary food source for sportfish in many North American and European lakes during the latter half of the previous century. Through vertical migration in the water column, mysids escape predation from planktivorous fish, including those species whose diets they were meant to supplement. The mysids themselves exert strong predation pressure on large zooplankton, often causing severe (>60%) reductions in cladoceran density and biomass within one to two decades after their introduction (Ricciardi et al., 2012). These reductions may lead to declines in the growth, abundance, and productivity of pelagic fishes. Mysid invasions can also alter the cycling of heavy metals and other contaminants. Vertical migrations of mysids increase the transfer of heavy metals between benthic and pelagic communities; and by lengthening the food chain, introduced mysids may increase the biomagnification of polychlorinated biphenyls and mercury contaminants in fish (Rasmussen et al., 1990; Cabana et al., 1994). The potential for mysids to generate cascading effects in food webs is exemplified by the well-documented introduction of the opossum shrimp Mysis diluviana Audzijonyte and Väinölä, 2005 (formerly Mysis relicta) into Flathead Lake, Montana, as a prey item for another introduced species, kokanee salmon Oncorhynchus nerka (Walbaum, 1792). The opossum shrimp not only avoided predation by the diurnally feeding salmon, but also outcompeted it for dwindling food resources and thus caused the salmon population to crash. The loss of salmon was followed closely by declines in local populations of grizzly bears and bald eagles, which depended on salmon spawners as a food source (Spencer et al., 1991).

CAN THE IMPACTS OF ALIEN INVERTEBRATES BE PREDICTED? A more predictive understanding of the impacts of invertebrate invasions is needed for prioritizing management interventions (Ricciardi, 2003) and for interpreting water quality assessments accurately that are based on benthic invertebrate community composition and relative abundance (MacNeil and Briffa, 2009). Unfortunately, only a few generalizations can be made. Invertebrate taxa that cause the greatest ecological impacts tend to be functionally unique in the invaded environment (Ricciardi and Atkinson, 2004), such that they use critical resources differently than native species (e.g., by displaying a unique form or a greater rate

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of consumption) (Carlsson et al., 2004; Dick et al., 2013). They are typically predators or grazers that are unregulated by higher consumers because of their trophic position, recruitment rate, or antipredatory defenses. They generally have a greater fecundity and abundance than related native species (Keller et al., 2007; Gherardi, 2011), and—in the case of crustaceans—are more aggressive (Gamradt et al., 1997; Dick, 2008). An invader’s impacts can vary along environmental (e.g., temperature, oxygen, conductivity) gradients, such that they differ at various sites within a single heterogeneous aquatic system. Consequently, the invader may extirpate a native species at one site but coexist with it elsewhere (Jokela and Ricciardi, 2008; Kestrup and Ricciardi, 2009). For example, the impetus for introducing the opossum shrimp M. diluviana into lakes beyond its native range in North America was derived largely from the positive response of kokanee salmon to the mysid when it was introduced to Kootenay Lake, British Columbia. The hydrology of the lake was such that it provided upwelling currents that flushed out the mysid to make it available to salmon during daylight hours; thus, it was an inappropriate model to predict the outcome of such introductions in most deep lakes in which mysids can migrate to profundal areas (Martin and Northcote, 1991). The composition of the invaded community also influences impact. The effects of even highly invasive species may be modified by the presence of predators, pathogens, or competitively dominant species (Kuhns and Berg, 1999; Ward and Ricciardi, 2010; Kestrup et al., 2011). The context dependency of impact presents the largest impediment to risk assessment. However, ecologically damaging invaders often have a history of strong impacts in other invaded regions (e.g., Ricciardi et al., 2012). This history, if documented sufficiently, can be used to generate predictive models for those species, taking into consideration key environmental variables that have been identified as modulators of the invader’s impact (Ricciardi, 2003; Ward and Ricciardi, 2007). An intuitive, but largely untested, hypothesis is that introduced invertebrates that cause strong ecological impacts also cause strong socioeconomic impacts (e.g., on outdoor recreation, fisheries, water supply systems). Species that are otherwise rather innocuous to ecological communities may still pose health risks to humans and domestic animals, if they are vectors of disease. Even commercially valuable invertebrate species can cause unforeseen economic damage (Gherardi, 2011).

MANAGEMENT OF INVASIVE AQUATIC INVERTEBRATES Although the costs of prevention may be far outweighed by chronic costs of impacts, management has, traditionally, been reactive to invasion threats, usually responding after

Chapter | 5  Ecology of Invasive Alien Invertebrates

a problematic species has become well established. Con­ sequently, successful eradication attempts are extremely rare. Efforts to contain, reduce, or remove alien invertebrate populations have used biological, chemical, and mechanical controls (e.g., Pointier and David, 2004; Wittmann et al., 2012). The benefits of these efforts are often limited by the lack of early detection and the difficulties of managing an expanding population. Targeted harvesting and the cultivation of native fish predators can aid in reducing populations of particularly harmful invaders (Robinson and Wellborn, 1988; Hein et al., 2007; Aquiloni et al., 2010). Highly invaded systems contain greater numbers of ecologically and economically damaging species (Ricciardi and Kipp, 2008), an observation that justifies management efforts to reduce invasion rates even where numerous invasions have already occurred. Given the apparently increasing accumulation of alien species in large lakes and rivers (Figure 5.1), recent efforts to manage invertebrate invasions have focused on controlling major vectors, particularly ballast water transport. Substantive progress has been made in developing control strategies for ballast water-mediated introductions (e.g., Bailey et al., 2006); however, the invasion risk posed by resistant diapausing stages in ballast tanks remains a management challenge. Attention must also turn to fostering policies for largely unregulated vectors, such as those associated with live trade (e.g., for food markets, ornamental/pet trade, use as fishing bait) (Keller and Lodge, 2007). Lastly, public outreach, including education campaigns that target anglers, boaters, and pet owners, may aid in limiting the spread of alien invertebrates.

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Laxson, C.L., McPhedran, K.N., Makarewicz, J.C., Telesh, I.V., MacIsaac, H.J., 2003. Effects of the non-indigenous cladoceran Cercopagis pengoi on the lower food web of Lake Ontario. Freshw. Biol. 48, 2094–2106. Lee, C.E., Bell, M.A., 1999. Causes and consequences of recent freshwater invasions by saltwater animals. Trends Ecol. Evol. 14, 284–288. Lehtiniemi, M., Gorokhova, E., 2008. Predation of the introduced cladoceran Cercopagis pengoi on the native copepod Eurytemora affinis in the northern Baltic Sea. Marie Ecol. Prog. Ser. 362, 193–200. Leuven, R.S.E.W., van der Velde, G., Baijens, I., Snijders, J., van der Zwart, C., Lenders, H.J.R., bij de Vaate, A., 2009. The river Rhine: a global highway for dispersal of aquatic invasive species. Biol. Invasions 11, 1989–2008. MacNeil, C., Briffa, M., 2009. Replacement of a native freshwater macroinvertebrate species by an invader: implications for biological water quality monitoring. Hydrobiologia 635, 321–327. MacNeil, C., Dick, J.T.A., Platvoet, D., Briffa, M., 2011. Direct and indirect effects of species displacements: an invading freshwater amphipod can disrupt leaf-litter processing and shredder efficiency. J. North Am. Benthol. Soc. 30, 38–48. Martin, A., Northcote, T., 1991. Kootenay Lake: an inappropriate model for Mysis relicta introduction in north temperate lakes. Am. Fish. Soc. Symp. 9, 23–29. Marsden, J.E., Hauser, M., 2009. Exotic species in Lake Champlain. J. Gt. Lakes. Res. 35, 250–265. McMahon, R.F., 2002. Evolutionary and physiological adaptations of aquatic invasive animals: r selection versus resistance. Can. J. Fish. Aquatic Sci. 59, 1235–1244. Miller, A.W., Ruiz, G.M., Minton, M.S., Ambrose, R.F., 2007. Differentiating successful and failed molluscan invaders in estuarine ecosystems. Mar. Ecol. Prog. Ser. 332, 41–51. Mills, E.L., Leach, J.H., Carlton, J.T., Secor, C.L., 1993. Exotic species in the Great Lakes: a history of biotic crises and anthropogenic introductions. J. Gt. Lakes. Res. 19, 1–54. Mills, E.L., Scheuerell, M.D., Carlton, J.T., Strayer, D.L., 1997. Biological invasions in the Hudson river Basin. N.Y. State Mus. Circ. (57), 59. Morrison, W.E., Hay, M.E., 2011. Feeding and growth of native, invasive and non-invasive alien apple snails (Ampullariidae) in the United States: invasives eat more and grow more. Biol. Invasions 13, 945–955. Morton, B., 1997. The aquatic nuisance species problem: a global perspective and review. In: D’Itri, F. (Ed.), Zebra Mussels and Aquatic Nuisance Species. Ann Arbor Press, Chelsea, Michigan, pp. 1–53. Muskó, I.B., Balogh, C., Tóth, Á., Varga, É., Lakatos, G., 2007. Differential response of invasive malacostracan species to lake level fluctuations. Hydrobiologia 590, 65–74. Muskó, I.B., Bence, M., Balogh, C., 2008. Occurrence of a new PontoCaspian invasive species, Cordylophora caspia (Pallas, 1771) (Hydrozoa: Clavidae) in Lake Balaton (Hungary). Acta Zool. Acad. Sci. Hung. 54, 169–179. Oscoz, J.O., Tomas, P., Duran, C., 2010. Review and new records of nonindigenous freshwater invertebrates in the Ebro River basin (Northeast Spain). Aquat. Invasions 5, 263–284. Padilla, D.K., Williams, S.L., 2004. Beyond ballast water: aquarium and ornamental trades as sources of invasive species in aquatic systems. Front. Ecol. Environ. 2, 131–138. Panov, V.E., Krylov, P.I., Riccardi, N., 2004. Role of diapause in dispersal and invasion success by aquatic invertebrates. J. Limnol. 63 (Suppl. 1), 56–69.

Chapter | 5  Ecology of Invasive Alien Invertebrates

Phelps, H.L., 1994. The Asiatic clam Corbicula fluminea invasion and system-level ecological change in the Potomac River Estuary near Washington, DC. Estuaries 17, 614–621. Pointier, J.P., David, P., 2004. Biological control of Biomphalaria glabrata, the intermediate host of schistosomes, by Marisa cornuarietis in ponds of Guadeloupe: long-term impact on the local snail fauna and aquatic flora. Biol. Control 29, 81–89. Poirrier, M.A., 1990. Freshwater sponges (Porifera: Spongillidae) from Panamá. Hydrobiologia 194, 203–205. Rasmussen, J.B., Rowan, D.J., Lean, D.R.S., Carey, J.H., 1990. Food chain structure in Ontario lakes determines PCB levels in lake trout (Salvelinus namaycush) and other pelagic fish. Can. J. Fish. Aquatic Sci. 47, 2030–2038. Rasmussen, J.L., 1998. Aquatic Nuisance Species of the Mississippi River Basin. 60th Midwest Fish and Wildlife Conference Aquatic Nuisance Species Symposium, Cincinnati, Ohio, 16 p. Reynoldson, T.B., Smith, B.D., Maitland, P.S., 1981. A species of North American triclad (Paludicola; Turbellaria) new to Britain found in loch Ness, Scotland. J. Zool. 193, 531–539. Ricciardi, A., 2003. Predicting the impacts of an introduced species from its invasion history: an empirical approach applied to zebra mussel invasions. Freshw. Biol. 48, 972–981. Ricciardi, A., 2006. Patterns of invasion in the Laurentian Great Lakes in relation to changes in vector activity. Divers. Distrib. 12, 425–433. Ricciardi, A., Atkinson, S.K., 2004. Distinctiveness magnifies the impact of biological invaders in aquatic ecosystems. Ecol. Lett. 7, 781–784. Ricciardi, A., Avlijas, S., Marty, J., 2012. Forecasting the ecological impacts of the Hemimysis anomala invasion in North America: lessons from other freshwater mysid introductions. J. Gt. Lakes. Res. 38 (Suppl. 2), 7–13. Ricciardi, A., Kipp, R., 2008. Predicting the number of ecologically harmful species in an aquatic system. Divers. Distrib. 14, 374–380. Ricciardi, A., MacIsaac, H.J., 2011. Impacts of biological invasions on freshwater ecosystems. In: Richardson, D.M. (Ed.), Fifty Years of invasion Ecology: the Legacy of Charles Elton. Wiley-Blackwell, Oxford, pp. 211–224. Ricciardi, A., Neves, R.J., Rasmussen, J.B., 1998. Impending extinctions of North American freshwater mussels (Unionoida) following the zebra mussel (Dreissena polymorpha) invasion. J. Anim. Ecol. 67, 613–619. Ricciardi, A., Reiswig, H.M., 1994. Taxonomy, distribution and ecology of the freshwater bryozoans (Ectoprocta) of eastern Canada. Can. J. Zool. 72, 339–359. Ricciardi, A., Whoriskey, F.G., Rasmussen, J.B., 1997. The role of the zebra mussel (Dreissena polymorpha) in structuring macroinvertebrate communities on hard substrata. Can. J. Fish. Aquatic Sci. 54, 2596–2608. Robinson, J.V., Wellborn, G.A., 1988. Ecological resistance to the invasion of a freshwater clam, Corbicula fluminea: fish predation effects. Oecologia 77, 445–452. Sousa, R., Pilotto, F., Aldridge, D.C., 2011. Fouling of European freshwater bivalves (Unionidae) by the invasive zebra mussel (Dreissena polymorpha). Freshw. Biol. 56, 867–876.

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Chapter 6

Economic Aspects of Freshwater Invertebrates Vincent H. Resh Department of Environmental Science, Policy, and Management, University of California, Berkeley, California, USA

David M. Rosenberg Freshwater Institute, Winnipeg, Manitoba, Canada

Chapter Outline Freshwater Invertebrates in Commerce 93 Aquaculture93 Human Diets: The Invertebrate–Fish Connection 94 Freshwater Pearls and Insect Jewelry 95 Biomonitoring97 Nuisance Aquatic Insects 98 Trichoptera98 Ephemeroptera98 Diptera98 Benefits or Damages Caused by Introduced or Invasive Species100 Direct Market Economic Benefits and Damages 100 Benefits100 Damages101

Freshwater invertebrates serve directly as food for humans and indirectly as food for other organisms that we eat. Moreover, they provide both positive and negative economic consequences in terms of products that we use and the diseases that they transmit. In this chapter, we will concentrate on some of the main economic consequences of these organisms. Certainly they are used in commerce in a plethora of other ways, ranging from the esoteric, small-scale businesses (e.g., selling the eggs of some true bugs for pet turtle food) to large-scale and expansive projects (e.g., attempts to create specially designed habitats to conserve threatened or endangered freshwater invertebrate species).

FRESHWATER INVERTEBRATES IN COMMERCE Aquaculture Scores of species, mostly fish and plants but also some arthropods and molluscs, are cultivated in a variety of natural and specially

Indirect Nonmarket Economic Benefits and Damages 102 Benefits102 Damages102 Medicinal Leeches 103 Stresses to Livestock and Wildlife from Biting Flies 104 Diseases Vectored by Freshwater Invertebrates 104 Incidence104 Habitats of Human Disease Vectors 105 Insect Vectors of Human Disease 106 Snails as Intermediate Hosts of Human Disease 106 Crustaceans as Intermediate Hosts of Human Disease 107 Acknowledgments107 References107

designed systems for use as food (several crustaceans), in hobbies (food for tropical fish), and for bioremediation (freshwater mussels). Freshwater prawns and crayfish are the most widely cultivated freshwater invertebrates, and they are grown in many regions throughout the world. Farmed freshwater prawns are all in the genus Macrobrachium; and, until recently, the giant river prawn or Malaysian prawn, Macrobrachium rosenbergi, was the only species widely cultivated (New, 2002). However, a large aquaculture program in China now involves Macrobrachium nipponense, and some other species are grown at smaller scales. Likely, most prawn farming in the world today involves M. nipponense and is done in China, which is remarkable given that widespread prawn cultivation only began there in the early 2000s. Prawn aquaculture is complicated by the life cycle of the species (Resh et al., 1992). Adult prawns live in freshwater, but larvae require brackish water to develop and survive. In nature, they have amphidromous life cycles, which involve their breeding and egg hatching in freshwater, but the newly

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00006-1 Copyright © 2015 Elsevier Inc. All rights reserved.

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hatched larvae must return to an estuary or the ocean to survive and undergo embryonic development (Resh et al., 1990; Resh and deSzalay, 1995; Myers et al., 2000). The technology for prawn farming involves hatcheries that produce “post larvae” that require about one month’s residence in saline conditions. Individuals are then transferred to freshwater ponds when they grow to market size and are harvested, often by draining these ponds, which is an approach similar to that used for aquaculture of marine shrimp. Macrobrachium are highly aggressive, so stocking densities must be considerably lower than those used for marine shrimp (New, 2002). The consequent reduction in waste products is one reason is why freshwater-prawn farming is considered to be less environmentally damaging than the farming of marine shrimp. However, lower yields often mean that larger areas are placed in cultivation. The Chinese mitten crab, Eriocheir sinensis, has a similar life cycle to prawns and is cultured in China because of the high value of its roe (Rudnick et al., 2003; Rudnick and Resh, 2005). This species has invaded many parts of the world, causing damage to levees and fishing nets (Rudnick and Resh, 2002), and it has been suggested that the economic value of its roe may have resulted in its purposeful dispersal by humans (Rudnick et al., 2005). Crayfish are also extensively farmed worldwide, and China and the United States mainly culture Procambarus clarkii, the red swamp crayfish. In Europe, both native (Astacus astacus, the noble crayfish) and nonnative North American (Pacifastacus leniusculus, the signal crayfish) species are cultured. Management systems in these areas differ greatly (Holdrich, 1993; Ackefors, 2008). For P. clarkii, low stock densities and little active management are the norm. However, P. leniusculus are often more intensively managed, with brooding females kept in indoor tanks and hatchery-raised eggs and juveniles reared in outdoor tanks or small ponds. Juveniles are released into lakes and waterways, and this crayfish species is rarely grown to full size in captivity as is done with prawns and P. clarkii. Native Australian crayfish in the genus Cherax are also being exported for use in aquacultural systems in other parts of the world (Jones, 1990). These species seem to counter the disadvantage of P. clarkii, whose burrowing behavior and frequent escapes have caused damage in rice paddies and competition with native species (Rudnick and Resh, 2002). Predation of freshwater mussels may also be a future problem with escapees. In all aquacultural schemes involving freshwater crayfish and other crustaceans (e.g., river crabs and prawns), disease is a major economic issue. Inevitably, the less than optimal conditions present in aquaculture compared to their natural environment have resulted in higher levels of disease than typically found in nature. Viruses, bacteria, and parasites are a constant drain on the potential profits of freshwater invertebrate aquaculturalists (Holdrich, 1993;

SECTION | II  General Ecology and Human Impacts

Ackefors, 2008), which is also a problem in fish aquacultural operations.

Human Diets: The Invertebrate–Fish Connection Freshwater fisheries are important economically in commercial, recreational, and subsistence fishing activities. Such fisheries depend on the health of fish populations, which in turn partly depends on the invertebrate food available to them. Early compendia of freshwater fishes covering large geographic areas (e.g., MacPhail and Lindsey, 1970; Scott and Crossman, 1973) list many species of fish feeding on freshwater invertebrates (i.e., zooplankton and zoobenthos). More recent studies using both direct analysis of gut contents and indirect stable isotope analysis have also documented the importance of a variety of freshwater invertebrates in the diet of fish in rivers (e.g., Little et al., 1998; Pilger et al., 2010) and lakes (e.g., Vander Zanden and Vadeboncoeur, 2002). The importance of freshwater invertebrates as fish food has been amply shown by attempts to introduce nonindigenous species of invertebrates as fish food (Ricciardi, 2011). For example, predatory mysid shrimp (Mysis, Hemimysis, Limnomysis, Neomysis) have been stocked as a supplementary food source for fishes in North American and European lakes—but not always with the desired consequences (see below and Chapter 5 for details). The accumulation of contaminants in fish may be influenced by their diets (Little et al., 1998). Mercury is the single most cited compound in fish-consumption advisories in the United States and Canada, and most of the scientific effort in studying mercury contamination of aquatic food webs is prompted by human health risks (Wiener et al., 2007). Toxicological problems from mercury in aquatic ecosystems result from exposure to methylmercury, a highly toxic compound that can biomagnify to high concentrations in predators occurring at the top of food webs. Methylmercury in fish is obtained almost entirely by dietary uptake of invertebrates and the fish that feed at lower trophic levels. Freshwater invertebrates are important in the trophic transfer of methylmercury to fish. Zooplankton are eaten by many fish and by early life stages of some fish that become piscivores later in their life cycle. Moreover, benthic invertebrates are important in the diets of many species of fish, birds, and mammals, and some freshwater benthic invertebrates (e.g., crayfish) are consumed by humans, which provides a direct pathway for exposure to methylmercury (Wiener et al., 2007; see also below). Crayfish have long been economically important as specialty foods and cuisines in restaurants. Restaurants in North America use farmed or wild-caught crayfish, whichever is available at reasonable cost. For example, restaurants in

Chapter | 6  Economic Aspects of Freshwater Invertebrates

the Lake Tahoe area are being supplied with wild-caught P. leniusculus from the lake. These crayfish were introduced to the lake in the 1800s and could now number as many as 300 million individuals. This commerce resulted from a local entrepreneur convincing the Nevada State Assembly to allow harvesting of this invasive species in an attempt to clarify the shallow waters of the lake. They are popularly marketed as a “local” food, and their harvesting presumably may reduce their numbers in the lake.

Freshwater Pearls and Insect Jewelry Pearls have been a symbol of elegance and beauty for millennia (Figure 6.1). Produced in some form by almost all species of shelled mollusc, pearls of greatest value and widest use come from marine oysters or from freshwater mussels in the family Unionidae. The pearl, like the shell of the mollusc, is composed of calcium carbonate and results when a foreign object enters the interior of the mussel shell and cannot be expelled. The mussel coats the object with its shell-building material, nacre, to reduce irritation from the object. In natural pearls, this object enters as a parasite or, typically, a sharp object. However, in cultured pearls, the object is deliberately inserted into the tissue of the mollusc (Hua and Roubo, 2002). During the Middle Ages in Europe, dense mussel beds in much of Northern Europe were the source of the decorative pearls used by nobility in their garments and by the Church to enhance art objects. They were embroidered into cloth and placed as integral parts of a variety of decorative objects such as religious icons and items (Figure 6.2). The species most

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commonly used was Margaritifera margaritifera (Figure 6.1(a)). Even earlier, freshwater pearls were used in decorations in China and other parts of Asia, and were more highly valued than the marine pearls by royalty. The Hopewell Native American culture in the midwestern United States also used pearls in decorations as far back as 2000 years ago. Many species of freshwater mussels have been harvested for their pearls in North America (Sweeney and Latendresse, 1984). In the 1800s, the discovery of beautiful pearls in many species of mussels inhabiting the rivers of the midwestern United States led to a resurgence of wild mussel collections in the hope of making similar finds in other regions. The effect of these collections on native mussels was devastating (Neves, 1999; Anthony and Downing, 2001). The most valuable freshwater pearls have typically occurred in the wild, but high-quality ones are relatively rare in nature. Anecdotally, irregular-shaped but aesthetically appealing pearls are found in about 1% of natural mussels examined, whereas higher-quality rounded ones occur only in about 0.01% of mussels collected. Cultured pearls are a major aquaculture industry today in China (Hua and Roubo, 2002), but the industry’s origins extend over 2500 years ago when tiny Buddha images were inserted into freshwater mussels to produce “blister” pearls, which are typically attached to the shells (Fiske and Shepard, 2007). Japan and the United States also have had limited pearl-culture operations, but they never reached the technological advances or commercial scale seen in China today. Cultured pearls are often generically called “biwas” because of the extensive aquaculture operations done with the pearly mussel (Hyriopsis schlegeli) in Lake Biwa in Japan, and the

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FIGURE 6.1  (a) Freshwater pearl mussel (Margaritifera margaritifera) from the river Navarån, Västernorrland, Sweden. Photograph courtesy of Joel Berglund from Wikipedia Commons. (b) Natural pearls in the shell of Tampico pearl mussel (Cyrtonaias) in Texas, which is harvested noncommercially. (c) Market for Hyriopsis pearl mussels in China. (b) and (c) Photographs courtesy of Chris Barnhart.

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(a)

(b)

(c)

(d)

FIGURE 6.2  (a) and (b) Russian Orthodox icons decorated with freshwater pearls. (c) Strand of cultured freshwater pearls in jewelry. (d) Jewish prayer shawl decorated with freshwater pearls. (a)–(c) Photographs courtesy of Cheryl Resh; (d) photograph courtesy of Nathan and Martha Rosenberg.

wide assortment of beautiful colors of pearls from that area. The Lake Biwa pearl industry collapsed because of pollution from industry and human population stresses. Techniques in the culturing of freshwater pearls have varied over time (Akamatsu et al., 2001; Fiske and Shepard, 2007) but generally involve slightly opening the mussel shell and inserting small pieces of live mantle tissue into slits made into the mussel’s mantle. The insertion of this tissue begins the nacre production. As many as 50 grafts of mantle tissue are added to a single mussel, but newer approaches use fewer grafts, and these produce higher-quality pearls. Likewise, the shift to different species and even hybridized mussels has enhanced the quality of the pearls produced. The culturing of mussels in their freshwater

habitat generally lasts from 2 to 6 years, after which they are harvested. Two pearls often result from each insertion, resulting in 24–32 pearls extracted per mussel (Hua and Roubo, 2002). The natural hue of pearls produced in culture is typically a lavender color, but bleaching produces white pearls and dying has produced a variety of colors, even “chocolate pearls”! Freshwater pearls are fashionable for their beauty but also for economic incentives. For example, the cost of a freshwater pearl is only about 20% of a cultured marine pearl. Culturing of marine pearls also uses the nucleation process, in which hard pieces of shell are inserted into the oyster (Bondad-Reantaso et al., 2007). Today, freshwater mussel shells from the United States are used as nuclei in

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the culture of marine pearls in oysters throughout the tropical waters of French Polynesia (where pearls produced are known commercially as Tahitian black pearls), China, and Japan; and although it is a specialized industry, it can be highly profitable. Freshwater mussels were long collected for the pearls they contained; but at the beginning of the twentieth century, the production of pearl buttons from mussel shells was a large industry in the midwestern United States. Mussels were collected from many large rivers in Illinois, Iowa, New York, and New Jersey (Anthony and Downing, 2001). Europeans at this time tended to make pearl buttons from Indo-Pacific marine shells; but by the mid-twentieth century, the United States market became predominant. However, the pearl-button industry was doomed by lower-cost plastic buttons and changing fashions, in which buttons were replaced by lower-cost and more practical zippers. Mussel collecting continues today (although it is highly regulated in terms of species that can be collected and where collections can be made), but it is largely for preparing nuclei for the culturing of marine pearls in oysters. Unfortunately, illegal harvesting has led to the decline of some native mussel species. Freshwater insects are also used in decoration, but typically as images rather than actual specimens. For example, dragonflies are a common motif in jewelry, textiles, and tattoos. It may be that these insects (along with butterflies) provide bilateral symmetry when their wings are at rest, and consequently have aesthetic and artistic appeal. There is, however, at least one use of aquatic insects in jewelry: the West Virginia-based firm Wildscape produces caddisfly-case pendants from the cases of emerged Pycnopsche caddisflies that are reared at large scales in their laboratory (www.wildscape.com accessed 10.01.13.).

BIOMONITORING Freshwater invertebrates can have economic importance in the biomonitoring of water quality in lakes and rivers, in terms of both the resulting decisions (e.g., consequent interventions to improve water quality or abandonment of projects) and the size of the industry that monitors freshwater invertebrates (Carter and Resh, 2001). Biomonitoring with freshwater invertebrates can be done at several different spatial and temporal scales (Table 6.1). By far the most common scale is the community level. Benthic invertebrates are the most commonly used group for biomonitoring (Rosenberg and Resh, 1993). They are used by a number of countries as the basis for national river health programs (e.g., RIVPACS, the River Invertebrate Prediction and Classification Scheme in the UK, AUSRIVAS, the Australian River Assessment System in Australia), partial national programs (e.g., BEAST, Benthic Assessment of Sediment in the Laurentian Great Lakes, and for Canadian rivers in British Columbia, the Yukon Territories,

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TABLE 6.1  Scales of Biomonitoring Using Freshwater Invertebrates and Examples of Analytical Techniques Appropriate at Each Scale Scale

Examples

Biochemical

Enzyme activities; RNA, DNA, amino acids, protein; ion regulation

Physiological

Respiratory metabolism, scope for growth

Individual

Morphology, behavior, life history, bioaccumulation

Populations and species assemblages

Biotic indices, multivariate approaches

Community

Taxa richness, enumerations, diversity indices, similarity indices, biotic indices, functional feeding group ­measures, multivariate approaches

Ecosystem

Structure of food webs, productivity, decomposition

See Rosenberg et al. (2008) for references to examples. Adapted from Rosenberg et al. (2008).

and the eastern maritime provinces), and in state-sponsored programs (e.g., multimetric analyses in the United States) (Wright et al., 2000). The RIVPACS, AUSRIVAS, and BEAST programs are all examples of the reference condition approach to bioassessment (Bailey et al., 2004), which is a powerful way to examine the environmental effects of water quality and both human and natural influences. An example of the economic implications of bioassessment using freshwater invertebrates at a local scale is illustrated in the decisions being made by the owner of a base-metal mine on a tributary of the Fraser River in British Columbia, Canada. This owner wanted to know if his mine effluent was having a deleterious effect on a nearby stream. A bioassessment, using baseline data already generated by the BEAST program in the area, coupled with new benthic macroinvertebrate samples and attendant physical/chemical data required by the BEAST, was taken in areas downstream of the discharge of the effluent. When compared to the BEAST database of unimpacted sites in the same geographic area of the stream, the mine owner could determine if the community of benthic invertebrates in the stream differed from that expected under unimpacted conditions. If impact was evident, remediation of the effluent could be ordered, and recovery of the stream could be followed over time using similar procedures. Therefore, because the cost of the bioassessment and any remediation required would be borne by the mine owner, continued operation of the mine could be evaluated on the economics of the remediation costs versus profits from mining.

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NUISANCE AQUATIC INSECTS Mass emergences of some aquatic insect species (e.g., Trichoptera, Ephemeroptera, and Diptera) can deface human property, result in economic losses to businesses, pose traffic hazards, and cause allergic reactions. Perhaps the most common human response is repulsion at the aesthetic mess, including the smell of rotting carcasses, created by such mass emergences.

Trichoptera One of Canada’s earliest attempts at controlling mass emergences of caddisflies was the “Shadfly Project” connected with Expo’67, the World Exposition held on Île St. Hélène in Montreal. Trichoptera emergences from the St. Lawrence River are a fact of life in Montreal: “Montreal is the only major Canadian city completely surrounded by water and, according to the experts, is the shad fly capital of Canada. Shad flies are harmless insects that live near freshwater, but in May and June, hordes of these winged pests invade waterfront terraces—so be careful they don’t land in your drink” (Tourisme Montreal, 2006, p. 11). Approximately 16,000 kg of the insecticide DDD (dichlorodiphenyldichloroethane, a DDT derivative) was applied to the St. Lawrence River in the mid-1960s in an attempt to control the populations of nuisance Trichoptera (Graham, 2012). DDD residues are still detectable in the river today. Stiege (2004) examined the human health effects (workrelated allergies) of Trichoptera mass emergences in and around hydroelectric generating stations on the Winnipeg River in Ontario. She recommended changes in management practices to decrease exposure of Manitoba Hydro employees to caddisfly particulates entering the stations and potentially being inhaled. Similar health problems have occurred from massive caddisfly emergences from the Niagara River in New York. A recent publication of the Iowa State University Horticulture and Home Pest News (Gissel, 2012) documented, with photos, a May 2012 mass emergence of Trichoptera in Bettendorf, Iowa. The publication brightly noted: “Mass emergences of caddisflies, like the better known mayflies, are temporary and the annoyance will pass. In the meantime, look on the plus side: large numbers of caddisflies indicate a healthy river!” (Gissel, 2012, p. 1).

Ephemeroptera A spectacular photo of Hexagenia bilineata adults attracted to the headlights of a car on the Mississippi River bridge at Winona, Wisconsin, on July 8, 1966 is provided in Fremling (1968, Figure 1). The mayflies are piled over the front bumper of the car, up to its headlights. “The Hannibal CourierPost reported that on the night of 10th July, two motorcycles and four cars were involved in a series of accidents on the

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Mark Twain Bridge at Hannibal, Missouri. The vehicles, which had slid into each other on a 6-in carpet of mayflies, were removed, and a snow plow was used to clear the slippery bridge. Cinders were then applied to insure traffic over the bridge” (Fremling, 1968, p. 280). Fremling (1968) also listed a number of areas around the Great Lakes, the Illinois River, and the Mississippi River where mayfly abundances and mass emergences were drastically reduced because of water pollution in the nymphal habitats. A recent report in the Portage Daily Register, Wisconsin (http://m.wiscnews.com/portagedailyregister/news/article_ Oda668de-c7e1-11e1-9f42-001a4bcf887a.html accessed 20.10.12.), indicated a mass emergence of mayflies on a bridge over the Wisconsin River on July 6, 2012. The article also mentioned that large emergences in major communities like La Crosse (on the Mississippi River) have been occasionally detected on the National Weather Service radar screens. The South Basin of Lake Winnipeg in Manitoba is dotted by resorts and privately owned cabins. Mass emergences of Hexagenia limbata and Hexagenia rigida are a regular occurrence (Figure 6.3), usually in July (D.G. Cobb, pers. com., October 30, 2012). The extent of emergence varies from year to year. Carman (1999) reported an especially heavy emergence around Gimli, Manitoba, in July 1999. Local residents complained about the unpleasant odors resulting from piles of dead mayflies at the town dump and lagoon. The owner of a local restaurant even reported a decline in business as a result. More recently, pest control companies have been active in spraying cabins in the Gimli area to kill spiders that are feeding on the emerging mayflies (D.G. Cobb, pers. com., October 30, 2012). Cobb, a retired entomologist, also reported being approached for advice on timing by people planning an outdoor wedding, in an effort to escape the worst of the annual mayfly emergence! In Winnipeg Beach, a resort town near Gimli, a local art gallery is named the “Fishfly Gallery” in honor of the prominent summertime feature of the area. Likewise, a hotel along the shoreline of Lake Victoria, Uganda, is named the “Shad Fly Hotel” because of the massive emergence of Povilla mayflies!

Diptera The classic story of nuisance emergences of freshwater Diptera may be the Clear Lake gnat (Chaoborus astictopus) and the historical efforts to control it in this large California lake (e.g., Suchanek et al., 2002) (Figure 6.3(c)). Before pesticide applications to the lake began in the 1940s to control these nuisance midges, large piles of dead gnats appeared below street lights and gnat swarms were so thick they impeded vehicular traffic along the edge of the lake. Pedestrians wore kerchiefs to avoid inhaling the flies.

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(a)

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(b)

(c)

FIGURE 6.3  (a) Common swarming mayflies Hexagenia rigida from the Red River, Manitoba. (b) Car parked under a light during mass emergence of Hexagenia from Lake Winnipeg, Manitoba, Canada, July 2013. (c) Clear Lake gnats (Chaoborus astictopus) attracted to house lights. (a) and (b) Photographs courtesy of Donald Cobb and Margaret Friesen; (c) Photograph courtesy of Lake County (California) Mosquito Abatement District.

DDD application began in 1949 in an effort to boost tourism around the lake and improve the local economy. DDD applications to the lake continued until 1957, when the deleterious effects on the Western grebe became obvious. In view of this early knowledge, later application of DDD to control caddisfly emergences from the St. Lawrence River for Expo’67 (see above) is amazing! Methyl parathion was substituted for DDD to control the gnat from 1962 to 1975. The gnat’s population in Clear Lake is now much lower than before and is believed to be kept in check by two introduced species of fish (threadfin shad, Dorosoma petenense, and the Mississippi silverside, Menidia beryllina) that compete for zooplankton with the gnat larvae. Unfortunately, these fish also compete with economically important species used for recreational purposes. The more common nuisance dipteran may be chironomid midges, also in the insect order Diptera, whose nuisance mass emergences have been recorded from a variety of natural and man-made waters from all over the world (Ali, 1991). As with Trichoptera, these nuisance mass emergences can cause allergic reactions in humans. Nuisance mass emergences from man-made waters have been recorded from residential-recreational lakes in southern California (Mulla, 1974 (see his Figure 2); Ali and Mulla, 1979), water impoundments in southern California (Ali and Mulla, 1978) and Florida (Ali and Baggs, 1982), and concrete-lined flood control channels in Los Angeles and Orange County, California (Ali and Mulla, 1976;

Ali et al., 1976). The City of Sanford, Florida, reported an annual loss of $3–4 million because of midge emergences from adjacent Lake Monroe and a nearby water-cooling reservoir (Ali and Baggs, 1982). Chironomid mass emergences affect homes on residentialrecreational lakes by “…defacing of stuccos and interior walls, ceilings, draperies and other furnishings” (Mulla, 1974, pp. 172–173). Spiders follow these mass emergences and spider webs further deface buildings. “In one instance, large numbers of adult midges invaded surgical wards and patient rooms of a modern hospital, dead midges were swept from beds and equipment daily for a period of 2–3 weeks” (Mulla, 1974, p. 173). According to Ali and Mulla (1978, p. 122), massive emergences of midges from water impoundments in the Santa Ana River Basin, California, caused the following problems: “At times, from April to November swarms become so dense that outdoor business, residential or recreational activities are hampered. At night adults are attracted to lights, swarming around indoor and outdoor fixtures. Dead midges and spider webs, where they are trapped, deface stucco and other wall finishes requiring frequent washing and maintenance. Midges also invade manufacturing facilities, swarm inside and get imbedded in paint finishes, plastics, and other manufactured goods, causing substantial economic loss.” Mosquito abatement agencies in the San Francisco Bay Area have compiled data indicating that most complaints about mosquitoes that they respond to are actually the result

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of presence of adult chironomid midges, which are nonbiting. Moreover, more complaints are filed by residents of areas where higher-cost homes exist, presumably reflecting increased intolerance with nuisance insects as home investment increases!

BENEFITS OR DAMAGES CAUSED BY INTRODUCED OR INVASIVE SPECIES This section examines the economic effects of introduced or invasive species; a detailed treatment of environmental effects of invasive freshwater invertebrates is presented in Chapter 5. Introduced or invasive species can have positive economic effects (e.g., many of the species raised in aquaculture operations for food are not from the region) or they can have negative economic effects (e.g., species can destroy valuable crops or clog water-intake pipes). Unfortunately, the negative outcomes of invasive species are far more frequent than the positive outcomes. The discussion will consider two kinds of economic effects: (1) direct market economic benefits or damages, in which economic costs or benefits are obvious or can be assigned an actual dollar value; and (2) indirect nonmarket economic benefits or damages, in which changes to ecosystems are obvious but cannot be assigned a dollar value.

Direct Market Economic Benefits and Damages Benefits Freshwater invertebrates such as prawns, snails, and crayfish have been introduced to areas in which they are not naturally found as part of aquaculture operations to produce human food. The global value of aquaculture in 2007 has been estimated as US$89.5 billion (Bartley, 2011). In addition, several species of freshwater invertebrates, mainly curculionid beetles, have been introduced to control invasive nuisance aquatic plants. Crayfish introductions have been deliberate, for commercial reasons: (1) aquaculture and both legal and illegal stocking; (2) live food trade; (3) aquarium pet trade; (4) live bait; (5) snail and weed control; and (6) supplies for science classes (Gherardi, 2011). For example, the crayfish or yabby Cherax destructor was introduced to Australian farm dams for aquaculture in 1932. Procambarus “Marmorkrebs,” a parthenogenetic crayfish, is sold in human food markets in Madagascar and in aquarium shops in Germany (MacIsaac, 2011). This crayfish is also called “North American marbled crayfish” (Gherardi, 2011, p. 130). Procambarus clarkii has been extensively cultivated in its natural range in the southern United States since the 1950s, but because of its commercial value, it was introduced to several more US states and elsewhere in the world. It even was introduced to Africa

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in the 1960s and onward to control freshwater snails that carried schistosomiasis, and to Spain in 1973 for economic and social reasons. Some species of crayfish are highly valued as specialty gourmet foods (e.g., in Louisiana and Scandinavia). Some species are commercially harvested from wild stocks (e.g., Lake Tahoe, described above) or have become aquacultural commodities in countries such as Australia, China, and the United States (Gherardi, 2011). Orconectes rusticus (the rusty crayfish) is widely used as live bait, which is largely the reason for its spread in North America. Despite the many problems associated with crayfish as invasive species (see below), this group of freshwater invertebrates has been credited with many positive impacts on humans: (1) restoring of crayfishing as a cultural tradition in Sweden; (2) providing economic benefits in poor areas such as southern Spain; (3) inducing development of cultivation systems in China; and (4) increasing the volume of international trade from countries such as China and Spain (Gherardi, 2011). Freshwater snails have also been deliberately introduced, mainly for commercial purposes: (1) food (including aquaculture); (2) aquarium industry; (3) biological control; (4) aesthetics; and (5) biological research (Cowie, 2011). For example, the ampullarid snail Pomacea canaliculata (the golden apple snail) was introduced from South America to Southeast Asia in the 1980s for aquacultural programs to supplement local food and attempt to develop a gourmet export industry. This species was taken to Hawaii for the same reasons. Cipangopaludina chinensis and Cipangopaludina japonica, which are freshwater viviparid snails, were probably introduced to the United States from Asia, also as food (Cowie, 2011). When freshwater snails are moved around the world for commercial purposes or sold in pet stores for domestic aquaria, they often either escape or are purposely released into local waters (e.g., Pomacea spp. and Marisa cornuarietis (Ampullaridae), C. chinensis (Viviparidae), Helisoma spp. and Planorbis spp. (Planorbidae), Physidae, and Radix auricularia (Lymnaeidae)). Various species of Ampullaridae and Thiaridae have also been introduced to control snail vectors of human schistosomiasis (Cowie, 2011), although transfers of some species of the latter family have also been involved in the geographic spread of trematodes that are major human parasites (Resh et al., 1992). Many species of freshwater invertebrates have been used to help control aquatic invasive plants such as Eichhornia crassipes (water hyacinth) and Salvinia molesta (giant salvinia) (Evans and Strong, 2011; Hill et al., 2011; MacIsaac, 2011). Water hyacinth produces a large number of human impacts in that it: (1) reduces the quality and quantity of drinking water; (2) increases the production of vectors and consequently the disease incidence of malaria, encephalitis, and various types of filariasis (see below); (3) increases siltation and sedimentation of rivers, lakes, and impoundments; (4) reduces useful water surface area for fishing, recreation,

Chapter | 6  Economic Aspects of Freshwater Invertebrates

and water transport; (5) clogs irrigation canals and pumps; (6) interrupts hydroelectric power generation; and (7) enhances flood damage to road and rail bridges and impoundment walls (Hill et al., 2011). Several biological control agents have been used for controlling water hyacinth: Neochetina eichhorniae and Neochetina bruchi (Coleoptera: Curculionidae), Niphograpta albiguttalis and Xubida infusella (Lepidoptera: Pyralidae), Eceritotarsus catarinensis (Hemiptera: Miridae), Megamelus scutellaris (Homoptera: Delphacidae), and Orthogalumna terebrantia (galumnid mite). The weevil Cyrtobagous salviniae (Curculionidae) has been used to help control S. molesta, and M. cornuarietis (an ampullarid snail) has been used to help control Pistia stratiotes (water lettuce) (Cowie, 2011). These biological control agents have met with varied success, more so in tropical than in temperate areas (Evans and Strong, 2011; Hill et al., 2011).

Damages Unfortunately, the commercial damage done by introduced species often outweighs the economic benefits described above. For example, when the North American crayfish P. leniusculus was introduced to Europe, it reintroduced the “crayfish plague” disease (the oomycete Aphanomyces astaci) that had previously dramatically reduced production of native crayfish species (Garcia-Berthou and Moyle, 2011; Gherardi, 2011; MacIsaac, 2011). It was especially devastating when it carried the fungus to European and Scandinavian populations of the native and economically important crayfish A. astacus (Bartley, 2011). Because the North American species is a resistant carrier that outcompetes native European crayfish, the plague confers an additional competitive advantage by weakening the European species. The reoccurrence of the plague led to considerable economic damage in Scandinavia, Germany, Spain, and Turkey, with a loss of >90% in the production of A. astacus and Astacus leptodactylus (the narrow-clawed crayfish) (Gherardi, 2011). Moreover, annual production of A. leptodactylus in Turkey in the 1980s fell from 7000 to 2000 tons, nearly eliminating exports from there to Europe. Very few crayfish importations to Africa have been successful, but these imports have spoiled the collection of commercially valuable fish species and damaged fishing nets (Gherardi, 2011). Moreover, P. clarkii has become a pest of rice culture in various parts of the world (e.g., in Portugal it caused a >6% decline in profits). Burrowing by several crayfish species can be a problem in agricultural and recreational areas (e.g., lawns, golf courses, levees, dams, dykes, canal irrigation systems) and can cause bank destabilization in rivers and lakes. The latter has also been a concern in Chinese mitten crab introductions, whether these introductions were intentional or not (Rudnick et al., 2003). There has been a high failure rate in attempts to eradicate or control invasive crayfish species. These attempts are

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usually costly (Gherardi, 2011). For example, the reintroduction of Pacifastacus fortis (the Shasta crayfish) to California cost $4.5 million, and the unsuccessful eradication of P. leniusculus in Scotland cost ∼100,000 pounds. Invasive crayfish can also have adverse effects on human health (Gherardi, 2011). For example, they can potentially transfer contaminants such as metals to their consumers. P. leniusculus and P. clarkii may transfer cyanobacterial toxins to humans. Procambarus clarkii may be an intermediate host of many helminth parasites of vertebrates and a vector for human tularemia (caused by Francisella tularensis bacterium). Introduced snail species have also had negative economic consequences. For example, anticipated markets in Asia for the introduced golden apple snail P. canaliculata never developed, so snails were discarded into local waterways (Bartley, 2011). The snails became established in rice fields, where they fed on young rice plants. In the Philippines, not only was 20–40% of the value of rice production lost, but these snails also were implicated in the reduction of native biodiversity, including the snail preferred for human food, Pila luzonica. In Thailand, introduction of P. canaliculata changed the steady state of wetland ecosystems (see below) (Evans and Strong, 2011). The changes led to eutrophication, among other effects, which posed a threat to local agricultural, fishing, and tourism industries. Pomacea canaliculata and Pomacea insularum brought into Hawaii as potential human food have become serious pests of taro crops (Cowie, 2011). Golden apple snails also threaten rice crops in other parts of the United States. Pomacea spp. have been implicated in the vectoring of the rat lungworm Angiostrongylus cantonensis, which can cause eosinophilic meningoencephalitis in humans (Cowie, 2011). Introduced freshwater lymnaeids are vectors of cattle liver flukes (Fasciola hepatica) and Potamopyrgus antipodarum (the New Zealand mud snail) is a vector of fish and bird parasites in New Zealand and possibly in other areas to which they have spread (McDowall, 2011). Finally, the costs of eradication, control, or prevention of snail invasive species can be costly (Cowie, 2011). For example, control of Pomacea spp. in Asia entails the use of pesticides, various cultural methods, and hand collection and destruction of their egg masses. All of these procedures add to the cost of agricultural operations. Damage caused by one of the most costly examples of an invasive species, the zebra mussel Dreissena polymorpha, has already exceeded $100 million (Johnson, 2011). This species causes two general categories of impacts: (1) mechanical or physical effects of biofouling (e.g., in municipal drinking water and industrial cooling systems); and (2) changes in the environment caused by the invasion and associated impacts on recreational and commercial exploitation of resources (e.g., declines in fish abundance or accumulation of shell debris or macrophytes on recreational beaches).

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The zebra mussel and the quagga mussel Dreissena bugensis have also infested water sources used by aquaculture in North America (Bartley, 2011), and farms with high infestations have had to be closed. Farm and water management protocols and monitoring are used in high-risk areas to prevent further spread of these species, with attendant costs to the aquaculture operation. Costs of attempts to control or eradicate zebra mussels (e.g., outreach efforts to change the behavior of boaters and fishers; chlorine additions at municipal and industrial facilities) are already high (Johnson, 2011). Moreover, climate change may lead to increased control costs, as warmer temperatures will result in faster growth around facilities like water intake pipes that must remain open for a variety of human needs (Dukes, 2011). Dreissenids also play a role in the transmission of disease, such as botulism (caused by the bacteria Clostridium botulinum) outbreaks in water birds in the lower Great Lakes (Mills and Holeck, 2011). This subject is covered in detail in Chapter 5.

Indirect Nonmarket Economic Benefits and Damages Benefits It is hard to ascribe positive effects to the introduction of the zebra mussel. However, this species and other invasive mussels have improved water clarity in many of the lakes that they have invaded (MacIsaac, 2011). In fact, Dutch researchers have introduced zebra mussels into eutrophic lakes to improve water clarity and reduce algal biomass. However, the perceived drawbacks in most of the Northern Hemisphere exceed potential benefits, and zebra mussels are not intentionally stocked into lakes there. In fact, the opposite is true in that a number of North American jurisdictions have banned the live importation of zebra mussels. Extensive accumulations of mussel shells, such as from zebra mussels, enhance the structural complexity of lake beds, especially those composed of mud or sand, and consequently improve invertebrate species diversity and abundance (MacIsaac, 2011). Many taxa of invertebrates also benefit from the accumulation of organic wastes in the feces and pseudofeces of the mussels. Moreover, invasive crayfish can be prey for fish, birds, and mammals, thus providing a new resource for higher trophic levels (Gherardi, 2011).

Damages Introduced or invasive species can act as “ecosystem engineers” in their new habitats, causing gross changes in ecosystem states. For example, the introduction of golden apple snails to Thailand resulted in wetlands previously dominated by macrophytes becoming dominated by phytoplankton (Evans and Strong, 2011), and threatened local wetland plant communities and rice crops (see above).

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After consuming macrophytes, the snails excreted previously unavailable phosphorus nutrients to the water, which created increases in eutrophic conditions. The increase in nutrients and decreases in light penetration, with the latter caused by reduced shading by macrophytes, allowed phytoplankton to become the dominant primary producer in the ecosystem. Moreover, these ecosystem-level changes disrupted interactions among native species. Introduced snails can also seriously affect native biota. For example, P. canaliculata and P. insularum have been implicated in the decline of native freshwater snail species in Southeast Asia and Florida (Cowie, 2011). Other ampullarids have had the same effect in the Caribbean islands. In North America, the New Zealand mud snail, P. antipodarum, is outcompeting native stream snails. In Florida, the introduced ampullarid M. cornuarietis dramatically reduces freshwater plants, destroying habitat for native animals associated with these plants (Cowie, 2011). In the Caribbean, M. cornuarietis and Pomacea glauca, introduced to control water lettuce, may also have affected native vegetation. Procambarus clarkii had similar ecosystem-engineering effects as P. canaliculata on wetlands in Europe (Zedler, 2011). By grazing and shredding macrophytes, this crayfish removes plants, increases turbidity, and converts marshes into open water. The energy base consequently shifts to phytoplankton, filamentous algae, and detritus. The crayfish O. rusticus can, in some North American lakes to which it has spread, directly or indirectly reduce abundance and species diversity of macrophytes through herbivory, destruction, or increased turbidity (MacIsaac, 2011). These two species of crayfish have also had negative effects on other invertebrate and vertebrate taxa (MacIsaac, 2011; Zedler, 2011). P. leniusculus, from North America, preys on the young of the European crayfish A. astacus, which is one of the reasons for the reduction of native species where the nonnative species is found (Bartley, 2011). In contrast, the zebra and quagga mussels D. polymorpha and D. bugensis cause system-wide increases in water clarity and light penetration, leading to reduced phytoplankton offshore and increased growth of macrophytes inshore, i.e., a shift from planktonic to benthonic processes (Johnson, 2011; Mills and Holeck, 2011). In their role as ecosystem engineers, these dreissenids also caused considerable declines in abundances of the burrowing amphipod Diporeia in the lower Great Lakes (Mills and Holeck, 2011). Prior to the invasion, Diporeia composed 60–80% of the benthic biomass in these lakes and was a critical food for lake whitefish, Coregonus clupeaformis. The loss of Diporeia has implications for the capacity of the Great Lakes to support fish populations that depend on this food resource, and for how energy will flow to these fish. For example, crustacean zooplankton abundance in Lake Huron declined sharply because of the focusing of predators on the remaining zooplankton (MacIsaac, 2011). Four other species of fish also steeply

Chapter | 6  Economic Aspects of Freshwater Invertebrates

declined in Lake Michigan since 1999 (MacIsaac, 2011), and dreissenid invasion of the Great Lakes also decimated many species of native unionid mussels (Johnson, 2011; MacIsaac, 2011; Mills and Holeck, 2011; Primack, 2011). Other species of introduced or invasive species of aquatic invertebrates have also caused dramatic ecological effects. For example, because the invasive zooplankters Bythotrephes longimanus (the spiny water flea) and Cercopagis pengoi (the fishhook water flea) have long tail spines that make them unpalatable to planktivorous fish, they escape substantial predation (Ricciardi, 2011). Also, Bythotrephes reduced abundance and species richness of zooplankton in several European and North American lakes (Ricciardi, 2011) and changed the food web in the Great Lakes by direct predation on other zooplankton, thus affecting sport and commercial fishing (MacIsaac, 2011). Cercopagis is suspected of causing the decline of dominant species of zooplankton in Lake Ontario (Ricciardi, 2011). Mysis diluviana (the opposum shrimp, formerly M. relicta) was introduced into Flathead Lake, Montana, and Kootenay Lake, British Columbia, to enhance kokanee salmon (Oncorhynchus nerka) production (MacIsaac, 2011; Ricciardi, 2011). These introductions, however, produced the opposite effect (see Chapter 5 for details). Introductions of mysids into other lakes have also caused a variety of ecological problems: (1) severe reductions in zooplankton; (2) enhanced bioaccumulation of mercury and polychlorinated biphenyls; (3) increased parasitism of fishes by nematodes, cestodes, and

(a)

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acanthocephalans; and (4) declines in growth, abundance, and productivity of pelagic fishes (Ricciardi, 2011). In conclusion, introduced or invasive species can have economic benefits or produce economic damages. On balance, however, the prospect of short-term economic gain needs to be balanced against the possibility of major, long-term negative consequences from localized ecological changes, changes in ecosystem state, and the economic implications of these changes. In anticipating or rectifying problems that result from introduced species, detailed life-history information on species that are candidates for introduction or for species that have already invaded an area in which they normally do not occur is a critical research need. It is also a need that will become increasingly important in the future.

MEDICINAL LEECHES The attachment of blood-sucking leeches to humans brings about almost universal disgust. The memorable scene from the classic movie The African Queen in which Humphrey Bogart is covered with leeches is horrifying to everyone who sees the movie, as are the personal experiences of leeches attaching to humans when swimming in ponds and slow-moving rivers. However, this disgust was not always the case. For perhaps 2500 years, some species of leeches have been used for medicinal purposes, with attachment and blood-sucking being done purposefully (Whittaker et al., 2004; Elliott and Kutschera, 2011, Figure 6.4(a)).

(b)

FIGURE 6.4  (a) Engraving illustrating the story of the king who was so fat that he tried to become thinner using medicinal leeches. From Histoires Prodigieuses, by Pierre Boaistuau, 1560. (b) A surgeon extracting a guinea worm from a man’s leg, in an engraving by J. Luyken. Both plates courtesy of Wellcome Library, London.

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All leeches are predators or parasites, and both their sense organs and their flexible bodies allow them to pursue their prey effectively. Several species of leeches have been used for medical purposes, but the one most commonly used is the European medicinal leech Hirudo medicinalis. This leech, originally described by Carl Linneus in 1758, can reach 20 cm in length when fully grown. Although generally believed to feed almost exclusively on the blood of mammals, this species also feeds on the blood of fish, aquatic birds, and amphibians (Elliott and Kutschera, 2011). The medicinal leech finds its prey using heat detection and chemosensory stimuli. In feeding, leeches pierce the skin and inject both anticoagulants and anesthetics before sucking out the blood of their victims. Large adults can consume 10 times their mass in a single feeding of blood. In medieval and early modern times, medicinal leeches were used to remove blood from a patient to balance the four “humors” of ancient medical philosophy—blood, phlegm, and black and yellow bile (Whittaker et al., 2004). These humors had to be kept in balance for the human body to function properly. Likewise, if a sickness caused the skin to become red (such as from fever or inflammation), this inflammation must have arisen from too much blood in the body. Bleeding with leeches was also used to treat behavioral problems. A person who was “too strident” or “too sanguine” was considered to have an excess of blood, which caused this behavior. Phlebotomy with leeches was widely used throughout the eighteenth and nineteenth centuries in Europe and also in North America (where the leeches had to be imported), even though some studies conducted during this time questioned their value as medical treatments. Leeches are also used in modern medicine. They provide an effective way of reducing blood coagulation, stimulating circulation in reattachment operations for organs with critical blood flow (such as eyelids, fingers, and ears), treating abscesses, avoiding necrosis and gangrene, and healing venous diseases and thrombosis (e.g., Conforti et al., 2002). The saliva of medicinal leeches contains hirudin, which is presumed to be the most powerful anticoagulant known. Some reports of modern use of leeches in medicine even suggest that they can cure infertility! Some side effects of modern leech therapy are often alluded to in articles on specific treatments, such as allergic reactions, bacterial infections, and, not unexpectedly, prolonged bleeding. Leeches also have a place in modern biological research. Neurologists use them as experimental animals because of their simple central nervous systems and exceptionally large neurons (e.g., Lamb and Calabrese, 2011). The former and present popularity of leeches in medical treatments has diminished natural-occurring medicinal leech populations. Some researchers have suggested that human bloodletting did not contribute to their decline, because after use they were frequently discarded into ditches or ponds (Elliott and Kutschera, 2011). However, contemporary

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collecting for neurobiology and medicinal studies does not involve the return of leeches to their habitat. The use of watering troughs also may have displaced potential cattle and horse hosts from coming into contact with leeches, and drainage of ponds and other medicinal-leech habitats likely has also contributed to their decline. In any event, the medicinal leech is currently listed as a Threatened Species by the Union for the Conservation of Nature (Elliott and Kutschera, 2011).

STRESSES TO LIVESTOCK AND WILDLIFE FROM BITING FLIES In addition to being important as pests and disease vectors of humans, adults of black flies in the family Simuliidae, whose larvae and pupae live in flowing waters, can be important pests of livestock and wildlife in areas of the United States, Canada, and many other parts of the world. Intense attacks by black flies can result in a variety of effects on cattle, such as: toxic shock reactions to the saliva of black flies; accidents occurring when animals try to escape attack, which can kill livestock; reduced milk production; reduced animal growth rates; reduced conception rates; mortality of calves and other cattle not previously exposed to black flies; and increased susceptibility to pneumonia and other stress-related respiratory disease (http://www.agf.gov.bc.ca/cropprot/blackfly.htm; Bechinski and Klowden, undated: “Black flies—­ Biology and control,” www.extension.uidaho.edu/ipm/documents/­ blackflies.pdf both accessed 17.12.12.). For example, along the Saskatchewan River in Saskatchewan, 1100 cattle were killed by Simulium arcticum during an outbreak from 1944 to 1947 (The Canadian Encyclopedia: http://www.thecanadianencyclopedia.com/articles/ black-fly accessed 17.12.12.). Outbreaks of the same species along the Athabasca River in northern Alberta in 1971 caused an average weight loss of 45 kg per animal and the death of 973 cattle in one region. In Idaho, Simulium vittatum is a severe pest of sheep, cattle, and horses, although animal deaths are rare (Bechinski and Klowden, undated; see above). In contrast, poultry and hogs are less frequently affected. Outbreaks of S. vittatum have been recorded in the 1930s, 1970s, 2001, and 2004. In Canada, past black fly control attempts led to the illadvised application of chlorinated hydrocarbon pesticides to large rivers (e.g., see Flannagan et al., 1979). More reasonable, smaller-scale control methods are described in Bechinski and Klowden (undated; see above).

DISEASES VECTORED BY FRESHWATER INVERTEBRATES Incidence Diseases that are transmitted by invertebrates that live in freshwater have been a major force in shaping both the size

Chapter | 6  Economic Aspects of Freshwater Invertebrates

of the human population and the development of our civilization. Certainly, far more people have been killed by diseases transmitted by freshwater invertebrates than by all of the wars that have been fought in the history of the world. For example, during the Vietnam War, many times more casualties resulted from malaria than from combat on both sides of the conflict (Resh, 2009). A variety of human diseases are transmitted by invertebrate vectors that have life cycles associated with various types of water bodies, including lakes, ponds, rivers, streams, reservoirs, and irrigated fields. Of these diseases, malaria, schistosomiasis, lymphatic filariasis (elephantiasis), onchocerciasis (river blindness), Japanese encephalitis, and dengue generally are considered to be the most important in terms of their infection rate, morbidity (i.e., either the incidence or the prevalence of a disease), or mortality (i.e., the number of deaths resulting from a particular disease). Human diseases that are vectored by freshwater invertebrates have been reviewed in detail by Resh (2009); the main vectors and the diseases that they transmit are summarized below. Diseases vectored by freshwater invertebrates have had, and continue to have, a devastating toll on human life. For example, the World Health Organization in their annual malaria reports emphasize that malaria is still endemic in over 100 developing countries and with over two billion humans at risk. Around 80–90% of malaria cases occur in the poorest countries of the world in Africa, but malaria is also a major problem in parts of South America, India,

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and Southeast Asia. Between 100 and 200 million people are estimated to be infected worldwide, and 1–2 million deaths generally occur per year (even higher during epidemic years). However, the widespread distribution and use of pesticide-impregnated bed nets reduced malaria deaths to below one million in 2011. An annual estimate of US$1.7 billion in lost productivity and treatment costs apply to sub-Saharan Africa alone.

Habitats of Human Disease Vectors Three important categories can be distinguished that provide habitat for vectors and intermediate stages of the causative agents of human diseases: (1) natural water bodies, (2) human-made water bodies, and (3) water bodies that form in human settlements and household environments. Some freshwater vectors of human disease can occur in all of these habitat categories. In terms of natural water bodies, streams and rivers are sources of the black flies that serve as vectors of onchocerciasis, and lakes and ponds provide habitats for snails that are intermediate hosts of schistosomiasis Although mosquito vectors do occur in natural systems as eggs, larvae, and pupae, their densities tend to be far higher when they occur in human-made or settlement habitats, probably because of reductions in predation and competition compared to those in natural systems (Resh, 2009). Likewise, standing water in barrels and pots in human settlements, habitats where predation and competition are also lacking,

(a)

(b)

(c)

(d)

FIGURE 6.5  (a) An adult female Anopheles gambiae mosquito taking a blood meal and possibly vectoring malaria. (b) Fully grown larvae of this mosquito species. (c) An adult Simulium damnosum female black fly taking a blood meal and possibly vectoring river blindness. (d) Fully grown larva of that species. (a)–(d) Photographs courtesy of WHO/TDR/Stammers.

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are important habitats for certain disease-transmitting mosquitoes. The creation of human-made water bodies such as impoundments and irrigation ditches often results in hydrological changes that favor intensified vector breeding. These changes can result in increased Anopheles mosquito populations that may transmit malaria or in appropriate snail populations that may increase prevalence of schistosomiasis. Likewise, shifts in species composition may occur that reduce numbers of predators or competitors and allow vectors to increase in number. Recent studies have also suggested that habitat disturbance may increase the prevalence of Buruli ulcer in ponds in Africa (Merritt et al., 2005, 2010).

Insect Vectors of Human Disease Biting flies (order Diptera) are the most important of the aquatic vectors of human disease. Of the flies, mosquitoes (family Culicidae) are the most important vectors, both because of the high mortality and morbidity of the many diseases they cause and also because of the range of diseases for which they can serve as vectors (Spielman and D’Antonio, 2001, Figure 6.5(a) and (b)). For example, certain species of mosquitoes can transmit many types of nematodes, protozoans, or viruses (Table 6.2). However, it should be remembered that although about 3000 described species of mosquitoes take blood for nourishment, most are nuisance biters (and some not even of humans) and do not transmit human diseases. Other Diptera are also important in disease transmission to humans and their animals. Black flies (family Simuliidae) in the Simulium damnosum species complex transmit onchocerciasis, which is caused by a nematode roundworm, Onchocerca volvulus (Remme, 2004; Yaméogo et al., 2004, Figures 6.5(c) and (d) and 6.6(b)). In contrast to mosquitotransmitted diseases, in which the vector occurs in still or at least very slow-moving water, onchocerciasis is vectored by black flies, whose immature stages occur attached to vegetation in fast-flowing streams (Resh et al., 2004). The disease has been successfully controlled in West Africa through insecticide applications and antihelminthic drugs by the World Health Organization Onchocerciasis Control Program (Lévêque et al., 2003; Resh et al., 2004). The disease also occurs in some parts of South America, where the nematode parasite was introduced through transport of African slaves and when a native black fly was a competent vector in transmitting the worm and infecting other humans. Many species of biting flies that occur in moist or semiaquatic habitats as larvae or that are common riparian-­dwelling species as adults also transmit diseases. For example, some sand flies (family Ceratopogonidae) can transmit the protozoan causing leishmaniasis, tsetse (Glossinidae) transmit the protozoan causing African sleeping sickness, and some deer and horse flies (Tabanidae) can transmit Loa loa, an eye worm affecting humans (Resh, 2009).

SECTION | II  General Ecology and Human Impacts

TABLE 6.2  Examples of Human Diseases Vectored by Mosquitoes Disease Vectored

Parasite Transmitted

Lymphatic filariasis

Brugia and Wucheria nematodes

Malaria

Plasmodium protozoans

Yellow fever

Flavovirus

Dengue

Flavovirus

St. Louis encephalitis

Flavovirus

Japanese encephalitis

Flavovirus

Murray Valley encephalitis

Flavovirus

Russia spring–summer encephalitis

Flavovirus

Omsk hemorrhagic fever

Flavovirus

West Nile fever

Flavovirus

Kyasanur forest disease

Flavovirus

Louping ill

Flavovirus

California encephalitis

Bunyavirus

La Crosse encephalitis

Bunyavirus

Rift Valley fever

Bunyavirus

Eastern equine encephalitis

Togavirus

Western equine encephalitis

Togavirus

Venezuelan encephalitis

Togavirus

Ross River fever

Togavirus

Adapted from Resh (2009).

Even biting flies that do not transmit disease can be major nuisances. In North America, horse flies, deer flies, and black flies are irritating pests, oftentimes with painful bites that produce allergic reactions and infections that result from scratching the bites (Resh, 2009). There is also an economic cost associated with them in terms of lost tourist revenue and lower human-population densities in areas where they are a problem. The effects of biting flies on livestock and wildlife are discussed above.

Snails as Intermediate Hosts of Human Disease Freshwater snails are the intermediate hosts of a variety of trematode flukes and some nematodes (roundworms) that cause many human diseases (Ross et al., 1997, Figure 6.6(c) and (d)). The most important of these diseases is schistosomiasis (sometimes referred to as Bilharzia) (Mahmoud, 2001). This parasitic trematode (blood fluke) expelled in human wastes in ponds and other still-water habitats must

Chapter | 6  Economic Aspects of Freshwater Invertebrates

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(a)

(b)

(c)

(d)

FIGURE 6.6  (a) X-ray of a knee showing a calcified Guinea worm, Dranunculus medinensis. (b) Onchocerca volvulus, the adult nematode worm that causes onchocerciasis. (c) The cercaria of S. haematobium that penetrates human skin and causes schistosomiasis. (d) S. haematobium, the adult trematode worm that causes schistosomiasis. (a) Photograph courtesy of Welcome Library, London. (b) Photograph courtesy of WHO/TDR/OCP; (c) and (d) Photographs courtesy of WHO/TDR/ Stammers.

be ingested by a particular species of snail to continue its life cycle (Secor and Colley, 2005). The ecological requirements of these snails are a key determinant in the distribution and prevalence of this disease. Schistosomiasis is a major public health problem in the world, and blood flukes in general are important parasites of cattle and other large animals that humans depend on for survival. Although schistosomiasis commonly occurs in developing countries, an infection related to this disease that occurs in developed countries is cercarial dermatitis or “swimmer’s itch” (Verbrugge et al., 2004). This disease occurs worldwide and affects people that are swimming, wading, or working in littoral areas of both marine and freshwater habitats. Infection results when cercaria meant to penetrate the skin of birds such as ducks or small mammals such as muskrats instead penetrate human skin, eliciting an immune response in the individual.

Crustaceans as Intermediate Hosts of Human Disease Another human disease that has had devastating effects in Africa and parts of the developing world is Guinea worm or dracunculiasis (Resh, 2009, Figure 6.6(a)). The large nematode Dranunculus medinensis (∼1 m long) causing this disease releases larvae from an adult female worm (usually embedded in a human leg) through a skin lesion, when an infected human comes in contact with a pond or well. A larva is then ingested by a water flea (Cyclops), in which it develops and becomes infective. When a person drinks this infected water,

the Cyclops is dissolved by the acidity in the drinker’s stomach. The nematode larva is then activated, migrates through the subcutaneous tissue, and stays within the new host for about a year. It then emerges and starts the life cycle again. The treatment for Guinea worm that has been used for thousands of years is to slowly wind the emerging worm around a stick over a period of several days (Figure 6.4(b)). If removal is too rapid and the worm dies, septicemia (blood poisoning) may result. This treatment is immortalized in the staff of Asclepius wound with a serpent, which is also the symbol of the modern-day healer or physician. An extensive Guinea worm eradication program sponsored by the Carter Center in the United States (www.cartercenter.org) has been highly successful and has eliminated the disease in most parts of the world, reducing human cases by over 99% (Resh, 2009).

ACKNOWLEDGMENTS We thank C. Barnhart, D. Cobb, D. Hua, M. Friesen, B. Nelms, C. Resh, M. Rosenberg, N. Rosenberg, and D. Strayer for assistance with photographs for our plates. P. Blanchfield, D. Bodaly, D. Cobb, T. Marcese, and S. McTaggart provided information on various economic aspects of freshwater invertebrates.

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Chapter | 6  Economic Aspects of Freshwater Invertebrates

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Chapter 7

Free-Living Protozoa Genoveva F. Esteban Centre for Conservation Ecology and Environmental Science, Faculty of Science and Technology, Bournemouth University, Poole, Dorset, UK

Bland J. Finlay The River Laboratory, Queen Mary University of London, Wareham, Dorset, UK

Alan Warren Department of Life Sciences, Natural History Museum, London, UK

Chapter Outline Introduction113 The Nature of Protozoa as a Group 113 The Nature of Protozoan “Species” 114 Structure and Function of the Protozoan Cell 115 General Characteristics 115 Locomotion115 Food Intake 116 Structural Elements 116 Reproduction116 Biological Diversity of Free-Living Protozoa 116 Functional Groups of Free-Living Protozoa 116 Ameboid Protozoa 118 Flagellated Protozoa 120

INTRODUCTION The Nature of Protozoa as a Group Protozoa are microscopic eukaryotic organisms with ­animal-like features. Each protozoon (the singular form of protozoa) typically exists as a single, independent cell, and all free-living protozoa fit within the definition of phagotrophic (i.e., they ingest particles from the surrounding environment) microbial eukaryotes (see below). They range in size over some four orders of magnitude, from around 2 μm up to 10 mm or more (some deep-sea protozoa reach 10 cm), a length range that exceeds that of vertebrates; the ciliated protozoa alone display a size range comparable to that of all terrestrial mammals (Fenchel, 1987). In some species, independent cells unite to form groups or colonies (e.g., colonial choanoflagellates, colonial peritrichs) (Lee et al., 2000). There has never been

Ciliated Protozoa 122 Other Types of Protozoa 125 General Ecology of Protozoa 126 Functional Roles of Free-Living Protozoa 126 Symbiotic Associations 127 Protozoa and Ecosystem Function 127 Life in Low-Oxygen Environments 128 Groundwater Protozoa 128 Collecting and Culturing Protozoa 129 Collecting Protozoa and Assessing Diversity 129 Culturing Protozoa 130 Acknowledgment131 References131

unanimous agreement as to where the boundaries of the protozoa should lie, largely because of the extraordinary diversity of their lifestyles (Margulis et al., 1990). Many species cause disease (e.g., malaria), others thrive as commensals in the digestive tracts of ruminants and woodeating insects (e.g., oxymonad flagellates; Brugerolle and Radeck, 2006), and many plant-like flagellates (e.g., Euglena) have at various times been referred to as protozoa, algae, or plants. Indeed, the systematics of microbial eukaryotes have become so confused that many practitioners eschew the terms “protozoa” and “algae” in favor of the all-embracing term “protists.” Here, we are concerned principally with free-living protozoa; and the definition of this group is not taxonomic, but one that is based on its key function in the natural environment. Specifically, protozoa are capable of phagotrophy— the ability to capture and ingest food particles (Fenchel,

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00007-3 Copyright © 2015 Elsevier Inc. All rights reserved.

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1987). They are found in all types of habitats where free water is available—in fresh waters, in the sea, and in soils and sediments. The need for free water relates to the fact that active protozoa have a naked membrane through which water would be easily lost. However, many protozoa produce resting stages (cysts or spores) protected by a secreted wall, which is usually waterproof; such resting stages may be found in dry habitats as well as wet ones, and even floating in the air. Like most microorganisms, protozoa have very large population sizes, and they are the most abundant group of phagotrophic organisms in the biosphere. Biodiversity at the level of protozoa has characteristics that are not shared by macroscopic animals and plants. Protozoa, like microorganisms in general, are more abundant in habitats where biological productivity is high. One milliliter (1 ml) of open water of a deep lake can contain up to 1000 flagellates, which feed on the planktonic bacteria. The soft sediments of biologically productive ponds and lakes also support large numbers of protozoa (Finlay and Maberly, 2000). Many sediment-dwelling (i.e., benthic) protozoa can live in the absence of oxygen—they are anaerobic. Many of these anaerobic protozoa contain symbiotic bacteria (i.e., bacteria that live in the cytoplasm of the protozoon), which produce the greenhouse gas methane. The ability of protozoa to thrive in organically polluted environments has also benefited mankind. Protozoa living in sewage treatment plants, especially in activated sludge plants, are largely responsible for the clarity of the effluent. Much of the dissolved organic matter in sewage is consumed by bacteria; the protozoa consume the bacteria, and by doing so they control their abundance. This control maintains the physiological capacity or vigor of the bacterial population. The protozoa also secrete substances that flocculate bacteria, removing them from suspension. The end result is that much of the dissolved organic materials end up as particulate material (protozoa and flocs), which is removed by sedimentation, resulting in a relatively clean effluent to be discharged. One milliliter of activated sludge usually contains 100,000 protozoa (Finlay et al., 1988). Most aquatic free-living protozoan species are probably globally ubiquitous—they are usually found in a specific type of habitat wherever that habitat occurs worldwide. The protozoa living in peat soils and penguin guano in the Antarctic are apparently morphologically identical to those living in moorland peat soils and cow pats in Britain. A very useful consequence of this is that identification keys to the protozoa have general applicability worldwide. Another consequence of the ubiquity of protozoa is that the global number of species is relatively small when compared with, for example, insects. A significant proportion of local protozoan species richness, at any moment in time, is rare or

SECTION | III  Protozoa to Tardigrada

cryptic (i.e., dormant) and awaiting the arrival of conditions suitable for growth and reproduction.

The Nature of Protozoan “Species” There is no universal agreement on what constitutes a protozoan “species.” The most widely used concept is the “morphospecies” because it is relatively easy to discriminate a great diversity of protozoa using body form alone. This concept is especially useful because morphology is closely related to ecological function. In many free-living protozoa, the structure of the feeding apparatus and the size and shape of the cell determine how the protozoon functions in the natural environment; therefore, the cell form largely determines the ecological niche that the protozoon occupies. Discriminating species on the basis of form might then be equivalent to discriminating them according to the ecological niches they occupy. We could say that if a protozoon looks the same in different places, then it is the same in different places. However, there are problems with such a simple concept. It is known that a morphospecies can be composed of ecologically distinct populations, such as those adapted for maximum growth rate at different temperatures. Therefore, although it is possible to collect apparently identical representatives of a morphospecies in different corners of the world and one might be encouraged by the belief that they were filling exactly the same niche in these different places, in reality they could be phenotypically quite different. The same problem applies with respect to genetic differences. It is possible to find genetic differences, especially sequence differences in ribosomal RNAs, in morphologically identical isolates collected from different places. The significance of these genetic differences is not known; nor is it known if genetic and phenotypic divergences are correlated. It appears, however, that there is no correlation between genetic divergence and geographic distance. Protozoa spend most of their time as asexual organisms, but several are known to reproduce sexually, if only periodically. Sexual reproduction is likely to be present in the majority, since sexuality is thought to be the ancestral condition for eukaryotes. In some well-studied cases (e.g., the ciliates Tetrahymena and Paramecium), morphologically indistinguishable biological species (reproductively isolated gene pools, also known as sibling species) have been investigated thoroughly. Different apparent “sibling species” within a morphospecies may be genetically identical to each other or extremely divergent (at least with respect to ribosomal RNAs), and there is no apparent correlation between genetic isolation and genetic divergence. Again, there is no good evidence for biogeography of protozoan sibling species, and members of the same sibling species can be found on different continents. It is possible that sibling species carry unique phenotypic traits

Chapter | 7  Free-Living Protozoa

that equip each species for a particular niche, but this has not been demonstrated. The real problem with using a biological species concept for protozoa is that it is not practical for all but a few easily cultivated protozoa. The majority of protozoan species have never been cultivated, and neither the frequency nor the character of their sexual behavior (if any) is known or is ever likely to be known. Most people who study protozoa in the natural environment use the morphospecies concept because it is practical, it embodies the close link between form and function, and it is the morphospecies that fills the niche that can be discerned by the human observer. The morphospecies may contain much genetic variation and be capable of expressing a wealth of phenotypic variation, but it is the best tool available for ordering the diversity that lies within the protozoa. A particularly acute problem with organisms of this size is that morphological resolution is a function of the quality of the microscope being used (Mitchell and Meisterfeld, 2005). Thus, better-quality microscopes enable the discrimination of finer detail. Even individual protozoa in a population might be discriminated one from the other and could, therefore, be referred to as separate morphospecies. Modern optics have undoubtedly lent impetus to the business of describing many new species, some of which are undoubtedly legitimate, whereas many have been established on morphological criteria that are trivial or have no conceivable functional significance. An additional problem with the morphospecies concept is that it is often difficult to decide exactly where a species begins and ends. Although most observations are made of the vast number of individuals that cluster around the central tendency of any population, the individuals at the tail ends of the distributions abut and often overlap those of other species. Morphological variation then appears to be continuous across many species. Examples of this have been described in some of the large spongiose spumellarian radiolarians. The major features (e.g., skeletal and cytoplasmic morphology) that are used to discriminate species intergrade to such an extent that it is not possible to unambiguously ascribe individual radiolarians to nominal species. However, the evolutionary process in radiolarians and other protozoa probably works in the same way as it does for other organisms such that it maintains phenotypically discrete species in a niche space that is essentially continuously variable. The phenotypic traits that sustain these discrete species in protozoa are presumably quite diverse in character and possibly not totally comprehensible. Among the more accessible of these traits are the morphological characters that we use to separate species. However, the likely limitation of these is that they enable us to perceive only the fairly coarsely resolved, sometimes overlapping entities that we call morphospecies.

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STRUCTURE AND FUNCTION OF THE PROTOZOAN CELL General Characteristics Being unicellular, protozoa lack tissues and organs, since these are defined as aggregations of differentiated cells (Hyman, 1940). Difference in size among protozoa means difference in individual cell size. In contrast, body size in multicellular organisms is a function of the number of cells they possess. Differences in protozoan cell size are reflected in certain cellular adaptations; for example, larger protozoa generally have more organelles (e.g., more mitochondria) and larger (and in many cases more) nuclei (Fenchel, 1987). In some protozoa the functions of food capturing, locomotion, and perception, transmission, and response to stimuli are performed by relatively undifferentiated protoplasm; in other cases there is a remarkable degree of functional differentiation. Such differentiations in a unicellular organism are called organelles, which are responsible for all of the functions for their life. These include the cilia and flagella for locomotion and food capture, various other food-catching devices, contractile fibrils for a variety of movements, and surface differentiation and skeletal secretion for protection and to confer shape and rigidity (Hyman, 1940). Protozoa in general have contractile vacuoles to regulate the amount of water inside the cell; they can also have sensory structures such as photoreceptors (Euglena). Most protozoa exhibit a fixed shape and size that are characteristic for each species, but some ameboid protozoa (see below) lack definite form except when a shell is present. There is a great variety of shapes among the protozoa, but spherical, oval, and elongated forms, often flattened, are most common. Larger cells are flattened, allowing transport (in aerobic protozoa) of oxygen by molecular diffusion to the whole of the cell’s inside (Fenchel, 1987).

Locomotion Protozoa move in the environment in three different ways: ameboid movement, flagella, and cilia. The ameboid movement is typical of ameboid protozoa (see below) and some other forms. Movement is achieved by cytoplasmic protrusions known as pseudopodia. Cilia (in ciliated protozoa) and the flagella (typical of flagellates and some ameboid protozoa) propel the organism through the water by their beating, or they are used to generate water currents to draw food particles. Cilia and flagella are structurally very similar, both formed by microtubules that depart from basal bodies (the kinetosomes). However, although similar in structure, cilia and flagella differ in function. Flagella are generally present in very low numbers (one or two); some parasitic flagellates can have up to 10 or 15, and exceptionally (e.g., in the symbiotic trichonymphids) hundreds per

SECTION | III  Protozoa to Tardigrada

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cell, whereas cilia are present in large numbers (in general, tens or hundreds, but some ciliated protozoa have cilia in their dispersive forms only, like in the suctoria). Flagella move by undulatory waves starting at the base of the flagellum (Fenchel, 1987). In contrast, a cilium has one bend only at any one time; cilia beat by metachronal waves caused by an effective force that bends the cilium at the base while the remainder of it is straight, and a recovery wave that causes the cilium to draw back to its initial position close to the cell surface.

Food Intake Some protozoa (e.g., Euglena) are autotrophic (=phototrophy)—they have chloroplasts, which enables them to fix CO2 into organic carbon compounds. Other protozoa have endosymbiotic algae or functional chloroplasts that have been sequestered from ingested algae. The resulting consortia are known as “mixotrophs” because they are capable of both phagotrophy and autotrophy, i.e., they can ingest particles as well as carry out photosynthesis (Esteban et al., 2010). Under some circumstances, especially in laboratory cultures and in habitats with much organic pollution, many protozoa can also be osmotrophic, whereby at least some of the carbon they require diffuses into the cell from the surrounding water. However, most protozoa have mouths and specialized feeding organelles, and their capacity to capture particles (phagotrophy) is what separates protozoa from other unicellular organisms. Phagotrophic protozoa take in organic molecules either as soluble molecules that pass through the membrane or in particulate form by formation of a food vacuole within which the food particle is digested (Warren, 2014). The organic molecules taken in are partly converted to body structures and partly broken down in respiratory processes, which in aerobic protozoa involve mitochondria, to release energy in a form that can be used to drive active processes in the cell.

Structural Elements Protozoa may contain structural elements deposited as an internal skeleton or as a thickened pellicle under the surface membrane, or they may secrete a protective shell, or theca, adhering tightly to the outside of the cell or a looser protective lorica (Figure 7.1) (Sleigh, 2006). Secreted coverings in many protozoa can be temporal or permanent. They may consist of organic material alone or of an organic matrix in which minerals or foreign bodies (such as sand grains) can be embedded. The main minerals found in shells and covering plates (like in some flagellates) are silica and calcium carbonate (for example, in the marine coccolithophores), but others may occur. In dinoflagellates (e.g., Ceratium), a hardened armor of plates covers the cell. They can be

variously ornamented with spines and other types of projections. Other protecting cases observed in protozoa consist of gelatinous envelopes of different shapes that attach to the surface of debris or zooplankton. Many protozoa secrete protective coverings when they pass into a dormant state, for example, resting cysts.

Reproduction Protozoa can reproduce or by both means. Asexual reproduction takes place by binary fission (see below), budding (for example, in suctorian ciliates, which produce an internal bud that gives rise to a dispersive larva), or multiple division. Sexual reproduction is widespread, and all degrees of difference between the two fusing cells are found. The sexual process of “conjugation” (when cells fuse and exchange genetic material), for example, is unmistakable in the ciliates. However, the asexual process of binary fission, in which a parent cell grows and divides into two equal-sized daughter cells, is the most frequently observed reproductive process in protozoa. Many protozoa have complicated life cycles that involve an alternation of sexual and asexual generations. The timing of sex appears to be crucial for the adaptive response in such protozoa, and the process of sexual recombination, generating high genetic diversity at the beginning of the growing season or during the initial stages of colonization, followed by asexual (clonal) reproduction and clonal selection, is a powerful tool for generating local adaptation. Sexual processes do not necessarily result in an increased number of individuals and so are not, strictly speaking, methods of reproduction, but rather methods of genetic recombination.

BIOLOGICAL DIVERSITY OF FREE-LIVING PROTOZOA Functional Groups of Free-Living Protozoa Protozoan morphology (especially of the food-capturing organelles) is closely linked to the way a protozoon functions as a grazer. Therefore, classification of free-living protozoa into broad morphological groups simultaneously allocates them to broad functional groups. The three broad morphological–functional groups, i.e., the ameboid, the ­ flagellated, and the ciliated protozoa, each have their own strengths as a phagotroph. Representatives of all three may feed on the same type of microbes in the same place (e.g., in an aquatic sediment), but they will differ in the mechanics and efficiency of capture of any particular food particle. A filter-feeding flagellate will have a relatively large filter area, a high volume-specific clearance, and competitive superiority over filter-feeding ciliates when grazing on planktonic bacteria. A helioflagellate (with very fine pseudopodia called

Chapter | 7  Free-Living Protozoa

(a)

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(b)

(c)

(d)

(e)

(f)

(g)

FIGURE 7.1  Free-living protozoa. (a) Amoeba proteus with ingested diatoms inside a food vacuole. (b) Arcella sp., a disk-shaped testate ameba; the active cytoplasm and pseudopodia can be observed gathered around the circular aperture of the test. (c) Acineta sp., a suctorian ciliated protozoon with four (two on view only; the other two are hidden behind them) clumps of tentacles for sucking out the prey’s cell content. (d) The ciliated protozoon Euplotes daidaleos with endosymbiotic algae of the genus Chlorella (circular green particles in the picture). (e) Transmission electron micrograph (shadowing staining; after Esteban et al., 2012) of the choanoflagellate Sphaeroeca sp.; note the long whip-like flagellum and the collar surrounding its base to trap the particles drawn with the flagellum. (f) The ciliate Nassula tumida, a proficient predator of filamentous cyanobacteria, with a filament in the process of being ingested through the ciliate’s mouth. (g) The ciliate Vorticella sp. attached to the surface of the diatom Fragillaria.

axopods) and a suctorian ciliate (with sucking tentacles) will both practice diffusion feeding, but the former will be adapted for snaring bacteria, whereas a diffusion-feeding suctorian will specialize in trapping flagellates and ciliates (Figures 7.1 and 7.2). Almost any free-living protozoon can be placed without difficulty in one of these three broad morphological– functional groups. It must be stressed, however, that these groups are not concordant with any system of classification of protozoa published in recent years. Moreover, in most cases they cannot be aligned with the independent lineages that are emerging in the molecular phylogenies (e.g., those based on sequence variation in ribosomal RNAs) that reflect the main episodes in the history of eukaryotic evolution. Heterotrophic flagellate groups,

such as the diplomonads and trichomonads, appear in the early emerging lineages, but other flagellates (e.g., choanoflagellates, chrysomonads, dinoflagellates, and haptomonads) are classified within recently diverging lineages. The ameboid protozoa too are scattered across many lineages. The naked amoebae without mitochondria (the pelobionts) diverge early and close to the diplomonads, whereas the vahlkampfiids, slime molds, and various other groups of naked and testate amoebae appear in other independent lineages that are evolutionarily quite distant from each other. The morphological–functional group of the ciliates is the only one that remains intact, as a monophyletic group, in current molecular phylogenies. The process of distilling the vast quantity of molecular information generated in recent years has resulted in

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SECTION | III  Protozoa to Tardigrada

FIGURE 7.2  A selection from the variety of form and function in ameboid protozoa and slime molds. (a) An actinophryid heliozoon (Actinophrys; diameter ∼0.1 mm) ingesting a flagellate. (b) A benthic foraminiferan (Rotalia) trapping diatoms and bacteria in its reticulopodial net. (c) A naked ameba (Amoeba) using pseudopodia to trap a flagellate. (d) A polycystine radiolarian (Heliosphaera; spherical body ∼0.3 mm) with symbiotic dinoflagellates and an entrapped tintinnid ciliate. (e) Polymorphic life cycles of dictyostelid slime molds (e.g., Polysphondylium; outer circle) and myxomycete slime molds (e.g., Physarum; inner circle). (f) Testate ameba (Assulina; ∼0.08 mm) with its prey (an algal cell) caught on a sticky filopodium.

some entirely new phyletic assemblages. These include: (1) the stramenopiles, a group containing organisms as morphologically and functionally dissimilar as chrysomonad flagellates and diatoms; and (2) the alveolates, a group that embraces the dinoflagellates, the ciliates, and a large group of exclusively intracellular parasites (the apicomplexans) (Adl et al., 2012). In the next sections, we focus on the broad morphological–functional groups of free-living protozoa.

Ameboid Protozoa Rhizopod amebae use pseudopodia (cytoplasmic protrusions) for locomotion and feeding. There are two large groups (Figures 7.1 and 7.2): the “naked amebae” (e.g., Acanthamoeba, Vannella, Amoeba, and Vampyrella) and the shelled “testate amebae” (e.g., Arcella, Nebela, and Euglypha). Most rhizopod amebae feed nonselectively by engulfing diatoms and other algae, unicellular and filamentous

Chapter | 7  Free-Living Protozoa

cyanobacteria, detritus, and bacteria. However, there are notable variants: Vampyrella dissolves a hole in the cell wall of a green alga or desmid and then enters through the hole to digest the cytoplasm of the prey. Some are opportunistic pathogens of man, for example some species in the genera Acanthamoeba and Naegleria. Naked amebae occur in the plankton, but these species are usually very small (1 mm, but most (e.g., Actinophrys) are 40–100 μm. Depending on their size, different species feed with little apparent selectivity on algae, flagellates, ciliates, and rotifers. Small heliozoan species account for most of the biomass of ameboid protozoa in the freshwater plankton. Most, however, are attached to or loosely associated with sediment and other submerged surfaces (Page and Siemensma, 1991). Slime molds may be regarded as protozoa because they spend most of their active lives as amebae or as naked ameboid (and often macroscopic) plasmodia (Figure 7.2). They have been known by many names, including mycetozoa (“fungus animals”). This results from the observation that they never develop hyphae, but they produce fruiting bodies (sporangia) supported on cellulose-rich stalks. There are two large groups: the cellular slime molds (dictyostelids), such as Dictyostelium, and the acellular slime molds (myxomycetes), such as Physarum. Two smaller groups are the acrasids (e.g., Acrasis) and the protostelids (e.g., Cavostelium). The dictyostelids are typically phagotrophic amebae; but when starved, they aggregate, form a migrating slug, and then produce a fruiting body from which spores are released. These later germinate to produce amebae. In the myxomycetes, individual amebae (sometimes thousands of them) with or without flagella coalesce to form a distinctive (often brightly colored) multinucleate plasmodium that gives rise to fruiting bodies. Slime molds are common in damp forest soils, tree bark, and dead or dying wood, and abundance and local species richness are greater in deciduous than in coniferous forests. There is extreme patchiness in the distribution of dictyostelid species in forest soils, and coexistence of multiple species in the same small soil samples is rare. Most myxomycete species are believed to

SECTION | III  Protozoa to Tardigrada

be cosmopolitan. The phagotrophic ameboid stages of slime molds may be quantitatively important grazers of bacteria, fungi, and the other primary decomposers of organic matter in soil. Other ecological interactions of slime molds may be at least as complex as their life cycles (e.g., the migrating slugs of dictyostelids appear capable of repelling grazing nematodes).

Flagellated Protozoa There is little consensus on how to classify the flagellates, and they are here divided into broad functional groups (Figures 7.1 and 7.3). Heterotrophic, nonphotosynthetic flagellates are fundamentally important because they are abundant (there are seldom fewer than 100/ml, sometimes 1000/ml, even in the plankton) and because their grazing activities are largely responsible for controlling the abundance of bacteria in aquatic environments. In some taxonomic groups, all species are exclusively heterotrophic (e.g., choanoflagellates and bodonids); others contain many mixotrophs (e.g., the euglenids and chrysomonads), whereas the haptomonads and cryptomonads are dominated by phototrophs and only a minority are capable of phagotrophy. In the past 25 years, a large diversity of heterotrophic flagellates has been discovered. Most of these are choanoflagellates, chrysomonads, euglenids, or bodonids. Some of the more easily recognized species (e.g., Rhynchomonas nasuta) have been recorded from a wide range of habitat types in freshwater, marine, and terrestrial environments (Patterson and Larsen, 1991; Lee and Patterson, 1998). Many heterotrophic flagellates are anaerobes, including intestinal parasites of humans (e.g., Giardia intestinalis), but free-living “diplomonads” (e.g., Hexamita and Trepomonas) and Retortamonas are relatively common bacteria feeders in organically enriched anoxic waters and sediments. Some anaerobic flagellates have hydrogenosomes, the anaerobic derivatives of mitochondria. Almost all of these are endosymbionts or internal parasites of animals. They are also known as parabasalians and include many of medical or economic importance, e.g., the human parasite Trichomonas vaginalis, and Trichomitopsis, which degrades cellulose in the hindgut of termites and other wood-eating insects and also supports endosymbiotic methane-producing bacteria (termites may be responsible for a globally significant flux of methane to the atmosphere) (Fenchel and Finlay, 1995). Most choanoflagellates are small (103 times their own body volume each hour! Ingestion rates (biomass consumed per animal per unit time) also are very high for rotifers. An adult rotifer may consume food resources equal to 10 times its own dry weight per day. If their assimilation efficiencies (i.e., assimilation divided by ingestion) are between 20% and 80%, rotifers can convert a good deal of their food to animal biomass that may be passed on to the next trophic level. Although microcrustaceans generally have higher clearance rates than rotifers (∼10–150 and 100–800 μl/animal/h for cladocerans and copepods, respectively), rotifers can exert greater grazing pressure on phytoplankton than some small cladocerans. In one study in a small eutrophic lake, K. cochlearis populations accounted for about 80% of the community grazing pressure on small algae during the year. This study also showed that K. cochlearis had clearance rates about 5–13 times higher per unit biomass than the cladoceran B. longirostris. Therefore, under certain conditions, rotifers may be important competitors to small, ­filter-feeding microcrustaceans and are important in nutrient recycling in aquatic systems. Furthermore, rotifers can alter the species composition of algae in certain systems. Studies have shown that intense feeding by Brachionus rubens Ehrenberg, 1838 can cause a shift in the dominant algal species from Scenedesmus to the spined algae, Micractinium. Apparently this shift is based on the inability of B. rubens to consume algae with protective spines. Although generally not ingested, cyanobacteria are increasingly recognized as playing an important role in determining zooplankton species composition. Large cladocerans are more sensitive to cyanobacteria (e.g., Anabaena spp.) than rotifers. The mechanism for this differential sensitivity to cyanobacteria toxicity is based on different tendencies to ingest filamentous cyanobacteria and different physiological tolerances to their toxins. Thus, having the ability to reproduce rapidly, rotifers may account for 50% or more of the zooplankton production, depending on the prevailing conditions. This production, in turn, can be an important food source for other rotifers, Asplanchna, cyclopoid and calanoid copepods, malacostracans (Mysis), zebra mussels, aquatic mites, insect larvae (Chaoborus) and adults (Buenoa), and small fishes. Abundance and species composition of rotifers often reflect the trophic status of lakes. For example, numerous

Chapter | 13  Phylum Rotifera

studies have reported changes in the maximal, totalpopulation density of several orders of magnitude when lakes were subject to intense eutrophication. Individual species sometimes undergo dramatic population changes during those periods. This was observed in the zooplankton of Lake Constance, where the density of Asplanchna increased its maximum population level 280-fold over a period of 28 year. However, in other lakes, dramatic population declines have been seen. In the years in which in Lake Washington had elevated concentrations of dissolved phosphorus, low water transparency, and high algal densities, K. cochlearis was abundant; however, as these water quality parameters improved, the population of this rotifer declined dramatically. Overall, there was at least a 20-fold increase and then a decline during a period of 15 years. Studies of the interactions of rotifers with other organisms will probably continue to receive attention in the future, especially predator–prey interactions, exploitative and interference competition among rotifers and other herbivorous zooplankton, life history strategies, and the toxic effects of cyanobacteria, dinoflagellates, and diatoms. Moreover, little detailed work has been done to examine the concept of functional complementarity in rotifer communities. This hypothesis argues that, within the constraints of niche requirements, a decline in the population levels of some species is offset by a rise in others, a concept termed compensatory dynamics (Fischer et al., 2001). The idea that the GR′ of communities can deviate widely over a season (see above) and that some rotifer communities appear to exhibit compensatory dynamics needs to be explored in greater detail.

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extinction after 2–3 weeks (Figure 13.36). Daphnia is unaffected by the presence of rotifers. Of course, protist–rotifer competition for bacterial food also can be important. Population growth of certain rotifers also is inhibited by Daphnia through interference competition. For example, in the presence of Daphnia, K. cochlearis suffers mechanical damage (i.e., is killed, wounded, or loses its eggs) when swept into the branchial chamber of the daphnids; the rate at which rotifers were killed is apparently proportional to daphnid body length. On occasion, Keratella may be found in the guts of Daphnia and Cypris (Figure 13.37), indicating a surprising pathway for trophic interactions (Gilbert, 2012). These microcrustaceans have the greatest impact on Keratella populations when larger than 2 mm; similar effects have been reported for other cladocerans (e.g., Scapholeberis kingi). Of course, interference and exploitative competition may occur simultaneously.

Predator–Prey Interactions Predation is another important regulatory factor in rotifer population dynamics, as rotifers are prey for several aquatic predators including protists, other rotifers, insects, cladocerans, copepods, and planktivorous fish. From many studies, we know that predation affects rotifer population dynamics both directly, by contributing to mortality, and indirectly, as a selective force shaping rotifer morphology, physiology, and behavior. One area that deserves additional study is the trophic interactions of microbes, protists, and

Competition with Other Zooplankton As noted before, rotifers, cladocerans, and copepods often compete for limited food resources and, in general, rotifers are relatively poor exploitative competitors because their clearance rates are usually many times lower than those of daphnids. In addition, rotifers also have a more limited size range of particles that they can ingest compared to cladocerans and are less resistant to starvation. Thus, cladocerans generally have broader food niches than rotifers in terms of food type and size, and through direct competition may suppress rotifer population growth. However, this outcome may be reversed when a sufficient quantity of suspended sediments is present in a lake. Exploitative competition between rotifers and daphnids is readily demonstrated when these zooplankton are grown in single and mixed cultures. Brachionus calyciflorus and Daphnia pulex both grow well on the alga Nannochloris oculata in single species cultures. When both species are present, however, Daphnia removes an increasingly larger proportion of algal cells until the rotifers gradually starve to

FIGURE 13.36  Competition between Brachionus calyciflorus and Daphnia pulex. Brachionus and Daphnia were grown in single species (closed symbols) and mixed-species (open symbols) batch cultures at 20 °C, daily renewed with 5 × 106 Nannochloris cells per milliliter. Population size (Y axis) is the number of individuals in the 80-mL culture. Error bars (±1 SE) are visible when they exceed the size of the symbols. Figure redrawn from Figure 1: Gilbert (1985), with kind permission of the author and the Ecological Society of America.

SECTION | III  Protozoa to Tardigrada

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rotifers within the microbial loop (Figure 13.38) (see also Parasitism on Rotifers below). Most rotifers are transparent and quite small; some are smaller than many ciliates (ca. 60–250 μm long). Although these features benefit planktonic rotifers by reducing their visibility to fish, a small body size renders rotifers more vulnerable to those invertebrate predators that are tactile feeders. Many rotifers produce a thickened integument (lorica) and/or spines and other projections, or carry their eggs, all of which have been shown to reduce the ability

FIGURE 13.37  Fecal pellet from Cypris pubera cultured with Keratella tropica. Visible in this pellet are at least five loricas of K. tropica (numbers). Original photomicrograph courtesy of J.J. Gilbert, Dartmouth College; see also Gilbert (2012).

of predatory zooplankton to prey upon them (Wallace and Smith, 2009). On the other hand, small size also appears to be a deterrent to predation. In laboratory cultures, Sarma and Nandini (2007) demonstrated that, despite its small body size (70 μm), Anuraeopsis fissa was not consumed at the same rate as two alternative brachionid prey by either A. brightwellii or A. sieboldii. In fact A. sieboldii ignored A. fissa, although it was consumed by A. brightwellii. Rotifers have other means of reducing predatory pressures. Spines are produced in some rotifers (e.g., B. calyciflorus, Figures 13.13 and 13.39) in response to a build-up of soluble substances released by invertebrate predators such as Asplanchna and several genera of copepods Epischura, Mesocyclops, and Tropocyclops. This phenomenon is another type of chemical signaling, except that the communication is between predators and prey rather than among conspecifics as in quorum sensing. Polymorphic spine production has been observed in B. calyciflorus, Brachionus urceolaris Müller, 1773, F. longiseta, K. cochlearis, Keratella slacki Bērziņš, 1963, and K. testudo. The importance of spined morphotypes is a significant reduction in capture and ingestion by invertebrate predators by making the rotifer more difficult to manipulate and swallow. The presence of spines on B. calyciflorus is a good example of the phenomenon (Figure 13.13) that works as follows. When disturbed by a potential predator, B. calyciflorus will retract its corona and, by doing so, increases the hydrostatic pressure within its pseudocoelom.

(a)

(c)

(b)

(d)

FIGURE 13.38  Examples of rotifers in the microbial loop. (a) Heliozoan with ingested Lecane. (b) Ciliate (Frontonia) with ingested Lecane. Photomicrograph courtesy of John Maccagno. (c) Asplanchna with two Keratella within its gut. (d) Several individuals of an unidentified microbe within the lorica of a dead Euchlanis. This culture crashed within three days; all of the dead rotifers were infested like this one. Key: r = rotifers; bar = 100 μm.

Chapter | 13  Phylum Rotifera

The elevated pressure causes the posteriolateral spines to swing forward and outward (Figure 13.39), making it more difficult for a predator to manipulate. For Asplanchna, this change is sufficient to prevent ingestion after capture. After a period of time, during which Asplanchna attempts to swallow B. calyciflorus (usually >60 s), the predator will release (reject) the prey, which then swims away unharmed. In this example, Asplanchna releases a biochemical cue (an allelochemic, called a kairomone) that initiates a developmental change in subsequent generations of B. calyciflorus (spine production), reducing the effect of predation. Some forms of K. cochlearis also possess posterior spines that make them much more likely to be rejected after capture by Asplanchna girodi than unspined forms. Unfortunately, the cost of spine production in rotifers, either in terms of developmental costs of producing the spines or increased energetic demands on swimming by increased mass, has not been fully resolved (Gilbert, 2013). Sometimes when Asplanchna or a related species (Asplanchnopus) swallows a spined form, the spines become lodged in the gut of the predator and may even puncture the delicate tissues. Presumably, both predator and prey die when that happens. Similarly transparent mucus sheaths produced by some planktonic, colonial rotifers (e.g., Conochilus, F ­igure 13.19(c), and Lacinularia) deter predation; the sheaths make the effective size of the prey too large for the invertebrate predator (e.g., Asplanchna and predatory copepods) without making them more visible to fish. The tubes and sheaths of sessile rotifers also work as refugia (Figures 13.8(a),(d) and 13.19(a)). Some rotifers escape predators by making rapid jumps using a variety of appendages: setous arm-like appendages (Hexarthra, Figure 13.14), paddles (Polyarthra, Figure 13.40), and long setae (Filinia, Figure 13.41). However, apparently Filinia also can use its long setae as foils to ward off predators and not just as lever-arms to initiate a jump.

FIGURE 13.39  Extension of the posterolateral spines in Brachionus calyciflorus. Here the spines are extended due to preservation in formalin. In life retraction of the corona causes an increase in the pressure within the pseudocoelom, which results in an outward flexing of the spines.

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Some rotifers assume a passive posture, displaying what has become known as the dead-man response, rather than fleeing when predators attack (Asplanchna, Brachionus, Keratella, Sinantherina, and Synchaeta). This simple behavior is

FIGURE 13.40  Polyarthra. This rotifer possesses long paddle-like appendages that are used in making rapid jumps to escape predators. Photomicrograph courtesy of Martin V. Sørensen, University of Copenhagen.

FIGURE 13.41  Filinia. This rotifer possesses long spines that it uses as foils to rapidly fend off predators. Photomicrograph courtesy of Martin V. Sørensen, University of Copenhagen.

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members of the Scenedesmaceae (Chlorophyta). For example, Scenedesmus obliquus exhibited a logistic dose– response to dilutions of test medium in which B. calyciflorus was incubated, thus indicating the presence of grazingreleased biochemicals, termed infochemicals (Verschoor et al., 2004). Algal defenses such as these can influence the long-term stability of both rotifer and algal populations and, in doing so, steady the population fluctuations often seen in bi- and tritrophic food chains. Rapid evolution also has been documented in long-term chemostats in which rotifers were fed algae from a stock composed of multiple clones. In the presence of rotifer grazing, there is strong selection for algal genotypes that are more digestion resistant; however, this defense comes at a cost of slower population growth (Yoshida et al., 2004).

Parasitism on Rotifers FIGURE 13.42  A few individuals of a colony of Sinantherina socialis. Here all the animals have retracted their corona, a behavior that exposes the warts on their anteroventral surface.

merely a retraction of the corona into the body and passive sinking. Contraction of the corona stops the animal from swimming, which eliminates the vibrations it produces that may be detected by the predator. In addition this behavior may make the rotifer more difficult to grasp in its turgid state. In Sinantherina spinosa (Thorpe, 1893), this passive posture exposes a group of small spines on its anteroventral body surface that may function in defense against planktivorous fish. To date, only one species of rotifer, S. socialis, has been shown to be unpalatable to small zooplanktivorous fishes (Felix et al., 1995) and certain invertebrates (Walsh et al., 2006). While neither the nature nor the location of the unpalatability factor(s) is known, this colonial species probably possesses a chemical that is held in gland-like structures, called warts, located at the anterior end of the animals (Figure 13.42). Defenses against predators such as spines, mucus sheaths, thickened loricas, and escape movements are energetically demanding. Some species, such as Synchaeta pectinata, are not well defended against predators, but have evolved very high maximal population growth rates that offset mortality from predation. Avoiding potential predators in space and/or in time is another simple yet effective defense mechanism against predators. Some rotifers occupy the habitat at a different time of year from that of an important predator, migrate vertically or horizontally in the habitat, or live in zones with low oxygen concentration, thereby evading predatory pressures altogether. Some algae initiate colony formation as an antipredator defense when rotifers feed on them. This occurs in

The importance of parasites in controlling population density in rotifers has not been examined thoroughly, although a few studies have correlated parasitic infection with a decrease in population density of planktonic species. In certain cases, parasites apparently caused the demise of an entire population in a lake within a few days. At least one virus causes high mortality in B. plicatilis aquaculture systems. The s­ porozoan parasite Microsporidium (Plistophora) frequently infects planktonic rotifers possessing thin loricas, such as members of the genera Asplanchna, Brachionus, C ­ onochilus, ­Epiphanes, Polyarthra, and Synchaeta. Water temperature seems to be an important mediating factor in the spread of the parasite, as infection rates drop off at water temperatures below 20 °C. In infected rotifers, the pseudocoelom of the animal becomes nearly filled with cysts (Figure 13.43). Several workers have described endoparasitic fungi that attack soil rotifers of the genera Adineta and P ­ hilodina. Some fungi form peg-like adhesive appendages on both small conidia (spores, ca. 30 μm long) and long vegetative hyphae. Once the adhesive pegs attach to a rotifer, they germinate and rapidly colonize the pseudocoelom (Figure 13.44). Other fungi produce spores that initiate parasitic attack when ingested. Another avenue of infection occurs via hypodermic injection of a vegetative cell into the host. Once inside the host, these fungal cells grow into ­assimilative hyphae, producing more infective cells either inside or outside the rotifer. However, bdelloids apparently free themselves from fungal parasites through anhydrobiosis (Wilson and Sherman 2010, 2013). Rotifers also can ingest the oocysts of parasitic protists important to human health such as Cryptosporidium and Giardia. However, it is not known whether they can significantly reduce the numbers of these parasites in natural conditions. Rotifers also act as vectors of disease agents such as white spot syndrome virus, which have been reported to attack shrimp in aquaculture systems.

Chapter | 13  Phylum Rotifera

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FIGURE 13.45  Phoretic association between Brachionus and Daphnia. Photomicrograph courtesy of Elizabeth J. Walsh, University of Texas at El Paso.

FIGURE 13.43  Photomicrograph of Brachionus sp. infected with a sporozoan parasite within its body cavity.

termed phoresis (G., to carry), to either the daphnid or to the rotifer have not been completely explored.

COLLECTING, CULTURING, AND PREPARATION FOR IDENTIFICATION Collecting

FIGURE 13.44  An example of a fungal parasite of rotifers. Four rotifers trapped by adhesive pegs on the vegetative hyphae of Cephaliophora. Bar = 100 μm. (Redrawn with permission of George Barron and the Canadian Journal of Botany 61(5):1345–1348.)

Rotifers as Parasites A few rotifers have been described as being parasitic on algae, sponges, other rotifers, freshwater oligochaetes, snail eggs, crustaceans, and fishes. However, we really know too little about these associations to classify all of them as parasitic. Members of the genera Brachionus, Limnias, Pleurotrocha, Proales, and Ptygura have been known to make temporary attachments to either invertebrates or vertebrates. In fact, it is not uncommon to find the carapace of Daphnia colonized by numerous individuals of B. rubens ­(Figure 13.45). The consequences of this phenomenon,

Collecting rotifers does not require complex or expensive equipment (Wallace et al., 2006; Wallace and Smith, 2009). One can almost always collect several species of planktonic rotifers by towing a fine-mesh (25–50 μm) net through any body of water. It is important to note that nets with larger mesh sizes (≥63 μm) tend to miss small-bodied forms (Chick et al., 2010). Productive lakes and ponds usually provide especially good sampling sites, but because fine-mesh nets can clog easily in these habitats, the water may need to be prefiltered through a net with a larger mesh size. More elaborate equipment such as closing nets and the Clarke-Bumpus sampler work well, but they are not necessary unless required by a specific sampling protocol. Water collected by discrete sampling devices (e.g., Van Dorn sampler, Kemmerer bottle, plankton trap, submersible pump) is then filtered. In weedy areas, a dip net or flexible collecting tube, called a water core, are very useful. Another simple method to collect rotifers is to submerge a 3- to 4-l (1-gallon) glass jar in a weedy region and to arrange loosely a few aquatic plants in it before retrieval. In the laboratory, place the jar near a subdued light source such as a north-facing window or a low-intensity lamp. Rotifers that swim to the surface on the lighted side may be removed using a transfer pipette. Certain aquatic plants such as Elodea, Myriophyllum, and filamentous algae are

SECTION | III  Protozoa to Tardigrada

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good substrata to examine for the presence of sessile rotifers, but Utricularia usually provides the richest diversity (Edmondson, 1959; Wallace, 1980). Plants with highly dissected leaves may be examined in small dishes using a dissecting microscope. Broad-leaved plants must be cut into strips and examined on edge. The upper few centimeters of moist sand taken just above the water line along a lake or a marine shoreline or the hyporheic interstitial zone of a streambed usually provides several species of rotifers. Unfortunately, very few studies have been conducted on rotifers from the psammon, in part because of the difficulty in separating the organisms from the sand. Recent studies have revealed a remarkable diversity (35–85% of the fauna) and abundance (up to 105 individuals/l) of rotifers in this habitat. Under certain conditions, rotifers may be found at depths of up to 60 cm into the interstitial. Sediments collected from the bottom using a dredge, coring apparatus, or suction device usually provide several species. The upper few centimeters of sediments from a core normally contain diapausing embryos that can be induced to hatch within a few days when several milliliters of sediment are incubated at ambient spring or summer temperatures. Dry sediment from desert rock pools also provide material when rehydrated in culture fluid and incubated for several days. Do not overlook laboratory aquaria as potential sources of material. We have found some unusual species in aquaria that had remained almost unattended for months. One might try adding a small amount of sediments from several sites as a way of adding variety to the rotifer community within the aquarium. Sessile rotifers may be present if the aquarium contains aquatic plants recently collected from the field. However, if you attempt to keep sessile forms, be sure to remove snails from the aquarium. Aquatic plants from hobby shops may bring new rotifer taxa to your aquarium, but the plants themselves may be alien species. Exercise care so that they are not released into the environment. Populations of rotifers may be maintained or even increased by adding a small amount of food once or twice a week (see Culturing), but keep the aquarium aerated. Aquarium filters that use fibrous materials to remove suspended materials may reduce the rotifer population. Once the collection has been made, it should be examined alive as soon as possible. It is generally a good idea to place live samples in jars over ice for the return trip to the laboratory, although we have found that a few species suffer when cooled (i.e., collections made from warm waters). Anesthetizing rotifers in the field has proved to be difficult (see below). Three preservatives are commonly used to preserve rotifers: formalin and Lugol’s iodine (I2KI) at concentrations of 5% or less, and ethanol at about 30–50%. (Higher concentrations of ethanol are used in preservation for DNA

analysis.) Lugol’s has two advantages over formalin. It is less toxic, and it stains the specimens slightly, which makes the animals more visible during sorting. Unfortunately, fixation deforms the specimens, making them difficult, if not impossible, to identify. On the other hand, in the genus Lecane, fixation with formalin is advisable so that one can study the morphology of the lorica. Dyes such as Rose Bengal are commonly used to stain preserved specimens. However, preservation sometimes causes rotifers to stick together and to other zooplankton. This preservation artifact has been misinterpreted as a behavior, as is seen in case of live B. rubens, which attaches to free-swimming Daphnia (Figure 12.45).

Culturing Laboratory Culture Many species of rotifers have been cultured for research, most notably in the genus Brachionus, both freshwater (B. calyciflorus and B. rubens) and saline (B. plicatilis). Because of their extensive use, these species have been referred to as the white mice of the rotifer world. The culture systems can be quite simple, using only small vessels such as depression slides, watch glasses, plastic tissue culture plates (Figure 13.46(a)–(b)), and small beakers or flasks (Figure 13.46(c)). However, larger systems are often used (Figure 13.46(d)), and, in aquaculture, very large systems are employed (Figure 13.46(e)–(f)). Wallace et al. (2006) provides a summary of procedures for culturing rotifers for research or aquaculture settings. Carlson (2000) provides directions for construction of a simple system for the hobbyist, which can be scaled up for research or large-scale culture (see Lawrence et al., 2012). Culture vessels that have had contact with formalin should not be used, as a residue of this toxin is thought to remain attached to both glass and plastic. Some species easily adapt to artificial conditions, attaining densities of >105 individuals per liter in a few weeks, whereas others seem impossible to keep even for short periods. Most cultures need regular maintenance a few times per week (e.g., changing the medium, feeding, cleaning the vessels). However, some species require little care. For example, some bdelloid species (e.g., Habrotrocha rosa, which is found in pitcher plant traps) are easily cultured in dilute suspensions of powdered baby food or crushed dried pet food. In this case, decomposition of the food source provides bacteria for the rotifers. In a similar manner, the techniques used to culture protists (e.g., making extracts and infusions of various grains, hay, manure, soil) have been adopted for the culture of some rotifer species with great success. Such cultures may be ignored for days and perhaps weeks at a time without loss of the culture. Nevertheless, most researchers grow the food needed to culture their

Chapter | 13  Phylum Rotifera

267

(a)

(b)

(c)

(d)

(e)

(f)

FIGURE 13.46  Different scales in culturing rotifers. Small-scale laboratory cultures: (a) Plastic tissue culture plates are common for small studies. Here, a 12-well plate (effective volume ≤5 ml) is shown, but 48-well plates (∼1 ml) are used to culture individual animals. (b) Plastic tissue culture flasks are used to achieve slightly larger populations (ca. 250 ml). Medium-scale laboratory cultures. (c) In this two-stage culture system, algae are grown in plastic bags on one shelf (numbers indicate relative age; oldest = 1), which is slowly supplied to the small columns (∼575 ml) on the lower shelf by gravity feed or a pump. Larger laboratory systems: (d) A single-stage culture system in which algae are grown in 250-l plastic columns and then a starter population of rotifers is added. Aquaculture scales: (e) High-density mass culture (>1 m3) achieve up to 4000 individuals/l (Nagasaki Prefectural Institute of Fisheries). (f) Mass culture pools (50 m3) achieve up to 400 individuals/l (Japan Sea Farming Association). Photographs (e) and (f) courtesy of Atsushi Hagiwara, Nagasaki University.

target species separately and feed it to the culture at regular intervals, usually 2–3 times per week. Although more costly, commercial products such as Roti-Rich® (Florida Aqua Farms, Inc.) and Sparkle® and related products (INVE Aquaculture®) give excellent results, especially where large numbers of rotifers are required. However, do not overlook the prospect of large populations suddenly arising in fish tanks. One of us (R.L.W.) followed the population dynamics of the sessile rotifer C. vorax (ca. 200–1100 μm) on the sides of a 115 L (50-gal.) aquarium for nearly 7 years; during that time, the density varied from 10 cm in height or diameter) and occur in slow-flowing streams and ponds. Pila leaves streams to forage for plants, using a “lung” for respiration. Both physids (Kesler et al., 1986) and planorbids (Calow, 1973b, 1974a) prefer detritus (Table 18.4). For example, although widely-spread species like P. gyrina and Planorbella (formerly Helisoma) trivolvis Say did not prefer detritus over periphyton in laboratory experiments, another physid, Aplexa hypnorum Linnaeus, much more common in wooded ponds with a rich detritus food base, did (Brown, 1982). Originally, snail algivores were considered indiscriminate grazers, taking all components of the periphyton (Hunter, 1980; Russell-Hunter, 1983). However, limpets and planorbids are selective (Calow, 1973a,b), and L. peregra grazes selectively on filamentous green algae (Lodge, 1985). Planorbis vortex L. ingests diatoms in greater quantities than found in the periphyton, but is still predomnantly a detritivore. Gastropods locate macrophytes through distant chemoreception (Croll, 1983). For example, L. peregra is positively attracted to Ceratophylum demersum L., because of dissolved organic materials excreted by the macrophyte (Brönmark, 1985). Similarly, Potamopyrgus jenkinsi (Gray 1843) orients toward both plant and animal extracts (Haynes and Taylor, 1984), while Biomphalaria glabrata Say either orients toward or away from specific macrophytes (Bousefield, 1979).

Chapter | 18  Introduction to Mollusca and the Class Gastropoda

Effects of Snail Feeding

401

of grazing may stimulate production, higher snail densities decrease both biomass and production (McCormick and Stevenson, 1989; Swamikannu and Hoagland, 1989). Grazing may not, however, have as much of an impact on shaded streams, where light may be the primary limiting factor (Hill and Harvey, 1990). Even in unshaded steams, there may be little overall effect of grazers, because the loss of the algal over story to grazing is compensated for by the competitive release and increased growth of adnate forms (Hill and Harvey, 1990). Snail grazers selectively remove larger filamentous green algae, and leave smaller, adnate species behind (Table 18.5). Under slight gastropod grazing pressure, periphyton assemblages are dominated by filamentous green algae, but more intensely grazed assemblages are dominated by more tightly adhering species or by toxic species such as cyanobacteria. Snail grazers may, in fact, indirectly facilitate macrophytes by eating the periphyton that limits macrophyte growth. For example, the growth of C. demersum increased when gastropod grazers were present (Brönmark, 1989a), although increased growth also occurred when plants were exposed only to snail-conditioned water, indicating increased nutrient recycling was the cause (Underwood, 1991). Similarly, when sunfish depress snail abundances and increase periphyton abundance in fish enclosures (Table 18.5), they may indirectly depress macrophytes (Martin et al., 1992). The molluscivorous tench can have similar cascading effects on European gastropods, periphyton, and macrophytes

Freshwater snails are grazing omnivores that consume primarily periphyton or macrophytes. Exceptions include members of the predominantly marine Buccinidae, Marginellidae, and Acochlidiida and freshwater Glacidorbidae, which are predators (Strong et al., 2008). The Viviparidae and Bithyniidae are in part ctenidial suspension feeders, and the Ampullariidae feed on bryozoans and planorbid eggs (Strong et al., 2008). Although diet studies exist for several species, there is little thorough coverage of taxa. Rollo and Hawryluk (1988) showed that L. elodes Say and P. gyrina modified their feeding rate when supplied reduced-quality diets, to maintain growth and reproduction. Dillon (2000) reviewed diet studies for freshwater snails. Almost all experimental manipulations have indicated snail grazers can decrease periphyton standing crops (Table 10.3; see review in Brönmark, 1989b). For example, Physa at high densities reduces algal biomass by 97%, and richness by 66% (Lowe and Hunter, 1988). In some cases snails also increase algal production, perhaps by decreasing total biomass and lowering algal competition for light or nutrients, removing senescent cells, or increasing rates of nutrient cycling. Pulmonate gastropods can also alter the quality (e.g., nitrogen to carbon ratios and chlorophyll-a levels) of periphyton (Hunter, 1980), as can caenogastropods in streams (Steinman et al., 1987). Although low levels

TABLE 18.5  The Effects of Experimental Manipulations of Gastropods on Periphyton Biomass, Production, and Assemblage Structure Decreased Algal Species Biomass

Increased Algal Production?

Favored Adnate Species?

Elimia or Juga (N = 11)

73%

17%

100%

Mulholland et al. (1983), Steinman et al. (1987), Lamberti et al. (1987), Marks and Lowe (1989), McCormick and Stevenson (1989), Hill and Harvey (1990), Mulholland et al. (1991), Tuchman and Stevenson (1991), Hill et al. (1991), Rosemond et al. (1993)

Theodoxus (N = 1)

Y

N

Y

Jacoby (1985)

Amnicola (N = 1)

Y

N.M.

Y

Kesler (1981)

Physella (N = 3)

100%

N.M.

100%

Doremus and Harman (1977), Lowe and Hunter (1988), Swamikannu and Hoagland (1989)

Promenetus (N = 1)

N

N

N

Doremus and Harman (1977)

Lymnaea, Physella, and Helisoma

Y

Y

Y

Hunter (1980)

Group

Studies

Caenogastropods

Pulmonates

Percentages refer to the number of studies noting an effect. For Single studies, Y = effect, N = no effect, N.M = not measured.

SECTION | IV  Phylum Mollusca

402

(Figure 18.16). Thus, interactions between gastropod predators, snails, periphyton, and macrophytes may be very complex in natural systems.

Dispersal A tropical family of gastropods, the Neritidae, includes several freshwater genera that occur in coastal streams (Figure 18.13; Pace, 1973; Ford and Kinzie, 1982; Schneider and Frost, 1986; Pyron and Covich, 2003; Blanco and Scatena, 2005). Growth rates and fecundity were studied by Shigemiya and Kato (2001) in Japan. The life histories of these snails includes reproduction in freshwater with eggs masses deposited on

stone surfaces. Following hatching, veliger larvae are washed downstream to estuarine reaches where pediveligers settle (Shigemiya and Kato, 2001). Small adults remain in the estuary or marine environment or migrate upstream to freshwater flowing environments (Shigemiya and Kato, 2001). Migration patterns vary among taxa and location with seasonal migrations and migrations that coincide with increased discharge events (Blanco and Scatena, 2005). Neritid snails have been studied in streams of Central America (Schneider and Frost, 1986), the Caribbean (Pyron and Covich, 2003; Blanco and Scatena, 2005, 2006; Ferney and Blanco, 2012), and Pacific islands of Hawaii (Way et al., 1993), Japan (Nishiwaki et al., 1991), and French Polynesia (Resh et al., 1990). Genetic studies of neritids demonstrated considerable marine dispersal of larvae among rivers on the same island, indicating constant recolonization of streams (Cook et al., 2009).

Population Regulation 6QDLOELRPDVVJ'0P



6QDLOV

     

3HULSK\WRQµJ$)'0FP

 

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FIGURE 18.16  Cascading effects of a molluscivorous fish (the tench) on gastropods, periphyton, and macrophytes, based on a field manipulation (from Brönmark and Vermaat, 1998). Note how snails are depressed, periphyton increases, and macrophytes are shaded and decrease.

Food Quality One of the first demonstrations of population regulation under field conditions was with L. elodes (Eisenberg, 1966, 1970). When the density of adult snails in pens in a small pond was increased, adult fecundity declined, as did juvenile survival. With addition of a high-quality resource, spinach, an increase in the number of eggs per mass occurred. Evidently, the availability of micronutrients in periphyton was the crucial variable (Eisenberg, 1970). Brown (1985) provided additional evidence by transferring juvenile L. elodes among a series of ponds differing in periphyton productivity. There was an exponential increase in growth with increasing pond productivity, and snails in the most productive pond laid nine times as many eggs as snails in the two less productive ponds. A number of field studies also provide indirect evidence for the importance of resource abundance. For example, populations in more eutrophic habitats have more generations a year, more rapid shell growth, and lay more eggs (Burky, 1971; Hunter, 1975; McMahon, 1975; Eversole, 1978). Highly eutrophic sites may however be detrimental to gastropods, as gastropod diversity declined over a 50-year period as Lake Oneida, New York, became highly productive (Harman and Forney, 1970). In lotic systems, manipulation of periphyton resources and the abundances of caenogastropod grazers have indicated both that food resources can control snail density and size distributions, and that the snails can in turn control their food resources (Hill et al., 1991; Rosemond et al., 1993).

Parasitism Although less studied than the role of periphyton, the parasitic larvae of trematode worms may also impact snail population dynamics and evolution (Holmes, 1983; Dillon, 2000). The adult worm produces eggs that are expelled in the feces of

Chapter | 18  Introduction to Mollusca and the Class Gastropoda

the final host and hatch into an infectious larval stage called a miracidium. Once inside the snail host, miracidia asexually produce several stages (redia and sporocysts), which eventually produce thousands of cercaria that infect the final host or another intermediate host. Infections either accelerate or decelerate snail growth (Brown, 1978; Anderson and May, 1979; Holmes, 1983, see discussion in Minchella et al., 1985). Immediately after infection or exposure, snail egg production rates increase dramatically (Minchella and Loverde, 1981), evidently to lessen the eventual costs to fitness. As parasites consume the ovitestis and hepatopancreas, however, both growth and egg production of “patent” snails (those infections in the final stage where cercaria are emerging from snails) drop below that of uninfected snails (Minchella et al., 1985). Infected snails are castrated, and frequently have increased mortality (Davies et al., 2002) and reduced growth (Hay et al., 2005). Snail invaders may also have dramatic impacts on final host populations. V. georgianus and Bythinia tentaculata have invaded upper midwestern lakes in the United States in recent years, and digenetic trematodes they harbor have subsequently caused massive die-offs in final waterfowl hosts (Hoeve and Scott, 1988). The role of trematodes in controlling snail populations is unclear, as prevalence (percent of population infected) varies considerably (Brown, 1978; Holmes, 1983). For example, prevalence in L. elodes in Indiana ponds varied from as low as 4% to as high as 49% (Brown et al., 1988). Prevalence was higher in less productive ponds, evidently because food limitation caused longer snail life-cycles that increased chances for snails to be located by miracidia. Life-table models predicted that the number of offspring produced per adult in the next generation declined by 14% to 21% in parasitized populations of L. elodes. In the case of Physa acuta, trematode infections cause increases in grazing rates, and the trematodes, therefore, indirectly affect periphyton biomass and composition (Bernot and Lamberti, 2008). Trematode parasites and their snail hosts are also extremely interesting from a coevolutionary viewpoint (Holmes, 1983; Minchella et al., 1985). Because invertebrates cannot easily acquire resistance to parasites, frequency-dependent selection may operate to ensure the fitness of any genotype less vulnerable to a particular

403

trematode (Holmes, 1983). Parasites may also cause a shift in investment of resources from costly reproduction to growth and maintenance, and thus even increase survivorship of infected snails (see also Baudoin, 1975; Minchella et al., 1985; Sandland and Minchella, 2004). One evolutionary model, called the “red queen” hypothesis, suggests that gastropods are constantly evolving new phenotypes merely to stay ahead of their trematode parasites. For example, the New Zealand mud snail Potamopyrgus has both sexual morphs (able to produce genetically more variable offspring through recombination) and parthenogenetic morphs. The frequency of sexual reproduction is positively correlated with trematode prevalence, as one would expect if sexual morphs have an advantage (Lively, 1987). Sexual morphs are also more common in shallow water, where waterfowl (the final hosts) occur (Lively and Jokela, 1995). Infected snails are also more likely to forage during the day than uninfected snails or gravid females, suggesting parasites may modify snail behavior so they are more likely to be consumed by waterfowl, to complete the life-cycle (Levri and Lively, 1996). Experimental studies have suggested trematode populations are locally adapted to better infect their own host populations than snails from different lakes (Lively, 1989). Gene flow has not swamped such local adaptation, even though rates of gene flow are greater among parasite populations than among their snail hosts (Dybdahl and Lively, 1996). However, sexual reproduction may not be favored in all snail-trematode systems. In Campeloma, trematode metacercaria actually feed on sperm, and parthenogenetic morphs thus have an advantage (Johnson, 1992). Caenogastropod taxa (e.g., Melanoides, Oncomelania, Semisulcospira, Tricula) tend to be intermediate hosts for trematode parasites that infect birds and mammals (Dudgeon, 1999). The systematics and ecology of these taxa are well-studied largely because of directed research toward parasitology.

Production Ecology Average standing crop biomass, productivity, and turnover times differ between caenogastropods and pulmonates (Table 18.6). Standing crops are greater on the average for

TABLE 18.6  Comparison of Average Standing Stocks, Productivity, and Turnover times for Populations of Pulmonate and Prosobranch Snails Mean Biomass ± SE (gCm−2, N)1

Mean Production ± SE (mgCm−2day−1)

Mean Turnover time (days) ± SE (N)1

Pulmonata

0.98 ± 0.50 (6)

5.71 ± 2.44 (10)

98.0 ± 9.46 (10)

Caenogastropoda

4.64 ± 1.80 (4)

3.56 ± 0.51 (5)

Taxon

1N = Number

of species averaged. Reported in Russell-Hunter and Buckley (1983).

385.3 ± 33.5 (4)

404

caenogastropods than for pulmonates, although caenogastropod populations studied to date have not included genera with smaller individuals such as Amnicola and Valvata. Even if pulmonates have lower standing stocks, their rapid growth and short life-cycles still result in higher average production rates and shorter turnover times (Table 18.6). For example, two pleurocerid grazers in an Alabama stream, Elimia cahawbensis (Lea 1861) and Elimia clara (Anthony 1845), have considerable biomasses of 2–5 g ash-free dry mass (=AFDM) per m2, but slow growth rates and long life-cycles result in relatively low rates of secondary production (0.5–1.5 g AFDM m2). The combination of high biomass and low production results in a low production to biomass ratio of 0.3 (Richardson et al., 1988). Caenogastropod detritivores may, however, have higher production rates. Viviparus subpurpureus (Say 1829) and C. decisum, with high densities and short life-cycles in Louisiana bayous, have high standing crop biomasses (10–20 g AFDM/m2) and production rates (20–40 g AFDM/ m2/yr), among the highest known for freshwater molluscs (Richardson and Brown, 1989).

Ecological Determinants of Distribution and Assemblage Structure Snails in freshwater ecosystems are nonrandom species assemblages (Pyron et al., 2009; Hoverman et al., 2011) that are strongly influenced by predators and competitors (Turner et al., 2007). Lodge et al. (1987) concluded that at large, biogeographic scales, the variables that controlled assemblage structure were colonization ability and water chemistry; while disturbance regime, competition, and predation were stronger explanatory variables at local scales. Although water chemistry variables contribute to freshwater snail distribution and abundance patterns, a minimal level of water hardness (calcium concentration) is required to support snail physiology. Briers (2003) showed that calcium requirement of British freshwater gastropods was a strong predictor of range size. Species that were classified as calciphiles that require high levels of environmental calcium had smaller range sizes than species able to use a wider range of environmental calcium. Dussart (1979) studied snail assemblages (B. tentaculata, Gyraulus albus Müller, Planorbis planorbis Linnaeus, and L. peregra) in northwest England and found that substrate composition was more important than water chemistry in determining abundance patterns. Local environmental variables (substrate composition, water conductivity) were predictors of local assemblage composition for assemblages in Indiana in the United States (Pyron et al., 2009). Smaller drainage water bodies had higher abundances of pulmonates than caenogastropods (Brown et al., 1998; Pyron et al., 2009). Brown et al. (1998) predicted that abundance differences are due to physiological adaptations of pulmonates. Caenogastropods occur in

SECTION | IV  Phylum Mollusca

larger rivers because of their ability to withstand competition, and inability to withstand harsh physicochemical variation in smaller streams.

Watershed Connections and Chemical Composition Although relatively few studies have examined the structure of freshwater snail communities (Hoverman et al., 2011), some have predicted species richness using environmental and biotic variables (Lodge et al., 1987). The number of lake connections (inlets and outlets that act as dispersal corridors) was correlated with species richness in northern highland lakes of Wisconsin and Michigan in the United States (Lewis and Magnuson, 2000). Species richness of pulmonates in individual watersheds increased with watershed size (Dillon and Benfield, 1982). Species turnover among lakes was important in the structure of snail assemblages of lakes in a Michigan preserve (Hoverman et al., 2011). Water hardness and pH are often considered major factors determining the distributions of freshwater snails (Russell Hunter, 1978; Okland, 1983; Pip, 1986). However, in lake districts with adequate calcium (above about 5 mg/l CaCO3), or in the normal (nonacidified) range of pH, relationships between physicochemical parameters and gastropod diversity are less clear (Lodge et al., 1987). For example, species in New York lakes overlap broadly in the ranges of physicochemical variable they occur in (Harman and Berg, 1971). Physicochemical parameters therefore set the limits for gastropod distributions, but are not as important in explaining the relative abundance patterns and densities of gastropods in most hardwater, circum-neutral lakes.

Biogeographic Factors On a biogeographic scale, a factor determining gastropod distributions is dispersal ability. For example, relatively isolated lakes in upper watersheds in Wisconsin have fewer species than lowland lakes that are well connected by river corridors to other lakes (Lewis and Magnusson, 2000). Studies have also indicated diversity increases with the area of lakes and ponds (Lassen, 1975; Browne, 1981; Brönmark, 1989a; Dillon, 2000). Since immigration rates generally increase and extinction rates decrease with habitat size, larger habitats, all else being equal, usually support more species (MacArthur and Wilson, 1967). After dispersal, successful colonization depends on the presence of suitable substrates. For example, a significant relationship exists between the number of gastropod species and the number of substrates in lakes and streams in New York in the United States (Harman, 1972). In fact, substrate preferences determined in the laboratory are good predictors of the types of ponds in which snails are common, such as algivores in open ponds and detritivores in wooded ponds (Brown, 1982). Gastropod diversity is also

Chapter | 18  Introduction to Mollusca and the Class Gastropoda

positively related to macrophyte biomass, probably because macrophytes increase surface area for periphyton colonization (Brown and Lodge, 1993). Regional patterns for species richness at broad continental scales are shown in Figure 18.12. In tropical freshwaters, caenogastropods have higher species richness than pulmonates (Dudgeon, 1999). For example, 89% of freshwater gastropods in Java are caenogastropods and 11% are pulmonates (van Benthem Jutting, 1956). Canadian gastropods have the opposite pattern, with 16% caenogastropods and 84% pulmonates (Dudgeon, 1999).

Flow and Hydroperiod Disturbance may also determine the assemblage of snails. In temporary ponds, diversity is lowered by frequent drying; and in habitats that go hypoxic, diebacks of macrophyte and gastropod populations will also occur (Lodge and Kelly, 1985; Lodge et al., 1987). Disturbance may also limit some species from disturbance-prone areas (wave-swept shores, littoral zones of reservoirs, etc.). In streams, current velocity may affect distribution and growth. For instance, adult Elimia avoid high-flow areas, perhaps because increased flow makes movement more difficult and lowers grazing rates or increases metabolic rates—both factors lowering adult size (Johnson and Brown, 1997). Juveniles are more common in fast-flowing areas because they are smaller and can exploit the boundary layer.

Predation Predation is predicted to have a strong effect on local assemblages, and competition is predicted to be a strong influence in temporary ponds (Brown, 1982). The strength of competition in local assemblages appears to be slight compared to effects of disturbance regime and predation (Lodge et al., 1987). Brown et al. (1998) predicted that pulmonate taxa have increased competitive ability and caenogastropod taxa have decreased competitive ability at sites where food is a limiting factor. Snail assemblage composition is not random as well across gradients of aquatic habitat permanence and predation risk. In northwestern Pennsylvania, vernal ponds have short hydroperiods but few predators; these systems are dominated by P. vernalis. Permanent ponds have the highest biomass of invertebrate predators, and are dominated by Physa ancillaria (Say 1825). Lakes have the highest biomass of fish predators, and are also dominated by P. ancillaria. Physa acuta is the only species that is a habitat generalist, evidently because of its high degree of phenotypic plasticity (Turner and Montgomery, 2009).

Competition In large lakes, biotic interactions such as interspecific competition or predation are important in determining gastropod

405

diversity and abundance (Lodge et al., 1987). In regards to competition, some evidence suggests it is rare in freshwater snails. For example, pulmonate pond snails in the midwestern United States overlap little on food and habitat dimensions of the niche, although some species do compete (Brown, 1982). Other studies also indicate differences in resource utilization. For example, the ancylid, A. fluviatilis prefers diatoms and is found on the top of cobble where periphyton is abundant, whereas the co-occurring planorbid, Planorbis contortus L. is a detritivore, and occurs more often under stones where detritus accumulates (Calow, 1973a,b; 1974a,b). Similarly, P. columella (Say 1817) and P. vernalis possess differences in their gut and radulae allowing coexistence in New England ponds (Kesler et al., 1986). Furthermore, niche partitioning may also be facilitated by differences among snails in enzymatic activity (Calow and Calow, 1975; Brendelberger, 1997) or radular morphology. There is indirect evidence for competition. For instance, there are often fewer coexisting congeners in field samples than would be expected, based on mathematical simulations (Dillon, 1981, 1987). Second, competition has been inferred from changes in relative abundance of gastropod species through time, that is, by apparent competitive exclusion (Harman, 1968). Third, niche overlap can in fact be higher in more diverse snail assemblages found in large lakes. For example, the abundances of gastropod species in lakes are often positively associated in macrophyte beds, and experiments show most species have similar preferences for macrophytes (Brown, 1997). Finally, pulmonates are common in ponds or in vegetated areas of lakes, while caenogastropods are rare in ponds and common in lakes and rivers. One explanation could be competitive exclusion of pulmonates from lakes by caenogastropods, or exclusion of caenogastropods from ponds by the same mechanism. However, additional explanations include greater vulnerability of caenogastropods to hypoxic conditions in ponds, poorer dispersal abilities of caenogastropods, or the fact that thinshelled pulmonates are more vulnerable to shell crushing fish common in lakes or rivers (Brown et al., 1998).

Snail Response to Predators Adaptations to predation by aquatic organisms include morphological and chemical defenses, modified behavior, and life-history modifications (Kerfoot and Sih, 1987). The study of predation using freshwater snails as a model has been instructive due to multiple predators that specialize on freshwater snails and influence populations, and the large variation in body size and degree of shell development (Rundle and Brönmark, 2001). Predators include fishes, decapod crustaceans, and leeches (Brönmark and Malmqvist, 1986; Brown and Strouse, 1988; Covich et al., 1994). Predators that crush shells have preferences for thin-shelled snails (Stein et al., 1984; Slootweg, 1987; Osenberg and Mittelbach, 1989; Alexander and Covich, 1991a; Nyström et al., 1999).

406

Behavioral responses to predators include reduced activity, remaining under cover, and crawling out of the water (­Alexander and Covich, 1991b; Turner, 1997). Snails with pseudo-lungs (the pulmonates), such as Physa, can partially avoid predation by leaving the water (Crowl and Covich, 1990; Mower and Turner, 2004). Gastropods that possess escape behaviors to avoid predators may experience reduced selection for morphological variation, compared to taxa that cannot escape by leaving the water (Ross et al., 2014). Exceptions to this pattern are species like Campeloma, a caenogastropod with reduced availability of shape variation. Species like Campeloma likely avoid predation by large body size, thick shell, presence of an operculum to cover the shell opening when the foot is withdrawn, and burrowing in sediments. Gastropods with taller shell spires are more susceptible to predation than gastropods with disc-like shells (Cotton et al., 2004). Thin-shelled species that are more vulnerable to predation thus use behavioral avoidance mechanisms, and thickershell species (e.g., Campeloma) tend to move under cover or have a mixed response (Rundle and Brönmark, 2001;­ Brönmark et al., 2011). Physid gastropods have been excellent models for investigating predator–prey interactions. P. gyrina respond in different ways to crayfish and fish predators, using covered habitats in the presence of fish, or crawling above the water line when crayfish predators are active (Turner et al., 1999). The increased use of covered habitats, or habitats near the water surface, can also decrease periphyton biomass in those habitats (Turner et al., 2000). Smaller physids also tend to use behavioral avoidance like crawling out more than larger conspecifics that have thicker shells (DeWitt et al., 1999). Exposure to chemical cues from predators can also alter physid shell morphology. Elongate shells with narrow apertures decrease risk of crayfish predators, whereas a more rotund shell spreads crushing forces over a greater area and reduces risk from fish predators (DeWitt, 1998). These anti-predator responses in physids also have a spatial and temporal component, decaying over time or space. Snails do not respond to fish predators at greater than 1 m distance, nor to chemical cues older than about 40 h (Turner and Montgomery, 2003). Physids are also much more likely to respond to predation on conspecifics or congenerics than more distantly related prey (Turner, 2008). Nor are these responses confined to physids alone. Hydrobiids of the genus Amnicola use periphyton-rich sand substrates in the spring, but switch to less-periphyton rich macrophytes in the summer, when crayfish predators are more abundant, resulting in a depression of snail growth rates (Lewis, 2001). The lymaneid Radix also responds morphologically to predator cues, with a rotund shell again more resistant to fish predation (Lakowitz et al., 2008). Small lymnaeids or thinshelled snails are also more vulnerable to crayfish predation (Brown, 1998; Nystrom and Perez, 1998). Physa acuta in

SECTION | IV  Phylum Mollusca

particular has been used as a model for predator effects (Crowl and Covich, 1990; DeWitt et al., 1999; Turner et al., 1999). Physa and many other gastropods detect predators by chemicals in the water (Covich et al., 1994). In the presence of crayfish—a size-selective shell-entry predator—Physa had fast growth and narrow apertures (Auld and Relyea, 2008). Auld and Relyea (2008) found increased shell thickness and delayed reproduction as a response to predator cues. However, increased shell thickness only occurred in Physa that had available mates. Auld and Relyea (2011) demonstrated that increases in shell thickness, mass, and shell dimension of snails induced by predator cues resulted in lower predation rates. Hoverman et al. (2005) also found delayed reproduction and fecundity in Planorbella trivolvis that were exposed to crayfish predators. Indeed, Lodge et al. (1987) argued that predators determine the composition of gastropod assemblages in lakes. Indirect evidence includes the fact that gastropods have a number of anti-predator adaptations. These include thick shells to protect against shell-crushing predators (Vermeij and Covich, 1978; Stein et al., 1984; Brown and DeVries, 1985; Brown, 1998; DeWitt, 1998; Krist, 2002), as well as escape behaviors such as shaking the shell, crawling above the water to protect against shell-invading invertebrate predators (Townsend and McCarthy, 1980; Brönmark and Malmqvist, 1986; Brown and Strouse, 1988; Alexander and Covich, 1991a,b; Covich et al., 1994), or seeking underwater refugia (Turner, 1997). Molluscivores are common in lakes and rivers. For example, the pumpkinseed sunfish, Lepomis gibbosus L., and the redear or shell-cracker sunfish, L. microlophus (Guenther 1859), specialize on gastropod prey, and have pharyngeal teeth adapted to crush shells. Crayfish will select snails instead of grazing on macrophytes if given a choice (Covich, 1977), using their mandibles to chip shells back from the aperture, and selecting species with thinner shells (Brown, 1998). Although Osenberg (1989) argued that lentic snail assemblages are limited by food resources, most experimental evidence supports a strong role for predators in determining snail diversity and abundance (Table 18.7). For example, the central mud minnow can significantly lower the density of relatively thin-shelled snails in permanent ponds. When fish densities were manipulated in pens, the number of eggs and juveniles of L. elodes was significantly less in the presence of fish. The small, gape-limited fish feed on eggs and juveniles and may restrict L. elodes from permanent habitats such as large marshes or lakes. Pumpkinseed sunfish also strongly prefer large, weak-shelled gastropod species in laboratory experiments, and thinshelled species also decline dramatically in pumpkinseed enclosures in lakes (see references in Table 18.7). Thus, most thin-shelled pulmonates should occur in lakes only in macrophyte beds, where they have a refuge from visual predators. Moreover, sandy areas should be dominated by

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TABLE 18.7  Summary of Experimental Field Manipulations Testing the Role of Snail Predators in Controlling Their Prey Study

Prey

Predator

Conclusions

Brown and Devries (1985)

Lymnaea elodes

Mud minnows

Decreased abundance of eggs and juveniles of thin shelled L. elodes.

Sheldon (1987)

Assemblage

Sunfish

Removal of fish increased snails which decreased macrophytes through herbivory.

Kesler and Munns (1988)

Assemblage

Belastomatid bugs

Decreased snail abundances in New England pond.

Osenberg (1989)

Assemblage

Littoral sunfish

Fish controlled only rare, large snail species, competition considered more important.

Hershey (1990), Merrick et al. (1991)

Assemblage

Lake trout

Trout removed better competitor (L.elodes), favored poorer competitor, Valvata.

Martin et al. (1992)

Assemblage

Smallmouth bass Largemouth bass

Fish decreased snails 10-fold, increased periphyton biomass 2-fold, decreased algal cell size, decreased macrophytes.

Brönmark et al. (1992)

Assemblage

Pumpkinseed sunfish

Fish decreased snails, increased periphyton, favored adnate algae.

Bronmark (1994)

Assemblage

Tench

Fish decreased snails, increased periphton, decreased macrophytes.

Lodge et al. (1994)

Assemblage

Orconectes rusticus

Crayfish decreased snails and macrophytes, had no effect on periphyton.

Daldorph and Thomas (1995)

Assemblage

Sticklebacks

Fish decreased thin-shelled snail species, increased periphyton.

Brönmark and Weisner (1996)

Assemblage

Piscivores and molluscivores

Molluscivores did decrease snails, but piscivores did not control increase in snails indirectly.

“Assemblage” refers to a natural community of gastropods.

thicker-shelled pulmonates like Planorbella, or by caenogastropods. Indeed, gastropod distributions among habitats within Indiana and Wisconsin lakes follow these patterns (Lodge et al., 1987; Brown and Lodge, 1993; Lodge et al., 1998). In fact, fish predators may indirectly benefit poorer gastropod competitors by preferentially removing dominant species. Lake trout, for example, preferentially consume larger lymnaeids, releasing valvatids from competition in Arctic lakes (Hershey, 1990; Merrick et al., 1991). However, experimental manipulations suggest such indirect effects do not extend through four trophic levels. Piscivores in Swedish lakes cannot depress predators to the extent that interactions cascade down the food web to facilitate snails and negatively impact periphyton (Brönmark and Weisner, 1996). Invertebrate, shell-invading predators may also limit snail populations, or cause shifts in gastropod relative abundances. Although some leeches, such as Nephelopsis obscura Verrill, have fairly low feeding rates (Brown and Strouse, 1988), crayfish can eat over a 100 snails per night. The crayfish Orconectes rusticus (Girard 1852) significantly reduced snail abundances in enclosure experiments in Wisconsin lakes (Table 18.7), and an inter-lake survey

indicated that snail abundances were negatively correlated with crayfish catches. Crayfish also affect gastropod habitat selection. For instance, even though cobble habitat is preferred by many snails (see earlier discussion), it also provides refugia for crayfish from fish predators. Crayfish predation overrides the effects of rich food resources (Weber and Lodge, 1990). Crayfish predators also shift size distributions of P. virgata upwards in Oklahoma streams, due possibly both to size selective predation and to the snail’s diversion of energy from reproduction to growth to reach a size-based refuge from predation (Crowl, 1990; Crowl and Covich, 1990). Interestingly, only snails in vulnerable size classes respond by crawling out of the water after predation by crayfish occurred (Alexander and Covich, 1991b). Other invertebrate predators may also be important. For example, belostomatid bugs can eat as many as five snails per bug per day in the laboratory (Kesler and Munns, 1989). Fish predators may alter snail foraging behavior as well. In experiments, Physa selects refugia in the presence of sunfish predators, only leaving them when sufficiently starved (Turner, 1997). This behavioral interaction may provide an additional explanation for why periphyton biomass increases when fish predators are present. For example, use

408

of open habitats by snails decreases with increasing predation risk to sunfish with the result that periphyton biomass also increases (Turner, 1997). Interestingly, such indirect facilitation of periphyton does not occur upon experimental addition of crayfish, probably because crayfish are also omnivorous and graze on periphyton (Lodge et al., 1994). In fact, behavioral avoidance of predators may be a more successful strategy than thicker shells. Thus, P. acuta, although it is smaller and has a thinner shell than L. elodes, suffers less from fish predation because it resorts to habitat refugia in the presence of molluscivorous fish while the lymnaeid does not (Mower and Turner, 2004). Anti-predator adaptations can include multiple predator responses in a species. Trait compensation and co-specialization were demonstrated in Physa spp. where more vulnerable, smaller snails used stronger anti-predator behavior than large snails (DeWitt et al., 1999). Larger snails with reduced aperture size (a response to predation) exhibited less anti-predator behavior. The cues that snails release as a response to predation can affect species interactions in local communities. Turner et al. (2000), for example, found that nonlethal effects of predators on snail behavior resulted in reductions in snail grazing on periphyton. Parasites can also have indirect effects on communities. Bernot and Lamberti (2008) tested this hypothesis in an experiment by infecting P. acuta with a trematode and quantifying that snail grazing on algae increased compared with uninfected snails. The algal assemblage changed from diatoms and cyanobacteria to one dominated by Cladophora. These changes in the algal assemblage were predicted to have effects on ecosystem function.

Flexibility in Shell Architecture Evolutionary responses of shell architecture thus occur due to selective pressures of predation (DeWitt et al., 2000; Covich, 2010) and local and regional environmental variables. DeWitt et al. (2000) identified morphological responses in Physa that differ with local fish and crayfish predation. Increased shell thickness in pleurocerid taxa that are susceptible to predation by large-bodied fishes (freshwater drum, suckers) is presumably an additional response to ecological selective pressures. Whether similar morphological responses to predators occur in other taxa is unknown, but likely. Fish, crayfish, and other invertebrates are potential predators of freshwater gastropods and likely influence shell evolution (Vermeij and Covich, 1978; DeWitt et al., 2000; Turner and Montgomery, 2009). Local environmental variables, including flow (Holomuzki and Biggs, 2006; Minton et al., 2008), resource availability (Hoverman et al., 2005), and substrate composition (Dunithan et al., 2012), also influence morphological variation in freshwater gastropods. Morphological variation in a mollusc resulting from flow variation was first described by Ortmann (1920) for

SECTION | IV  Phylum Mollusca

unionid mussels, called Ortmann’s law (Goodrich, 1937; Minton et al., 2010; Dillon, 2011; Dillon et al., 2013). Similar morphological patterns were identified at a smaller scale of hundreds of meters for the pleurocerid Elimia potosiensis Pilsbryi (Minton et al., 2010). Brönmark et al. (2011) demonstrated that shell shape in Radix balthica Linnaeus was also flexible. Snails from populations without fish had narrow shells with well-developed spires. Snails from populations exposed to fish had more rotund shells with a low spire, characteristics that increase survival rate from predators that crush shells. Brönmark et al. (2011) used common garden experiments where snails were exposed to fish cues, resulting in a rounder body shape compared to control snails. The round body shape characteristic was thus phenotypically plastic and responded to predator cues, reinforcing the evolutionary importance of phenotypic plasticity. Parasite infection can also influence body size or shell morphology. Although many additional studies exist (Żbikowska and Żbikowska, 2005), we present only a few examples here. Krist (2000) experimentally induced shell size variation with infections from the digenean Proterometra macrostoma Horsfall in Elimia livescens Menke. The shell morphology of the mud snail Zeacumantus subcarinatus (Sowerby 1855) was altered by infection of trematode parasites (Hay et al., 2005). These and others suggest that morphological modifications of shell shape with parasite infection are a selective host response rather than a parasite modification (Levri et al., 2005; Lagrue et al., 2007). This phenotypic plasticity in shell morphology in response to biotic and abiotic environmental factors has also become highly controversial because it calls into question the identification of some taxa, including federally listed species (http://fwgna.blogspot.com, accessed 2/20/2013). This contentious issue will likely demand resolution soon.

Conservation Ecology The need for conservation of freshwater snails worldwide is as urgent as for North American taxa (Lydeard et al., 2004; Brown et al., 2008). Similar to declines seen in unionid mussels, freshwater snail diversity has also declined in the United States in the last 100 years (Neves et al., 1998; Stewart, 2006). The quite diverse gastropod assemblages in rivers in the southeastern United States have been seriously affected by habitat alteration. Pleurocerids are quite common in shallow riffles, where water is warm and welloxygenated, and rough substrates provide refugia from flow and sedimentation. Impoundments often change these habitats to hypoxic, cold water habitats with little periphyton food. Some gastropod species are threatened because they also limited to geographically isolated springs, whose existence is increasingly threatened by rising demands for groundwater.

Chapter | 18  Introduction to Mollusca and the Class Gastropoda

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FIGURE 18.17  (a) Relative diversity of families of freshwater snails. (b) Fraction of snail species at risk of extinction that belong to each family. Remaining refers to remaining families (Valvatidae, Potamiopsidae, Neritinidae, Ampullaridae, and Acroloxidae) that are not as diverse. At-risk is defined as having a global ranking of G2 or less, indicating only a few populations remain. Note that the two most diverse families, Pleuroceridae and Hydrobiidae, also have the most at-risk species, because their populations are often fragmented between different rivers or springs.

North American pleurocerid and hydrobiid gastropods are highly endangered (Lydeard and Mayden, 1995; Neves et al., 1998; Brown et al., 2008; Perez and Minton, 2008). In the United States, ∼74% of the species of Hydrobiidae and ∼45% of the Pleuroceridae (the two largest snail families in North America; Figure 18.17) are considered at risk, with a Nature Conservancy rank of ≤G2, indicating these species have few remaining populations and therefore face imminent risk of extinction (Figure 18.17). The pleurocerid Elimia has experienced the greatest number of extinctions, all in river systems in Alabama and Georgia (Table 18.2), followed in order by the pleurocerid genera Leptoxis, Gyrotoma, and Athearnia. A smaller number of hydrobiids have become extinct, but the fact that many species in the western and southeastern United States are found at only a few isolated springs puts them at considerable risk. At the current time, 21 species of gastropods are listed as endangered or threatened (Table 18.8). Nine of the species are hydrobiids from western states or Alabama. Five of the species are pleurocerids found in the Mobile River watershed in the Southeast. Three species are viviparids from Alabama watersheds. However, the only listed physid has been recently recognized to be a junior synonym of P. acuta (Rogers and Wethington, 2007), and two more of the species are currently under study to be de-listed, based on the discovery of additional populations (see footnotes in Table 18.8). Because the families Pleuroceridae and Hydrobiidae have the most at-risk species in North America, we will briefly discuss their ecology.

Ecology of Pleuroceridae Pleurocerids are perennial, with life-cycles ranging from 2–10 years (Richardson et al., 1988; Huryn et al., 1994; Dillon, 2000; Brown and Johnson, 2004). Life-history

variation in the pleurocerid Elimia is affected by both local and landscape-level factors (Huryn et al., 1995). Food availability limits individual growth rates in Elimia although considerable urban development and nutrient enrichment might be occurring now in streams in the southeastern United States. High current velocity also alters pleurocerid abundances, feeding, and growth. For example, flow refugia are important during spates (Stewart and Garcia, 2002), and Elimia orients upstream to minimize drag on the shell (Huryn and Denny, 1997). Johnson and Brown (1997) found that adult Elimia density and size decreased in higher flow. Considerable variation in shell anatomy occurs in pleurocerids, and the role of shell shape in the ability to withstand high flows is in need of further study. Pleurocerids are major players in stream ecology within the southeastern United States. Periphyton abundance is regulated by nutrients (bottom-up) and snail grazing (topdown) in these streams (Rosemond et al., 1993; see earlier discussion). Snails reduced periphyton production by half under ambient nutrient conditions and by two-thirds when nutrients were added. The role of competitive interactions within the fairly diverse pleurocerid assemblages in southeastern rivers has received less study, although Brown et al. (1998) studied competition between the pulmonate P. trivolvis and the pleurocerid Lithasia obovata Say and found the pleurocerid was a better competitor. Similarly, two co-occurring species of Elimia in Alabama streams were both affected by competition, but neither was competitively dominant, allowing co-existence (Cross and Benke, 2002). Lotic gastropods also might be important competitors with insects (Hawkins and Furnish, 1987; Harvey and Hill, 1991). If competition is important, the distribution of pleurocerids in rivers could be due to niche partitioning. For example, pulmonates are common in headwaters or littoral

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410

TABLE 18.8  Federally Listed Species of Gastropods in the United States as of Early 2008, Along with the States in which they are Found Family

Species

States

Assiminidae

Assiminea pecos (E)

Texas, New Mexico

Hydrobiidae

Jutumia kosteri (E), Pyrgulopsis bruneauensis (E), Pyrgulopsis neomexicana (E), Pyrgulopsis New Mexico, Idaho, Tennessee, Alabama ogmoraphe (E), Pyrgulopsis pachyta (E), Pyrgulopsis roswellensis (E), Taylorconcha serpenticola (T), Tyronia alamosae (E)

Lymnaeidae

Lanx sp. (E)

Idaho

Pleuroceridae

Athernia anthoni (E), Elimia crenatella (T), Leptoxis taeniata (T), Leptoxis plicata (E), Leptoxis ampla (T)

Tennessee, Alabama, Georgia

Physidae

Physa natricina (E)

Idaho

Valvatidae

Valvata utahensis (E)

Idaho

Viviparidae

Campeloma decampi (E), Lioplax cyclostomatiformis (E), Tulotoma magnifica (E)

Alabama

“E” refers to species listed as endangered and “T” to species listed as threatened.

zones of the Salt River in Kentucky, but are replaced by pleurocerids in higher-order reaches (Brown et al., 1998). Similarly, in the Ohio River, L. obovata is common in shallow littoral areas because its relatively large foot adapts it to wave action, while the thick shell of Pleurocera canaliculum Say protects it from most fish, and this snail occurs at all depths (Greenwood and Thorp, 2001). The heavily armored shells of most pleurocerids reduce mortality from predators (Vermeij and Covich, 1978; Stein et al., 1984; Hawkins and Furnish, 1987), although darters feed on juvenile Elimia when they are abundant in spring and summer (Haag and Warren, 2006). Crayfish also may affect pleurocerids, as E. livescens reared in effluent from crayfish-fed conspecific snails alters shell anatomy to deter crayfish from chipping the shell at the aperture (Krist, 2002).

Ecology of Hydrobiidae The ecology of hydrobiids has received less study than pleurocerids, but factors that influence the abundance of hydrobiids include substrate size, stream shading, water velocity, and flood frequency (Richards, 2004). Martinez and Rogowski (2011) found that density of a springsnail in Arizona springs varied with water depth, distance from a springhead, and pH. Hydrobiids are relatively resistant to flow in large rivers because they burrow into sediments (Holomuzki and Biggs, 1999; Holomuzki and Biggs, 2000). Hydrobiids are annual and recruitment is continuous in warm springs or seasonal in cold systems (Hershler, 1984). Native North American hydrobiids are sexual, sexual dimorphism is pronounced (females larger than males), and sex ratios commonly are skewed toward females (Hershler, 1984).

Hydrobiids can occur at high densities, and community– level interactions seem possible but remain virtually unexplored. In the western United States, the high diversity of hydrobiids may be due the absence of other gill-breathing snails (i.e., viviparids, pleurocerids). Sympatry of congeneric hydrobiids in most springs is rare, but habitat segregation facilitates coexistence in large springs with greater habitat heterogeneity (Hershler, 1984). The role of predators in controlling hydrobiid populations is also relatively unknown, although Cyprinodon pecosensis, Gambusia affinis, and cichlids are known predators (Hershler, 1984).

Conservation and Propagation Successful conservation of extant populations, propagation of at-risk species, and re-introduction of locally extirpated species to stream and spring systems will require more basic research on the life histories and ecology of pleurocerids and hydrobiids (Johnson and Brown, 1997; Brown et al., 2008). For pleurocerids, little is known of reproductive patterns and longevities for Lithasia, Leptoxis, Pleurocera, and Io in rivers in Tennessee and Alabama where they are common. Studies that assess the relative roles of periphyton and spates in determining snail population dynamics are needed, as well as more studies of the role of competition and predators. More information is also needed on successful propagation and re-introduction techniques. Captive rearing of pleurocerids is less complicated than for unionid mussels because of the simpler life-cycle, and snails can be induced to reproduce relatively easily in the laboratory (Figure 18.18). Hydrobiids are difficult to identify, and future studies should use molecular and protein markers, shell morphology, and soft anatomy. Biogeographic studies should address how drainage patterns influence spring

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411

species, subspecies, and unique population units of pleurocerids and hydrobiids merit government protection (Perez and Minton, 2008).

Invasive Species

FIGURE 18.18  Close-up of two Leptoxis foremani ovipositing on the walls of a glass aquarium in the laboratory. The egg masses are the brown circles just above the water. Aquatic gastropods are easier to propagate in the laboratory because they do not have a larval stage dependent on fish for dispersal, like endangered unionid mussels. Photograph courtesy of Paul Johnson.

habitats, as well as basic information on life history, feeding ecology, population dynamics, and habitat requirements of hydrobiids. Finally, further collaboration is needed among geneticists, taxonomists, and agency staff to develop guidelines for use of genetic data when deciding whether at-risk species, subspecies, and unique population units of pleurocerids and hydrobiids merit protection under state and federal laws (Perez and Minton, 2008). Successful conservation of extant populations, propagation of at-risk species, and re-introduction of locally extirpated species to stream and spring systems will require more basic research on the life histories and ecology of pleurocerids and hydrobiids (Brown and Johnson, 1997; Brown et al., 2008). For pleurocerids, little is known of reproductive patterns and longevities for Lithasia, Leptoxis, Pleurocera, and Io in rivers in Tennessee and Alabama where they are common. Studies that assess the relative roles of periphyton and spates in determining snail population dynamics are needed, as well as more studies of the role of competition and predators. More information is also needed on successful propagation and re-introduction techniques. Captive rearing of pleurocerids is less complicated than for unionid mussels because of the simpler life-cycle, and snails can be induced to reproduce relatively easily in the laboratory (Figure 18.18). Hydrobiids are difficult to identify, and future studies should use molecular and protein markers, shell morphology, and soft anatomy. Biogeographic studies should address how drainage patterns influence spring habitats, as well as basic information on life history, feeding ecology, population dynamics, and habitat requirements of hydrobiids. Finally, further collaboration is needed among geneticists, taxonomists, and agency staff to develop guidelines for use of genetic data when deciding whether at-risk

Conservation of native snails also requires understanding threats from invasive snail species. Invasive snails affect native snails directly through competition for food (Carlsson et al., 2004) or indirectly through changes in ecosystem function or parasite populations. Thirty-seven nonnative freshwater gastropod species representing nine families are found in North America (NatureServe.org, accessed 2007). Approximately three dozen examples of invasion and transport snails among states are recognized in the United States (Brown et al., 2008). These species have frequently reduced the abundance of native snails and altered ecosystem function, as can be illustrated by two well-known invasive snails: apple snails and the New Zealand mud snail. Apple snails (family Ampullaridae), especially P. canaliculata, originated in temperate Argentina, and have been introduced to various Asian countries (Dudgeon, 1999) and to US states along the Gulf of Mexico as well. Although they take two years to mature in their native habitat, they take only 10 months in Hawai’i and as little as 2 months in Asia, where they lay their characteristic, calcareous egg masses above the water line to avoid predation (Lach et al., 2000). Apple snails are voracious herbivores that can shift wetland food webs away from a macrophyte base to turbid, phytoplankton-based systems (Carlsson et al., 2004). Pomacea canaliculata has also damaged endangered species and ecosystems (Hall et al., 2003; Carlsson et al., 2004). The channeled apple snail lays bright pink egg masses just above the water’s surface (Figure 18.19). The channeled apple snail aggressively feeds on macrophytes and can convert rapidly to bare substrates (Carlsson et al., 2004). In comparison, the invasive ampullarid Bellamya (Cingopaludina) japonica von Martens has invaded eastern states but does not have as much impact as the invasive apple snails because it filter-feeds and increases water clarity rather than feeding on macrophytes. Ladd and Rogowski (2012) showed that the prosobranch Melanoides tuberculata Müller impacts native snail communities in desert spring systems by direct displacement and introducing foreign trematodes. More work is needed on how invaders interact with native gastropod communities and on mechanisms to control the spread of invasive snails. However, the most prolific invader is the New Zealand mud snail, P. antipodarum (family Hydrobiidae). This species was evidently held in check in its native range by trematode infections. Trematode larvae in New Zealand are locally adapted to parasitize host mud snail populations (see discussion above), and may again explain why sexually reproducing individuals are maintained in populations,

SECTION | IV  Phylum Mollusca

412

FIGURE 18.19 The channeled apple snail (P. canaliculata) is an ampullarid native to South America that has invaded Florida in recent years. The pink egg cases have a calcareous shell to limit dessication. Photograph courtesy of Katasha Cornwell.

due to their genetically variable offspring, or why parthenogenetic morphs cycle in abundance as trematodes adapt to parasitize them (Jokela et al., 2009). In the 1980s, P. antipodarum from New Zealand colonized the Snake River in Idaho, in the United States, where it now reaches densities up to 800,000/m2 (Figure 18.20), with the highest secondary production values ever recorded for an aquatic invertebrate (Hall et al., 2006). Brenneis et al. (2011) found that invasive New Zealand mud snails altered food webs. New Zealand mud snails consume up to 93% of primary production and alter energy flow (Hall et al., 2006). Their invasive success stems from their rapid growth, early maturity, high reproductive rate, resistance to stresses like desiccation during inadvertent transport between sites, and lack of biological enemies in their invasive range (Alonso and Castro-Diez, 2008). Resource consumption rates are no higher than native species, but enclosure experiments suggest asymmetric competitive effects, with resident species at a disadvantage (Riley et al., 2000). Richards (2004) found that the endangered hydrobiid Taylorconcha serpenticola was negatively affected by P. antipodarum. This invasive snail also depressed the abundance of other macroinvertebrates in western US rivers (Kerans et al., 2005). The New Zealand mud snail continues to expand its distribution, with additional populations reported recently in estuarine bays in Oregon and in Lake Ontario. Similarly, P. canaliculata has recently invaded Florida and several other southeastern states, is larger than the native P. paludosa, and may be replacing some populations of the native ampullarid snail. Other taxa that are extremely vulnerable include the springsnails of Australia and western United States and

FIGURE 18.20  The invading New Zealand mud snail (Potamopyrgus antipodarum) reaches high densities in the Snake River in the Western United States, and has impacted other populations of native hydrobiids. Photograph courtesy of D.L. Gustafson.

Mexico (Lydeard et al., 2004). Many of the possibly susceptible hydrobiid taxa are, unfortunately, not yet described. They are vulnerable because of invasive species and habitat destruction, including by cattle grazing and numerous human impacts on small spring ecosystems. Habitat modification likely results in population declines, changes in habitat use, and extirpations of freshwater gastropods. Similarly, Van Bocxlaer et al. (2012) found that gastropods in Lakes Malawi and Tanganyika shifted habitat use to shallower water or have been extirpated, probably as a result of increased surface runoff, eutrophication, and surface-water warming.

COLLECTING, CULTURING, AND SPECIMEN PREPARATION Collecting A number of sampling techniques exist (reviewed in ­Russell-Hunter and Buckley, 1983; Dillon, 2000), although some are not quantitative. The least quantitative technique, but one that often gives large numbers of individuals and a good idea of species composition, is sweep netting with a net of 1 mm mesh (Brown, 1979, 1997). In soft sediments or sand, quantitative samples can be collected with Ekman grabs or corers. When sampling macrophytes, the sampler must collect both plants with attached snails, and the substrate with any bottom-dwelling species. Examples of such samplers are described in Gerking (1957), Savino and Stein (1982), and Lodge et al. (1994). In coarse sand, Ponar dredges are the best alternative. In cobble, little recourse to direct counts of given areas by visual search is available. You can estimate densities by covering the rocks with aluminum foil and then estimating the area from weight to area regressions for the foil.

Chapter | 18  Introduction to Mollusca and the Class Gastropoda

The best sorting technique for gastropods is hand sorting. Large adults can be removed visually and samples washed through a graded series of sieves (≥0.5 mm mesh) to remove mud but retain smaller snails. Place those samples in shallow water on flat white trays, tease the vegetation apart, and then examine the whole tray in a systematic fashion to remove small gastropods and egg cases.

Culturing Temperate gastropods will grow well at 15–20 °C, while subtropical species grow better at 20–25 °C. Any hard substrate with a dense periphyton covering can be added for food, although artificial foods such as lettuce, cereal, or spinach are sometimes used. Provide food as needed to avoid fouling containers. Detritivores should be fed leaf litter colonized with bacteria and fungi (i.e., held for at least two weeks in a pond or stream). Avoid crowding of snails, as growth and reproduction are sensitive to density. An approximate rule is one snail per liter. Water should be recirculated through a gravel or charcoal filter, or at least be changed weekly. Caenogastropod and pulmonate snails should be paired with conspecifics so that mating can occur. Culturing of “weedy” species like physids is best done at low temperatures to retard egg production, and constant removal of egg cases is necessary to prevent population explosions. Adequate lighting (with a 12L:12D cycle) is necessary to promote periphyton growth in aquaria (“gro lites” work well).

Specimen Preparation and Identification Identification of freshwater gastropods requires patience and some familiarity with general differences among gastropod groups. For example, individuals should be collected alive if possible so that the presence of an operculum can correctly indicate the specimen is a caenogastropod. In general, gross shell characteristics can initially be used before determining whether an operculum is present. However, some confusion between pulmonate and pleurocerid snails, for example, is a common mistake if novice students use preserved specimens and do not correctly note the presence or absence of an operculum. While there is undoubtedly much eco-phenotypic variation in shell morphology, understanding the basic shell shapes and morphology is still very important in using a dichotomous key, at least at higher levels. If at all possible, a range of shell sizes should be collected, as size is used as a classification trait in several instances. In the case of hydrobiids, little recourse exists to dissecting the snail and observing penial morphology under a dissecting microscope. We hope progress will be made in the future in using molecular characteristics in identification

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of gastropods, to help solve the problem with phenotypic plasticity or convergence of shell shapes. However, at the current time, no sophisticated (e.g., using both molecular and morphological traits) overall study of species determination has been done for caenogastropods. Although pulmonate species are somewhat easier to discern, there is still controversy about higher taxonomic levels (genus and above).

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Chapter | 18  Introduction to Mollusca and the Class Gastropoda

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Chapter | 18  Introduction to Mollusca and the Class Gastropoda

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Chapter 19

Class Bivalvia1 Kevin S. Cummings Illinois Natural History Survey, Center for Biodiversity, Champaign, IL, USA

Daniel L. Graf Biology Department, University of Wisconsin-Stevens Point, Stevens Point, WI, USA

Chapter Outline Introduction424 Introduction to the Class Bivalvia 424 General Phylogenetic Relationships 424 Evolution and Phylogenetics 425 Diversity of the Nearctic Unionoidea 426 Family-Level Relationships 426 Species Diversity and Distributions 428 Population Genetics, Phylogeography, and Molecular Evolution 431 Diversity of the North American Sphaeriidae 431 Supra-Specific Relationships 432 Species Diversity and Distributions 433 Polyploidization433 Diversity of Corbicula and Dreissena in North America 434 General Biology 434 External and Internal Anatomy 434 Shell and Mantle 434 Shell Growth and Microstructure 435 Internal Shell Structure 436 External Shell Structure 438 Mantle Cavity and Apertures 440 Ctenidia441 Feeding Structures and Mechanisms 441 Mechanics of Suspension Feeding 442 Clearance Rates and Retention Efficiencies 442 Particle Sorting and Labial Palps 445 Ingestion, Digestion, and Assimilation 445 Deposit Feeding 448 Foot and Locomotion 449 Physiology451 Circulation and Respiration 451 Osmoregulation and Excretion 452 Neural Systems and Sense Organs 453

Reproductive Anatomy 454 Gonads, Gametes, and Fertilization 454 Brooding457 Larval Morphology 459 General Ecology 460 Life Cycles 460 Unionids461 Sphaeriids and Corbicula466 Age and Growth 467 Population Demography 468 Biotic Interactions 468 Predation468 Parasites470 Diseases471 Ecosystem Processes 471 Habitat and Abiotic Interactions 472 Macro- and Microhabitat Characteristics 472 Substrate, Sediment, and Stream Flow Preferences473 Effects of pH 474 Depth Effects 475 Oxygen and Air Exposure 475 Temperature476 Changes in Hydrology 476 Floods476 Impoundments477 Channelization477 Pollution and Freshwater Bivalves as Biomonitors 478 Sewage478 Metals479 Non-Essential Metals 479 Essential Elements 480 Pesticides and PCBs 481

1 This chapter was slightly rearranged by James H. Thorp to fit the format and section titles of the fourth edition and full citations were added from a previous Elsevier Website, but the text and figures are taken largely intact from Cummings, K.S., Graf, D.L., 2010. Mollusca: Bivalvia. Chapter 11. In: Thorp, J.H., Covich, A.P., (Eds.), Ecology and Classification of North American Freshwater Invertebrates, third ed. Academic Press, Boston, MA, pp. 309–384. Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00019-X Copyright © 2015 Elsevier Inc. All rights reserved.

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Endangered Species and Conservation 481 Consumptive and Commercial Impacts 482 Invasive Bivalves (Corbicula, Dreissena, and Sphaeriids) 483 Zebra Mussels and Benthic Invertebrate Interactions 484 Collecting, Curation, and Rearing 484 Collecting484

INTRODUCTION Introduction to the Class Bivalvia The freshwater bivalves of North America are represented by two native groups, the freshwater mussels (Unionoidea; Appendix 19.1) and the pill, fingernail, and pea clams (Sphaeriidae), as well as two widely publicized invasive genera, Corbicula and Dreissena (Figure 19.1). These mollusks2 have interesting and important ecological interactions with their environments, not the least of which is their relationship to humans. As regards freshwater mussels (also called pearly mussels or naiads), humans have been among their most potent enemies, but today considerable resources are dedicated to reversing the wave of decline and extinction that these mollusks face. Conversely, the minute sphaeriids have received comparatively less attention; however, the review of the ecology and classification of these bivalves by Mackie (2007) has been a timely and valuable addition to the malacological community. Our emphasis with this chapter will be on the native lineages, although relevant comparative information about the exotic species and genera is also provided.

General Phylogenetic Relationships The taxonomic nomenclature of North American freshwater bivalves is a potential source of confusion, and we have tried to be conservative in our use of names, following the taxonomy of Graf and Cummings (2007) and Lee (2004). Freshwater mussels are discussed at various systematics levels of generality. The order Unionoida, superfamily Unionoidea, and family Unionidae, are colloquialized to unionoids, unionoideans, and unionids, respectively. Only two families occur in North America (the other being the Margaritiferidae), and those two comprise the Unionoidea. We use Unionoida when referring to the global order of six families. Older North American literature often referred to the Unionacea, which can usually be regarded as synonymous with Unionoidea (-oidea being the preferred superfamilial suffix rather than -acea). It has recently been proposed that 2 Members of the phylum Mollusca are referred to as either mollusks (preferred by the authors of this chapter) or molluscs (preferred by many other authors, especially in Europe); either spelling is correct.

Survey Methods 484 Season, Permits, and Gear 485 Data Recording 485 Curation486 Culture, Propagation, and Rearing 486 References490

using the name Unioniformes rather than Unionoida might eliminate some of the confusion over so many similar-looking names and bring the ordinal suffix more in line with those of fishes and birds (Nelson et al., 2004). For this work, we have decided to keep the conjugation of the Unionoida in agreement with the rest of the bivalve orders. There is less confusion with sphaeriid nomenclature because there is only a single family. Based upon recent analyses, we regard each of these higher taxonomic units to represent clades rather than mere assemblages of convenience. Thus, they are referred to in the singular. The bias of this chapter is toward the freshwater mussels. They are the numerically dominant group, with 10 times more species, and considerably more data are available relative to the sphaeriids. Both of these lineages of bivalves share many of their ecological habits with each other and their marine counterparts. The stereotypical freshwater bivalve lives in the sediment at the bottom of lakes and streams. The entire bivalve body is enclosed in a calcareous shell composed of two halves (valves, hence the name), and the only soft protrusions from this rocky exterior are the soft anterior foot for burrowing and the posterior apertures of the mantle. These mollusks are predominantly filter-feeders, drawing water through their enlarged ctenidia for both gas exchange and to capture particulate matter from the water column. All native, North American freshwater bivalves are adapted for life in flowing water, having given up a planktonic larval stage for parental care in the form of larval brooding. Freshwater mussels have taken this specialization further by producing parasitic larvae infecting fishes (and, in at least one case, an amphibian) as hosts. It is on their host that the mussel larvae, known as glochidia, undergo metamorphosis. The mussels tend to have long lives (up to 102 years) and generally produce 104–106 larvae per breeding season, each with a low probability of survival. Sphaeriid species, in contrast, are often adapted to ephemeral habitats, produce orders of magnitude fewer direct-developing offspring, and tend to have shorter life histories. However, among the species of both groups, there is considerable variation in strategies and a wide range of ecological and diversity patterns can be observed in the North American species. We have endeavored to describe the morphology and general life histories from both groups of native bivalves

Chapter | 19  Class Bivalvia 

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Toxolasma

Potamilus

Megalonaias

Utterbackia Alasmidonta

Amphinaias

Sphaerium Elliptio

Corbicula

Pisidium

Dreissena

FIGURE 19.1  Conchological variation in the shells of North American freshwater bivalves. All shells are relative sizes.

and to provide discussions of the ecological functions and evolutionary diversification of these mollusks. The last few decades have provided a wealth of new information, especially from the perspectives of conservation biology and systematics, and new data is arriving at a pace faster than we can incorporate it. To facilitate further research on the freshwater bivalves, we have included a key to the genera of the Nearctic fauna and copious photographs to facilitate identification (Editor’s note: now see Chapter 11 in Volume II for information on identifying Nearctic bivalve mollusks).

Evolution and Phylogenetics Two distinct, ancient lineages of freshwater bivalves occur in the freshwaters of North America: the Palaeoheterodonta (represented by the Unionidae and Margaritiferidae) and the Heterodonta (Sphaeriidae). Corbicula and Dreissena, two invasive genera, are heterodonts as well. Both of these ancient clades (subclasses) originated in the Paleozoic (Moore, 1969; Cope, 1996), and over the last few 100 million years, they have evolved to fill various niches. The Palaeoheterodonta arose as a marine taxon, with the freshwater mussels (order Unionoida) appearing

perhaps as early as the Triassic (Haas, 1969a; Newell and Boyd, 1975; Watters, 2001). The marine palaeoheterodont lineages (order Trigonioida) were hit hard by the extinction event marking the close of the Cretaceous, and today only a single genus, Neotrigonia, remains as the extant sister group to the modern Unionoida (Allen, 1985; Darragh, 1998). Recent freshwater palaeoheterodonts (i.e., freshwater mussels) are represented worldwide by six families, two of which (Unionidae and Margaritiferidae) are widespread across the northern continents, including North America (Graf and Cummings, 2007). In contrast, the heterodonts are currently the dominant group of marine bivalves. The order Veneroida, in particular, diversified during the Mesozoic and Cenozoic eras (Stanley, 1968). Seven veneroid families have invaded estuarine and freshwater environments, including the Sphaeriidae (Deaton and Greenberg, 1991; Bogan, 2008; Bogan and Roe, 2008). Sphaeriids are worldwide in distribution, including North America (Burch, 1975a). The family is represented in the fossil record back to the Cretaceous (Keen and Casey, 1969), but its nearest marine relative is unknown. Traditionally, sphaeriids were grouped with the Corbiculidae based upon their shared adaptation to freshwater and conchological similarities

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(Keen and Casey, 1969). However, more recent molecular work has shown independent radiations of those families from the marine veneroids (Park and Foighil, 2000). Although the native freshwater bivalve species coalesce to only three families, the various subfamilial lineages (down to below the species level) reflect the complex history of connectivity between North America and other landmasses and among more localized areas of endemism. Freshwater mussels and pill clams share many of the general characteristics of bivalves, and both are found in freshwaters around the world, but that is the limit of their similarity. The species of these two groups have very different life histories, ecologies, and dispersal abilities, and as a result, the diversity patterns of the Unionoidea and Sphaeriidae provide an interesting contrast. We will begin with a discussion of the diversity of the freshwater mussels. The Sphaeriidae is less well-studied (historically), but recent phylogenetic work has shed new light on the group. Finally, the systematics of the recently introduced lineages of Corbicula and Dreissena will be addressed.

Diversity of the Nearctic Unionoidea A number of cladistic studies have reviewed and refined our understanding of the phylogenetic relationships among the Nearctic genera of the Unionidae (Lydeard et al., 1996; Hoeh et al., 2001; Roe and Hoeh, 2003; Campbell et al., 2005; Graf and Cummings, 2006) and Margaritiferidae (Huff et al., 2004; Walker et al., 2006). That research has built upon previous traditional arrangements relying on morphological characteristics (Ortmann, 1912, 1916; Thiele, 1934; Modell, 1964; Haas, 1969a; Heard and Guckert, 1971; Burch, 1975b; Smith, 2001) and phenetic studies based upon allozymes and immunological assays (Davis and Fuller, 1981; Davis, 1984; Hoeh, 1990). Most of the more recent cladistic studies have relied upon nucleic acid sequences (most often mtDNA), but a few have made an effort to determine morphological synapomorphies (Lydeard et al., 1996; Graf and Foighil, 2000; Hoeh et al., 2001; Roe and Hoeh, 2003). This work is ongoing, and the currently accepted taxonomy of North American freshwater mussel genera and species will likely be changed and improved as the efforts of the freshwater malacological community progress. At the family-group level, however, a number of patterns have been repeated in the various analyses of molecules and morphology, making it possible to draw some robust conclusions about the pattern of evolution of the Unionidae and Margaritiferidae.

Family-Level Relationships Over the last 100 years, the family-level taxonomy of these species has been shuffled and reshuffled as new data, methods, or theories have been discovered. The available

family-group names applied have been relatively stable, but the various groupings of genera and species have not. It would perhaps be instructive for the student of the history of freshwater malacology to review the arc of progress that has led us from the arrangement proposed by Charles Torrey Simpson (1900, 1914) to that listed in Table 19.1. ­However, for those interested in freshwater mollusks, it should be sufficient to point out that (1) pre-cladistic systems were based upon flawed methodologies, either upon authoritarian essays or studies making no distinction between synapomorphy and plesiomorphy (derived vs. ancestral homologies); and (2) cladistic analyses have been reasonably consistent with regard to the patterns of phylogeny recovered. We will dwell upon the current system of the Nearctic Unionoidea. A thorough review of the history of freshwater mussel classification is available elsewhere (Roe and Hoeh, 2003). Globally, the Unionidae is the largest family of freshwater bivalves with more than 670 currently recognized species, making it among the largest bivalve families in marine as well as freshwater environments (Graf and Cummings, 2007). Unionids possess parasitic glochidia and their larvae are brooded in either the lateral (outer) pair of ctenidial demibranchs or in all four. The principle diagnostic character for the family is the presence of TABLE 19.1  Family-Level Classification of North American Freshwater Bivalves (Lee, 2004; Graf and Cummings, 2006) Order Unionoida   Superfamily Unionoidea Rafinesque, 1820   Family Unionidae Rafinesque, 1820   Subfamily Unioninae Rafinesque, 1820   Tribe Anodontini Rafinesque, 1820   Subfamily Ambleminae Rafinesque, 1820   Tribe Amblemini Rafinesque, 1820   Tribe Lampsilini von Ihering, 1901   Tribe Pleurobemini Hannibal, 1912   Tribe Quadrulini von Ihering, 1901   Tribe Gonideini Ortmann, 1916   Family Margaritiferidae Haas, 1940 Order Veneroida   Family Sphaeriidae Deshayes, 1854   Subfamily Sphaeriinae F.C. Baker, 1927   Subfamily Euperinae Heard, 1965   Family Corbiculidae gray, 1847 [introduced]   Family Dreissenidae gray, 1840 [introduced]

Chapter | 19  Class Bivalvia 

427

a supra-anal aperture (Graf and Cummings, 2006). The North American unionid assemblage is currently estimated at ∼300 species (Williams et al., 1993; Graf and Cummings, 2007; Bogan, 2008; Bogan and Roe, 2008). The North American Unionidae is represented by two subfamilies: Unioninae and Ambleminae (Figure 19.2) (Graf, 2002b; Graf and Cummings, 2006). The Unioninae is Holarctic in distribution, with only a few species spreading south into Central America, Africa, or southeastern Asia. The subfamily is diagnosed by large (200–380 μm), subtriangular, hooked-type glochidia. One tribe of the Unioninae, the Unionini (e.g., Unio), is strictly Old World in distribution. The other tribe, the Anodontini, is Holarctic in distribution, with representatives on both sides of the continental divide in North America (Graf and Cummings, 2007). Anodontines are generally without schizodont pseudocardinal hinge teeth, and the laterals, if present, tend to be rudimentary. However, some taxa (e.g., Lasmigona) have secondarily derived hinge teeth. Species are bradytictic (long-term brooding), and the marsupial demibranchs bear lateral, accessory water-tubes and have ventral margins enhanced with additional tissue to allow for greater expansion (Graf and Foighil, 2000). The area Unionini 7 3 10† Anodontini 12 11

Lampsilini

7

Pleurobemini Quadrulini

10†

Amblemini 9†

9†

Gonideini Old World unionids

1

4 2

8† 5

10† Margaritiferidae 6 Etherioidea Neotrigonia

FIGURE 19.2  Phylogeny of the Unionoidea, showing evolutionary transformations of key morphological characters. Taxa with representatives in North America are in bold-faced type. Synapomorphies are coded white, gray and black to indicate the progression from plesiomorphic to derived transformations. Characters: (1) inhabits freshwater; (2) unhooked, parasitic glochidia; (3) hooked, subtriangular glochidia; (4) brooding; (5) interlamellar spaces of the ctenidia divided by perforated septa into water-tubes; (6) reduction of water-tubes; (7) interlamellar spaces divided by complete septa; (8) brooding in only the inner demibranchs (endobranchy); (9) brooding in all four demibranchs (tetrageny); (10) brooding in only the outer demibranchs (ectobranchy); (11) supra-anal aperture; and (12) bradytictia. (Adapted from Graf & Ó Foighil, 2000 and Graf & Cummings, 2006).

from the Arctic south to Mexico is inhabited by 44 species of the Anodontini (Unioninae) (Graf and Cummings, 2007). The subfamily Ambleminae is much more diverse than the Unioninae in North America (253 species) (Graf and Cummings, 2007). The subfamily may be endemic to North America, although the phylogenetic relationships among New and Old World unionid lineages have thus far been approached with too few taxa and characteristics to derive any meaningful conclusions (Claassen, 1994). Based upon the current evidence, only a single amblemine species occurs west of the Rocky Mountains; the remaining species are found east of the Continental Divide. Gonidea angulata in the Pacific Basins has been argued to share a more recent common ancestor with certain eastern Asian taxa than any of the eastern Nearctic tribes, but this has not been supported by molecular analysis (Graf, 2002b). The four eastern tribes of the East (Amblemini, Quadrulini, Pleurobemini, and Lampsilini) have consistently been recovered as monophyletic, and the clade has been informally named the “Amblemini Tribe group.” The group is diagnosed by the presence of complete (i.e., imperforate) septa dividing the interlamellar spaces of the demibranchs. The species of the Ambleminae possess either the plesiomorphic unhooked or the axehead-type glochidia, are either tachytictic or bradytictic, and may brood their larvae in either all four demibranchs or only the outer (lateral) pair of demibranchs (Graf and Cummings, 2006). These characteristics associated with reproduction and brooding were given great weight in historical arrangements of the Unionoida (Ortmann, 1912; Heard and Guckert, 1971; Davis, 1984; Lydeard et al., 1996; Graf and Foighil, 2000). Because nearly the full range of global variation was observed within the North American assemblage, it has been assumed that the worldwide Unionidae could be accommodated in the system derived for the North American taxa—thus, the source of the modern confusion over the relationships between New and Old World mussels. While these reproductive characteristics have been shown to be misleading for diagnosing the intergeneric relationships of the Ambleminae (indeed, only the Lampsilini has been well characterized by morphological characteristics), they are conservative among congeneric species (Graf and Cummings, 2006). This, however, may simply be due to the over-emphasis of reproductive characteristics in our current generic definitions, and forthcoming revisions based upon combined molecular and morphological evidence may render this generalization inaccurate (Campbell et al., 2005). The interrelationships among the lineages of the Amblemini Tribe group are equivocal, and we have left the tribes unresolved in the phylogeny in Figure 19.2. The Amblemini, Quadrulin, and Pleurobemini are recognized largely by molecular characteristics (Davis, 1984; Lydeard et al., 1996; Campbell et al., 2005). The Lampsilini (122 Nearctic species + 28 in Mesoamerica), in contrast, is well diagnosed by a suite of morphological and behavioral

428

characteristics (Graf and Foighil, 2000). The marsupium is restricted to only a portion of the outer demibranchs. Among the core lampsilines, it is restricted to a posterior section, it is capable of great expansion, and the shells of the females are more inflated as well (i.e., sexually dimorphic). In the most derived genera of the tribe, there are also a number of adaptations for host attraction, including larvae packaged into conglutinates or super-conglutinates and mantle lures mimicking host prey items (Zanatta and Murphy, 2006; Barnhart et al., 2008). While these sex-oriented traits are a boon for those making an argument for conserving the Nearctic freshwater mussel assemblage, most unionids (worldwide) apparently lack elaborate host attraction adaptations. However, new discoveries of interesting variations on the typical model are still being discovered (Vicentini, 2005; Barnhart et al., 2008). Among the other three amblemine tribes, the taxonomic and morphological diversity is generally lower than seen among the lampsilines, and the genera have been based largely upon shell characteristics and the number of brooding demibranchs (Ortmann, 1912; Simpson, 1914). Those tetragenous lineages with shell sculpturing are generally placed in either the Amblemini (3 species), or Quadrulini (26 species), depending upon their mitochondrial lineage. The Pleurobemini (100 species) are generally ectobranchous with only a few genera brooding in all four demibranchs (Graf and Foighil, 2000; Graf and Cummings, 2007). Existing hypotheses of the interrelationships among the members of the Amblemini Tribe group remain to be robustly tested. The sister group to the Unionidae is the Margaritiferidae (Roe and Hoeh, 2003; Graf and Cummings, 2006). Margaritiferids differ from unionid species in lacking any kind of posterior mantle fusion, the diaphragm dividing the infrabranchial from the suprabranchial chamber is grossly incomplete, and the interlamellar spaces of the ctenidia are not divided by vertical septa (Graf and Cummings, 2006). There is variation among the species with regard to the development of interlamellar junctions (Smith, 2001). The Margaritiferidae has a patchy, Holarctic distribution (including southeastern Asia), with only about a dozen species worldwide (Ziuganov et al., 1994; Smith, 2001; Huff et al., 2004; Graf and Cummings, 2007). Based upon the morphological simplicity exhibited, the family may represent living fossils from the early diversification of the Unionoida. However, an equally parsimonious interpretation is that margaritiferids are derived and degenerate (Graf, 2002b; Graf and Cummings, 2006). Together, the Unionidae and Margaritiferidae form one of two superfamilies of the Unionoida: the Unionoidea (Graf and Cummings, 2006). Historically, the Hyriidae of South America and Australasia were grouped with the Unionoidea based upon the shared characteristic of parasitic glochidia (Parodiz and Bonetto, 1963). However, more recent phylogenetic work (utilizing larval and adult morphology as well as molecular characteristics) has suggested that glochidia are the primitive larval condition among the Unionoida (i.e., not useful for forming groups among unionoids), and

  SECTION | IV  Phylum Mollusca

hyriids were recovered as part of a clade found on the southern continents (superfamily Etherioidea) (Graf, 2000; Graf and Cummings, 2006). Other etherioidean lineages have parasitic larvae known as lasidia (Wächtler et al., 2001). Based upon current taxonomy and fossil distributions, it is possible that the extant lineages of the Unionoidea first arose on the northern continents (Laurentia) following the breakup of Pangaea early in the Mesozoic. Subsequently, as fragments of the southern supercontinent (Gondwana) have reconnected with the north, unionoideans have spread south into Africa and India, for example (Graf, 2000).

Species Diversity and Distributions The taxonomy of the Unionoida has been, and continues to be, in a state of flux. In our discussion above, we were able to largely ignore the historical dynamics of higherlevel nomenclature due to the relative conservatism of the family-group level names applied and the acknowledgment that previous arrangements lacked the rigor of a phylogenetic classification (and could therefore be ignored in the light of current information). While the road to our present understanding of species-level freshwater mussel diversity in North America has been as bumpy, it is more difficult to ignore. Freshwater mussels, because of their large size, conspicuous ecological roles, and high diversity, have attracted no small amount of interest over the last 200+ years. The first phase of taxonomic research was purely descriptive and lasted until the end of the nineteenth century. Rafinesque, Conrad, J.G. Anthony, and many others, but especially Isaac Lea, introduced over 1400 species names to describe the North American assemblage. A narrative phase began with the turn of the twentieth century and C.T. Simpson’s synopses of the global mussel fauna (1900, 1914). Until near the end of the last century, estimates of freshwater mussel species diversity in North America were continually revised as interest rose and fell, and as the philosophical climate changed. For example, the growing database of soft-anatomical and behavioral characteristics, the adoption of the International Code of Zoological Nomenclature (ICZN), the New Synthesis, and the modern renaissance in freshwater mussel conservation have all likely influenced the genus and species-level classifications of Nearctic unionoideans at different times—not to mention the backgrounds, biases, and personalities of the malacologists doing the work. It is significant, however, that of the several recent phylogenetic studies focused at the species level (Mulvey et al., 1996; Roe and Lydeard, 1998; Lydeard et al., 2000; Buhay et al., 2002; Mock et al., 2004; Jones et al., 2006b; Serb, 2006; Serb et al., 2003), it has been rare for the work to be translated into taxonomic revisions (Hoeh, 1990; Roe et al., 2001; Roe and Hartfield, 2005). Thus, much of what is currently understood of freshwater mussel species-level systematics and diversity is incomplete, especially with regard to molecular data (Campbell et al., 2005; Graf and Cummings, 2007).

Chapter | 19  Class Bivalvia 

429

Following Simpson, numerous workers have contributed to our present consensus of the species-level diversity and distributions of the Unionoidea in North America. Since 1900, there have been nine comprehensive treatments of all Nearctic mussel species and many more localized studies (Graf and Cummings, 2007). The overall trend from 1900 to the 1980s was to reduce the number of recognized (i.e., valid) species-level taxa (Figure 19.3). However, both the resurgence in conservation interest and the availability of new technologies for genetic research prompted new interest in endemic taxa with restricted ranges and a desire to reevaluate the North American freshwater mussel species. It has been pointed out that since the Endangered Species Act in the United States does not protect population-level lineages, freshwater bivalve taxonomists may feel they need elevate the status of certain populations to bring them under those legislative provisions (King et al., 1999). This may, in large part, explain the rise in recognized species diversity toward the end of the last century. From a low species count of 227 in 1975 (Burch, 1975b), more recent estimates range from 297 to 302 (Williams et al., 1993; Graf and Cummings, 2007; Bogan, 2008; Bogan and Roe, 2008). These estimates, however, must be regarded as historical, given the high rate of extinction reports for freshwater mollusks in general. Thirty-one species and three recognized subspecies are currently regarded as extinct (Table 19.2), and the conservation of the remaining malacofauna is an area of active research.

Alasmidonta mccordi Athearn, 1964 Alasmidonta robusta Clarke, 1981 Alasmidonta wrightiana (Walker, 1901) Elliptio nigella (Lea, 1852) Epioblasma arcaeformis (Lea, 1831) Epioblasma biemarginata (Lea, 1857) Epioblasma flexuosa (Rafinesque, 1820) Epioblasma haysiana (Lea, 1834) Epioblasma lenior (Lea, 1840) Epioblasma lewisii (Walker, 1910) Epioblasma personata (Say, 1829) Epioblasma propinqua (Lea, 1857) Epioblasma sampsonii (Lea, 1862) Epioblasma stewardsonii (Lea, 1852) Epioblasmaturgidula (Lea, 1858) Lampsilis binominata Simpson, 1900 Medionidus mcglameriae van der Schalie, 1939 Pleurobema altum (Conrad, 1854) Pleurobema avellanum Simpson, 1900

1500

Pleurobema bournianum (Lea, 1840) Pleurobema chattanoogaense (Lea, 1858)

1250

Pleurobema flavidulum (Lea, 1861) Pleurobema hagleri (Frierson, 1900)

1000 Species

TABLE 19.2  Extinct Species and Subspecies of Native North American Freshwater Bivalves (Based on Turgeon et al., 1998; Bogan, 2006)

Pleurobema hanleyianum (Lea, 1852) 750

Pleurobema johannis (Lea, 1859) Pleurobema murrayense (Lea, 1868)

500

Pleurobema nucleopsis (Conrad, 1849) Pleurobema rubellum (Conrad, 1834)

250 0 1750

Pleurobema troschelianum (Lea, 1852) Pleurobema verum (Lea, 1861) 1800

1850

1900 Year

1950

2000

FIGURE 19.3  The accumulation of available species-group level nomina and changes in the number of valid species recognized over time for the North American freshwater mussel assemblage. The data for this graph were derived from the MUSSEL Project Database (Graf & Cummings, 2007). Circles indicate the dates of key works that dealt with the entire Nearctic freshwater mussel assemblage: Adams & Adams (1957) (reference not listed), Lea (1870) (reference not listed), Simpson (1900, 1914), Frierson (1927), Haas (1969b), Burch (1975a), Turgeon et al. (1998), Williams et al. (1993), and Graf and Cummings (2007).

Theliderma tuberosa (Lea, 1840) Epioblasma florentina florentina (Lea, 1857) Epioblasma torulosa torulsa (Rafinesque, 1820) Epioblasma torulosa gubernaculum (Reeve, 1865)

  SECTION | IV  Phylum Mollusca

430

The historical species group-level diversity of the Unionoidea is not uniformly distributed on the North American continent. Instead, there are species (and genera) with ranges restricted to single biogeographical provinces and species with wide ranges, occurring in multiple provinces. The observed distribution patterns of freshwater mussels are very much a product of their history. Because of their complex life cycles and reliance upon freshwater fishes for dispersal, unionoidean species have no capacity for overland dispersal (van der Schalie, 1939; Johnson, 1970; Graf, 1997c; Graf, 2002a). Even if a freshwater mussel were passively dispersed from one basin to another, for that event to lead to colonization, at minimum the mussel would have to be a gravid female and a suitable host population would need to have already been established. Thus, geographical disjunctions within freshwater mussel taxa across drainage divides are strong evidence for historical confluence (van der Schalie, 1945). Areas of endemism reflect both geological age and isolation; that is, the populations were isolated long enough for speciation to occur and/or isolated enough to be protected from extirpation throughout the rest of their range (Brown and Lomolino, 1998). This argument, of course, does not apply to human-mediated introductions of freshwater mussels via transportation of host fish. Sinanodonta woodiana, for example, has been introduced widely through the movement of exotic fishes (Watters, 1997).

Based upon patterns of freshwater mussel species diversity and endemism, the North American continent can be divided into biogeographical provinces (Figure 19.4) (Johnson, 1980; Parmalee and Bogan, 1998; Watters, 2000; Graf and Cummings, 2007). Various degrees of connectivity (as determined by species shared between provinces) have been used to test hypotheses of past confluence, but most of these hypotheses have dealt with historical biogeographical hypotheses operating on scales of 103 years to explain current species distributions. Older events correlated with the origins of lineages above the species level are less well understood. The primary barrier to freshwater mussel distribution is the continental divide separating the Pacific from the Atlantic drainages. Nine species are known from the Pacific basins of North America, whereas 295 are known from east of the Rocky Mountains. Only two species are shared between the two regions (Figure 19.4). The shared species indicate areas of severed lotic connections due to geological changes in the Rocky Mountains (Johnson, 1980). The rich mussel fauna of the eastern portion of North America provides ample species-level distribution patterns that can be employed to test hypotheses of biogeographical processes, and the influence of the relatively recent, endPleistocene glacial maximum on mussel distributions has been given a great deal of attention (Johnson, 1980; Graf, 1997b, 2002a). The southern extent of Pleistocene glaciation reached approximately to the modern-day Missouri and

Canada 15 (0)

Gr. Lakes 46 (0)

Pacific 9 (7)

Missouri 33 (0)

N. Atlantic 23 (3)

Up. Miss. 51 (0) Ohio 80 (2) Ozark 77 (11)

Species diversity in each ecoregion S. Atlantic 54 (33) Cumberland 103 (30)

W. Gulf 53 (19) Low. Miss. 57 (2)

Mobile 84 (41)

E. Gulf 52 (29)

≥81 61–80 41–60 21–40 ≥ 20

Florida 14 (7)

FIGURE 19.4  Freshwater ecoregions of North America, showing the species diversity (endemism) of each region. (Adapted from Parmalee & Bogan, 1998; Watters, 2000; and various other literature sources.)

Chapter | 19  Class Bivalvia 

Ohio Rivers. All mussel species (and all other organisms) found north of the last ice advance (Wisconsinan) dispersed there from southern refugia within the last 15,000 14C-years. Those species present in the Missouri, Upper Mississippi, Canadian, and Great Lakes drainages dispersed from protected highlands on the Cumberland and Ozark Plateaus, whereas those on the Northern Atlantic Slope (including the St. Lawrence River) originated east of the Appalachian Mountains on the Southern Atlantic Slope (Johnson, 1980). Like the Rocky Mountains to the west, the Appalachians present a hard barrier to freshwater mussel dispersal, although not as stringent, with various historical points of connection in the north and south (Johnson, 1970, 1980).

Population Genetics, Phylogeography, and Molecular Evolution Population genetic and phylogeographic studies using allozymes, mitochondrial DNA, and microsatellites have been used to address the genetic structure of freshwater mussel populations in both the interior basin and the Atlantic slope regions of North America (Kat, 1982a, 1983b; Kat and Davis, 1984; Berg et al., 1998; King et al., 1999; Turner et al., 2000; Burdick and White, 2007; Elderkin et al., 2007; Zanatta et al., 2007; Zanatta and Murphy, 2007, 2008). The common theme of this research is that, for species found north of the Pleistocene glacial maximum, the recovered patterns are consistent with the rapid northward expansion of populations from southern refugia. For example, nuclear allele and mtDNA haplotype diversity is generally lower in northern populations, and southern populations located in hypothesized refugia frequently harbor unique alleles and haplotypes (Burdick and White, 2007; Elderkin et al., 2007; Zanatta and Murphy, 2008). The population structure of unionoidean species is influenced by other factors as well, including life-history traits (Hoeh et al., 1998). For example, freshwater mussels of the Atlantic Slope with anadromous/ salt-tolerant hosts display higher levels of gene flow between populations in disjunct river basins than those with more territorial hosts (Kat and Davis, 1984), and lower-than-expected levels of heterozygosity among populations of Margaritifera has been argued to result from the meta-population dynamics of their patchy distribution (Curoleet al., 2004). These types of studies examining the intra- and inter-population genetic diversity of freshwater mollusks are vitally important to understanding the Nearctic species diversity. At the genome level, freshwater mussels are unique for the pattern of doubly uniparental inheritance (DUI) of mitochondria that they present. DUI has been extensively documented among marine mytilid mussels (true mussels) (Zouros et al., 1992, 1994) and among veneroids as well (Passamonti and Scali, 2001), but neither of these taxa have maintained sex-specific mitochondrial genomes as conservatively as the Unionoidea. Among animals generally,

431

mitochondria are passed strictly maternally. The plastid contributions from the sperm to the zygote are not observed in adult somatic or germ tissue, nor in the subsequent generation. In bivalves that exhibit DUI, like freshwater mussels, the paternal mitochondria from the sperm are, in male zygotes, sequestered into the germ-line cells in the developing embryo. Thus, while female freshwater mussels are homoplasmic, bearing only maternal-lineage mitochondria in both their somatic and germ tissue, male germ tissue is homoplasmic for paternal lineage mitochondria. Male somatic tissue is heteroplasmic (Breton et al., 2007). The result of DUI is that these two, nonrecombining mitochondrial genomes can evolve independently, as evidenced by their divergent DNA sequences (Hoeh et al., 1996). Freshwater mussels differ from marine mussels in the degree to which these two parental lineages of mitochondria are maintained. Whereas mytilid paternal lineages are occasionally feminized, resetting the divergence between maternal and paternal sequences (generally referred to an F-type and M-type mtDNA), the Unionoidea seems to have maintained perfect DUI for 1010 years, based upon the phylogenetic distribution of the phenomenon (Curole and Kocher, 2002; Walker et al., 2006). F-type and M-type mitochondria of all freshwater mussel species belong to parental-lineage specific clades. The utility of these two genomes for species phylogenies and the differing patterns of their molecular evolution are areas of active research. M-type mtDNA seem to be evolving at a rate orders of magnitude faster than that of the F-type (Liu et al., 1996; Walker et al., 2006), but there are only limited gene arrangement differences at the genome level (Curole and Kocher, 2002; Breton et al., 2007).

Diversity of the North American Sphaeriidae The evolutionary patterns of the Sphaeriidae provide interesting contrasts to the phylogenetic and species diversity of the Unionoida. While in general sphaeriid genera and species tend to have larger geographical ranges than those of freshwater mussels—Pisidium casertanum, for example, is cosmopolitan in distribution (Clarke, 1973; Burch, 1975a)—the actual diversity is much lower. Whereas worldwide ∼840 freshwater mussel species are currently considered valid, only as many as 200 sphaeriids are recognized in four genera (or five, depending upon the treatment of Musculium; see below) (Korniushin, 2006; Bogan, 2008). The Nearctic sphaeriid assemblage consists of 40 species including five exotics (Grigorovich et al., 2000) in three (or four) genera (Turgeon et al., 1998), with the majority of species occurring in the formerly glaciated areas of the north and the Rocky Mountains, where mussel diversity is low. At the genome level, sphaeriid species are unique among Nearctic freshwater bivalves for their copious chromosomal material, most species being polyploid and showing evidence of gene duplication (Lee, 1999; Petkeviciute

  SECTION | IV  Phylum Mollusca

432

et al., 2007). These interesting biological traits have made sphaeriids a boon for life-history studies (Holopainen, and Hanski, 1989; Guralnick, 2004), but they have also made the evolution of these miniature bivalves difficult to understand and classify.

Musculium

8

7

Sphaerium

Supra-Specific Relationships The relationships of the Sphaeriidae to other families of the Veneroida are currently unknown due to the lack of a convincing sister group. Traditionally, the Sphaeriidae were combined with the Corbiculidae into the superfamily Corbiculoidea based upon shared life-history characteristics such as occurrence in freshwater, parental care in the form of larval brooding, direct development of juveniles, and superficially similar conchological characteristics (Thiele, 1934; Keen and Casey, 1969; Boss, 1982; Morton, 1996; Mansur and Meier-Brook, 2000). However, cladistic studies of both morphological and molecular characteristics have demonstrated the independent origins of the Sphaeriidae and Corbiculidae from marine clams and that the hypothesized diagnostic characteristics of the Corbiculoidea are a suite of plesiomorphic and convergent traits (Park and Foighil, 2000; Giribet and Wheeler, 2002). While we have convincing evidence of which veneroid lineage the sphaeriids are not closely related to, the placement of the family remains to be resolved (Giribet and Wheeler, 2002; Taylor et al., 2007). Nevertheless, the Sphaeriidae can be diagnosed by their small size, occurrence in freshwater, the presence of an extended incurrent siphon, hermaphroditism, and parental care of direct-developing juveniles within the interlamellar spaces of the inner demibranchs (Lee, 2004) (Figure 19.5). Until the late 1990s, the relationships among Nearctic sphaeriid genera and species had been approached by classical, non-cladistic methods, and the taxonomy dated largely from the work of Herrington (1962), with a few subsequent modifications (Heard, 1965b; Clarke, 1973; Burch, 1975a; Turgeon et al., 1998; Korniushin et al., 2001). Four genera in three subfamilies have historically been recognized in North America: Sphaerium (eight Nearctic species) and Musculium (four species) comprise the Sphaeriinae, the Pisidiinae is equivalent to Pisidium (26 spp.), and the Euperinae has only a single Nearctic species of the genus Eupera. Modern cladistic analyses of morphological and molecular characteristics have supported different relationships among these species, based upon the character sets favored and hypothesized transformation polarities (Lee, 2004). Some strictly morphological analyses, emphasizing minute characteristics of the kidneys, support an evolutionary trend of miniaturization, leading to a monophyletic Pisidium sister to Musculium and a basal, paraphyletic Sphaerium (Mansur and Meier-Brook, 2000; Korniushin and Glaubrecht, 2002). Molecular studies of mitochondrial and nuclear DNA to date, however, have supported an evolutionary trend in

2

6

3

11 Pisidium

1

5

10

4 Eupera

9 FIGURE 19.5  Phylogeny of North American genera of the Sphaeriidae, showing evolutionary transformations of key characters. Synapomorphies are coded white, gray, and black to indicate the progression from plesiomorphic to derived transformations. Characters: (1) inhabits freshwater; (2) elongate incurrent siphon, well separated from excurrent siphon; (3) incurrent siphon fused in part of excurrent siphon; (4) incurrent siphon simplified; (5) hermaphroditism; (6) asynchronous brooding in interlamellar spaces; (7) brooding in brood sac; (8) synchronous brooding in multiple brood sacs; (9) adventitious maculations on the inner shell surface; (10) byssus in the adult; (11) smaller eggs with less yolk. (Adapted from Lee, 2004.)

brooding morphology, with a paraphyletic Pisidium (synchronous brooding) and a clade composed of asynchronous brooding species of Sphaerium and Musculium (Cooley and Foighil, 2000; Lee and Foighil, 2003). The morphological analysis of Lee (2004) reconciled these conflicting results somewhat, arguing that previous studies were biased by the improper use of corbiculid species to polarize character transformations and supporting a monophyletic Pisidium (diagnosed by characteristics of the shell and siphons) sister to a clade of asynchronous brooders. Unfortunately, combined analyses of sphaeriid interspecific relationships using both molecular and morphological characteristics are currently unavailable. Based upon the work done to date, the Sphaeriidae can conservatively be regarded as composed of two, monophyletic subfamilies: Euperinae and Sphaeriinae (Figure 19.5). The Euperinae is basically Gondwanan in distribution, with two genera, Eupera and Byssanodonta. Only a single species, Eupera cubensis, occurs in North America, extending from the Neotropics into the southeastern United States (Heard, 1965b). The subfamily Euperinae is diagnosed by the retention of unfused siphons and synchronous brooding within the interlamellar spaces (both plesiomorphies) and the presence of a byssal gland in the adult (Lee, 2004). The Sphaeriinae has reduced the size of its ova, providing less yolk to the developing embryo. The offspring are retained, however, within specialized brood-sacs and their

Chapter | 19  Class Bivalvia 

nourishment is provided by their mother or perhaps through sibling sacrifice (Lee, 2004). Within the Sphaeriinae, two major clades (corresponding to genera) are recognized, each representing different trends in morphological evolution. In Pisidium, the trend has been toward reduction, in both overall size and the elaboration of the incurrent siphon. Broods are synchronous (as with Eupera), but the offspring are supplied with a brood-sac within the interlamellar space of the inner demibranch (similar to Sphaerium) (Lee, 2004). In Sphaerium (+Musculium), brooding is asynchronous, with multiple broods in different stages of development each housed in their own brood-sac, and the incurrent and excurrent siphons are fused along at least part of their lengths. Given the morphological diversity among the sphaeriid species thus far analyzed, it would be surprising if the simplicity of our current understanding of the evolution of these lineages was not altered with additional taxon and character data.

Species Diversity and Distributions Forty valid species (including five introduced taxa) of sphaeriids are recognized to occur in North America, although some estimates suggest a higher diversity (Guralnick, 2005; Bogan, 2008), and this tally has remained relatively stable for the last 30 years. Burch (1975a) listed 38 species following Herrington (1962). One new species was subsequently described (Taylor, 1987; Turgeon et al., 1998), and another introduced species can be added to the list (Grigorovich et al., 2000; Korniushin, 2006). But this taxonomic stability should not necessarily be regarded as a consensus in support of the accuracy of the current species-level classification. It is merely the result of limited interest and effort by the freshwater malacological community to challenge it. While sphaerids are commonly used in comparative life-history studies, these studies are complicated because these species are minute, morphologically variable, and geographically ubiquitous. While the conservation crisis in freshwater mussels and the problems of exotic nuisance bivalves have generated considerable interest and mobilized research resources, the taxonomy of the Sphaeriidae has been placed on the backburner in North America. Where genetic and morphological diversity has been systematically sampled along a habitat gradient in a single genus, it has been demonstrated that the current estimates of sphaeriid species diversity and taxonomic concepts are inadequate (Guralnick, 2005). Shortcomings of the sphaeriid species-level taxonomy notwithstanding, some generalizations can be made about the distribution and diversity of these bivalves in North America. Based upon their distributions, sphaeriid clams are fully capable of passive overland dispersal. A single pea clam, transported from one water body to another by an aquatic bird, insect, or human conveyance, can found a new breeding population, given its potential for self-fertilization,

433

ovovivipary, and short generation times (Mackie, 1979a; Grigorovich et al., 2000). Thus, the presence of a disjunct spaheriid population in a particular stream, lake, or pond is merely indicative of currently suitable habitat conditions rather than historical connectivity. The overwhelming majority of sphaeriid species have ranges that occupy portions of formerly glaciated, boreal North America, including the Rocky Mountains (95%; 38 species), and more than half (55%; 22 species) are strictly boreal/mountain in distribution. Thus, fewer than half the species of the Nearctic sphaeriid assemblage have any portion of their known distributions south of the Pleistocene glacial maximum (Herrington, 1962; Clarke, 1973; Burch, 1975a). These freshwater bivalves provide an interesting contrast to freshwater mussels, which have no overland vagility and that reach their highest diversities in the southeastern portion of the continent.

Polyploidization The species of the Sphaeriinae are characterized by high and variable levels of polyploidy. Of the global sphaeriid species that have been karyotyped, only two have been found to be diploid: Sphaerium rhomboideum (2n = 44) and Sphaerium corneum (2n = 36 and 30) (Keyl, 1956; Petkeviciute et al., 2007). The remaining species, are all highly polyploid (up to 13n), with 2n × 1n chromosome counts as high as 247 (Lee, 1999; Petkeviciute et al., 2007). The fact that polyploidy is common across both Pisidium and Sphaerium (+Musculium) suggests that it is derived from the common ancestor of the modern species, and this is supported by the shared presence of divergent clades of alleles of single-copy nuclear genes across individuals of different species (Lee and Foighil, 2002). However, both of the diploid species of Sphaerium are nested within the otherwise polyploid Sphaeriinae (Cooley and Foighil, 2000; Lee and Foighil, 2003), which suggests either multiple origins of polyploidy or two, independent reversions to diploidy from polyploid ancestors. It may be that the latent effects of genome duplication in the Sphaeriidae have contributed to much of the taxonomic confusion that also characterizes the group (Petkeviciute et al., 2007). Among the Bivalvia, polyploidy is best understood among species of the marine genus Lasaea (O’Rourke et al., 2004) and the invasive freshwater genus Corbicula (Lee et al., 2005; Hedtke et al., 2008); and in those genera, polyploidy is associated with asexuality. Corbicula in North America is androgenetic (offspring are clones of their fathers) and triploid (Hedtke et al., 2008). In these freshwater lineages, the sperm are unreduced, bearing two flagella (Lee et al., 2005). However, in the case of the Sphaeriidae, it is unknown whether any species reproduce asexually.

  SECTION | IV  Phylum Mollusca

434

Diversity of Corbicula and Dreissena in North America A great deal of freshwater malacological energy has been expended to document the historical invasion, biology, and effects on native species of exotic bivalve genera in North America (Britton, 1979; Britton and Morton, 1982; Nalepa and Schloesser, 1993). The Asian clam (Corbicula spp.) was first noticed in western North America in the first quarter of the twentieth century, crossed the Continental Divide in the 1950s, and then spread widely over the eastern United States and Mexico (Counts, 1981; McMahon, 1999). The literature on Corbicula has until recently recognized a single, polymorphic species, Corbicula fluminea, with two morphs: Form A with white nacre, and Form B, with purple nacre (Britton and Morton, 1986). However, these two forms are morphologically and allozymically distinct (Hillis and Patton, 1982), and genetic characterization of these forms comparing mitochondrial DNA from North American Corbicula with Asian specimens in their native ranges have found that these two forms represent two different, clonal species: C. fluminea (Form B) and Corbicula leana (Form A) (Veinott and Cornett, 1996; Waller, 1998) (Table 19.3). Corbicula leana is the more widespread form, while C. fluminea is limited to the southwestern United States. Dreissena is represented in North America by two exotic, Eurasian species: the zebra mussel, Dreissena  polymorpha, and the quagga mussel, Dreissena bugensis. The historical range of D. polymorpha includes the Black and Caspian seas in Eastern Europe, and the construction of canals connecting the various river systems facilitated the bivalve’s expansion throughout the Ponto-Caspian region during the twentieth century (Bij de Vaate et al., 2002). In

TABLE 19.3  Exotic Species of Freshwater Bivalves in North America (Grigorovich et al., 2000) (Nomenclature for Corbicula species Follows Siripatttrawen et al. (2000) and Hedtke et al. (2008) Family Species Sphaeriidae

Pisidium amnicum (Müller, 1774) Pisidium henslowanum (Sheppard, 1825) Pisidium supinum Schmidt, 1850 Pisidium moitessierianum Paladilhe, 1866 Sphaerium corneum (Linnaeus, 1758)

Corbiculidae

Corbicula fluminea (Müller, 1774) [form B] Corbicula leana Prime, 1864 [form A]

Dreissenidae

1986, the zebra mussel was first documented in Lake St. Clair, from which it spread rapidly eastward to the Hudson River and southwest to the Mississippi Basin (Hebert et al., 1989). The presumed vector for the introduction of North American zebra mussels was anthropogenic transport in ships ballast (Carlton, 1993). The second species, D. bugensis, from the same region of eastern Europe, reached Lake Ontario via the same mechanism by 1991 (May and Marsden, 1992; Rosenberg and Ludyanskiy, 1994). Quagga mussels have spread somewhat but retain a more limited distribution than zebra mussels in only the lower Great Lakes and the St. Lawrence River (Mills et al., 1996). Based upon recent molecular analyses of Dreissena species throughout their native ranges, it has been suggested that D. bugensis from the Black Sea Basin may only be a local variety of Dreissena rostriformis of the Caspian Sea. However, these two forms are morphologically distinct and the populations examined bear diagnostic mitochondrial DNA haplotypes (Mills et al., 1996; Therriault et al., 2004). The exotic species of Dreissena and Corbicula are a nuisance to both humans and the native freshwater bivalves of North America.

GENERAL BIOLOGY External and Internal Anatomy The body plans of the various North American freshwater bivalves share a number of typical molluscan characteristics with the gastropods (see Chapter 18), in addition to characteristics diagnostic of bivalves generally. The typical freshwater bivalve body plan consists of a visceral mass and foot, surrounded by a pair of calcified valves (the shell) and the lobes of the mantle that secrete them. The shell provides attachment points for a number of prominent muscle systems, and the two valves are articulated by an elastic dorsal ligament. The visceral mass bears the labial palps on either side of the mouth and a pair of ctenidia extending into the posterior mantle cavity. The ctenidia serve not only as gills (i.e., surfaces for gas and ion exchange) but are also used in filter feeding and to brood developing larvae. Internally, the visceral mass houses the gut, gonads, heart. and nephridial system, including an open hemocoel that facilitates circulation and provides hydrostatic support. Many of these characteristics are of taxonomic significance, the native and exotic lineages of the Unionoida and Veneroida presenting various diagnostic modifications. Moreover, these freshwater bivalves often exhibit convergent morphological and physiological adaptations to life in freshwater not seen in their marine relatives.

Dreissena polymorpha (Pallas, 1771)

Shell and Mantle

Dreissena bugensis Andrusov, 1897

The external anatomy of a freshwater bivalve is dominated by the two valves of the shell. Adult size varies from small

Chapter | 19  Class Bivalvia 

members of the Sphaeriidae 20 cm long (Figure 19.1). Indeed, freshwater bivalve specimens are generally represented in malacology collections by their shells alone, and thus understanding the morphology of the shell (or conchology) is crucial for proper identification. In general, freshwater bivalve shells vary in lateral outline from elongate to ovate to rhomboid to trigonal. The nature of each valve of the bivalve shell is that of a spiral, homologous to that of a gastropod. A bivalve, though, expands the diameter of its aperture very quickly relative to its rotation about the coiling axis, nearly obliterating any sense of its coil (Savazzi, 1990). The only vestige of this growth pattern is the umbo or beak of each valve. The umbos represent the oldest parts of the shell (i.e., the apex of the spiral), and they are frequently eroded in older specimens of larger freshwater mussels. The prominence of the dorsal umbos is a focus for terminology regarding the symmetry of the shell and its polarity. Equivalve refers to the bilateral symmetry of the valves, while equilateral (perhaps somewhat confusingly) denotes a central position of the umbo along the anterior–posterior axis. The (shelled) gastropods provide a useful contrasting perspective on the typical characteristics of the bivalve shell and mantle. The typical snail has only its visceral mass and mantle cavity covered by the mantle and a single, coiled shell riding dorsally upon an exposed head and foot. Among bivalves, the mantle is expanded, with lateral lobes covering both sides of the mollusk. Each lobe of the mantle secretes a single, calcified valve, and the two valves are connected dorsally by an elastic ligament to form a bivalved shell. The mollusk is completely contained within its shell except for protrusible elements of the mantle edge and the foot while burrowing. As a result, bivalves have undergone decephalization, and sensory organs have been concentrated along the mantle edge.

435

outer layer, the periostracum. In general, the deposition of CaCO3 is facilitated by an organic matrix of proteins and polysaccharides (like chitin) secreted by the mantle (Marin et al., 2000; Marie et al., 2007). The term conchiolin refers to the insoluble residue remaining after demineralizing the shell (Crenshaw, 1990). New periostracum originates in the groove between the outer and middle folds of the mantle edge; while within the periostracal groove, the fibrous proteins of the periostracum undergo sclerotization or “tanning” to insolubulize and harden the proteins by cross-linking them. As the growing sheet of periostracum is extruded from the groove and around the outer fold of the mantle, an inner, untanned layer is added by the adjacent mantle epithelium. After the periostracum reflects around the edge of the mantle and joins the leading edge of the shell, the inner layer provides the site of nucleation for the inorganic CaCO3 crystals of the shell (Saleuddin and Petit, 1983; Checa, 2000). The secretion of the inorganic CaCO3 component of the shell begins with the precipitation of prismatic aragonite crystals perpendicular to the inner surface of the periostracum. The mantle epithelium lies in close proximity to edge of the shell and the new periostracum, creating a narrow extrapallial space (Checa, 2000). Ions, including calcium and bicarbonate, are transported from the hemolymph across the epithelium into the extrapallial space, supersaturating the aqueous medium; other ions include Na, K, and Mg, as well as various organic compounds. Calcium is obtained from dietary sources, but the primary source is the external medium (Wilbur and Saleuddin, 1983); among freshwater mussels, individual calcium content is correlated with environmental concentrations (Mackie and Flippance, 1983). Bicarbonate largely originates from CO2 derived from metabolic processes (Dettman et al., 1999), as catalyzed by carbonic anhydrase. CO2 + H2O ↔ H2CO3− ↔ HCO3− + H+

Shell Growth and Microstructure The shell of a bivalve is composed largely of calcium carbonate (CaCO3). As the mollusk grows, the shell increases in size with secretion of new shell material (both organic and inorganic) along the leading edge of the mantle. Valve thickness increases by the addition of either nacre (“mother of pearl”) or more complex cross-lamellar microstructures (Sphaeriidae, Corbiculidae, and Dreissenidae). The particular crystalline arrangement of the calcium carbonate precipitated is a function of the organic matrix secreted by the epithelium of the mantle. Among bivalves, the edge of the mantle consists of three folds: the inner, middle, and outer; the outer is adjacent to the shell. The mantle edge is the leading frontier of new shell formation (Figure 19.6). The addition of calcium carbonate to the shell begins with secretion of the proteinaceous

Ca2+ + HCO3− ↔ CaCO3 + H+ Parallel aragonite fibers grow from spherulites on the inner surface of the periostracum through precipitation in the extrapallial space. Nucleation crystals may be facilitated Prismatic layer

Shell margin

Nacreous layer Outer periostracum Inner periostracum

Ma

ntle

Growth lines

FIGURE 19.6  Diagram of the secretion of new shell along the leading edge of the mantle (after Checa & Rodríguez-Navarro, 2001).

  SECTION | IV  Phylum Mollusca

436

by Ca-binding residues in the organic matrix (Wilbur and Saleuddin, 1983; Marie et al., 2007). This chemical process is in equilibrium, and the survival of larger crystals is favored over smaller ones. As the aragonite prisms self-assemble in three dimensions, they abut each other and constrain further lateral growth (Checa and Rodríguez-Navarro, 2001). The thickness of the prismatic layer is limited by contact with the mantle epithelium (Checa, 2000). The result is the evolution of parallel aragonite crystals up to 200 μm in length aligned perpendicular to the inner surface of the periostracum (Checa and Rodríguez-Navarro, 2001). Away from the frontier of new shell formation, the mantle epithelium leaves a wider extrapallial space, and parallel sheets of nacre are deposited perpendicular to the prismatic crystals. Nacre is formed of flat round or polygonal aragonite crystals that self-organize in a discontinuous conchiolin matrix (Checa and Rodríguez-Navarro, 2001) (Figure 19.7). Accretion of nacre continues over the entire mantle surface to add thickness to the shell. In large freshwater bivalves, the total thickness of the shell can be measured in millimeters. The organic matrix of the bivalve shell serves to retard dissolution of the shell. Proteins involved in secretion of the valves are closely related to the mucins that cover the mollusk generally (Panha and Phansuwan, 1996; Marin et al., 2000). Freshwater bivalves generally have a well-developed periostracum to protect their shells from their hypo-osmotic medium. Internally, shell dissolution is controlled by biological mechanisms to maintain a saturated extrapallial fluid and by buffering the pH with bicarbonate (or perhaps ammonium) (Wilbur and Saleuddin, 1983). Notice in the chemical reactions above that production of bicarbonate and

calcium carbonate releases protons to the medium. This can lead to a drop in pH when the shell is closed to the external medium and a loss of shell mass. In temperate regions, shell growth stops below 10 °C (Dettman et al., 1999; Hua and Neves, 2007). This process results in a pattern of concentric annuli marked by a buildup of excess periostracum visible on the external surface of the valves as well as in internal sections. These annuli have been used to age freshwater bivalves, each ring indicating a single winter’s cessation of growth (Heard, 1977; Neves and Moyer, 1988; Ziuganov et al., 1994). However, various other disturbances can halt shell growth, leaving a visible “growth ring,” and so these methods have proven contentious. Nacre has gemological value. Historically, freshwater mussel shells were harvested for their natural pearls and to make buttons. Current commercial value is derived from the cultured marine and freshwater pearl industries (Anthony and Downing, 2001). Natural freshwater pearls form occasionally (one in thousands) (Anthony and Downing, 2001) when a foreign particle (e.g., a sand grain) intrudes between the mantle and shell. The irritant serves as the nucleus upon which nacre can accumulate. It has also been documented that natural pearl formation can be caused by the secretion of nacre around invading parasitic worms or larval trematodes (Hopkins, 1934). Cultured freshwater pearls can be made by surgically implanting bits of foreign (conspecific) mantle or other materials (Panha and Phansuwan, 1996). This practice is thousands of years old in the Far East (Hua and Gu, 2002), and has also been tested with some success on North American freshwater mussels (Hua and Neves, 2007). North American mussels are also currently harvested so that spherical bits of shell can be used to nucleate cultured marine pearls (Anthony and Downing, 2001). Nacre is of such high economic value that efforts are underway to culture mussel epithelial cells. Even in vitro, tissue cultures can create conditions for the precipitation of aragonite crystals (Barik et al., 2004).

Internal Shell Structure

10 FIGURE 19.7  Electron micrograph showing the interface between vertical prismatic aragonite and horizontal blocks of nacre (from Checa, 2000).

Besides the association of the leading edge of the mantle with the growing shell frontier, the mantle (or pallium) is bound to the inner shell surface through various muscle attachments, leaving a number of conspicuous scars. Running concentrically around the inner surface, parallel with the valve margin, is the pallial line, the site of attachment for the pallial muscles. Dorsally, anterior and posterior adductor muscles connect the two valves, providing the contractile force to close the valves. Closing the valves stretches the dorsal ligament, and relaxing the adductors allows the ligament to pull the valves back to gaping. The anterior and posterior pedal retractors (distally attached to the foot) are anchored on the shell. These muscles are often closely associated with the adductors, and indeed they often leave

Chapter | 19  Class Bivalvia 

437

inseparable scars. Immediately posterior to the anterior adductor (often in close association) is the pedal protractor muscle, and the pedal elevator muscles attach beneath the umbos; the scars of the latter muscles are most conspicuous on the shells of the Unionidae, leaving a series of minute

pits in the beak cavity. Whereas the adductor muscles connect the lateral halves of the bivalve, the others are laterally paired, and the whole complex has been referred to as the DVM (dorsoventral musculature) in the systematic literature (Figure 19.8).

Umbo Growth lines

Pustules

External left valve Umbo Pseudocardinal tooth

Ligament

Lateral tooth Posterior pedal retractor scar

Anterior adductor scar Protractor scar

Posterior adductor scar

Pallial line

Left mantle lobe (cut away) Anterior retractor muscle

Umbo cavity Internal right valve Posterior retractor muscle Supra-anal aperture Posterior adductor muscle

Anterior adductor muscle

Excrurrent aperture

Pedal protractor muscle Labial palp

Incurrent aperture Papillae

Foot

Outer demibranch Inner demibranch Right mantle lobe Internal soft anatomy FIGURE 19.8  Gross Anatomy of a freshwater mussel, Theliderma intermedia. Internal anatomy adapted from Ortmann (1912).

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438

Among the species of the Unionoidea, the inner surfaces of the valves are nacreous, generally pearly white, but occasionally other colors such as copper, salmon, blue, or violet are found depending upon the species. Besides the various muscle attachment scars, the morphology of the inner valves is dominated by the hinge teeth. The two valves articulate along their dorsal margins, immediately ventral to the external, opisthodetic ligament that extends posteriorly from the umbos along the dorsum of the shell. While certain lineages among the Nearctic assemblage have secondarily reduced or lost their hinge teeth (e.g., Anodontini), the typical unionoidean hinge bears schizodont dentition. The anterior teeth are positioned just below and in front of the umbos, often consisting of two peg-like pseudocardinal teeth in the left valve and a single, interlocking tooth in the right. The lateral teeth (or perhaps pseudolateral) run posteriorly from beneath the umbos, parallel to the ligament. Among Nearctic forms bearing hinge dentition, two elongate laterals are generally found in the right valve, with a single interlocking tooth in the left (Figure 19.8). However, large individuals may present additional teeth to these combinations. The function of the hinge dentition is to resist twisting of the valves along the plane of articulation. The hinge teeth of the Sphaeriidae are more typically heterodont, rather than schizodont. There are both anterior and posterior lateral teeth in front of and behind the umbo, respectively, and the peg-like cardinal teeth are position directly below the umbo. The number of cardinal teeth in the left valve has taxonomic value: one in Eupera (Euperinae)

and two in Pisidium and Sphaerium (both Sphaeriidae); all sphaeriids have one cardinal tooth in the right valve (Burch, 1975a; Lee, 2004). The ligament is external in species of the Sphaeriidae, but in most species it is obscured by a thin layer of calcification. The hinge structure of the introduced species of Corbicula is similar, but the larger shell can accommodate three cardinal teeth in each valve. The shell microstructure of the species of the Sphaeriidae and Corbiculidae is similar as well, appearing porcelaneous and lacking the iridescent, pearly luster seen in freshwater mussels. Rather than sheets of nacreous aragonite, the calcium carbonate is arranged in what is referred to as complex crosslamellar structure (Burch, 1975a; Boss, 1982). Both groups bear minute pores that are visible on the internal surface of the shell, and the pallial line bears a weak posterior pallial sinus to allow for retraction of the siphons.

External Shell Structure The characters of the bivalve shell are paramount for species-level identification. Besides the general characteristics described above, the major native lineages (Unionoidea and Sphaeriidae) differ markedly in their external shell anatomy, as do the invasive species. Freshwater mussels, in general, are relatively elongate-ovate in outline and inequilateral, with prosogyrous umbos positioned in the anterior section of the shell. However, there is great variation in form, and species may be round, trigonal, or rhomboid in outline (Figure 19.9). Significantly, shell morphology is known to

(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

(i)

FIGURE 19.9  Shell outlines of freshwater bivalves. Shell shape descriptions: (a) rhomboidal; (b) triangular or trigonal; (c) round; (d) quadrate; (e and f) oval or ovoid; and (g) elliptical. Posterior ridge morphology: (h) convex; (i) concave (from Burch, 1975b).

Chapter | 19  Class Bivalvia 

vary ecophenotypically, and this has caused considerable historical inflation of diversity estimates. However, environmental effects are often predictable in their influence, summarized as Ortmann’s Law of Stream Position (Ortmann, 1920). Specimens collected in large river environments will tend to be less elongate, more laterally obese, and have their umbos positioned more anteriorly. Conversely, headwaters forms will be more elongate and laterally compressed, with centrally placed umbos (Figure 19.10). The upstream form is better suited for burrowing, whereas downstream, the preferred shell shape is that of an anchor (Watters, 1994b). The ecophenotypes will grade into each other over the course of a large river system. Sexual dimorphism in freshwater mussel shells can also be striking, especially in genera like Lampsilis and Epioblasma. In the former, the shells of females are more obese and blunted posteriorly, and in the latter genus, there is often a conspicuous difference in shell outline between males in females (Figure 19.11). There can also be noticeable allometric variation as bivalves increase in size. For example, individuals become more elongate (i.e., decrease their height:length ratio) as they grow, allowing proportional increase in the surface area of the ctenidia as the volume of the mollusk’s body increases. The external surface of a freshwater mussel shell generally bears conspicuous growth rings marking periods of

439

decreased shell position. These rings are often taken to be annual. The disk of the shell may be sculptured with ridges or pustules of various sorts, but even species with unsculptured shells may have distinctive beak sculpture on the umbos retained from the juvenile shell (Figure 19.12). Shell sculpturing (of the umbos and the wider disk) is formed by the calcified layers of the shell. The periostracum itself may also bear color patterns, including rays or mottling (Figure 19.1). However, coloring is often highly variable and environmentally influenced, and it may be missing altogether from long-dead shells. The shells of sphaeriids are characterized by their small size, explaining their common labels as pill, pea, or fingernail clams. Specimens seldom exceed 25 mm in length; and in certain species, individuals 25 mm), the periostracum is yellow to chestnut brown, and the concentric external sculpture is more pronounced. Zebra and quagga mussels of the genus Dreissena are distinctive for their superficial similarity to mytilid mussels, with the umbo positioned on the highly reduced anterior end, the internal ligament and the external pigmentation bearing dark stripes (hence their common names). In zebra mussels, the ventral margin is flattened to facilitate epibenthic attachment to hard substrates with their byssus.

Mantle Cavity and Apertures The shell/mantle complex of a bivalve is large enough to cover more than just the visceral mass and foot; those structures are concentrated anteriorly. The posterior portion of the space nominally forms a chamber known as the mantle cavity. This space, though completely covered by the mantle, can be thought of as “outside” the animal. That is, the anus, nephridiopores, and gonopores empty into this space, their products carried away by the current of water generated by the ctenidia. This current not only flushes the mantle cavity of waste and gametes but also re-supplies oxygen and food. The surfaces of the ctenidia serve not only as gills for gas exchange but also to filter food particles from the incoming water. The internal spaces of the ctenidia serve also as brooding chambers for developing embryos and larvae. Given the biological importance of ctenidia to the bivalve lifestyle, it is not surprising that those organs are large and prominent features of bivalve anatomy (Figure 19.8). The ctenidia divide the mantle cavity into two chambers: the infrabranchial space and the suprabranchial space. Cilia on the outer surfaces of the ctenidia create a water current through the mantle cavity for feeding and gas exchange. Water is drawn from the surrounding medium through a posterioventral incurrent aperture or siphon to the infrabranchial space, passes through the sieve-like surfaces of the demibranchs, and travels dorsally through the water tubes to the suprabranchial space. The anus, gonopores, and nephridia empty into the suprabranchial space, and their products are flushed to the external medium via the excurrent aperture (or siphon). There has been some confusion among freshwater malacologists regarding the proper anatomical terms for the posterior apertures of the mantle cavity of freshwater mussels. The structures are frequently referred to as siphons, a term borrowed from marine and freshwater heterodont clams, like the sphaeriids or Corbicula. In those taxa, the tissue of the inner folds of the opposing mantle lobes have become fused and

  SECTION | IV  Phylum Mollusca

developed into retractile, tubular structures. In many marine taxa, these structures are developed even further (Yonge, 1982). Among those species possessing siphons (including unionoids of the families Iridinidae and Hyriidae) (Graf and Cummings, 2006), the diaphragm dividing the infrabranchial space from the suprabranchial space is formed by both the ctenidia and the mantle. It is a “complete” diaphragm. Among most species of North American freshwater mussels (family Unionidae), there is only minimal posterior fusion between the left and right lobes of the mantle. There are no protrusible siphons, and the diaphragm dividing the infrabranchial from the suprabranchial space is composed solely of the ctenidia (Figure 19.13). The ascending lamellae of the outer (lateral) demibranchs are fused to the adjacent mantle along their lengths, and the ascending lamellae of the inner (medial) demibranchs are fused to each other posterior to the visceral mass. The distal ends of the ascending lamellae of the inner demibranchs may be fused to the visceral mass before they join with each other, or the association may be limited to the proximal attachment point of the ctenidia. There is no mantle fusion between the incurrent and excurrent apertures, and the separation is achieved only by the association of the posterior mantle with the ctenidia, forming a slightly incomplete diaphragm (Davis, 1984; Graf and Cummings, 2006). There is no mantle fusion ventral to the incurrent aperture. Posterior to the excurrent aperture, however, unionids have a short stretch of mantle fusion formed by only the inner folds of the adjacent mantle edges, but this closure reopens to create a third posterior aperture. The function of this supra-anal aperture is unknown. The margins of the incurrent aperture (and sometimes the excurrent) are adorned

FIGURE 19.13  Posterior apertures of the Margaritiferidae (left) and Unionidae (right): EA, excurrent aperture; IA, incurrent aperture; SA, supra-anal aperture. The arrow indicates mantle fusion between the inner folds of adjacent mantle lobes.

Chapter | 19  Class Bivalvia 

with crenulations or papillae of variable complexity among both males and females (Ortmann, 1912). These structures serve a tactile sensory function as well as serving as loose sieves guarding the entrance to the infrabranchial chamber. In addition, females of certain genera have further elaborations of the posterior mantle associated with reproductive behaviors (Ortmann, 1911; Kraemer, 1970). Among the margaritiferid freshwater mussels, the diaphragm and posterior apertures are even less developed. There is no fusion of the posterior mantle, and the ctenidia, while fused to each other to their distal extremity, are free of the mantle at their posterior ends (Ortmann, 1912). This grossly incomplete diaphragm is augmented by pallial ridges that help provide somewhat better separation between the infrabranchial and suprabranchial chambers (Smith, 1980; Graf and Cummings, 2006). The Sphaeriidae, in contrast, present more extensive patterns of mantle fusion, typical of heterodont bivalves, and variations are diagnostic of the three sphaeriid genera (Lee, 2004; Korniushin, 2006). In Eupera, both the incurrent and excurrent apertures of the mantle are developed into tubular, contractile siphons that are free of each other along their lengths, and the ventral mantle remains fused for some distance approaching the foot. Thus, the three mantle openings are the excurrent siphon, the incurrent siphon, and the pedal slit, which allows for protrusion of the foot. In Sphaerium (+Musculium), the two siphons are further joined to each other for at least part of their length, though the pedal slit is generally not as well developed. In Pisidium, the incurrent siphon is reduced to a simple aperture that may or may not retain the ventral fusion that forms the pedal slit. The presence of siphons to manage the in- and out-flow of water through the mantle cavity increases the efficiency of the pumping mechanism (Morton, 1983).

Ctenidia All North American freshwater bivalves (both native and invasive) possess lamellibranch gills, modified from the primitive protobranch ctenidia to serve as filter-feeding organs. Protobranch ctenidia (as found in certain marine bivalves like Nucula, Yoldia, and Solemya) resemble the primitive molluscan ctenidia seen in the other classes and primarily function for gas exchange (Brusca and Brusca, 2003). Primitive, bipectinate ctenidia have a central axis, and from both sides of this axis extend a series of lateral filaments. The arrangement is analogous to that of a feather or a double-sided comb. Lamellibranch ctenidia can be derived from the protobranch type by lengthening and then reflecting the filaments. The ctenidial axis is aligned roughly parallel with that of the body, and so the filaments first descend ventrally from the axis and then reflect distally to ascend dorsally. Moreover, adjacent filaments are fused to each other into ascending and descending lamellae. In schematic cross-section, each lamellibranch ctenidium

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appears as a “W,” highly modified from the primitive, bipectinate condition. Each “V” of the “W” is referred to as a demibranch. The demibranchs are thus paired, but they are typically paired according to their positional homology rather than upon a shared ctenidial axis: the outer (lateral) demibranchs and the inner (medial) demibranchs. Rather than actually being V-shaped in cross-section, the ascending and descending lamellae of each demibranch are in close proximity to each other along their entire heights. Among the Nearctic unionids, the interlamellar spaces of the demibranchs are divided into a series of tubes by interlamellar septa; the spacing between the septa is generally 15–20 filaments, depending upon the taxon and sex in brooding species (Ortmann, 1911; Tankersley, 1996). These septa provide not only structural support but also the water tubes that they create (closed at the ventral end) are employed to brood developing larvae (see below). The individual filaments of each lamella are joined to each other by inter-filamental junctions. The tissue joining the filaments is perforated by water canals that, when open, allow communication between the infrabranchial chamber of the mantle cavity and the interlamellar space within each demibranch. Between adjacent filaments, lateral cilia beat to draw a water current from the infrabranchial space, through the ctenidia to the suprabranchial space. The flow rate of this current is determined by muscular control of the size of the ostia, the suprabranchial openings of the water canals. Elongate bands of muscles in the anterior–posterior direction alternate with rows of ostia along the length of the ctenidium. These muscles are attached to calcified, chitinous rods that run in the dorsal–ventral direction in each filament, and contraction reduces the spaces between adjacent filaments, analogous to an accordion. Thus, the ostia in the inter-filamental space are reduced to closed slits. When the muscles relax, the filaments move away from each other and the ostia open to allow water to flow from the infrabranchial space, through the water canals to the water-tubes, then to the suprabranchial space of the mantle cavity. A second set of muscles controls the internal ostia that connect the water canals to the watertubes. It has been shown experimentally that the application of serotonin (a neurotransmitter) can increases the flow of water through the mantle cavity by both relaxing the musculature associated with closure of the ostia (expanding the size of the ostia 2–3×) and by increasing the rate of beating of the lateral cilia (15–20+ Hz) (Gardiner et al., 1991).

Feeding Structures and Mechanisms The importance of the suspension-feeding lifestyle and its influence upon the evolution of the Bivalvia has historically attracted a good deal of study, as reviewed by Morton (1983). The mechanics and dynamics of bivalve suspension feeding is an area of active research, and the introduction of new methods such as in vivo endoscopic studies to

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the long-employed practice of in vitro studies of isolated ctenidia has opened up new questions about the process and provided novel insights. Different methods have returned different observations, and there is disagreement over distinguishing results from methodological artifacts. For explorations of this debate, see Jørgensen (1996) and Ward (1996). Our discussion of feeding incorporates the results of these endoscopic analyses of both freshwater bivalves specifically and bivalves generally.

Mechanics of Suspension Feeding Bivalve feeding is based upon the general model of active suspension feeding seen in other invertebrate groups (LaBarbera, 1984; Levinton et al., 1996). A water current through the mantle cavity is generated by ciliary action of the ctenidia. Environmental water is drawn in via the incurrent aperture to the infrabranchial chamber. The current passes through pores into the interlamellar spaces of the demibranchs, then to the suprabranchial space and out the excurrent aperture. Food (and other) particles suspended in the afferent through-current are captured by the ctenidia, partially sorted, and passed anteriorly for further sorting and ingestion. Much of this process is similar among the suspension-feeding bivalves, but in many freshwater lineages the mechanics are somewhat disrupted during the brooding period in females when some or all of their demibranchs may be gravid with developing larvae (Allen, 1914; Tankersley, 1996). The lamellae of the demibranchs are composed of individual filaments and each filament bears lateral cilia that protrude into the inter-filamental spaces. It is the beating of the lateral cilia that provides the motive force for the water current through the mantle cavity. Although the basal tissues of the adjacent filaments are fused together, these inter-filamental junctions form an incomplete barrier. Ostia between the filaments connect the infrabranchial chamber of the mantle cavity with suprabranchial chamber via the interlamellar spaces of the demibranchs. The diameters of the ostia determine the rate at which water can flow through the mantle cavity (Tankersley, 1996), and experiments with various neurotransmitters have suggested that the size of the ostia is under neuromuscular control. Relaxing the musculature of the lamella opens the gill ostia to their widest diameter. The through-current generated by the lateral cilia brings food particles into close proximity to the frontal surfaces of individual filaments. Filter feeding models derived from in vitro observations hypothesized that either or both mucus and the frontolateral cilia of the filaments act as a mechanical sieve to capture food particles from the incoming water (Morton, 1983). Frontolateral cilia are fused into compound cirri that flex into the inter-filamental space to form a grid with a mesh size smaller than 1 μm (Morton, 1983; Silverman et al., 1999). However, more recent models emphasize the

  SECTION | IV  Phylum Mollusca

utility of the highly viscous aqueous medium at the low Reynolds numbers of those minute scales (Jørgensen, 1996; Ward, 1996). Numbers of cilia per cirrus range from >10 in some species of unionids to >40 in Dreissena (Silverman et al., 1995, 1996b, 1997). Among freshwater mussels, captured particles are entrained in mucus and passed ventrally by the frontal cilia of both the ascending and descending lamellae (Tankersley, 1996). A ciliated food groove occurs at the ventral margin of the inner (medial) demibranch and transmits the mucus-bound particles anteriorly, toward the labial palps and mouth (Ortmann, 1912; Atkins, 1937). Particles reaching the ventral edge of the outer (lateral) demibranch are passed to the surface of the inner demibranch (Tankersley, 1996). The viscosity of the mucus keeps the captured particles from being resuspended in the surrounding water currents (Ward, 1996). From the ventral food grooves, the mucous-bound particles are transferred to the labial palps for sorting. The labial palps are large triangular or crescent-shaped flaps extending from either side of the mouth and in close association with the anterioventral margins of the inner (medial) demibranchs of the ctenidia. Each palp consists of two lamellae (an outer one continuous with the upper lip of the mouth and a lower lamella extending from the lower lip), the inner surfaces of each lamellae being grooved and bearing ciliated tracts (Morton, 1983). During feeding, the inner demibranch lies between the two lamellae of its associated labial palp, and the ciliated tracts of the palps receive the captured mucus-bound particles passed forward by the ventral food groove of the inner demibranch. The viscous complex of mucus and captured particles is broken down by the labial palps into a suspended slurry (Tankersley, 1996; Ward, 1996) and then passed to the mouth for ingestion.

Clearance Rates and Retention Efficiencies Freshwater bivalves differ in both the amount of water driven through the mantle cavity per unit time (clearance rate) and the effectiveness with which different particles are removed from that current (retention efficiency). Both of these properties have been found to vary across taxa, different environmental parameters, and under different experimental methods. Unfortunately, there are relatively few observations for freshwater bivalves and little standardization across labs, and as a result it is difficult to discuss absolute values. At this time, we can make only a few generalized statements about how much water is actually cleared by a freshwater bivalve and what particles are being removed through the action of the ctenidia. Two separate classes of approaches have been undertaken to determine the clearance rates of bivalves. Direct methods involve measuring the actually volume of water that leaves the excurrent aperture per unit time. These techniques generally involve some amount of apparatus

Chapter | 19  Class Bivalvia 

Where V is the volume of the suspension, n is the number of bivalves, t is time, Co is the initial concentration of the suspension, and Ct is the final concentration. Indirect approaches require a known (or assumed) retention frequency (often assumed to be100%) in order to relate clearance rate with filtration rate (Winter, 1978). Filtration, or clearance rate, is generally presented as the volume of water cleared per dry mass of bivalve (soft-parts) per unit time, although it is not uncommon to replace mass with shell length or merely “clam.” Many studies relate the filtration rate to size using the following relationship: b Filtration Rate = a(size) The slope, a, is taken to be the absolute filtration rate, and the exponent, b, reflects allometric variation with size. For b 1, the rate increases with size. For most bivalves, the filtration rate decreases with size, and typically b ranges are 0.6–0.9 (Winter, 1978; Kryger and Rissgard, 1988; Petkeviciute et al., 2007). Whereas the small size, hardiness, and ubiquity of sphaeriid and corbiculid species recommend them for laboratory studies, relatively few observations exist for unionoidean freshwater mussels. Zebra mussels, too, are well studied in this regard. More research into both how best to measure these rates as well as a broader taxon sampling are necessary. In general, a gram (ash-free dry mass) of native North American freshwater bivalve can clear 1–2 l of water per hour (Figure 19.14), but there is a wide variance associated with these as estimates. Compared with marine bivalves or freshwater corbiculids and dreissenids, these values are low by up to a factor of 3 (Kryger and Rissgard, 1988). Taxonomic differences aside, almost any change in water quality can affect clearance rates (Morton, 1983). Environmental variables like low winter temperatures or summer anoxia can both lower clearance rates, and as the concentration of suspended particles increases, the clearance rate also decreases (Paterson, 1984; Burky et al., 1985; Lauritsen, 1986; Vanderploeg et al., 1995). While the clearance rate steadily deceases with increasing particle concentration, the amount of material captured from the throughcurrent increases, but only to a point (Figure 19.15).

Dreissena FR = 6.82•DM0.88

Anodonta FR = 1.10•DM0.78

0.1

0.01

Sphaerium FR = 2.14•DM0.92

0.001

0.01

0.1

1

10

Dry mass (soft parts) (g) FIGURE 19.14  Filtration rate is related to dry mass by the allometric function, FR = aDMb, where a is the absolute filtration rate and b is the effect of body size. Dreissena has a filter absolute filtration rate per body mass that is greater than either Sphaerium or Anodonta. However, these rates are all generally below those obtained for similarly-sized marine bivalves. (Data adapted from Kryger & Riisgård, 1988).

50 12 40

30

8

20 4

Pseudofeces produced 10

0

Particles cleared for 1g clam (µg/h)

Filtration Rate = V/nt · ln Co /Ct

1 Filtration rate (l/h)



10

Filtration rate for 1mg clam (ml/h)

impinging upon the bivalve (i.e., tubing inserted into the open aperture) (Allen, 1914; De Bruin and Davids, 1970; Kryger and Rissgard, 1988; Vanderploeg et al., 1995), and as such they can be inappropriate for smaller bivalves like sphaeriids. An indirect estimate of any suspension feeder’s clearance rate can be calculated by examining the rate of removal of particles from an experimental suspension using the following equation (Morton, 1983; Kryger and Rissgard, 1988):

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0 0

20

40

60

Particle concentration (mg/l)

FIGURE 19.15 In Sphaerium striatinum, filtration rate (closed circles) decreases steadily as the concentration of particles increases, while the amount of particles cleared per unit time (open circles) increases to a point and then decreases. The peak in the amount of material cleared coincides with the concentration of particles above which pseudofeces are produced. These data are from Hornbach et al., 1984a, who used 2.02 μm latex beads as particles. Vertical bars represent 95% confidence intervals.

Eventually, the ctenidia can no longer process the incoming material or are held up by rate-limiting processes like particle sorting by the labial palps or ingestion (Winter, 1978; Levinton et al., 1996), and excess particles accumulating in the infrabranchial chamber are rejected as pseudofeces (Hornbach et al., 1984a; Burky et al., 1985; Way, 1989).

  SECTION | IV  Phylum Mollusca

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Under experimental conditions, optimal particle concentrations approximate the natural amount of suspended solids in the bivalve’s environment (Hornbach et al., 1984a), and it is easy to understand how anthropogenic increases in suspended silt could affect filtering abilities in freshwater bivalves (Aldridge et al., 1987). The size of the suspended particles also influences filtration rates, as materials of different effective diameters have different retention efficiencies. Suspension-feeding bivalves are generally regarded to retain particles larger than 4 μm with maximum efficiency, but retention rates drop off quickly in freshwater mussels for particles smaller than 1–2 μm (Figure 19.16) (Winter, 1978; Jørgensen et al., 1984; Vanderploeg et al., 1995). Unionoidean freshwater mussels are capable of retaining (Silverman et al., 1997) and ingesting (Nichols and Garling, 2000) bacteriumsized particles at the smaller end of the spectrum, but not as effectively as either Corbicula or Dreissena do (Figure 19.17) (Silverman et al., 1995, 1996a). The minimum size of the particles that can be retained by a bivalve is correlated with the number of laterofrontal cilia (responsible for particle capture; see above) per compound cirrus. In the study by Silverman et al. (1997), among the ctenidia of unionids found in ponds in lakes, cilia/cirrus counts were as low as 11–16, and those bivalves cleared 4 μm, but efficiencies drop off below 2 μm. However, Lampsilis, when tested with a smaller size range of natural particles was able to retain sizes  protease > lipase > glucosidase (Christian et al., 2004). Similar enzymes are present in juvenile mussels, although at lower activities (Areekijseree et al., 2006). Freshwater mussels have also been reported to have cellulase activity (Haag et al., 1993; Areekijseree et al., 2002), as have other freshwater bivalves (Farris and Van Hassel, 2007). It is unclear whether cellulase is a component of the crystalline style (Alyakrinskaya, 2001) or produced by some other organ. The intestine (midgut and hindgut) descends from the posterioventral portion of the stomach, and coils in the visceral mass, surrounded by the gonads (Figure 19.19). The proximal portion is in close association with the style sac and the crystalline style, which protrudes into the stomach and may be more than half the length of the mussel (Nelson, 1918; Alyakrinskaya, 2001). The distal portion (the rectum) of the intestine pierces the pericardium and the ventricle of the heart before terminating in a simple anus on the posterior face of the posterior adductor muscle. The hind portion of the intestine of freshwater mussels is unique for the possession of a large typhlosole (Jegla and Greenberg, 1968). The course of the intestine is lined with cilia, mucus is secreted, and waste is consolidated into fecal pellets (Purchon, 1968). The intestine (and stomach) may be the site of extracellular digestion and absorption. The rupture of fragmentation spherules may release digestive enzymes, and they have also been implicated as having a role in the dissolution of the crystalline style (Morton, 1983). The association of the intestine with the pericardium indicates an osmoregulatory role. The role of roving amebocytes in freshwater bivalve digestion and assimilation is poorly understood. These motile cells also have a function analogous to the leukocytes of vertebrates in combating infections, as well as a clot-forming role in the hemolymph. Amebocytes have also been implicated as having an osmoregulatory function. It has been hypothesized that one of the roles of amebocytes in bivalves is to aid in digestion and assimilation by taking up food particles in the gut (stomach and intestine) through phagocytosis and then actively migrating via the circulatory system to bring catabolic products directly to the tissues

(Narain, 1973). Amoebocytes in the gut may provide the only absorptive surfaces for assimilation outside of the digestive diverticula, but this has been disputed (Purchon, 1968; Morton, 1983), and more work is clearly necessary. Because most digestion and absorption largely takes place within blind tubules shunted from the gut rather than in the lumen of the stomach or midgut, digestive activity is cyclical to accommodate the two-way traffic of food and waste to and from the stomach (Morton, 1973). That is, while bivalves have a complete, one-way gut, the dynamics of the digestive process has analogies to the incomplete guts of taxa like cnidarians or platyhelminths. During periods of starvation, the crystalline style is dismantled (Allen, 1914), and the digestive cells of the digestive diverticula are in a holding phase (Morton, 1983). During active feeding, the style matrix is reassembled by the style sac within hours (Nelson, 1925). As ingestion exceeds the capacity of the diverticula, even otherwise suitable food materials are rejected to the intestine. During field examinations of gut contents, as much as 50% of the material in the intestine is undigested and undamaged, including algae and invertebrates such as microcrustaceans and rotifers (Allen, 1914, 1921; Nichols and Garling, 2000). Digested material is readily identified by the granular appearance of the fragmentation spherules. It is difficult to determine assimilation efficiencies of bivalves given the sorting of filtered and ingested particles. At higher food concentrations, much material that is cleared is never ingested, and much that is ingested may never be digested (Morton, 1983). Given the disparity between what is ingested and what is assimilated, it has been necessary to rely upon analyses comparing stable isotope ratios (δ13C, δ15N) of freshwater bivalve tissues with potential food sources (Christian et al., 2004; Nichols and Garling, 2000; Raikow and Hamilton, 2001). These studies upon freshwater mussels indicate that the carbon and nitrogen they assimilate is derived from the small (5 l) of water, the rinse water should be filtered through a fine mesh net or sieve. This procedure should be repeated four to nine times, depending on the amount and composition of the material. It reduces the amount of mineral and coarse

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organic material. The method is less appropriate to obtain quantitative results. Some groups can best be observed alive. After collection, the annelids should be placed in a small container (cooled, except for collections from warm water habitats) for return to the laboratory. Additionally, live samples often yield whole individuals or zooid chains of naidids that often are lost when fixed. Observation of live individuals can provide a clear view of internal structures and can be a learning experience for students.

Preparation for Identification Samples can be fixed, preferably immediately after collection, particularly in warm weather conditions or when samples contain larger amounts of degradable material. The samples and local conditions decide the usage of preservatives or fixatives. Small annelids can be preserved with ethanol. Larger annelids are fixed using a 10% endsolution of formalin, preferably buffered with something like borax or calcium or magnesium carbonate. Bouin’s fluid is less widely used than formalin, but it is one of the best fixatives to use if staining and sectioning of annelid tissue is planned. Except for the larger worms (Lumbriculidae, Sparganophilidae, and Lumbricidae), the aquatic oligochaetes should be identified using whole slide mounts and a compound microscope. The mounting media can vary, but could include Canada balsam or Amman’s lactophenol.

INTRODUCTION TO THE INLAND WATER POLYCHAETA General Introduction Inland water (fresh and brackish water) polychaetes are, like all polychaetes, easy to recognize by the paddle-like appendages that protrude from the lateral sides of each body segment. These appendages are covered by numerous tiny hairs that contribute to movement and, in some groups, create water currents for feeding. These numerous hairs give the polychaetes their name (poly = many; chaeta = hairs). A literature review for this chapter of Polychaeta (Annelida) including Aphanoneura (the oligochaete-like Aeolosomatidae and Potamodrilidae) living in freshwater yielded records of 168 species, 70 genera, and 24 families representing all of the major polychaete clades, but less than 2% of all species (Glasby and Timm, 2008). Worldwide, the number of real freshwater polychaetes is very low. It is estimated that fewer than 50 species seem to be restricted to freshwater habitats (Pennak, 1989). Of the 85 families of polychaetes known to occur worldwide, freshwater polychaetes can be found in the families of Nereididae, Sabellidae, Spionidae, and Histriobdellidae, as well in the former

SECTION | IV  Phylum Annelida

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FIGURE 20.10  Distribution of species of Namanereis (Nereididae) and Stratiodrilus (Histriobdellidae) across Gondwana before its breakup in the mid to late Jurassic. Reproduced with kind permission of Chris Glasby.

oligochaete family, Aeolosomatidae. Most of these belong to the Nereididae, a primarily marine family. The Nereidae are known to be tolerant for high salinity fluctuations. The family Nereidae (Phyllodocida) includes 55 freshwater species, over half of which are in the subfamily Namanereinae. Fourteen other families are represented by a single species and genus. The freshwater polychaete fauna is more diversified in warmer regions than in temperate zones. More than half the nonmarine nereidids occur in the tropical and subtropical regions of the western Pacific. Higher numbers of nonmarine species are also found in Asia and South America. One remarkable feature of freshwater polychaetes is their small size. This could also be reason that they are sometimes overlooked. Polychaetes are an ancient group dating back to the Middle Cambrian (540 million years ago), and possibly earlier. However, because they do not fossilise well— usually only the jaws, chaetae, tubes and burrows leave imprints—there are large gaps in the fossil record. Still, at least three distinct processes appear to account for the colonization of inland waters: (1) invasion of a clade prior to the break-up of Gondwana, as in Aphanoneura, Namanereis, Stratiodrilus, and Caobangia; (2) relatively recent stranding of individual species (relicts); and (3) the temporary visitation of euryhaline species (Glasby and Timm, 2008). In the latter category are several marine species

that have extended their distribution range into brackish and freshwater but are unable to reproduce there. Again, the largest numbers stem from the families Nereidae and Sabellidae (Figure 20.10). The prevailing theory is that most freshwater or brackish polychaete habitats were once geographically connected or are presently connected to the ocean (Hartman, 1959). In addition, almost all freshwater polychaetes have been collected within a range of 32 km of the ocean (Pennak, 1989), supporting a theory of recent evolution into freshwater. More than half of all species and genera inhabit lakes and rivers, followed by lagoons, estuaries (which have a high proportion of euryhaline species), and inland seas. Less common, atypical polychaete habitats include subterranean waters, the hyporheic zone of rivers, and plant container habitats (phytotelmata) (Glasby and Timm, 2008). In general, the non-Clitellata can be grouped into the Polychaeta and Aphanoneura, as described next.

General Systematics Polychaeta Three major clades of the Polychaeta (sand worms, tube worms, or clam worms) are recognized: Scolecida, Aciculata (among them Amphinomida, Eunicida, and

Chapter | 20  Introduction to Annelida and the Class Polychaeta

Phyllodocida), and Canalipalpata (with Terebellida, Spionida, Pogonophora, and Sabellida). Polychaetes, like other members of the Annelida, have two pre-segmental regions (the prostomium and peristomium), a segmented trunk (metastomium), and a post-segmental pygidium. The nuchal organs, a pair of chemosensory structures on the posterolateral margin of the prostomium, are apparently the only synapomorphy of the Polychaeta that distinguishes them from other Annelida (Rouse and Fauchald, 1995). Nuchal organs vary from welldeveloped posteriorly projecting loops to inconspicuous pits or grooves. The largest nuchal organs are found in the Amphinomidae and the Euphrosinidae, where the ciliated folds on the caruncle represent the nuchal organs; in many other families nuchal organs are not easily seen. Polychaetes exhibit a wide range of feeding strategies, ranging from those that are carnivorous predators, deposit feeders, suspension feeders, herbivores, and opportunistic feeders. A few species are parasitic, and some are commensal. Still, most inland water species are deposit feeders or scavangers (Nereididae) or suspension feeders or grazers (Sabellidae). Polychaetes are tolerant of reduced oxygen levels or of toxic pollutants, like heavy metals. Therefore, they can be useful pollution indicators, as are the oligochaetes.

Aphanoneura The Aphanoneura, which are head-crawling or suctionfeeding worms, are regarded as aberrant canalipalpatans (a taxon of Polychaeta). They include the Aeolosomatidae, formerly aligned with the Oligochaeta, and the Potamodrilidae (Fauchald and Rouse, 1997). Aeolosomatids are minute oligochaete-like ciliated worms with no clear affinity to other polychaetes; they mostly inhabit freshwater. Like the similar potamodrilids, they lack head appendages and parapodial lobes, but may be distinguished from them by the presence of colored epidermal glands all over the body surface. The Aeolosomatidae is the only Aphanoneura family that has epidermal glands present as discrete colored spots.

Phylogenetic Relationships For a long time the most commonly used division of the polychaetes was the separation of the ‘Errantia’ from the ‘Sedentaria.’ This division was based on a system of convenience without any evolutionary argumentation (Fauchald and Rouse, 1997). Recent cladistic analyses of Annelida and other groups have provided a new division of polychaetes into two main clades (Rouse and Fauchald, 1997): Scolecida and Palpata. The small group of Scolecida, with less than 1000 named species, composes burrowing worms with bodies reminiscent of earthworms. The majority of polychaetes belong to the Palpata. The Palpata are divided

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FIGURE 20.11  Aeolosoma hemprichi. http://animalkingdom.su/books/ item/f00/s00/z0000048/st052.shtml.

into Aciculata and Canalipalpata. Half of the polychaete species belong to the Aciculata—the group that largely comprises the former taxonomic group Errantia. The group includes major subgroups, such as Phyllodocida and Eunicida, which tend to be mobile forms with well-developed eyes and parapodia with internal supporting chaetae, or aciculae, for rapid locomotion. The Canalipalpata, a group with more than 5000 named species, is distinguished by having long grooved palp structures that are used for feeding. Canalipalpata is divided into Sabellida, Spionida, and Terebellida. Members of most of these groups live in tubes and use their palps to feed in various ways. The families Aeolosomatidae (Figure 20.11) and Potamodrilidae were placed in the subclass Aphanoneura by Timm (1981). Brinkhurst (1982) then elevated this subclass to a class, maintaining parity with the class Oligochaeta, while noting that the Oligochaeta and Hirudinea both are often considered to be subclasses of the Clitellata (Brinkhurst and Wetzel, 1984). Singer (1978) addressed the biology, ecology, physiology, and systematics of the aeolosomatids and discussed their phylogenetic affinities with other annelids. More recently, polychaete systematists have included the aeolosomatids in the Polychaeta (Fauchald and Rouse, 1997; Rouse and Fauchald, 1997). Suffice it to say that the taxonomy of the species of Aeolosoma currently needs attention. Members of the family Aeolosomatidae are widely distributed, although the difficulties encountered in the study of this family have restricted the documentation of species. Potamodrilidae is represented by a single monotypic genus, Potamodrilus.

Distribution and Diversity Of the subfamily Namanereidinae (Nereididae), most of the 37 species (including three species groups) in three genera (Lycastoides, Namanereis and Namalycastis) are reported

SECTION | IV  Phylum Annelida

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from habitats with reduced salinity, freshwater, or even semi-terrestrial environments (Glasby, 1999). These include Namanereis catarractarum from water-filled tree holes in Papua New Guinea and moist leaf axils of Pandanus trees in Fiji. Another polychaete, Namanereis cavernicola, occurs in a freshwater pool in a Mexican cave 1650 m above sea level and 176 km from the Pacific coast. Also, Namanereis araps Glasby, 1997 has been collected from groundwater in the Sultanate of Oman. The Aeolosomatidae have a worldwide distribution and mainly are freshwater inhabitants, but can also be found in shallow brackish waters (Jørgensen and Jensen, 1978). They have been collected from ponds, lakes, and rivers from decaying plant materials (Niederlehner et al., 1984).

GENERAL BIOLOGY OF POLYCHAETA External Anatomy The head of a polychaete is composed of the prostomium, peristomium, and pharynx. The prostomium is the most anterior pre-segmental part of the body to the mouth, which may or may not be retractile and often bears antennae, eyes, tentacles, and palps. The antennae have sensory organs, and the palps may be sensory or can be used as feeding appendages. Some species have one or two pairs of eyes on the prostomium. The peristomium, which is the first distinct postprostomal region or segment around the mouth, includes the tentacular cirrus (especially in ciliary feeders, which may bear a crown of tentacles that can be opened like a fan or withdrawn into the tube) and the proboscis. The latter may bear chaetae, pals, and sometimes chitnous jaws. The pharynx, which is mostly eversible, is the anterior part of digestive tract for feeding and is sometimes used for burrowing. The body or trunk is segmented. Each segment generally has its own local nerve center called a ganglion and a pair of nephridia for excretion (Figure 20.12). Each segment also has a pair of flatlike projections, the parapodia, on both sides of each segment. The parapodia are used for locomotion and gas exchange. Parapodia are unjointed segmental extensions

of the body wall found in many polychaetes, though many lack these features. They are also absent in Clitellata and Echiura. Parapodia are equipped with musculature derived mainly from the circular muscle layer and usually carry chaetae. Parapodia vary in structure but basically can be considered to consist of two elements: a dorsal notopodium and a ventral neuropodium. In addition to bundles of chaetae, notopodia and neuropodia can also have a variety of cirri and gills. The movement of the parapodia is controlled by oblique muscles that run from the midventral line to the parapodia in each segment. They are most elaborate in actively crawling or swimming species, where they form large fleshy lobes that act as paddles. Parapodia of burrowing or tubicolous polychaetes can be slightly raised ridges carrying hooked chaetae called uncini. Each parapod bears chitinous bristle-like chaetae that are used in locomotion, feeding, and tube-building. The chaetae can vary strongly and can be simple, compound, capillary, limbate, bifurcate, trifurcate, pinnate, harpoon, pectinate, or spatulate. The tail (posterior body section) is truncated or tapered and contains a dorsal or terminal anus. Cirri may also be present. Some polychaetes have gills. For example, the marine genus Amphitrite has three pairs of branched gills and long extensible tentacles, and the marine genus Arenicola, the burrowing lugworm, has paired gills on certain segments. The Aeolosomatidae have a large, lobe-like prostomium that is almost completely ciliated ventrally and has lateral ciliated grooves that have been interpreted as nuchal organs. The muscular pharynx lies in the peristomium. The constrictions along the trunk give the appearance of external segmentation, but they actually represent a chain of zooids produced by fragmentation (paratomy). Parapodia are absent, and chaetae are usually present as four bundles per segment and are rarely absent. They comprise usually only capillaries, or sometimes sigmoid hooks or, rarely, only hooks. The body is often brightly colored due to the epidermal gland cells. Each gland cell consists of a vacuole, filled with red, green, bluegreen, yellow, or sometimes colorless liquid; the function of the gland cells is unknown (Bunke, 1988). The Potamodrilidae are similar to the Aeolosomatidae. In the Potamodrilidae, the prostomium fused to the peristomium, flattened, is frontally blunt, and is the peristomial part limited to lips. Antennae and palps are absent, and nuchal organs present as paired sensory papillae. All segments are similar and segmented, with parapodial structures that lack tentacular, dorsal, and ventral cirri. Also absent are gills, epidermal papillae, pygidial cirri, lateral organs, and dorsal cirrus organs. All are hair-shaped chaetae.

Internal Anatomy

FIGURE 20.12  Anatomy of a polychaete segment.

Besides being segmented, the body wall of polychaetes is characterized by the presence of both circular and longitudinal muscle fibers surrounded by a moist, cellular cuticle

Chapter | 20  Introduction to Annelida and the Class Polychaeta

that is secreted by an epidermal epithelium. Polychaetes have, like all annelids, a brain or cerebral ganglion that originates and usually resides in the head. The brain varies in structure, with more mobile active forms having the most complex brains, and sessile or burrowing forms having simple brains with little differentiation. The brain is connected to the ventral nerve cord by the circumpharyngeal connectives, which run down each side of the pharynx. The ventral nerve cord runs the length of the body and is usually composed of a pair of cords that are bound together. The nerve cord varies in thickness and dilates into a ganglion in each segment; from there, pairs of segmental nerves pass out to the body wall, muscles, and gut. The polychaetes’ digestive system consists of a foregut, a midgut, and a hindgut (Figure 20.13). The foregut includes a stomodeum, pharynx, and anterior esophagus. It is lined with cuticle, and the jaws, where present, are constructed of cuticular protein. The more anterior portions of the midgut secrete digestive enzymes, but absorption takes place toward the posterior end. A short hindgut connects the midgut to the exterior via the anus, which is on the pygidium. There are six major kinds of sensory structures found in polychaetes. These include palps, antennae, eyes, statocysts, nuchal organs, and lateral organs. Palps and antennae are located on the head of many polychaetes. In some groups both are sensory, while in others the palps are used for feeding. Satocysts are balance sensory receptors. Nuchal organs are ciliated, paired, chemosensory structures, innervated from the posterior part of the brain. They are present in nearly all polychaetes, and Rouse and Fauchald (1997) suggested that they may represent an apomorphy for Polychaeta. This has been challenged by other authors, who suggest that nuchal organs may be an apomorphy for Annelida as a whole and have been lost in Clitellata/Oligochaeta

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(Purschke et al., 2000). Polychaetes also have various epidermal sensory cells that may be responsive to light or touch (such as lateral organs). Most polychaetes have two fluid systems: the coelom and the circulatory system; and both (if present) are involved in excreting waste products. To achieve this excretion, there must be ducts to the exterior, and these are generally referred to as nephridia. Ducts, known as gonoducts or coelomoducts, are also required for the transfer of gametes that develop in the coelom to the outside of the body on maturity. The two different kinds of ducts are often simply referred to as segmental organs, since determining what kind of duct is present is problematic. The circulatory system is closed in most polychaetes and Echiura, as it is in many clitellates. In some polychaete groups, however, the closed circulatory system is limited to major blood vessels and the distal capillary vessels are missing. A circulatory system is absent in many small polychaetes. The polychaete excretory organs consist of protonephridia and mixed proto- and metanephridia in some taxa. Most polychaetes, however, have metanephridia, with one pair per segment; their interior end (nephrostome) opens into a coelomic compartment. The coelomic fluid passes into the nephrostome, and selective resorption occurs along the nephridial duct. The families of Aeolosomatidae and Potamodrilidae have a more simple internal anatomy. As in oligochaetes, their building plan consists of a double-tube structure. The outer longitudinal muscles are grouped in bundles. The stomodaeum, at the anterior end of the middle gut, has a structure resembling a ventral buccal organ. For example in Aeolosoma hemprichi, it comprises the widened part of the gut in segments two to five. For the remaining species, the gut consists of a straight tube. A

FIGURE 20.13  Dorsal view of a polychaete internal building plan. Hartman, 1959; Kasprzak, 1984; Rouse and Pleijel, 2001.

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diaphragm or gular membrane is absent. The number of nephridia is limited to the first segments, specifically segments one and two in Potamodrilus and segments two to five in A. hemprichi. A connection between the nephridia and coelomoducts is absent. The circulatory system is closed, and a heart-like organ is absent. In Potamodrilus, the female gonads are situated in segment five and male gonads in segment six.

Physiology The guts of most polychaetes are adapted anatomically and physiologically to obtain the maximum extraction of the very small proportion of organic matter from the mineral material which is often their diet. This process of food intake was well described for the marine worm Arenicola marina by Kermace (1955) as follows. The diet contains a large inorganic indigestible portion and as is to be expected. In as far as the digestive processes of other soil eating animals are known, e.g., oligochaetes, the polychaetes share the characteristic that a large amount of digestion occurs intracellularly in wandering amoebocytes, thus leaving the gut lumen free to deal with the vast quantities of indigestible siliceous material. Therefore, the gut wall consists of four layers; (1) the outer peritoneal covering, (2) a longitudinal muscle layer, (3) a circular muscle layer and (4) an inner epithelium. There is a system of ciliary tracts, which in the glandular region of the esophagus is associated with the longitudinal ridges of the lining epithelium; and which consists of the ciliated ventral groove and lateral tracts arising from it in the post-cardiac region of the stomach, the intestine and the rectum. There are isolated patches of cilia on the walls of the cardiac region of the stomach. Movement of the cilia in these tracts is believed to keep the contents of the stomach mixed and in suspension, thus ensuring that the food particles are brought into contact with the entire epithelium. The cilia also assist the passage of food through the gut, although this is mainly brought about by the rhythmic contractions of the body wall impinging upon the latter. The esophagus is capable of a considerable amount of muscular activity and initiates some of the gut movements. Parts of the blood system of the animal lie in close proximity to the gut and as the blood system plays a part in digestion. The food passes through the trunk gut, i.e., the proboscis, esophagus, stomach and intestine, to the rectum in about 14 min in an actively feeding worm. Food is taken into the esophagus from the base of the funnel of the burrow by the constantly everting and inverting proboscis, without any preliminary sorting. It is temporarily stored in the muscular region of the esophagus, but eventually it passes to the stomach through the glandular region of the former, where mucus and a little enzyme material is poured onto it by the unicellular glands in the walls. At the junction of the esophagus and the stomach there are the openings of the esophageal pouches, the secretion from which contains mucus and Qestive enzymes. While in the stomach food particles in the sand are digested both intra- and extra-cellularly, the epithelial cells engang

the food particles thus leaving the lumen of the gut filled with the large residual mass of indigestible, mostly siliceous, material. These food particles are taken from the epithelial cells by amoebocytes, which while digesting them, wander to all parts of the body in the blood and coelomic fluid. The indigestible remains of these particles are deposited by the amoebocytes in the epidermis, peritoneal cells, intravasal tissue, coelom and in the lumen of the gut…. pH values are given for different parts of the gut, only that of the stomach (pH 5.4–6.0) differing appreciably from neutrality. Changes in pH affect the viscosity of the mucus, which is least viscous in the stomach where the pH is lowest. This aids the cilia and the movements of the gut in keeping the contents of this region in suspension. Elsewhere in the gut, the mucus is more viscous and acts as a lubricant, protecting the gut wall from the abrasive action of the sand grains. An actively feeding worm defaecates at intervals of approximately 45 min.

Diffusion of gas through the body wall accounts for part of the gas exchange. Gills are common among polychaetes and mostly associated with the parapodia. Most polychaetes have a well-developed blood–vascular circulation system. The circulation is relatively simple, as blood flows in the anterior direction through the dorsal vessel, which goes over ventral vessels on the anterior side of the body. In most segments, lateral vessels transport part of the blood toward the ventral vessel. The ventral vessel carries the blood posteriorly. Smaller vessels carry blood to the parapodia, nephridia, body wall, and gut. Polychaete blood is mostly colorless or has respiratory pigments such as hemoglobin, chlorocruorin, or hemerythrin. Coelomic fluid that contains wastes passes through a funnel into the tubule of the metanephridia. This funnel penetrates the membrane between two segments, and then the remaining part of the metanephridia opens at the base of the neuropodium on the ventral side of the segment anterior of the funnel. The osmoregulatory ability of polychaetes is great, especially in the Nereididae. In these tolerant species, the blood and tissue fluids are isotonic with the surrounding environment. In freshwater, however, these species remain at an internal salt concentration higher than the surrounding freshwaters. Under these conditions, the nephridia produce a urine that is hyposmotic to the coelomic fluid. The chloragogen tissue, coelomocytes, and gut wall play a supporting role in excretion.

Reproduction Although many marine polychaetes reproduce asexually, all freshwater species reproduce sexually. Most polychaetes have separate sexes, but some crawling and swimming species are hermaphroditic. A free-swimming larval stage is typical of marine forms, but is absent in all freshwater species. The freshwater polychaetes have no permanent sex organs, and they usually have separate sexes. The reproductive system is simple with gonads that appear as temporary swellings of the peritoneum and shed their gametes into the

Chapter | 20  Introduction to Annelida and the Class Polychaeta

coelom. The gametes are then carried to the outside through gonoducts, through the metanephridia, or by rupture of the body wall. Fertilization is external. Generally, aeolosomatids reproduce by asexual transversal fission (paratomy). Reproduction begins when the worm reaches a determined number of metameres (depending on the species). This stage gives rise to the clonal production of a chain of filial zooids that detach themselves from the parental zooid in a few days (fragmentation). The doubling time of Aeolosomatidae is short and usually lies between one and four days (Inamori et al., 1990). This mode of reproduction can lead rapidly to high population densities. Sexual reproduction has been reported in only a few species (Christensen, 1984). Only one species, Aeolosoma singulare, is known to reproduce exclusively by sexual reproduction (Marotta et al., 2003).

GENERAL ECOLOGY AND BEHAVIOR OF POLYCHAETA Polychaetes are often divided into two groups based on their activity: sedentary (benthic) polychaetes and errant (freemoving) polychaetes. Sedentary polychaetes spend much or all of their lifetime in tubes or permanent burrows and many of them, especially those that live in tubes, have special adaptations for respiration and feeding. Errant polychaetes include different locomotion groups: free-swimming pelagic forms, active burrowers, crawlers, and tube worms that temporarily leave their tubes for feeding or breeding. Most of these, like species of the genus Nereis, are predatory and equipped with jaws or teeth. Predaceous polychaetes have an eversible, muscular pharynx armed with jaws that can be thrust out with surprising speed to capture prey.

Distribution and Habitat Selection The regions supporting the highest diversity of freshwater polychaetes are, in order, the Palaearctic, Neotropical, Oriental, Nearctic, Australasian, and Afrotropical regions. They rarely occur on oceanic islands (only Nereididae). They are essentially absent from the Antarctic region, except for Namanereis quadraticeps, a circum-subantarctic species found in the freshwater seep zones of the upper shores. Three further species are found in the Arctic. The species Marenzelleria wireni, Manayunkia speciosa, and Chone sp. have occasionally been found in tundra lakes. Note that all three arctic freshwater polychaetes are also widespread south of the Arctic. M. wireni is a tolerant species that occurs in a wide range of conditions. It is found in both fresh and marine waters, although it usually inhabits the less brackish part of estuaries of rivers. It also tolerates a wide range of oxygen levels and survives temporarily without oxygen. Chone sp. and M. speciosa are more restricted to freshwater habitats.

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The species of the family Namanereidae are highly variable in their habitat (Williams, 2004). The occurrence in brackish water areas is common, although the range of salinity tolerated is high. The species generally prefer shallow waters and soft, calcareous fine-grained substrate, often rich in decaying plant detritus. However, as predatory worms, this detritus is not a food source. Aeolosoma maritimum is the only marine representative of the family Aeolosomatidae. They occur in the mesopsammal of a sandy beach at the Gulf of Tunis. The salinity of the habitat ranged from 29.2‰ to 34.6‰. Potamodrlilus fluviatilis is a typical rheophilic inhabitant of the mesopsammal. Its epidermal glands enable this species to stick itself to the substrate in high current velocities.

Physiological Constraints The optimal temperature for growth and reproduction of A. hemprichi and Aeolosoma variegatum is 20–30 °C, and at 10 °C reproduction stops (Kamemoto and Goodnight, 1956). Various authors recorded mass presences of Aeolosomatidae, especially A. hemprichi, in wastewater treatment plants for sludge reduction (Liang et al., 2006).

Feeding Behavior Polychaetes have diverse feeding habits, depending on their lifestyle. Most errant polychaetes are typically predators or scavengers, and most sedentary polychaetes feed on suspended particles, or are deposit feeders that consume sediment particles. Some species dip into the substrate with feeding tentacles, scoop up some muck, and draw it into their mouths to digest any edible particles. This occurs predominately in the tube-living polychaetes, which are primarily filter feeders. By moving their appendages within these tubes, they create a water current that draws smaller particles to their mouths. The predaceous polychaetes are often active hunters, swimming through the water and grasping prey, such as small worms or algae, with an extendible, jaw-like structure. Little is known about the specific feeding habits of namanereidid polychaetes, but it is usually assumed they most are predatory or omnivorous. Most predaceous species have a muscular, eversible pharynx equipped with a pair of toothed, opposing jaws. The jaws of some phyllodocid polychaetes are used to capture and hold prey or to tear off pieces of algae and decaying matter, while those of some Eunicida are used for scraping food particles from hard ­substrates. Aeolosomatidae feed on plant tissue, detritus, protozoa, bacteria, and algae. The particles are swept toward the mouth by ciliary currents set up at the anterior end and by a type of suction feeding. Aeolosoma suck fine material by placing their mouth on the substrate.

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When lifting the center of the prostomium by muscular contraction, a partial vacuum is created that dislodges particles from the substrate. Potamodrilidae worms mainly feed on dead organic plant material. Their eversible pharynx bulb also provides a comparable kind of suction apparatus.

Locomotion Polychaetes’ locomotion traits, which involve extension/ retraction of parapodia and chaetae, are burrowing, slow creeping (six to eight segments in one ‘wavelength’), fast crawling (14 segments), and swimming. Some Aeolosoma species swim with the aid of beating the cilia at the head end of the body. Aeolosoma forms a resistant mucous cyst stage that can easily be transported from one place to another. Potamodrilidae do not use body muscles to move; the only active movement is provided by moving cilia on the head.

Predators and Parasites The freshwater polychaete Manayunkia speciosa serves as the obligate intermediate host for the myxosporean parasites Ceratomyxa shasta and Parvicapsula minibicornis, which adversely affect the survival and freshwater production of juvenile salmon in the Klamath River and elsewhere in the Pacific Northwest of the USA.

Invasive Tendencies An interesting example of an invasive freshwater polychaete is Hypania invalida. This species is native in the Ponto-Caspian range, in both fresh and brackish waters. It tolerates a wide range of salinity (0–12 S), temperature (2–25 °C), and depth (from the shore line up to more than 400 m depth). H. invalida penetrates the European waters from three directions, using northern (the Volga catchment), central (Dnepr and Vistula catchments), and southern (the Danube catchment) corridors (Bij de Vaate et al., 2002).

COLLECTING POLYCHAETES AND SPECIMEN PREPARATION Freshwater polychaetes are collected in much the same manner as other annelids. As with most invertebrates, searching individually for the small freshwater polychaetes in the field is not productive. If representative samples of a polychaete fauna are required, it is necessary to collect a sample of the substrate (e.g., sand, mud, algae). A grab, shovel, or corer (a cylinder that is pushed into the sediment) are mostly used to collect sediment. To facilitate sorting, Rose Bengal, which stains all living material

pink, can be added to the sample. The sediment is carefully washed through a sieve, and the specimens are picked off the sieve with a forceps or the material is poured into a white tray. Next, the worms and other organisms are separated from the sediment. At this point, it is advisable to use a dissecting microscope to ensure all specimens in the sample are found. Polychaetes should initially be preserved in 5–10% formalin (2–4% formaldehyde) or in other histological fixatives, such as Bouins fluid. Afterward, specimens should be transferred to 70% ethanol, although material preserved in high-strength (94%) alcohol is normally adequate, especially if bulk samples are elutriated to remove sediment before preservation. As color, presence of eyes, and other features are best recognized in live specimens, specimens should be observed live whenever possible. Free-living or swimming species can be collected using a hand net or by collecting macrophytes and algae, which can be placed immediately into a bag and sealed. These samples are sorted in the laboratory in the same way the sediments were sorted. Quantitative studies of polychaete communities often involve the use of quadrats, corers, or comparable equipment that collects a specific volume of sediment. The sediment with the often-hidden fauna should then be placed in a container and returned to the laboratory for examination. All samples should be labeled using waterproof paper and ink. When the entire sample is fixed for later sorting, such labels should be placed within the sample. Aeolosomatids can be very small and pelagic and can be collected using plankton nets. When studying swamps, aeolosomatids can be collected by squeezing moss tufts. When studying live specimens, one can use narcotics to relax the animal. Magnesium chloride is an effective narcotic for invertebrates at a concentration of about 7% in freshwater. Also carbonized drinking water is a cheap and effective option. Another advantage of the latter is a quick recovery from the anesthetic.

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Brinkhurst, R.O., 1982a. British and Other Marine and Estuarine Oligochaetes. Freshwater Biological Association. 127 pp. Brinkhurst, R.O., 1982b. Evolution in the Annelida. Can. J. Zool. 60, 1043–1059. Brinkhurst, R.O., Gelder, S.R., 1989. Did the lumbriculids provide the ancestors of the branchiobdellidans, acanthobdellidans and leeches? Hydrobiologia 180, 7–15. Brinkhurst, R.O., Wetzel, M.J., 1984. Aquatic Oligochaeta of the world, supplement: a catalogue of new freshwater species, descriptions, and revisions. Suppl. Can. Tech. Rep. Hydrogr. Ocean. Sci. 44, 1–101. Brusca, R.C., Brusca, G.J., 2003. Invertebrates, second ed. Sinauer Associates, Sunderland, Massachusetts. Bunke, D., 1988. Aeolosomatidae and potamodrilidae. In: Higgins, R.P., Thiel, H. (Eds.), Introduction to the Study of Meiofauna. Smithsonian Institution Press, Washington, DC, pp. 345–348. Chekanovskaya, O.V., 1962. Aquatic Oligochaeta of the USSR. Keys Fauna USSR 78, 1–513. Christensen, B., 1984. Asexual propagation and reproductive strategies in aquatic Oligochaeta. Hydrobiologia 115, 91–95. Chua, K.E., Brinkhurst, R.O., 1973. Evidence of interspecific interactions in the respiration of tubificid oligochaetes. J. Fish. Res. Board Can. 30, 617–622. Eibye-Jacobsen, D., Kristensen, R.M., 1994. A new genus and species of Dorvilleidae (Annelida, Polychaeta) from Bermuda, with a phylogenetic analysis of Dorvilleidae, Iphitimidae and Dinophilidae. Zool. Scr. 23, 107–131. Erséus, C., 1987. Phylogenetic analysis of the aquatic Oligochaeta under the principle of parsimony. Hydrobiologia 155, 75–89. Erséus, C., 2005. Phylogeny of oligochaeteous Clitellata. Hydrobiologia 535, 357–372. Fauchald, K., Rouse, G., 1997. Polychaete systematics: past and present. Zool. Scr. 26, 71–138. Gelder, S.R., 1996. A review of the taxonomic nomenclature and a checklist of the species of the Branchiobdellae (Annelida: Clitellata). Proceedings of the Biological Society of Washington, 109, 653–663. Gelder, S.R. Monograph of the Branchiobdellida (Annelida: Clitellata) or Crayfish Worms, in preparation. Glasby, C.J., 1999. The Namanereidinae (Polychaeta: Nereididae) part 1 taxonomy and phylogeny, part 2 cladistic biogeography. Rec. Aust. Mus. Suppl. 25, 1–144. Glasby, C.J., Timm, T., 2008. Global diversity of polychaetes (Polychaeta; Annelida) in freshwater. Hydrobiologia 595, 107–115. Hartman, O., 1959. Capitellidae and Nereididae (marine annelids) from the Gulf side of Florida with a review of freshwater Nereidae. Bull. Mar. Sci. Gulf Caribb. 9, 153–168. Inamori, Y., Kuniyasu, Y., Hayashi, N., Ohtake, H., Sudo, R., 1990. Monoxenic and mixed cultures of the small metazoa Philodina erythrophthalma and Aeolosoma hemprichi isolated from a waste-water treatment process. Appl. Microbiol. Biotechnol. 34, 404–407. Jamieson, B.J.M., 1974. The zoogeography and evolution of Tasmanian Oligochaeta. In: Williams, W.D. (Ed.), Biogeography and Ecology in Tasmania. Junk, The Hague, pp. 195–228. Jamieson, B.G.M., 1988. On the phylogeny and higher classification of the Oligochaeta. Cladistics 4, 367–410. Jørgensen, K.F., Jensen, K., 1978. Mass occurrence of the oligochaete Aeolosoma hemprichi Ehrenberg in activated sludge. Biokon Reports 7, 9–11.

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Kamemoto, F.I., Goodnight, C.J., 1956. The effects of various concentrations of ions on the asexual reproduction of the oligochaete Aeolosoma hemprichi. Trans. Am. Microsc. Soc. Invertebr. Biol. 75, 219–228. Kasprzak, K., 1984. The previous and contemporary conceptions on phylogeny and systematic classifications of Oligochaeta (Annelida). Ann. Zool. Warszawa 38, 205–223. Kaygorodova, I.A., Dzyuba, E.V., Pronin, N.M., 2012. Leech-like parasites (Clitellata, acanthobdellida) infecting native and endemic eastern Siberian Salmon fishes. Scientific World J. 2012, 1–8. Kermace, D.M., 1955. The anatomy and physiology of the gut of the polychaete Arenicola marina (L.). Proc. Zool. Soc. Lond. 125, 347–381. Liang, P., Huang, X., Qian, Y., 2006. Excess sludge reduction in activated sludge process through predation of Aeolosoma hemprichi. Biochem. Eng. J. 28, 117–122. Marotta, R., Ferraguti, M., Martin, P., 2003. Spermiogenesis and seminal receptacles in Aeolosoma singolare (Annelida, Polychaeta, Aeolosomatidae). Italian J. Zool. 70, 123–132. Martin, P., Martinez-Ansemil, E., Pinder, A., Timm, T., Wetzel, M.J., 2008. Global diversity of oligochaetous clitellates (“Oligochaeta”; Clitellata) in freshwater. Hydrobiologia 595, 117–127. Martin, P., 2001. On the origin of the Hirudinea and the demise of the Oligochaeta. Proceedings of the Royal Society of London, Series B 268, 1089–1098. Niederlehner, B.R., Buikema Jr., A.L., Pittinger, C.A., Cairns Jr., J., 1984. Effects of cadmium on the population growth of a benthic invertebrate Aeolosoma headleyi (Oligochaeta). Environ. Toxicol. Chem. 3, 255–262. Omodeo, P., 2009. Evolution and biogeography of megadriles (Annelida, Clitellata). Italian J. Zool. 67, 179–201. Parish, J., 1981. Reproductive ecology of Naididae (Oligochaeta). Hydrobiologia 83, 115–123. Pennak, R.W., 1989. Fresh-water Invertebrates of the United States, third ed. John Wiley and Sons, Inc., New York, NY. 628 pp. Piguet, E., 1906. Observations sur les Naididtes. Rev. suisse Zool. 14, 185–315. Poddubnaya, L.L., 1984. Parthenogenesis in tubificidae. Hydrobiologia 115, 97–99. Purschke, G., Hessling, R., Westheide, W., 2000. The phylogenetic position of the Clitellata and the Echiura – on the problematic assessment of absent characters. J. Zool. Syst. Evol. Res. 38, 165–173. Rouse, G.W., Fauchald, K., 1995. The articulation of annelids. Zool. Scr. 24, 269–301. Rouse, G.W., Pleijel, F., 2001. Polychaetes. Oxford University Press, London. Rouse, G.W., Fauchald, K., 1997. Cladistics and polychaetes. Zool. Scr. 26, 139–204. Rousset, V., Pleijel, F., Rouse, G.W., Erseus, C., Siddall, M.E., 2007. A molecular phylogeny of annelids. Cladistics 23, 41–63. Sawyer, R.T., Dierst-Davies, K., 1974. Observations on the physiology and phylogeny of colour change in marine and freshwater leeches (Annelida: Hirudinea). Hydrobiologia 44, 215–236. Siddall, M.E., Apakupakul, K., Burreson, E.M., Coates, K.A., Erséus, C., Gelder, S.R., Källerjö, M., Trapido-Rosenthal, H., 2001. Validating Livanow: molecular data agree that leeches, branchiobdellidans and Acanthobdella peledina form a monophyletic group of oligochaetes. Mol. Phylogenet. Evol. 21, 346–351. Singer, R., 1978. Suction feeding in Aeolosoma (Annelida). Trans. Am. Microsc. Soc. 97, 105–111.

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Sket, B., Trontelj, P., 2007. Global diversity of leeches (Hirudinea) in freshwater. In: Balian, E.V., Lévêque, C., Segers, H., Martens, K. (Eds.), Freshwater Animal Diversity Assessment. Hydrobiologia. http://dx.doi.org/10.1007/s10750-007-9010-8. Sket, B., Trontelj, P., 2008. Global diversity of leeches (Hirudinea) in freshwater. Hydrobiologia 595, 129–137. Struck, T.H., Schult, N., Kusen, T., Hickman, E., Bleidorn, C., McHugh, D., Halanych, K.M., 2007. Annelid phylogeny and the status of Sipuncula and Echiura. BMC Evol. Biol. 57, 1–11. Timm, T., 1981. On the origin and evolution of aquatic Oligochaeta. Eesti NSV Eaduste Akad. Toim. Biol. Seer. 30, 174–181.

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Wang, H.Z., Liang, Y., 1997. Analyses of distribution and faunal relationship of inland-water microdrile Oligochaeta (Annelida) in the world and China. Acta Hydrobiol. Sin. 21, 91–102. Westheide, W., 1997. The direction of evolution within the Polychaeta. J. Nat. Hist. 31, 1–15. Wetzel, M.J., Kathman, R.J., 2002. North American Freshwater Oligochaeta. An introduction. 16 pp. Williams, D.D., 2004. Review of the polychaete genus Namanereis (Nereididae) in the Caribbean region, with a record of N. hummelincki from deep freshwater wells in Barbados. Caribb. J. Sci. 40, 401–408.

Chapter 21

Clitellata: Oligochaeta Tarmo Timm Institute of Agricultural and Environmental Sciences, Centre for Limnology, Estonian University of Life Sciences, Rannu, Tartumaa, Estonia

Patrick J. Martin O.D. Taxonomy and Phylogeny, Royal Belgian Institute of Natural Sciences, Brussels, Belgium

Chapter Outline Introduction529 General Systematics 529 Phylogenetic Relationships 531 Distribution and Diversity 533 General Biology 535 External Anatomy 535 Internal Anatomy 537 Physiology540 Reproduction and Life History 541 General Ecology and Behavior 542 Macrohabitat Distribution and Microhabitat Selection 542

INTRODUCTION General Systematics Oligochaeta is a paraphyletic stem group (traditionally treated as a subclass) of the class Clitellata that is separate from the advanced, sucker-bearing clitellates: Hirudinea, Acanthobdellida, and Branchiobdellida. They are sometimes called “earthworms” after their bigger and betterknown terrestrial representatives (the so-called megadriles). However, many of them are small worms, from about 1 mm to a few centimeters in length (microdriles), living in aquatic and terrestrial habitats (Figure 21.1). The nickname “sludge worms” has been coined for some species of the family Tubificidae abundant in polluted watercourses. Up to the 1960s, the freshwater polychaetes Aphanoneura (families Aeolosomatidae and Potamodrilidae) were also classified in the Oligochaeta (Brinkhurst and Jamieson, 1971). The Oligochaeta includes about 5000 valid nominal species distributed between 30 or so families. More than 1100 species of 13 families live in freshwater (Martin et al., 2008), while the remainder inhabits terrestrial soil or marine

Physiological Constraints 542 Feeding Behavior 543 Predators and Parasites 543 Other Ecological Aspects of Oligochaeta 544 Collecting, Culturing, and Specimen Preparation 544 Collecting544 Rearing545 Preparation for Identification 545 References548

sediments. Classifying the families into orders is difficult and problematic, especially in the case of aquatic forms. To date, there is no consensus on the classification that should be adopted in this group. One of the main factors contributing to this situation is the advent of molecular methods that enabled DNA sequencing of gene fragments. By giving an access to new characters, DNA sequences have led to a fundamental reappraisal of current classifications (see Adoutte et al., 2000). In 2006, Jamieson and Ferraguti (2006) proposed a revision of the phylogenetic classification of oligochaetes, which integrates the most recent molecular data available at that moment. However, such a classification remains uncompleted and unstable, due to constant progress in the molecular field. Molecular studies have also confirmed the long-suspected paraphyly of the Oligochaeta if the group does not include branchiobdellids and leeches, so that Clitellata has become synonymous with “Oligochaeta” (Martin, 2001; Siddall et al., 2001; Martin et al., 2008). Another complication in the nomenclature of aquatic oligochaetes arose after the family Naididae was included into Tubificidae as a subfamily (Erséus et al., 2002), while

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00021-8 Copyright © 2015 Elsevier Inc. All rights reserved.

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FIGURE 21.1  Examples of freshwater Oligochaeta (live pictures): (a) Stylaria lacustris (Naididae; note a budding zone in the midbody); (b) Ilyodrilus templetoni (Tubificidae; note the clitellum); and (c) Lumbriculus variegatus (Lumbriculidae). Scale: a–b = 1 mm, c = 5 mm. Photo by Patrick Martin. Copyright © 2015 Dr Patrick Martin. Published by Elsevier Inc. All rights reserved.

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the name Naididae appeared to be a senior synonym of the Tubificidae (Erséus et al., 2008). Translating phylogenetic analyses into biological nomenclature can make the latter impracticable for many purposes (Timm, 2012). For this reason, the senior author is in favor of maintaining paraphyletic taxa in this chapter, for the sake of a workable classification, namely the traditional family Tubificidae in its former sense (as the stem group for the Naididae, Pristinidae, and Opistocystidae), and the whole subclass Oligochaeta (hence clitellates exclusive of branchiobdellids and leeches).

Phylogenetic Relationships Oligochaeta have been treated as successors to marine polychaetes that adapted to life in freshwater and soil by developing a clitellum-secreted cocoon for protection their eggs and embryos, adopting hermaphroditism, concentrating the reproductive organs into a few segments (thereby testes always located anterior to the ovaries), and losing the parapodia, cirri, and nuchal organs. However, all these characters exist separately in some marine polychaetes and may have served as preadaptations. On the other hand, many extant polychaetes have invaded freshwater or soil without similar anatomical changes. Two more characters—the dorsal position of pharyngeal pad and the shift of brain backward from prostomium—are shared by oligochaetes and the terrestrial polychaete (?) Hrabeiella, once assumed to be their sister taxon (Purschke, 2003), a phylogenetic relationship not supported by later molecular studies (Jördens et al., 2004; Rousset et al., 2007). A recent phylogenomic study by Struck et al. (2011) has recovered the Clitellata within polychaetes with strong support, and thus rendered the Polychaeta paraphyletic. The name Pleistoannelida was coined for a major clade of annelids by Struck (2011), splitting into the sister taxa Errantia and Sedentaria sensu Struck et al. (2011). The Clitellata are now nested within the Sedentaria, a clade characterized by adaptations to a sedentary lifestyle by reductions of head and body appendages and the position of the chaetae being in closer proximity to the body wall than Errantia. Interestingly, the genetically most ancient extant oligochaete families, Capilloventridae and Randiellidae, live in marine habitats (Erséus, 2005), as do the vast majority of polychaetes. Brinkhurst (1984, 1994) developed an elegant theory of oligochaete phylogeny based on their reproductive system. According to this, the original number of gonads was four pairs (two pairs of both testes and ovaries) in segments X–XIII, as preserved in the typical representatives of the family Haplotaxidae. The haplotaxids reveal also the simplest male ducts. This theory treats the paired, simplepointed chaetae typical of the Haplotaxidae and many other, mostly burrowing oligochaetes as an ancestral character. However, it ignores the resemblance of the multiple hair and bifid chaetae typical of many aquatic oligochaetes to those

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present in polychaetes. Also, a certainly nonadaptive, rudimentary upper tooth occurs in the paired sigmoid chaetae of the Lumbriculidae and even some Haplotaxidae. As a rule, the Haplotaxidae are considered to contain the most primitive oligochaetes known (Kathman and Brinkhurst, 1998). However, this interpretation is now questionable. This family, with “simple” anatomy, was recently shown to have no basal position within the clitellates, and was confirmed as the closest taxon to the Crassiclitellata, although as a paraphyletic assemblage (Martínez-Ansemil et al., 2012). More generally speaking, the displacement of taxa long thought to represent successive grades of complexity at the base of a tree to much higher positions inside the tree is probably one of the salient results obtained with genetic data. This can be illustrated by the Metazoan example in Adoutte et al. (2000). Since then, this observation was repeatedly made in other groups; the Haplotaxidae is just another example of the obligation to reconsider our older evolutionary reasoning. According to alternative opinion shared, for example by Timm (1981), the “first oligochaete” likely had hair chaetae and sigmoid, bifid crotchets like many extant polychaetes and aquatic oligochaetes. This is supported by the independent, secondary loss of hair chaetae and the gradual loss of the upper tooth of sigmoid chaetae in many lower taxa. The weak points of this theory are the reduced number of gonads (mostly to two pairs) in the extant aquatic oligochaetes other than haplotaxids, as well as the trend toward paired sigmoid chaetae, which have independently arisen in several clades. The latter characteristic is established in the Lumbriculidae, Haplotaxidae, “true earthworms,” and some other taxa, and appears to be developing in the Enchytraeidae. In all probability, the “ancestral oligochaete” had a variable number of chaetae of different types and a set of four pairs of gonads. All modern combinations of chaetae and gonads can be derived from this condition by reduction of separate elements. A review of different theories, and a modern scheme of the phylogeny of Clitellata that takes into account gene sequences, was developed by Erséus (2005). Jamieson and Ferraguti (2006) generated their own classification, as a result of their synthesis of current morphological and molecular knowledge of oligochaetes. In a conservative approach, the classification below developed is an attempt to reflect the most consensual point of view in the community of oligochaete specialists, although it is clearly provisional, pending future revision. The main large groupings of Oligochaeta can be defined as follows (see also Figure 21.2): 1. T  he order Tubificida contains the sole clade with hair chaetae (although the latter often being lost). The number of gonads is usually two pairs, either in X–XI or XI–XII, while spermathecae are aligned to gonadal segments. The typical representatives are the burrowing Phreodrilidae and Tubificidae, and derived from the latter, phytophilous Naididae, Pristinidae, and Opistocystidae, with

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FIGURE 21.2  Location of reproductive organs in different families of Oligochaeta. Redrawn after Martin and Aït Boughrous, 2012.

Chapter | 21  Clitellata: Oligochaeta

2.

3.

4.

5.

prevailing vegetative reproduction and shifted position of the reproductive system. The hair chaetae, if present, are usually delimited to the dorsal bundles, while two small families, Capilloventridae and Parvidrilidae, contain also ventral hair chaetae. Male ducts are plesioporous and mostly equipped with atria, but are without them in the most primitive group, Capilloventridae, where also a “sterile” segment between the testicular and ovarial segment can occur. Spermathecae also lie further forward in Capilloventridae. The Tubificida are aquatic, either freshwater or marine. The order Enchytraeida contains the mostly terres trial (with few marine and freshwater genera) family Enchytraeidae, with simple-pointed chaetae in variable number per bundle, two pairs of gonads in XI–XII, plesioporous male ducts without true atria (but often with a complex penial apparatus), and spermathecae located far forward from the gonadal segments. Two small, aquatic families may be phylogenetically related to the root of Enchytraeidae: the freshwater Propappidae, differing from the enchytraeids by bifid chaetae and a sterile segment XII separating the testicular and ovarial segment; and the marine Randiellidae, with one to two pairs of testes and with very simple male ducts. The order Haplotaxida contains the freshwater family Haplotaxidae, typically with four pairs of gonads (but partially reduced in some species), very simple male ducts, spermathecae in variable number and position anteriad to the gonadal segments, and paired or single sigmoid, usually simple-pointed chaetae per bundle. The monospecific family Tiguassuidae (with single pairs of testes and ovaries) can also be related to Haplotaxida. The Haplotaxidae were treated as paraphyletic to most other oligochaete families by Brinkhurst (1984) but only to Crassiclitellata by Timm (1981). Jamieson et al. (1987), Erséus et al. (2010), and Martínez-Ansemil et al. (2012) also placed the Haplotaxidae closer to Crassiclitellata (and Lumbriculida) than to Tubificida. The order Lumbriculida appears paraphyletic, if  you exclude the advanced, carnivorous, leech-like Branchiobdellida, Acanthobdellida, and Hirudinea (which are treated in separate chapters of this book). They can be characterized by prosoporous or semiprosoporous male ducts. The freshwater family Lumbriculidae reveals strictly paired chaetae, either simple-pointed or with a rudimentary upper tooth, while the male ducts are equipped with atrium and prostate glands; there are three or two pairs of gonads, and spermathecae are aligned to gonadal segments. A puzzling, small family, Dorydrilidae, is similar to the lumbriculids in all features but their (secondarily?) plesioporous male ducts. The order Opisthopora is a large, monophyletic clade with opisthoporous male ducts and includes the numerous “megadrile” families of the mostly terrestrial “true

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earthworms” or the suborder Crassiclitellata (with characteristic multilayered clitellum), as well as the small, freshwater “microdrile” suborder Alluroidida, with the families Alluroididae and Syngenodrilidae. Their chaetae are paired, sigmoid, and simple-pointed (except the perichaetine chaetae of some tropical earthworms, which are numerous and not arranged into bundles). 6.  The family Moniligastridae (the only family of the order Moniligastrida) reveals basically microdrile internal anatomy but with testes closed in special capsulae together with male funnels, and terrestrial “earthworm” appearance and way of life. 7.  A monospecific, South American family, Narapidae, apparently belongs to its own order Narapida and consists of tiny riverine worms without chaetae, with testes in the V segment, and with ovaries and spermathecae in the VII segment; in addition, the plesioporous male ducts include atria and penes. Thus, the Clitellata may have derived from an aquatic, sediment-dwelling, polychaete-like ancestor characterized by possession of variable number and shape of chaetae and by plesiopore male ducts. A trend toward loss of hair chaetae and the upper tooth of the bifid sigmoid chaetae, as well as a trend toward paired chaetae or toward a gradual loss of chaetae, can be observed in many separate genera and families. Atria, prostatic glands, and penial structures seem to be convergently arisen in different clades, as well as the alignment of spermathecae to other genital pores.

Distribution and Diversity Oligochaeta is a cosmopolitan group, and so are some of its freshwater families. The Naididae is probably the most cosmopolitan freshwater oligochaete family, being present in all biogeographic regions, including sub-Antarctic islands, as is the subfamily Rhyacodrilinae within the Tubificidae. Species of the family Haplotaxidae are known on all continents except Antarctica. The family Tubificidae is most diverse in the Northern Hemisphere, although several endemic genera are known in Australia and South America. In the last few centuries, several anthropochorous tubificids of Holarctic origin have been introduced and became common in the southern temperate zone. Tubificids are not so diverse and common in the tropics, being represented there mainly by the architomic genera like Aulodrilus and Bothrioneurum. The only tubificid species reproducing exclusively by sexual means, and common also in tropical countries, are Limnodrilus hoffmeisteri and Branchiura sowerbyi. The first of them originates from the temperate zone but is able to reproduce also at persistently high temperatures, while the second one even breeds more effectively at temperatures over 20 °C. Humans have unintentionally spread these two taxa among many equatorial

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regions. Anthropochorous exchange of tubificids between Europe and North America has been intensive in the twentieth century, as demonstrated, for example, by the recent appearance of Potamothrix spp. and many other European species in the Great Lakes, as well as by the presence of Quistadrilus multisetosus, V ­ arichaetadrilus harmani, and several American Limnodrilus species in Western Europe. Many older exchanges between Nearctic and Palaearctic fauna may have occurred. The original homeland of the peregrine Tubifex tubifex is obscure, but it may have been derived from the Western Palaearctic, because this species is absent from the natural water bodies of extreme northeastern Asia. The families of Naididae and Pristinidae, with their prevailing asexual reproduction (and thus not so affected by lack of seasonally cooler temperatures), are common both in the tropical and in temperate zones. The gill-bearing genera Dero and Aulophorus are particularly diverse in the hot regions. However, many naidids common in the temperate climate (e.g., Stylaria lacustris) are lacking in the tropics. Most naidid species of the temperate zone of southern continents like Australia and South America were probably introduced from the Holarctic. The family Lumbriculidae is certainly Holarctic, with most endemic species living in the cool subarctic or alpine water bodies or in groundwaters. Only a few species are widely distributed in the summer-warm open waters; and only one of them, the architomic Lumbriculus variegatus, is not dependent on sexual reproduction. Humans have successfully distributed that oligochaete on several temperate regions of the Southern Hemisphere. The terrestrial/amphibious/marine family Enchytraeidae is cosmopolitan. Their intrusions from soil into fresh water occur independently on all continents. The small freshwater families of Propappidae and Dorydrilidae are Palaearctic in distribution, while the Parvidrilidae occur also in North America. In contrast, the family Opistocystidae is architomic and, therefore, able to live in tropical waters, where it is limited to the Americas and Africa. The family Capilloventridae was originally described from sea but later found also from Australian freshwaters. The family Phreodrilidae, an ecological equivalent of Tubificidae but of Gondwanan origin, occurs in Australia, South America, Asia (Sri Lanka and the Arabian Peninsula), and southern Africa, with a relict species (Astacopsidrilus naceri) in North Africa and some recent introductions in Ireland and Japan. Lake Baikal is an extraordinary hot spot of diversity for Oligochaeta as well as for some other animal groups. Indeed, it has even been treated as a separate freshwater zoogeographic region! This large rift lake has existed in one form or other for at least 20 million years. Its waters are typically cool and well supplied with oxygen down to its maximum depth (∼1.6 km) (Martin et al., 1998) for at least

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the last several million years. The Baikalian fauna is rather saturated; and, in light of this rift lake’s special physical conditions, the ecosystem is considered inhospitable for possible newcomers from the “common” Palaearctic water bodies (Semernoy, 2004). Most of the Baikalian oligochaete species and many genera are endemic; species flocks are even recognizable or suspected (Martin, 1996). The most diverse families are Lumbriculidae (particularly the genera Lamprodrilus, Rhynchelmis, and Stylodrilus), Tubificidae (Baikalodrilus, Isochaetides, Rhyacodrilus, etc.), and Naididae (Nais, Amphichaeta, and Chaetogaster). An evolutionary radiation is in progress there, which was originally based on a limited number of founding species coming from surrounding waters. This modern radiation, rather than the relict character of fauna as once assumed by Michaelsen (1902), may be the reason of the richness of the Baikalian oligochaete fauna. However, discovery of some rare species in Europe that are congeneric with Baikalian endemics (in the genera Pseudorhynchelmis and Rhyacodriloides) can be explained by their relict character. A possible source of the Baikalian cool-water oligochaetes, besides the open water bodies, may be the groundwater fauna, as suggested by the occurrence of Phallobaikalus gladiiseta in this lake, a representative of a primarily marine tubificid subfamily, the Phallodrilinae, which is known to be occasionally present in groundwaters as well (Creuzé des Châtelliers et al., 2009); unfortunately, the groundwater fauna is virtually unstudied in Siberia. The smaller lakes of the Baikalian Rift (Lake Hubsugul, Lake Oron, etc.) reveal a few endemic species related to the Baikalian ones. Several smaller Holarctic ancient lakes like Tahoe, Ohrid, Issyk-kul, and Biwa have each developed a small selection of their own endemic oligochaetes. The Laurentian Great Lakes of North America are comparable with the Baikal in their ecological conditions, but they have arisen only in the Holocene and are therefore too young to have formed an endemic fauna. On the contrary, they are rather hospitable for immigration of lacustrine oligochaetes from Europe. The oligochaete fauna of the great African lakes (Tanganyika, Malawi, etc.) is very scarce, although these lakes are hardly younger than Lake Baikal. This is partly due to an obvious lack of studies, but it is probably also because these lakes provide a less favorable environment for most aquatic oligochaete groups (too warm, profundal region anoxic) (Martin, 1996). Another special hot spot for the evolution of the Palaearctic freshwater Oligochaeta is the Ponto-Caspian Subregion (the Black, Azov, and Caspian Seas together with lagoons and estuaries). The great water bodies of this basin have been alternatively fresh, brackish, and saline, and they have been connected with each other or separated for long geological periods beginning with the Tethys Ocean. Many local endemic tubificid and naidid species have arisen there and are able to survive in both brackish and fresh waters. On the

Chapter | 21  Clitellata: Oligochaeta

other hand, at least three species of the otherwise strongly freshwater family Lumbriculidae (Stylodrilus cernosvitovi, S. parvus, and Trichodrilus pauper) have adjusted to the brackish water of the Caspian Sea. In the Holocene and after the retreat of the continental ice from northern Europe, several Ponto-Caspian species of the genera Potamothrix, Psammoryctides, etc. successively colonized the emerging new freshwater bodies—a process accelerated by humans in the last few centuries (Milbrink and Timm, 2001). Since 2000s, a cluster of endemic Potamothrix spp. (probably related to the Ponto-Caspian ones) was discovered in the well-oxygenated profundal of the deep mountain Lake Fuxian (southern China). They may represent relicts of the other, eastern end of the Tethys Ocean. Lake Fuxian is too warm and located too far south for colonization by any Lumbriculidae characteristic of Lake Baikal. Subterranean fauna of the regions not glaciated in Holocene is characterized by many small endemic oligochaetes, mainly tubificids and lumbriculids. These have been most extensively studied in Europe, but they have also been investigated in arid southwestern Australia, where many endemic phreodrilids occur. The Holarctic Parvidrilidae can be considered the most representative oligochaete family in groundwater, with all its nine constitutive species restricted in distribution to such habitats (Martínez-Ansemil et al., 2012). Occasionally, subterranean oligochaetes occur in springs together with epigean species. A succession of the oligochaete fauna along the river from headwaters downstream can be observed, with the most oxyphilous species living in the cool, stony rhitral and more tolerant, psammo- or pelophilous ones in the potamal sections of the river. Many rheophilous oligochaetes can exist also in the well-aerated surf zone of lakes. A few species, e.g., the genus Lamprodrilus (Lumbriculidae), occur only in standing water bodies. In North America, a particularly diverse, endemic oligochaete fauna occurs in the running waters of the western mountain ranges, as well as in the southeastern United States. The most diverse oligochaete fauna in the northern temperate zone, which consists mostly of the naidids, tubificids, and lumbriculids (phreodrilids can occur in similar habitats in Australia), occurs in the vegetation-rich shallows of lakes, small streams, and rivers where a successful bottom grab or pond net sample can often reveal 10–12 species. This number usually declines in deeper river habitats due to more monotonous (unvegetated) sediment, and in the lake profundal because of low dissolved oxygen levels. Lake Baikal serves as an exception to this latter pattern because its sediment is well oxygenated at all depths. Consequently, Lake Baikal has far more than 200 species of Oligochaeta, most of them endemic (Semernoy, 2004). For comparison, the well-studied temperate Lake Peipsi-Pihkva and Lake Võrtsjärv, in Estonia, have only 59 and 54 oligochaete species, respectively. Freshwater oligochaete fauna of the southern temperate zone

535

is characterized by many naidids and tubificids apparently introduced from the north and often dominating over the local ones. Phreodrilidae can also be diverse but are mostly in Australia. In tropical countries, the freshwater oligochaete fauna is usually represented by some genera of the families Naididae (particularly Dero, Aulophorus, and Allonais), Pristinidae, and a very limited number of Tubificidae. However, species of other families can sometimes dominate, such as Narapa bonettoi and Haplotaxis aedeochaeta on the unvegetated sand bottom in the Paraná River system, South America.

GENERAL BIOLOGY External Anatomy Oligochaetes are bilateral, segmented, vagile annelids sharing with polychaetes two types of chaetae (hairs and crotchets) and their location in four bundles, but lacking the parapodia and paired head appendages. The size of freshwater oligochaetes varies usually between 1 mm and several centimeters. Their body is mostly soft and smooth, but in some taxa it can be covered with armor formed by epidermal papillae and/or secretion and adhered particles. The first segment, or peristomium, surrounding the mouth, is always devoid of chaetae. Roman numerals are conventionally used for marking the segments, and Arabic numerals for the intersegmental furrows or dissepiments (e.g., 3/4 is the furrow between segments III and IV). In addition to a basic segmentation that corresponds to the succession of metameres, a secondary annulation is sometimes present in some of the anterior segments: the segments are divided into annuli that may make it difficult to determine the limits of the segments. A small tactile appendage, the prostomium, is attached to the peristomium above the mouth; sometimes it can be extended into an unpaired proboscis. A pair of pigmented eyespots can occur on the prostomium in some Naididae. Only a few oligochaetes have external gills: those in Branchiodrilus cover the anterior hair chaetae; segmental finger-like gills are present dorsally and ventrally on the hind body of B. sowerbyi and laterally in Phreodrilus branchiatus; and a limited number of gills and/or palps surround the anus in Dero, Aulophorus, and members of the family Opistocystidae. Chaetae (called also setae) are present in four bundles (two ventral, two dorsal or dorsolateral) on each segment except the first. They are sometimes absent on the genital and some other segments, but they are seldom entirely lacking. Two basic types of chaetae are present: sigmoid and hair chaetae (Figure 21.3). Short, mostly S-shaped (sigmoid) chaetae or crotchets are characterized by a more or less median swelling (the nodulus) and a bifid distal tip. The two teeth (or prongs) of the ectal end are called the upper (distal) and lower (proximal) teeth. The bifid chaetae, modified with presence of intermediate denticles or a wavy web between the two teeth, are called pectinate or comb-like chaetae. The teeth

536

FIGURE 21.3  Types of chaetae in Oligochaeta:(1–5) hair chaetae: 1, smooth; 2, pilose; 3, feathered; 4, serrated; 5, with supporting chaetae; (6–17) short chaetae: 6, common bifid crotchet with equal teeth; 7, with shorter lower tooth; 8, with shorter upper tooth; 9, simple-pointed; 10, pectinate; 11, palmate; 12, needle chaeta with equal teeth; 13, with shorter upper tooth; 14, simple-pointed crotchet without nodulus; 15, stick-shaped chaeta; 16, furrowed spermathecal chaeta; 17, obtuse penial chaeta; (18–27) chaetal bundles: 18, dorsal bundle with hair and bifid (or pectinate) chaetae; 19, dorsal bundle with a single hair and needle chaeta; 20, bundle of bifid crotchets; 21, paired simple-pointed crotchets; 22, bundle of penial chaetae with converged distal tips; 23, Mesenchytraeus-type; 24, Lumbricillus-type; 25, Enchytraeus-type; 26, Fridericia-type; 27, paired stick-shaped chaetae. Original drawing by Tarmo Timm.

can be of uneven length and thickness; in particular, the upper tooth may be reduced or even disappears. Chaetae in some species can be spade-like, brush-like, or of some other form. Most terrestrial groups bear simple-pointed chaetae (sigmoid, or straight and stick-shaped), and may lack a nodulus. In some aquatic taxa, the dorsal bifid chaetae are modified as straight, sharp-tipped needles, which are either bifid or simple-pointed. A trend expressed in several oligochaete groups is the assumed reduction of the originally indefinite number of chaetae per bundle to two (the paired chaetae) and, seldom, to one or no chaetae. The ventral chaetae located at genital pores can be modified as genital chaetae (see below).

SECTION | IV  Phylum Annelida

The second basic type of chaetae in many aquatic Oligochaeta of the order Tubificida is hair chaetae (also called simply “hairs”). They are long and gradually thinning distally, without any nodulus, and usually with a fine simplepointed tip. The hair chaetae can be irregularly covered by fine hairs (then called hirsute or pilose chaetae). The latter can be distributed in one or two longitudinal rows (on serrate or plumose hair chaetae, respectively). Most often, the hair chaetae look smooth in a light microscope. However, scanning electron microscope photos demonstrate a fine pilosity in many apparently smooth hair chaetae, as well as the presence of occasional fine intermediate denticles in apparently bifid crotchets. Lack of the hair and pectinate chaetae in ventral bundles was regarded as a character discriminating Oligochaeta from the nonoligochaete Aeolosomatidae bearing hair chaetae in all bundles. However, ventral hair chaetae are present in the recently discovered small oligochaete families Capilloventridae and Parvidrilidae. The function of the hair and pectinate chaetae is unclear and apparently not always vital, since they have been replaced by bifid crotchets in separate species of the same genus and sometimes even within a species. Chaetae can be more numerous and diverse in the anterior portion of the worm, while becoming gradually more scarce and uniform in the tail portion. The clitellum is an external, often conspicuous reproductive organ present during sexual maturity in all oligochaetes and other Clitellata (Figure 21.1(b)). This is a muff-like or saddle-shaped, glandular thickening of the mucous epithelium of body wall. It typically covers several segments, usually including those bearing male and female genital pores, but it may be located several segments posteriorly of the genital pores in some earthworms (suborder Crassiclitellata). Genital pores are usually present on the ventral side of these worms. Male pores are often prominent, lying on a glandular protuberance or (seldom) on an external penis. The pores are either on segments XII, XI, or X (or IX–X) in most aquatic oligochaetes but further back in terrestrial earthworms. Female pores (mostly one pair, but two pairs in some Haplotaxidae) are inconspicuous and located in intersegmental furrows either immediately posterior to the male pores or one segment caudally (but before the male pores in most “earthworms”). Spermathecal pores can lie immediately before and/or after the male and female pores, or several segments anteriorly. They are present either ventrally, laterally, or dorsally, and are sometimes unpaired. Spermathecal pores are often distinct like the male pores. Ventral chaetae at the male and spermathecal pores can be modified into genital chaetae (penial or spermathecal, respectively). Penial chaetae are usually blunttipped; when several are present in a bundle, their distal tips converge. They seem to be used for attachment and sperm transfer into the spermathecae of the concopulant.

Chapter | 21  Clitellata: Oligochaeta

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The penial chaetae (as well as unmodified ventral chaetae) are usually lacking at male pores when internal chitinous penial sheaths exist; these probably have a similar function in facilitating sperm transfer into the spermathecae during copulation. The spermathecal chaetae are mostly sharptipped, with a longitudinal furrow, and are often equipped with a distinct gland at their base. In all probability, they act as piercing chaetae, whose principal role is a mechanical or chemical stimulation, the latter via the inoculation of secretions into the partner’s body at copulation (Cuadrado and Martínez-Ansemil, 2001). In some rare cases, modified chaetae of spermathecal type can occur both at the spermathecal and male pores (e.g., in the tubificid genus Haber). A forward shift of the whole genital system occurs individually after regenerating the anterior end in those genera or species capable of asexual reproduction by architomy, or fragmentation.

Internal Anatomy Internally, the body cavity of each segment is usually separated from that of neighboring segments by muscular– epithelial dissepiments (septa); coelom of every segment is, in turn, divided into left and right parts by a mesentery. Coelomic corpuscles of various sizes and structures—the so-called coelomocytes—can float freely in the coelom fluid of several oligochaete taxa. The digestive tract of Oligochaeta is a strict longitudinal tube consisting of five to six consecutive sections. A simple mouth cavity is followed by a pharynx, which is characterized by a dorsal, muscular-glandular roof called pharyngeal pad. This pad can be everted for collecting small food particles. Bodies of paired pharyngeal glands can reach for several segments backwards and are then also called “septal glands” because they attach to the dissepiments. Both the pharyngeal pad and glands tend to degenerate in predatory oligochaetes like Chaetogaster and Phagodrilus whose pharynx is entirely muscular. A narrow esophagus follows the pharynx and enlarges into a midgut or intestine after several segments. Sometimes the beginning of midgut is dilated and then called stomach. The midgut nearly fills the coelom cavity of most body segments, narrowing whenever piercing a dissepiment. The midgut, as well as the posterior portion of esophagus, is covered with a layer of darker cells, the chloragogen tissue. The simple hindgut occupies a few hindmost segments. Different appendages and modified portions of the wall can occur at the esophagus or the beginning of intestine, in some taxa. The blood circulatory system always includes at least one dorsal and one ventral longitudinal vessel. The dorsal vessel pulsates, pumping blood forward. The two vessels are interconnected in some segments by pairs of transverse vessels (Figure 21.4). A pair or two of these transverse vessels in the forebody can be modified into muscular,

FIGURE 21.4  Scheme of blood circulation system of the tubificid Tubifex tubifex, by d’Udekem (1853), from Wesenberg-Lund (1939).

pulsating “hearts,” while transverse vessels in the posterior segments can be turned into blind appendages of the dorsal vessel (Figure 21.5). In addition, a blood sinus is common inside the intestinal wall; and in larger worms, other smaller vessels may be present. Dissolved erythrocruorin gives the blood a red or pink appearance in many oligochaetes when the worm is alive. The excretory system includes a number of paired metanephridia, or segmental organs, located in many segments except the genital and some anterior segments. A metanephridium consists of a ciliated funnel on the anterior surface of a dissepiment, with a duct piercing the dissepiment and winding in the cavity of the next segment before opening through a ventral pore. The central nervous system consists of a ventral nerve cord (with a pair of merged ganglia in every segment except the foremost ones), dorsal brain, and circumpharyngeal nerve ring linking the brain and cord. The brain is located slightly posterior to the prostomium, usually in the segments I–III. Three giant nerve fibers in the ventral nerve cord ensure a quick information exchange between the brain and tail and rapid contraction in response to mechanosensory

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SECTION | IV  Phylum Annelida

FIGURE 21.5  Branched, blind, lateral blood vessels on the posterior segments of Lumbriculus variegatus (live picture). Scale = 200 μm. Photo by Patrick Martin. Copyright © 2015 Dr Patrick Martin. Published by Elsevier Inc. All rights reserved.

irritation. External sense organs are represented by inconspicuous sensory papillae, which are more abundant on the prostomium. In armored species, the papillae are more prominent and penetrate through the armor. Light-sensible sensors are present in the body wall of most oligochaetes, but pigmented eyespots on the prostomium occur only in some Naididae. The presumably ancient condition of four pairs of gonads—with testes in segments X and XI and ovaries in XII and XIII—has been preserved in most Haplotaxidae. This number is reduced to one to two pairs of testes and one pair of ovaries in other families (Figure 21.6). Most often the single pairs of testes and ovaries are located in their original segments XI and XII, respectively (Enchytraeidae, Phreodrilidae, many Lumbriculidae, etc.), or in X and XI (Tubificidae and some others). Developing gametes usually float freely in the coelom of the gonadal segments. To provide sufficient space for them, the posterior (or also anterior) wall (dissepiment) of a gonadal segment will bulge into a neighboring segment(s) to form a sperm or egg sac. The posterior egg sac always surrounds the sperm sac, and the ripe eggs lie usually caudad from the sperm mass. These sacs can reach for many segments behind the clitellum. The accumulation of gonadal products in sacs gives a milky-white appearance in the region adjacent to the clitellum, and is the most conspicuous element of the reproductive system in superficial observation. Gonoducts are almost always paired like the gonads.

Male ducts include a ciliated sperm funnel at the posterior wall of the testicular segment and a sperm duct or vas deferens piercing the dissepiment and usually winding in the coelom of the next segment. Vas deferens can open through a simple male pore in the body wall, as in Haplotaxidae or Crassiclitellata. More often, the invaginations of body wall have formed various terminal elements of the male duct, called atria and penes in Lumbriculidae, Tubificidae, and some others, or penial bulbs in Enchytraeidae. The atrium is a spacious chamber with glandular walls, serving for collecting sperm before copulation. The bodies of the gland cells of the atrial wall are called prostatic glands when bulging into the coelom. They can cover the outer surface of atrium (diffuse prostate) or form aggregations sending their secretions into atrial lumen by a common narrow stalk (compact prostates; not to be confused with the homonymous prostates of many megadriles, which are different, separate organs located near the male genital pores). When the distal end of the atrium is modified for protrusion during copulation, it is called pseudopenis. The atrium can also terminate with a true penis, surrounded by a penial sac when not protruded (seldom is a penis permanently external, e.g., in Stylodrilus heringianus). Sometimes there is a narrow ejaculatory duct between the atrium and penis. The penis can be either completely soft or surrounded by a more or less chitinized penial sheath. Male pores, located near or instead of the ventral chaetal bundles of the respective segment(s), are

Chapter | 21  Clitellata: Oligochaeta

539

FIGURE 21.6  Reconstruction of reproductive organs of one side in the tubificid Embolocephalus velutinus. at = atrium; de = ejaculatory duct; ff = female funnel; mf=male funnel; o =ovary; pa = penial apparatus; pr = prostate gland; st = spermatheca; sts = spermathecal chaeta; t = testis; vd = vas deferens. From Holmquist (1978).

FIGURE 21.7  Types of male duct in Oligochaeta. From Martin and Aït Boughrous (2012).

usually the most prominent external genital organs beside the clitellum. Michaelsen (1929) coined three useful terms to describe the types of male gonoduct (Figure 21.7): plesiopore (with external pores in the segment following the testicular one, as in most aquatic oligochaetes), prosopore (with pores in the same segment with the testes, as in Lumbriculidae and many of their leech-like relatives), and opisthopore (with the male pore several segments backwards, as in the Crassiclitellata). Many Lumbriculidae display a semiprosopore condition when the first pair of vasa deferentia lack their own atria and join atria of the second pair (thus being, in fact, plesiopore). Prosoporous vasa deferentia of the testicular segment can either penetrate in the posttesticular

segment and make a loop before piercing again its dissepiment, or only cling to its anterior surface before joining the atrium. Sometimes the male pores of the left and right side can open into a common, median copulatory chamber, or there can be a single, unpaired atrium receiving all vasa deferentia. Female ducts consist only of a ciliated funnel at the posterior wall of the ovarial segment and a vestigial canal penetrating the dissepiment, with an inconspicuous female pore in the intersegmental furrow, in the line of ventral chaetal bundles. Spermathecae are another important element of the reproductive system because they retain the partner’s sperm after copulation and before egg laying. Their number and location vary among taxa, from one single spermatheca to many pairs, and from being in the

SECTION | IV  Phylum Annelida

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immediate neighborhood of the male and female pores and clitellum (e.g., in the segment preceding them in Tubificidae and Naididae, before or after them in Lumbriculidae) to many segments before them (e.g., in Enchytraeidae and Lumbricidae). They can be simple sacs inverted from the body wall, or be differentiated into spermathecal ampulla and external duct. The ampulla can bear diverticula, and the duct can be covered with a glandular layer or bear separate glands. The sperm contained in the ampulla or diverticula can be amorphous or arranged into bundles or even spermatozeugmata (characteristic of most Tubificinae, but also rarely present in the Limnodriloidinae, Phallodrilinae, and Rhyacodrilinae). A tubificine spermatozeugma consists of fertilizing and nonfertilizing spermatozoa, the latter forming a cortex for the former ones with some sort of cementing agent (Figure 21.8). In some groups, particularly among the Enchytraeidae, an internal duct connects the ampulla with the digestive tube, apparently for absorbing the remains of sperm not used for fertilizing eggs. External pores of the spermathecae can lie in different parts of the respective segment: ventrally, laterally, or dorsally within a segment or in an intersegmental furrow. Spermathecae can be lacking in parthenogenetic individuals of some taxa and also in taxa that attach external sperm packets (spermatophores) to the partner’s body surface. A forward shift of the whole genital system occurs individually after regenerating the anterior end in genera or species able to reproduce asexually by architomy, or fragmentation (e.g., Lumbriculus, Aulodrilus, Bothrioneurum,

and Cognettia). A limited, species-specific number of segments is always regenerating anteriorly on the anterior end, and the single or first pair of testes is then developing in the last regenerated segment (Hrabě, 1981). In paratomic families of Naididae and Pristinidae, such a relatively anterior position of the genitalia (with testes in IV, V, or VII) is genetically fixed. The extensive backward shift of the whole genital system in some of the paratomic Opistocystidae has no reasonable explanation.

Physiology The basic mode of locomotion in the sediment-dwelling oligochaetes is slow peristaltic crawling. Every external vibration or other tactile irritation will result in a momentous contraction of the whole animal. Most oligochaetes are thigmotactic, which is why tubificids gather into balls when extracted from sediment. Some species of Lumbriculus and Rhynchelmis (Lumbriculidae), as well as most Naididae, that spend at least some time among aquatic vegetation can swim short distances with wriggling or spiral movements. Most oligochaetes are negatively phototactic, with lightsensitive cells dispersed in the whole body wall. However, some Naididae have a pair of primitive eyespots, which they use for orientation. Oligochaetes transpire (exchange gases) through their mucous body wall and, in rarer cases, with external gills. Another important respiratory organ is the ciliated hindgut, which explains why burrowing tubificids expose their hind

FIGURE 21.8  Spermatozeugmata in spermathecae of Potamothrix hammoniensis. Scale = 100 μm. Photo by Patrick Martin. Copyright © 2015 Dr Patrick Martin. Published by Elsevier Inc. All rights reserved.

Chapter | 21  Clitellata: Oligochaeta

end in the water layer and even try to create vertical water circulation, with “swimming” movements of their tail. For a similar reason, other worms such as L. variegatus and Criodrilus lacuum stretch their tails horizontally along the air–water interference when living in anaerobic mud under a thin water layer. Gases and various materials are circulated internally, primarily by the vascular system and secondarily by coelomic fluid. Blood is especially rich in erythrocruorin in species such as Potamothrix hammoniensis or T. tubifex living in sediment that is poor in oxygen or even periodically anoxic. Most aquatic oligochaetes are more sensitive to the scarcity of oxygen and, therefore, inhabit cleaner and/or cooler water bodies. Digestion in oligochaetes depends in part on their internal microbial flora. Some bacteria can propagate in their gut and may contribute to the digestive process by helping decompose high-molecular organic compounds. Assimilation of dissolved organic material by the body wall has been demonstrated in some mud-dwelling tubificids. Metanephridia (or segmental organs) are the primary excretory organs. In addition, solid excretory products accumulate in the chloragogen cells covering the intestine. These cells darken over time and are partially eliminated by the seemingly drastic process of autotomy (pinching off) of the old tail. This is followed by regeneration of a new tail that replaces the old, waste-laden chloragogen cells with new tissue. Growth occurs primarily in the caudal end of oligochaetes. New segments are adding one by one to the posterior end (anterior to the pygidium), both in the case of regular growth and in tail regeneration. When the anterior of the worm requires regeneration, a small unsegmented outgrowth appears initially and then differentiates into a species-specific number of segments. The number of regenerated segments is often less than the number of lost segments. Genital segments usually do not regenerate (except in the species with asexual reproduction). For this reason, regeneration of anterior segments is limited.

Reproduction and Life History The individual lifetime of tubificids and lumbriculids in aquaria often lasts several years and may include several reproduction cycles. Some individuals of T. tubifex and Spirosperma ferox have reached an age of 10 or more years. The oldest recorded age was for an individual of C. lacuum (an aquatic “earthworm”), which lived 46 years in an aquarium, although without reproducing. The actual lifetime of the same species in nature is undoubtedly shorter due to predation. The facultative parthenogenetic species T. tubifex can survive and reproduce in aquaria for many years, even at a persistent room temperature, as can the architomic clones of the tubificids Bothrioneurum and Aulodrilus.

541

Some other species can live several years without reproduction unless “hibernation” enables them to undergo the normal sexual process (Timm, 1987). Most species reproduce once a year and then reabsorb the whole system apparatus (except gonads) and apparently become immature until the next sexual cycle. Oligochaetes are hermaphrodites, mostly protandric, with the partners exchanging sperm during copulation. The clitellum and some special cutaneous structures, sometimes in combination with penial chaetae (Cuadrado and Martínez-Ansemil, 2001; Caramelo and Martinez-Ansemil, 2012), help to hold the partners together during this process. The alien sperm is stored in spermathecae either as amorphous mass, organized into bundles of different complexity, or as spermatozeugmata consisting of two kinds of spermatozoa, the latter characteristic of the subfamily Tubificinae (see Jamieson and Ferraguti, 2006). When spermathecae are absent, the spermatophores are then attached to the outside of the partner’s body; this occurs in the tubificids Bothrioneurum and Paranadrilus, the crassiclitellate Criodrilus, and some others. Both the egg(s) and alien sperm will be laid into a cocoon secreted by clitellum. The mother worm sheds the cocoon by crawling backwards, after which the cocoon’s shell will harden and the ends will contract into a sealed plug. Eggs are fertilized within the cocoon, and development is direct. After some weeks, the young worms will leave the cocoon. “Microdrile” aquatic worms lay large, yolk-rich eggs. In contrast, the eggs of “megadriles” are smaller, and embryonic growth depends on nutritious fluid secreted into the cocoon by the mother’s clitellum. Spermatogenesis occurs earlier than oogenesis and at relatively lower temperatures. This explains the observation that in a moderate, seasonal climate many aquatic oligochaetes mature and copulate in winter or early spring and lay eggs in the beginning of the warmer period. Although oligochaetes may reproduce only once per year in seasonally cool aquatic systems, some species (e.g., L. hoffmeisteri) can mature rapidly in warmer water and produce several generations per year. Some species (e.g., T. tubifex and Ilyodrilus templetoni) can produce parthenogenetic eggs, as confirmed by lack of spermathecae in many individuals. Some rare taxa also lack the male apparatus, as in Tubifex pomoricus. Parthenogenesis is a way to avoid the temperature problem with spermatogenesis and to produce several generations a year, as in the case of T. tubifex and L. hoffmeisteri. However, fecundity gradually decreases over several parthenogenetic generations. Another form of asexual reproduction that completely avoids the resource- and time-consuming sexual process and egg-laying is vegetative reproduction, either by architomy or paratomy. Architomy means enhanced ability to regenerate a complete animal from separate pieces, after either a violent attack or spontaneous fragmentation. Architomy is almost obligatory in L. variegatus but common also in some Tubificidae (e.g., Bothrioneurum, Aulodrilus, Potamothrix

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bedoti) and Enchytraeidae (Cognettia and some others). At least in these tubificids, architomy is characteristic of warmer periods, whereas at lower temperatures, the process ceases and at least some part of the population becomes sexually mature. In Lamprodrilus mrazeki (Lumbriculidae), architomy occurs in cysts formed in summertime when pools dry up. A more advanced method of asexual reproduction is paratomy: the production of a chain of zooids by longitudinal budding. This characterizes whole families of Naididae, Pristinidae, and Opistocystidae. Paratomy can prevail during favorable seasons, while sexually mature individuals appear and produce cocoons mostly before a colder or drought period. The parents then usually die after forming cocoons.

GENERAL ECOLOGY AND BEHAVIOR Macrohabitat Distribution and Microhabitat Selection The most typical habitat of freshwater oligochaetes is the uppermost (several centimeters thick) layer of sediment where the worm occupies a “tube” (the genus name Tubifex is derived from this behavior). Tubes are slime-lined walls of a burrow in soft sediments from which the “head” and tail may protrude above the sediment surface. Oligochaetes may reside temporarily in a deeper, anoxic layer of muddy sediment while feeding on bacteria and avoiding predators. The egg cocoons are also often laid inside sediment. In the interstitial spaces of stony or gravel bottoms, especially in the well-aerated hyporheic zone of the torrent streams, the depth distribution of oligochaetes can reach several meters. This is a transition zone to the groundwater. The latter is inhabited partially by invaders from surface waters, partially by stenothermic species specific to this habitat, which are often endemic. Some oligochaetes, particularly of the family Enchytraeidae, can live equally in freshwater sediment and moist terrestrial soil. Several oligochaete groups have independently adapted to another habitat—the underwater “forests” of macrovegetation and algae. Here they crawl around on the plants and the sediment surface rather than residing in subsurface burrows. These include most Naididae, Pristinidae, and Opistocystidae, and at least some species of the lumbriculid genera Lumbriculus and Rhynchelmis. Some species can even swim short distances. For example, small naidids equipped with long hair chaetae (e.g., Stylaria and Ripistes) have been observed floating or swimming in the open water. Naidids, although able to swim (Dero, Aulophorus, Ripistes), also build temporary, slimy hiding tubes, which they attach to aquatic plants. Other naidids, e.g., Uncinais uncinata and Piguetiella blanci, prefer the surface of sand bottoms

SECTION | IV  Phylum Annelida

to the surface of plants. Some heavily armored (with papillae) Baikalodrilus spp. (Tubificinae) in Lake Baikal are assumed to crawl or roll on the sandy bottom on this lake (Semernoy, 2004). A few freshwater oligochaetes live on or inside other animals. Some Australian species of Astacopsidrilus (Phreodrilidae) live as commensals within the gill chamber of crayfishes. Chaetogaster limnaei (Naididae) has two subspecies, one as a commensal epibiont on aquatic snails and the other parasitizing their kidneys. All species of the tropical genus Allodero (Naididae) are known from the excretory organs of tree frogs.

Physiological Constraints Distribution of oligochaetes is limited primarily by availability of dissolved oxygen above and within the sediment. The abundance of erythrocruorin in blood is characteristic of mud dwellers like the tubificids and phreodrilids but not of the vegetation dwellers like the naidids and pristinids. Body length can limit the vertical distribution of tubificids in anoxic sediment because worms need to be able to reach with their tail end to the overlying aerated water. Oxygen deficiency also limits the distribution of oligochaetes into deeper zones of lakes. Oligochaete richness in Lake Baikal at all depths is greatly due to good aeration of its whole water mass and the uppermost sediment layer (Martin et al., 1998, 1999). Depth (water pressure) per se has no importance for oligochaetes. Most of them do not need light within their sedimentary burrows and even avoid it. Some vegetation-dwelling naidids equipped with eyes are an exception to this pattern. Many oligochaetes are eurythermic and able to survive both low (near-zero) and high (above 20 °C) temperatures; some are even adapted to seasonal freezing, like the tubificid Alexandrovia ringulata in Arctic tundra lakes. Most of the Lumbriculidae prefer persistent low temperatures, inhabiting either the Arctic, alpine, or underground water bodies. The groundwater species can be stenothermic. The annual temperature regime can delimit distribution of many surfacewater species via the reproduction. For example, the lumbriculid Lamprodrilus isoporus survives the warm summer but reproduces only in winter in Lake Peipsi (Estonia). In general, maturation, spermatogenesis, and copulation require lower temperatures than oogenesis and the laying and incubation of eggs. Asexual reproduction by paratomy or architomy is suppressed by temperatures lower than 10–15 °C. The thermal regime (and scarcity of dissolved oxygen at higher temperatures) may be the reason for the limited number of aquatic oligochaete genera in the tropical waters. For example, only L. hoffmeisteri, B. sowerbyi, some naidids, and pristinids were found in the tropical, persistently warm (19–31 °C) ponds in Mumbai, India. The presumably ubiquitous T. tubifex was lacking there but was found in the nearby Ooty Hills, where

Chapter | 21  Clitellata: Oligochaeta

the winter air temperatures range between −4 and + 15 °C (Naveed, 2012). In aquaria, T. tubifex has laid viable eggs at + 25 °C and even at +30 °C; however, these eggs may be parthenogenetic and not supporting a sustainable population. Water current supports the life of oligochaetes primarily because of its role in aeration. Many species characteristic of flowing water can also inhabit surf littoral of large lakes persistently washed by waves. Flowing water is avoided mostly by inhabitants of plant thickets but also by some openbottom species (e.g., the lumbriculid L. isoporus), possibly because of their vulnerability to drifting away in currents. Different aquatic oligochaete taxa are adapted to either fresh, brackish, or saline (marine) water. A few euryhaline species (Nais elinguis) can live both in freshwater and in marine littoral. Some freshwater species (e.g., P. hammoniensis and Psammoryctides barbatus) occur also in the brackish Baltic Sea at salinities up to 5–7 ppt. A salt tolerance up to 10 ppt was established for the freshwater species L. hoffmeisteri and I. templetoni taken from a tidal water estuary (Chapman and Brinkhurst, 1980). In the Ponto-Caspian basin, which has endured many salinity fluctuations during its geological history, many endemic species have arisen, thriving equally well in brackish water estuaries and freshwater bodies. An extreme example is the Caspian Sea (with average salinity of 11 ppt), where three species of the otherwise freshwater family of Lumbriculidae occur: the endemic Trichodrilus pauper and Stylodrilus cernosvitovi, and the otherwise freshwater S. parvus. Several genera of the basically marine tubificid subfamily Phallodrilinae (as well as of some other marine groups) include separate species in freshwater, e.g., Thalassodrilus hallae in the Great Lakes of North America and many representatives of Aktedrilus and Abyssidrilus in the European underground waters. The inland saline lakes are usually devoid of oligochaetes, except for some single species of Enchytraeidae. Tissue desiccation after drying of the surrounding water body is lethal for the most aquatic oligochaetes. The ubiquitous L. variegatus and T. tubifex have been observed dormant in a slimy cyst in the sediment of temporary pools. The ability of the hard-shelled cocoons of Naididae to survive in an aerial environment has been discussed but never proved. Some Pristina species and the phreodrilid Schizodrilus have been described living in humid soils, and the tubificids Rhyacodrilus falciformis and Bothrioneurum grandisetosum are known to thrive even in mesic soil.

Feeding Behavior Burrowing species ingest large amounts of fine sediment particles of suitable size, selectively digesting some organic components but defecating most of the ingested material. Tubificidae digest a species-specific selection of bacteria, whereas Enchytraeidae presumably prefer soil fungi but can thrive also on bacterial cultures or on the biofilm of sewage

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beds. Masses of tubificids inhabiting organic-rich but badly aerated sediment will often hold themselves in a vertical position, with anterior end (and mouth) in deeper sediment but the tail end held high over the bottom surface so they can simultaneously feed and respire. The phytophilous Naididae most often graze on various epiphytic algae, bacteria, and protists, digesting selectively some algae (particularly diatoms) but egesting others. Curiously, Ripistes parasita uses its extra-long hair chaetae for food filtration, while U. uncinata ingests sand grains, digesting their biofilm. A few oligochaetes have adapted to predatory feeding. Chaetogaster spp. (Naididae) have developed a voluminous pharynx able to suck in various smaller animals or diatoms. Phagodrilus and Agriodrilus (Lumbriculidae) feed mostly on other oligochaetes by grabbing them with their very muscular pharynx, much like some leeches. Haplotaxis gordioides (Haplotaxidae) is assumed to grip its prey oligochaetes with its extended prostomium and peristomium and then constrict the prey boa-like with its very long body and sickle-shaped ventral chaetae. The internal parasites of excretory organs (C. limnaei waghini and Allodero spp.) feed on epithelium cells of their hosts. The ectocommensal Astacopsidrilus spp. (Phreodrilidae) is probably a micropredator, cleansing microorganisms from the gill chamber of their crayfish hosts.

Predators and Parasites Oligochaetes serve as an easy and nutritious prey for predatory chironomid larvae, leeches, and other invertebrates. The numerous stiff and sharp chaetae in some species like Nais barbata or Vejdovskyella comata (Naididae) may provide some protection against predators. Oligochaetes are certainly an important food object for benthophagous fishes, but they are difficult to count or weigh in the contents of fish intestine because they are digested very rapidly, leaving only chaetae. For this reason, the role of Oligochaeta in the diet of fishes is often underestimated. Fishes can feed on the mud-dwelling tubificids not only by devouring them completely but also by picking their waving tails stretched out of sediment. Fortunately for the worms, their tails are soon regenerated (Wiśniewski, 1978). Sessile ciliates often attach to the posterior ends of tubificids, using them as a substrate that is mostly exposed to aerated water. Parasitic rotifers Drilophaga are known to attach to the body surface of oligochaetes, while the rotifers Albertia and Balatro can live either in the worm intestine or coelom. Various unicellular parasites (ciliates, gregarines, microsporidians, etc.), as well as nematodes and trematodes, are known from the interior of oligochaetes. In the 1980s, the actinosporean internal parasites of Oligochaeta were identified as a regular stage in the life cycle of Myxozoa, the economically important fish parasites (Kent et al., 2001). Another group of fish parasites,

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cestodes of the family Caryophyllaeidae, spend their larval (procercoid) stage mostly in the coelom of tubificids. Some of them (Archigetes) can reach sexual maturity and reproduce in oligochaetes too.

Other Ecological Aspects of Oligochaeta Competition between the tubificids and another abundant group of freshwater mud dwellers, the chironomid larvae, would seem likely but has never been proved. On the contrary, the years of maximum abundance of the Chironomus plumosus in Lake Võrtsjärv (Estonia) also coincided with the greatest densities of the dominating tubificid P. hammoniensis. This oligochaete may thrive well when fish feed primarily on this midge competitor. The ecologically similar mud-dwelling tubificids and some others (e.g., the lumbriculid S. heringianus) ingest large amounts of sediment, selectively digesting some species-specific strains of bacteria but accelerating the growth of some others. In this way, different species of oligochaetes can “feed” each other and thrive better in mixed populations. Indeed, mixed populations of these worms are common in nature. It has been demonstrated that T. tubifex and L. hoffmeisteri thrive better in mixed than in pure cultures. However, at least two ubiquitous aquatic oligochaetes, the sediment-dwelling T. tubifex and the phytophilous N. elinguis, are typically most abundant in the extreme trophic conditions (N. elinguis also in brackish water) where the other, related species are scarce or lacking. Their assumed sensitivity to competition is a possible explanation of this. Significant ecological impacts of the phytophilous Naididae grazing on periphyton communities are possible but have never been quantitatively evaluated. Neither has potentially significant impact on diatoms and other protists been demonstrated for the micropredatory Chaetogaster spp. The commensal of gastropods, C. limnaei limnaei, can render a service to its hosts as a cleaner, while their kidney parasite C. limnaei vaghini is certainly pathogenic. The burrowing activity of oligochaetes mixes and aerates the uppermost sediment layer and accelerates its oxidation while simultaneously returning biogenic elements and contaminants accumulated in sediment back to the water layer. The widespread assertion that Oligochaeta as a group are indicators of heavy organic pollution is based on the ability of two ubiquitous tubificid species, L. hoffmeisteri and T. tubifex, to develop en masse in α-mesosaprobic or even polysaprobic conditions. Many other tubificids and naidids thrive in either β-mesosaprobic or oligotrophic conditions. Their communities, rather than separate species, can indicate the trophic conditions in a water body (Milbrink, 1978; Uzunov et al., 1988). Organic enrichment per se is not dangerous for oligochaetes as long as there is good oxygen supply. However, they may be sensitive to

chemical pollution, particularly by heavy metals. T. tubifex is broadly used for the laboratory tests of water pollution biology (Rodriguez and Reynoldson, 2011).

COLLECTING, CULTURING, AND SPECIMEN PREPARATION Collecting Oligochaetes can be collected with any standard dip net, corer, or bottom grab used for sampling most zoobenthos. A great deal of material is obtained in the course of routine ecological surveys. Washing the sediment often causes maiming of these soft worms. Elutriation of the sediment (particularly in the case of coarse sediments) and using rayon rather than metal sieves can mitigate this danger. Sorting of sieve residue alive on an enameled dish, either in field or in laboratory, is preferable; however, without magnification, some smaller individuals can be missed. Unfortunately, postponing the sorting can cause selective mortalities of the species more sensitive to higher temperatures and lowered oxygen concentration. To save time in field, the sediment samples can be preserved as a whole and later washed and sorted in laboratory. In this case the quantitative account of specimens can be more accurate, and even the smallest individuals will be found, particularly if the samples are sorted under a dissecting microscope. Small amounts of a stain like phloxine B or rose bengal can be helpful for finding the small worms while sorting preserved samples. Such dyes should be avoided, however, if oligochaetes are intended for molecular analyses, as they might affect the process of DNA sequencing. An easy preservative for bulk samples is 10% buffered formalin (4% formaldehyde solution), although a lower concentration can also work. Higher concentrations make most animals brittle, while too low ones do not prevent rapid degradation of fragile worms. As formalin is an acidic solution, it may be useful to neutralize it by adding 20 g/l sodium bicarbonate crystals (baking soda) or blackboard chalk. This is vital for some other animal groups having calcified shells. Hence, it is advisable in the course of general surveys to use a buffered 7–10% formalin solution as the “default” fixative. One can carry 40% formalin in the field and then add the appropriate amount to the aqueous sample to produce a concentration of 10% or less formalin. Preservation of bulk samples with alcohol (ethyl or isopropyl) enables collectors to avoid inhaling toxic formalin vapor, but large amounts of 95% alcohol are required to reach a minimal concentration of 70% in a preserved sample, except if the sample is elutriated prior and its volume reduced to a minimum. Seventy percent ethanol is the best medium for fixing live oligochaetes or storing them when originally fixed in formalin. Care should be taken to avoid diluting the alcohol at sorting; the volume of alcohol must exceed at least tenfold

Chapter | 21  Clitellata: Oligochaeta

the volume of the worms fixed. It is also a good idea to change the fluid in the vial before final storage of a sample. In a crammed vial, the oligochaetes will macerate into a worthless pulp. The same can happen when a vial is not well sealed and a part of the fluid has evaporated. The worms provided for DNA sequencing must be fixed and preserved in strong alcohol (80–96%) but never in formalin or alcohol “denatured” with additives that might interfere with DNA. After fixation, worms intended for molecular analyses should be kept as soon as possible at low temperatures (−20 °C). When available, a portable cool box should be used to temporarily store alcohol-preserved samples in the field. The strong alcohol makes worms somewhat stiff and, therefore, less easy for subsequent mounting. Freezing the entire sediment sample in the field for later defrosting in the laboratory should be avoided because the dead worms will start decaying immediately when defrosted. The worms collected for histological studies could be fixed in some special media recommended in respective guides. Special methods can be used for qualitative research in some cases. Naididae and Pristinidae can rapidly propagate by paratomy in an aquarium with macrovegetation and then be found crawling on the glass walls. Tubificidae and others can be extracted from sediment rich in plant debris, using the wet funnel method common for the studies of Enchytraeidae. A sediment sample can then be placed on a screen over a container with pure water; after some time, the worms will leave the sediment and fall to the bottom of the container.

Rearing Many tubificids, as well as the lumbriculid S. heringianus and the “aquatic earthworm” C. lacuum, have been successfully cultivated for scientific purposes in lacustrine mud sieved to avoid any incidental species. They do not need additional feeding or even water aeration if they are held in a sufficient amount of mud under a thin layer of water. The addition of a moderate amount of easily decayed organic matter, like yeast, fish food pellets, algal culture, etc., will accelerate the growth of the population. Some other species, e.g., L. variegatus, do not thrive without such additional feeding. Many species raised in aquaria need at least seasonally lower temperatures for their sexual reproduction. This is not essential for Lumbriculus, Aulodrilus, and Bothrioneurum, which reproduce by fragmentation, nor is it necessary for the facultative parthenogenetic T. tubifex. The last species has been cultivated for laboratory material in pure sand enriched with yeast or lettuce. Rearing the phytophilous Naididae and Pristinidae is more complicated because their paratomic clones, although reproducing rapidly, tend to fade over time. Bacterial film from hay or lettuce infusion can serve as a suitable food for them.

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Aquatic oligochaetes can serve as a live food for fish fry and aquarium fishes. Successful attempts at commercial production of T. tubifex on a mixture of cow dung and sand, under permanently flowing water, were described by Marian et al. (1989). The tubificid worms sold in aquarium shops as “Tubifex” (in fact, mostly L. hoffmeisteri) are often not cultivated but collected in polluted open waters (­Naveed, 2012). L. variegatus, with a commercial name “black worm,” may be the most suitable object for cultivating in aquaria since it can reproduce by fragmentation for an unlimited time.

Preparation for Identification Immature Oligochaeta are often unidentifiable; when available alive, they can be cultivated in small aquaria with natural substrate at 4–12 °C until sexual maturation (Bülow, 1957). Of course, the maturation can take several months, and the results are not always positive. Cultivated individuals may be more suitable for reference collections because they tend to lack injuries (missing tails and broken chaetae) common after sampling in nature. A microscopic study of live individuals is recommended if possible, as is common in the studies on Enchytraeidae. The living worms are relatively transparent so that most internal organs are visible, as are their intestine peristaltics, blood vessels’ pulsation, and work of cilia in nephridia. Mobility of the live worms can impede continuous investigation, photographing, or even measuring any structures. A slight pressure applied to the worm under a cover slip can be helpful, but excessive pressure must be avoided, as this will cause the worm to burst, leaving only chaetae well exposed. Different methods of immobilization can be used for small oligochaetes, e.g., putting them under a cover slip in a drop of carbonated water. This will bring the worm to a standstill due to suffocation, but the worm can survive for tens of minutes. Any fixative makes the body wall opaque so that the internal organs become hardly distinguishable. Fixed oligochaetes can be studied in temporary mounts under a cover slip in glycerin, which makes their bodies soft and half-transparent. Glycerin is very soluble in water and alcohol, and temporary mounts can be made that will persist for several years (but not forever) as long as the cover slip edges are sealed. After sealing, the slide can be possibly stored in a vertical position in microscope slide boxes. The use of nail varnish for sealing the slides should be avoided, although its use is regularly advised in the literature. The varnish becomes fissured with time; being permeable to ambient air, the slide finally dries and the worm is lost. Some have been using synthetic media (e.g., DPX mounting medium) and also Canada balsam for sealing purposes. Type and reference specimens should be maintained in permanent whole mounts, e.g., in Canada balsam. For

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this purpose, the worms can first be transferred from glycerin into dilute alcohol, then dehydrated through a series of baths of increasing alcohol content (70%, 96%, 100%; 50:50 alcohol-xylene), shifted to xylene, put into a drop of Canada balsam, and finally placed under a cover slip. After some days of desiccation of the slip edges, this mount is ready and can survive 100 years, if not more. The xylene makes worms and their chaetae hard and brittle, but this will be no problem with specimens flattened under a cover slip in glycerin. The larger oligochaetes (and also leeches) can be fixed, for this purpose, at the same time by pressing them flat between two slides. Internal organs are better visible in the Canada balsam than in glycerin, particularly if the complete worms were slightly stained before mounting. The specimens can be stained (e.g., with Mayer’s paracarmin or Ehrlich’s hematoxylin, always followed by cautious differentiation) before dehydration. An opposite problem can appear when chaetae become even too translucent in the balsam because their refractive index is more similar to balsam (and xylene) than it is to glycerin or water. Use of differential interference contrast microscopy (DIC) seems to mitigate this problem. There exist several other clearing media for temporary investigation, as well as synthetic resins for making permanent mounts (e.g., Euparal, broadly used in entomology); however, the persistence of the latter over decades is not yet proved. Many researchers have used for preliminary studies the compound Amman’s lactophenol, as first employed by Brinkhurst (1960). It consists of glycerin, water, phenol, and lactic acid. Brinkhurst also recommended polyvinyl lactophenol (Jones, 1946) for making semi-permanent whole mounts of aquatic Oligochaeta. The polymerization of vinyl makes the mounting medium semi-rigid, in which the worm cannot be displaced anymore. Phenol-based media are good for routine identification since they make chaetae and other chitinous structures very distinct. Unfortunately, they are unsuitable for reference material, since all soft internal organs will merge in a continuous mass under a cover slip, and even chaetae may swell after some years. A closer description of internal reproductive organs, particularly the male ducts, is necessary for a detailed taxonomic study. Unfortunately, these structures are often overshadowed by amorphous masses of sperm and prostate glands in the whole mounts. For this purpose, the genital segments can be dissected, a part of the body wall removed, and the male ducts and spermathecae extracted under a dissecting microscope, using sharp razor blades and very fine needles. That is a difficult task because these organs are soft and can be easily lost. Moreover, it is even more difficult to mount dissected small objects. An excellent alternative consists of cutting the anterior part of the worm into two halves, along the sagittal plane, by means

SECTION | IV  Phylum Annelida

of a microscalpel, such as the ones used in ophthalmology (iris knife-type) (Figure 21.9). The gut is then removed in the sexual region of the body, taking care of genital organs adhering to the gut. Both dissected parts are then stained, dehydrated, and mounted in Canada balsam. The taxonomically valuable material is securely maintained when histological sections are made from the segments of interest, and internal organs are reconstructed after the sections. This time-consuming procedure is not described here. A routine identification can always proceed using either live or whole-mounted oligochaetes. Taxonomically puzzling specimens, if present, could be sent to an expert for a consultation. When beginning identification of an oligochaete, it is necessary to first establish the anterior end; it is usually thicker, bears the mouth and prostomium, and has more numerous and longer chaetae on several segments. The orientation of chaetae also provides a good clue: as a rule, their ectal tip is directed backwards. The tail is usually thinner and longer, and it lacks many taxonomic characters (if not bearing gills). The next step is to distinguish the dorsal and ventral side. Hair chaetae, when present, occur only dorsally in all common taxa. The prostomium also lies dorsally of the mouth; it can be extended into unpaired proboscis, or bear a pair of simple eyespots. In contrast, genital pores, if present, are almost always ventral. Next, the segments should be counted (bearing in mind that usually the chaetae begin mostly in II but never in the I segment); the total segment number is not as important as the presence (or lack) and position of the elements of the reproductive system (clitellum, genital pores, internal reproductive organs sometimes bulging from the body, and modified ventral chaetae). The latter is most probably located in the region of the segments X–XIII. The male pores are often prominent as tubercles in the anterior portion of the clitellum, while the spermathecal pores lie often (but not always) ahead of the clitellum. The female pores are usually inconspicuous. The sperm and egg sacs can reach for several segments rearward and will be obvious as massive (in live worms, milky-white) bodies above the intestine. The body wall is externally smooth in most oligochaetes but can be armored with cutaneous papillae and/or adhered particles in some species. The tail portion is usually devoid of specific characters, except the forms with external gills. Now the characteristics of the chaetae should be noted. Are they of one sort or different (bifid, simple-pointed, and hair chaetae)? How many chaetae are in each bundle: two, one, or in variable numbers? The chaetal number and diversity are usually largest in the most anterior (preclitellar) segments while decreasing in the tail portion. The absolute length of chaetae is usually less important than their relative lengths (and sometimes also their thickness) in separate body regions. Variations of shape must be observed in the hair (smooth, pilose, serrated?) as well

Chapter | 21  Clitellata: Oligochaeta

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D

E

F

G

H

I

J

K

FIGURE 21.9  Dissection of a tubificid worm: (a) and (b) separation of anterior part of the worm containing genital segments; (c)–(e) with the worm lying on its back, cutting along the sagittal plane; (f) separation of left and right halves; (g) and (h) removal of the gut in the area of genital organs. From Martin and Aït Boughrous (2012).

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as in the dorsal bifid chaetae (pectinate with intermediate teeth, palmate?). In the ventral (and also dorsal) bifid chaetae, the relative length of the upper and lower tooth is important, as are position of the nodulus (proximal, medium, or distal) and the thickness of chaetae; all these characters can successively change along the body length. The segments bearing male and/or spermathecal pores can lack ventral chaetae, or those can be replaced with modified genital chaetae (penial or spermathecal, respectively) in the sexually mature, maturing, or postreproductive individuals, even in those devoid of a clitellum. The bundles of penial chaetae, as well as chitinous penial sheaths present in many taxa, are often hidden inside the body and visible only in the living individuals or cleared whole mounts. The digestive tract is the most apparent internal organ, traceable even in whole mounts and in immature specimens. The pharynx roof (pharyngeal pad) in II–III is more or less thickened, covered with gland tissue, and fastened to the dorsal body wall with numerous muscles. A relatively thin esophagus follows through several subsequent segments, dilating into midgut (intestine) usually in some of the anteclitellar segments. Different glands, appendages, and modified parts can occur at the pharynx, esophagus, and anterior portion of the midgut, particularly in the Enchytraeidae. Dark chloragogen tissue covers the digestive tract, usually beginning with a certain esophageal segment. The dorsal and ventral blood vessels are usually well distinguishable in whole mounts but usually of inferior value for identification. The number and modifications of commissural vessels in some segments (as dilated “hearts,” or blind appendages, sometimes covered with chloragogen tissue) can be more important. Nephridia are visible only in living or well-cleared specimens; their structure and presence/absence in the anterior segments is important, especially in Enchytraeidae. Spermathecae are one of the most distinct elements of the reproductive system in whole mounts since they are usually not hidden by the clitellum. Their structure and segmental location, position of their pores on a segment (ventral, lateral, etc.), and arrangement of sperm in the spermathecae (amorphous, in bundles, or organized into spermatozeugmata) are important. Testes, ovaries, sperm sac, and egg sac do not provide many identification characters. Separate elements of the male ducts (funnels, vasa deferentia, atria, prostatic glands, penial apparatus) are most important for the oligochaete taxonomy but are often hardly distinguishable in whole mounts. Different special methods like DIC microscopy are helpful here, as is dissection or sectioning of the gonadal segments. It would be reasonable to maintain in alcohol any unidentifiable (potentially new for the region or science), sexually mature oligochaetes discovered in routine hydrobiological surveys for further study by an expert taxonomist.

Some individuals, or at least their tail portions, could also be fixed in strong alcohol for possible DNA sequencing.

REFERENCES Adoutte, A., Balavoine, G., Lartillot, N., Lespinet, O., Prud’homme, B., de Rosa, R., 2000. The new animal phylogeny: reliability and implications. Proc. Natl. Acad. Sci. U.S.A. 97, 4453–4456. Brinkhurst, R.O., 1960. Introductory studies on the British Tubificidae (Oligochaeta). Arch. für Hydrobiol. 56, 395–412. Brinkhurst, R.O., 1984. The position of the Haplotaxidae in the evolution of oligochaete annelids. Hydrobiologia 115, 25–36. Brinkhurst, R.O., 1994. Evolutionary relationships within the Clitellata: an update. Megadrilogica 5, 109–112. Brinkhurst, R.O., Jamieson, B.G.M., 1971. Aquatic Oligochaeta of the World. Oliver & Boyd, Edinburgh. 860 pp. Bülow, T. von, 1957. Systematisch-autökologische Studien an eulitoralen Oligochaeten der Kimbrischen Halbinsel. Kiel. Meeresforsch. 13, 69–116. Caramelo, C., Martínez-Ansemil, E., 2012. Morphological investigations of microdile oligochaetes (Annelida, Clitellata) using scanning electron microscopy. Turkish J. Zoology 36, 1–14. Chapman, P.M., Brinkhurst, R.O., 1980. Salinity tolerance in some selected aquatic oligochaetes. Int. Rev. gesamten Hydrobiol. 65, 499–505. Creuzé des Châtelliers, M., Juget, J., Lafont, M., Martin, P., 2009. Subterranean aquatic Oligochaeta. Freshwater Biol. 54, 678–690. Cuadrado, S., Martínez-Ansemil, E., 2001. External structures used during attachment and sperm transfer in tubificids (Annelida, Oligochaeta). Hydrobiologia 436, 107–113. Erséus, C., 2005. Phylogeny of oligochaetous Clitellata. Hydrobiologia 535/536, 357–372. Erséus, C., Källersjö, M., Ekman, M., Hovmöller, R., 2002. 18S rDNA phylogeny of the Tubificidae (Clitellata) and its constituent taxa: dismissal of the Naididae. Mol. Phylogenet. Evol. 22, 414–422. Erséus, C., Wetzel, M.J., Gustavsson, L., 2008. ICZN rules – a farewell to Tubificidae (Annelida, Clitellata). Zootaxa 1744, 66–68. Erséus, C., Rota, E., Matamoros, L., DeWit, P., 2010. Molecular phylogeny of Enchytraeidae (Annelida, Clitellata). Mol. Phylogenet. Evol. 57, 849–858. Holmquist, C., 1978. Revision of the genus Peloscolex (Oligochaeta, Tubificidae). Zool. Scr. 7, 187–208. Hrabě, S., 1981. The freshwater Oligochaeta (Annelida) of Czechoslovakia. Acta Univ. Carol. Biol. 9, 1–166 (In Czech, with English Summary). Jamieson, B.G.M., Erséus, C., Ferraguti, M., 1987. Parsimony analysis of the phylogeny of some Oligochaeta (Annelida) using spermatozoal ultrastructure. Cladistics 3, 145–155. Jamieson, B.G.M., Ferraguti, M., 2006. Non-leech Clitellata. In: Rouse, G., Pleijel, F. (Eds.), Reproductive Biology and Phylogeny of Annelida. Science Publishers, Enfield, pp. 235–392. Jones, B., 1946. Impregnating polyvinyl alcohol with picric acid for the simultaneous staining and permanent mounting of Acarina. Proc. R. Entomol. Soc. London Ser. A Gen. Entomol. 21, 85–86. Jördens, J., Struck, T., Purschke, G., 2004. Phylogenetic inference regarding Paregodrilidae and Hrabeiella periglandulata (Polychaeta, Annelida) based on 18 rDNA and COI sequences. J. Zool. Syst. Evol. Res. 42, 270–280. Kathman, R.D., Brinkhurst, R.O., 1998. Guide to the Freshwater Oligochaetes of North America. Aquatic Resources Center, Thompsons Station, Tennessee.

Chapter | 21  Clitellata: Oligochaeta

Kent, M.L., Andrée, K.B., Bartholomew, J.L., El-Matbouli, M., Desser, S.S., Devlin, R.H., Feist, S.W., Hedrick, R.P., Hoffmann, R.W., Khattra, J., Hallett, S.L., Lester, R.J.G., Longshaw, M., Palenzeula, O., Siddall, M.E., Xiao, C.X., 2001. Recent advances in our knowledge of the Myxozoa. J. Eukaryot. Microbiol. 48, 315–413. Marian, M.P., Chandran, S., Pandian, T.J., 1989. A rack culture system for Tubifex tubifex. Aquac. Eng. 8, 329–337. Martin, P., 1996. Oligochaeta and Aphanoneura in ancient lakes: a review. Hydrobiologia 334, 63–72. Martin, P., 2001. On the origin of the Hirudinea and the demise of the Oligochaeta. Proc. Roy. Soc. London Ser. B 268, 1089–1098. Martin, P., Aït Boughrous, A., 2012. Guide taxonomique des oligochètes aquatiques du Maghreb. Belgian national focal point to the global taxonomy initiative. Royal Belgian Institute of Natural Sciences, Brussels. 186 pp. Martin, P., Granina, L., Martens, K., Goddeeris, B., 1998. Oxygen concentration profiles in sediments of two ancient lakes: Lake Baikal (Siberia, Russia) and Lake Malawi (East Africa). Hydrobiologia 367, 163–174. Martin, P., Martens, K., Goddeeris, B., 1999. Oligochaeta from the abyssal zone of Lake Baikal (Siberia, Russia). Hydrobiologia 406, 165–174. Martin, P., Martínez-Ansemil, E., Pinder, A., Timm, T., Wetzel, M.J., 2008. Global diversity of oligochaetous clitellates (“Oligochaeta”; Clitellata) in freshwater. Hydrobiologia 595, 117–127. Martínez-Ansemil, E., Creuzé des Châtelliers, M., Martin, P., Sambugar, B., 2012. The Parvidrilidae – a diversified groundwater family: description of six new species from southern Europe, and clues for its phylogenetic position within Clitellata (Annelida). Zool. J. Linn. Soc. 166, 530–558. Michaelsen, W., 1902. Die Oligochaeten-Fauna des Baikal-Sees. Verhandlungen des. Naturwiss. Vereins Hamburg 9, 43–60. Michaelsen, W., 1929. Zur Stammesgeschichte der Oligochäten. Z. für Wiss. Zool. 134, 693–710. Milbrink, G., 1978. Indicator communities of oligochaetes in Scandinavian lakes. Verhandlungen Int. Ver. für Limnol. 20, 2406–2411. Milbrink, G., Timm, T., 2001. Distribution and dispersal capacity of the PontoCaspian tubificid oligochaete Potamothrix moldaviensis Vejdovský et Mrázek, 1903 in the Baltic Sea Region. Hydrobiologia 463, 93–102.

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Naveed, M.I., 2012. Preliminary studies on aquatic Oligochaeta in and around Chennai, Tamil Nadu, India. Turkish J. Zool. 36, 25–37. Purschke, G., 2003. Is Hrabeiella periglandulata (Annelida, “Polychaeta”) the sister group of Clitellata? Evidence from an ultrastructural analysis of the dorsal pharynx in H. periglandulata and Enchytraeus minutus (Annelida, Clitellata). Zoomorphology 122, 55–66. Rodriguez, P., Reynoldson, T.B., 2011. Pollution Biology of Aquatic Oligochaetes. Springer, Dordrecht, Heidelberg, London, New York. 263 pp. Rousset, V., Pleijel, F., Rouse, G.W., Erséus, C., Siddall, M.E., 2007. A molecular phylogeny of annelids. Cladistics 23, 41–63. Semernoy, V.P., 2004. Oligochaeta of Lake Baikal. Nauka, Novosibirsk (In Russian, with English translation of descriptions of the new species). 527 pp. Siddall, M.E., Apakupakul, K., Burreson, E.M., Coates, K.A., Erséus, C., Gelder, S.R., Källersjö, M., Trapido-Rosenthal, H., 2001. Validating Livanow: molecular data agree that leeches, branchiobdellans, and Acanthobdella peledina form a monophyletic group of oligochaetes. Mol. Phylogenet. Evol. 21, 346–351. Struck, T.H., Paul, C., Hill, N., Hartmann, S., Hösel, C., Kube, M., Lieb, B., Meyer, A., Tiedemann, R., Purschke, G., Bleidorn, C., 2011. Phylogenomic analyses unravel annelid evolution. Nature 471, 95–98. Struck, T.H., 2011. Direction of evolution within Annelida and the definition of Pleistoannelida. J. Zool. Syst. Evol. Res. 49, 340–345. Timm, T., 1981. On the origin and evolution of aquatic Oligochaeta. Eesti NSV Teaduste Akadeemia Toimetised, Bioloogia 30, 174–181. Timm, T., 1987. Aquatic Oligochaeta of the Northwestern Part of the USSR. Valgus, Tallinn (In Russian, with English Summary). 299 pp. Timm, T., 2012. About the scientific names of paraphyletic taxa. Turkish J. Zool. 36, 139–140. Uzunov, J., Košel, V., Sládeček, V., 1988. Indicator value of freshwater Oligochaeta. Acta Hydrochim. Hydrobiol. 16, 173–186. Wesenberg-Lund, C., 1939. Biologie der Süswassertiere. Julius Springer, Wien. 817 pp. Wiśniewski, R.J., 1978. Effect of predators on Tubificidae groupings and their production in lakes. Ekol. Pol. 26, 493–512.

Chapter 22

Clitellata: Branchiobdellida Stuart R. Gelder Department of Science & Mathematics, University of Maine at Presque Isle, Presque Isle, ME, USA

Bronwyn W. Williams Department of Zoology, Southern Illinois University, Carbondale, IL, USA

Chapter Outline Introduction to the Branchiobdellida 551 General Systematics 551 Phylogenetic Relationships 552 Distribution (Endemic and Alien) and Diversity 552 General Biology 553 External Anatomy 553 Internal Anatomy 553 Muscular System 553 Digestive System 554 Vascular System 554 Neural System 554 Secretory Glands 554 Excretory System 555 Reproductive System 555 Physiology555 Nutrition556

INTRODUCTION TO THE BRANCHIOBDELLIDA Branchiobdellidans are leech-like, obligate ectosymbionts (Figure 22.1) primarily of astacoidean crayfish (Figure 22.2). The branchiobdellidan–host symbiosis is common among freshwater habitats of the Holarctic, including North and Central America, the Euro-Mediterranean, and East Asia (Gelder, 1999; Fard and Gelder, 2011). Branchiobdellidan taxonomic diversity varies across its Holarctic range, with more than two-thirds of the 22 total genera and approximately 140 total species found in North and Central America (Gelder et al., 2002; Gelder, 2011). The evolution and ecology of branchiobdellidans have been poorly studied relative to those of their crustacean hosts, despite the common yet unusual symbiosis. No free-living members of Branchiobdellida are known, although the

Locomotion556 Reproduction556 General Ecology and Behavior 557 Habitat Selection 557 Population Abundance 558 Sympatric Crayfish Ectosymbionts and Interspecific Competition 559 Predators and Parasites 559 Branchiobdellidan–Host Relationship 559 Collecting, Culturing, and Specimen Preparation 560 Collecting Crustacean Hosts and Sampling Branchiobdellidans560 Culturing560 Preparation for Identification 560 References561

obligate nature of the symbiosis is based only on embryonic development. Branchiobdellidans display high morphological convergence, perhaps as a function of their shared symbiotic nature. However, ecological adaptability varies widely across the taxon.

General Systematics Branchiobdellidans currently comprise a single-family order, although their taxonomic ranking has ranged from subfamily to class (see Gelder, 1996a). The modern taxonomic arrangement of Branchiobdellida was initially proposed by Holt (1965) who, by elevating the group to a single-family order, ensured recognition of their independence from the equivalently ranked oligochaetes and leeches within Clitellata. Subsequent reassessment of the taxon (Holt, 1986) resulted in arrangement of the 18 then-known

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00022-X Copyright © 2015 Elsevier Inc. All rights reserved.

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FIGURE 22.1  Cirrodrilus cirratus from Japan; whole mount preserved specimen photographed by dark-field illumination, ×40. Gelder S.R., 2006. Invertebrate Biology cover. 125 (1); with publisher permission.

FIGURE 22.2  Xironogiton victoriensis adults and juveniles on the chela joint of Pacifastacus leniusculus (signal crayfish) photographed by dark-field illumination, ×10. Souty-Grosset et al., 2006; with permission of publisher.

branchiobdellidan genera into five families within the order. Furthermore, the term “branchiobdellidan” was first applied by Holt (1986) as the common epithet for all members of the order. Morphological differences among the five families were relatively small compared to those of oligochaetes and leeches. Consequently, the families were demoted to subfamilies with Caridinophilinae being suppressed following the transfer of its monotypic Caridinophilus unidens into the Bdellodrilinae (Gelder, in preparation). This resulted in the current taxonomic arrangement consisting of an order, Branchiobdellida, containing one family, Branchiobdellidae, with four subfamilies, Branchiobdellinae, Bdellodrilinae, Cambarincolinae, and Xironodrilinae. Since publication of the species checklist of Gelder (1996a), one new genus (Gelder, 2011) and three new species (Gelder and Ohtaka, 2000; Nesemann and Hutter, 2002; Williams and Gelder, 2011) have been described.

Phylogenetic Relationships The Branchiobdellida forms a monophyly within the clitellate annelids, although relative placement within Clitellata has been historically problematic. The taxon shares several morphological characters with leeches and Acanthobdella peledina (Grube, 1851) (reviewed by Brinkhurst, 1999) argued both as homology (Purschke et al., 1993; Brinkhurst, 1994) and analogy based on a shared symbiotic lifestyle

SECTION | V  Phylum Annelida

(Brinkhurst and Gelder, 1989; Holt, 1989). Furthermore, the presence of a semiprosopore male duct in Branchiobdellida is considered a synapomorphy for B ­ ranchiobdellida and Lumbriculidae. Molecular data support a close association among branchiobdellidans, A. peledina, and leeches ­(Martin et al., 2001; Siddall et al., 2001; Erséus and Kallersjö, 2004), although sister relationships among the three remain in debate. Likewise, the phylogenetic position of the three leech-like orders with respect to members of the Lumbriculidae remains uncertain. Initial assessments of phylogeny within Branchiobdellida were largely intuitive and reflective of taxonomic rankings (Holt, 1968, 1986). Subsequent attempts at phylogenetic reconstruction using morphology (Gelder and Brinkhurst, 1990; Cardini et al., 2000; Cardini and ­Ferraguti, 2004) and molecular data (Gelder and Siddall, 2001) resulted in poor resolution and support for relationships among many of the included taxa. A recent molecular study of North American branchiobdellidans recovered strong support for groupings that were inconsistent with current taxonomy (Williams et al., 2013). Monophyly was rejected for all four subfamilies and four of seven sampled non-monotypic genera, suggesting a need for reexamination of morphological characters used to define taxonomic rankings within the Branchiobdellida and the evaluation of alternative datasets.

Distribution (Endemic and Alien) and Diversity Branchiobdellidans are found in three separate Holarctic regions: North America, Europe and adjacent ­western Asia, and East Asia. In North America, the range of ­Branchiobdellida extends from southern British Columbia across the Prairie Provinces, Ontario, and Quebec to New Brunswick in Canada and south to Costa Rica (Gelder et al., 2002; Williams et al., 2009). All 107 known North American species are historically endemic to areas either east or west of the Great Divide. Although this is also true for 12 of the 15 North American genera, Cambarincola, Sathodrilus, and Xironogiton are represented in both eastern and western regions. Diversity varies widely across the continent, with the central Appalachian area (Kentucky, Tennessee, Virginia, and West Virginia, USA) exhibiting a greater number of genera and species than in any other part of the world. The distribution of North ­American branchiobdellidans spans large portions of the temperate Nearctic and northern Neotropical zoogeographic realms (sensu Bănărescu, 1990). Branchiobdellidans in the Palaearctic realm are restricted to two disjunct regions, the Euro-Mediterranean and East Asia, each with a distinct set of historically endemic species. One genus comprising eight species occurs in the Euro-Mediterranean region, and six genera totaling 37 species occur in the East Asia

Chapter | 22  Clitellata: Branchiobdellida

region, with Branchiobdella common to both regions. All six genera within the East Asia region are found in mainland countries, whereas only locally endemic Cirrodrilus species occur on Hokkaido and northern Honshu Island, Japan. Human-mediated transportation of crayfishes and shrimp for aquaculture, sport fishing, stocking, gastronomy, research, and teaching purposes has frequently led to the accidental importation of associated branchiobdellidans, extending both host and worm distributions intraand extra-continentally. Gastronomic demands have resulted in widespread commercial translocations of crayfishes, particularly Astacus astacus (Linnaeus, 1758) and Astacus leptodactylus Eschscholtz, 1823, around Europe. Several North American crayfishes, including the signal crayfish (Pacifastacus leniusculus (Dana, 1852)), red swamp crayfish (Procambarus clarkii (Girard, 1852)), and, to a lesser extent, species of Orconectes, have been introduced to Europe and East Asia, also to meet culinary demands. Consequently North American branchiobdellidans have been found on both introduced (Gelder, 2004; Gelder et al., 2012) and some adopted native crayfishes (Ďuriš et al., 2006). The introduction of P. leniusculus has resulted in the occurrence of Cambarincola okadai Yamaguchi, 1933 and Cambarincola gracilis Robinson, 1954 in France, Xironogiton victoriensis Gelder and Hall, 1990 in France, Hungary, Italy, Spain, and Sweden, and C. okadai, X. victoriensis, and Sathodrilus attenuatus Holt, 1981 in Japan. Interestingly, despite worldwide widespread exportation of P. clarkii (Australia being an exception), the sole extra-range report of Cambarincola mesochoreus Hoffman, 1963 is from northern Italy (Gelder et al., 1994, 1999). Holtodrilus truncatus (Liang, 1963) now occurs on Honshu Island, Japan following its introduction on Chinese shrimp, Neocaridina spp. (Niwa et al., 2005; Ohtaka et al., 2012). Because these shrimp are popular bait for freshwater sport fishing, it is likely that H. truncatus will quickly become widespread across the Japanese Islands.

GENERAL BIOLOGY Adult branchiobdellidans range from 0.8 to 12 mm in length. The head is distinct from the body and, by definition of the order, does not have a prostomium. The body is usually rod or spindle-shaped (terete), although some species have a characteristic pyriform or flask-shape with either ventral or dorso-ventral flattening. Segment number is constant in all branchiobdellidan species consisting of a head and 11 body segments. A single cerebral ganglion and 14 double-paired ganglia are arranged along the ventral nerve cord. The excretory, closed circulatory, and reproductive systems have each undergone reduction from the basic annelid metameric arrangement.

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External Anatomy In branchiobdellidans, a distinct head is followed by body segments designated 1 to 11 with Arabic numerals (Figure 22.3). In contrast, oligochaetes and leeches lack a head and, by convention, their segments are numbered with Roman numerals. Oligochaete numbering ignores the prostomium and starts with the peristomium (segment containing the mouth), whereas in leeches the prostomium is recognized as segment I and the peristomium as segment II. Comparative anatomical studies of clitellates have not always recognized these different but accepted numbering systems, and the nonconcordant interchangeability of Arabic and Roman numerals used for branchiobdellidans has frequently confused the location of homologous organs. As branchiobdellidans do not have a prostomium, the oligochaete system of segment numbering using Roman numerals is the most appropriate for comparative work with other clitellates (Figure 22.3). The anterior portion of the head consists of a peristomium divided into a dorsal lip often with lobes or tentacles (te), paired lateral lobes (pll), and a ventral lip (vl) usually with a median emargination. The remainder of the head rarely shows any apparent segmentation. Each body segment is divided into a major anterior and a minor posterior annulus, although in a few species the major annulus is subdivided, resulting in three annuli per segment. Dorsal ridges (dr) may occur on the major annulus, some supporting digitiform or multi-branched appendages (dp). Lateral segmental lobes, found on segments 8 and 9, vary from conical to plate-like extension (ll). Median apertures consist of the genital pore (gp) ventrally-located on segment 6, the spermathecal pore (spp) ventrally-located on segment 5, and one or two nephridial pores dorsally-located on segment 3 (np). The clitellum’s (cl) thickened epidermis covers segments 5 to 7, containing both isolated and groups of gland cells. Segment 11 is modified into the posterior, discshaped attachment organ often referred to as the sucker (pa) (Figure 22.3). The external surface of branchiobdellidans is covered by a thin, flexible cuticle that also lines the mouth, pharynx, and anus.

Internal Anatomy Muscular System Body musculature consists of outer circular and inner longitudinal layers. In the head these layers, together with radial muscles, are modified and attach to the oral region and pharynx to produce complex feeding motions. Longitudinal muscles from the posterior three segments terminating in various parts of the posterior disc enable the body to rotate around the attachment site (Schmidt, 1903). Dorsal ridges are permanent features produced by short lengths of modified longitudinal muscles called supernumerary muscles in

SECTION | V  Phylum Annelida

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FIGURE 22.3  Lateral view of generalized branchiobdellidan to show most external features. P, peristomium; H, head; 1–11, body segments; I–XV, oligochaete segment numbering system; aa, anterior attachment organ; cl, clitellum (shaded); dp, dorsal projections; dr, dorsal ridge; gp, genital pore; ll, lateral segmental lobes; np, anterior nephridial pore; pa, posterior attachment organ (sucker); pll, peristomial lateral lobe; spp, spermatheca pore; te, dorsal peristomial lip tentacle; vl, ventral peristomial lip. Gelder, in preparation; with permission. Copyright © 2015 Dr Stuart R. Gelder. Published by Elsevier Inc. All rights reserved.

FIGURE 22.4  Lateral view of generalized branchiobdellidan to show longitudinal supernumery muscles (orange), alimentary canal (green), and circulatory system (red). A, anus; adbv, anterior dorsal blood vessel; e, esophagus; gs, gastric sinus; i, intestine; j, jaw; lsbv, lateral segmental blood vessel; m, mouth; op, oral papillae; pdbv, posterior dorsal blood vessel; ph, pharynx; phs, pharyngeal sulcus; sm, supernumery muscles; st, stomach; vbv, ventral blood vessel. Gelder, in preparation; with permission. Copyright © 2015 Dr Stuart R. Gelder. Published by Elsevier Inc. All rights reserved.

a major annulus (Figure 22.4, sm). Valvassori et al. (1994) found body wall muscle cells comprising a medulla containing the nucleus and cytoplasm and a cortex of contractile fibers ranging from a complete cylinder (circomyarian) to an incomplete (polyplatymyarian) arrangement.

Digestive System The alimentary canal consists of a mouth (m) surrounded by oral papillae (op) followed by a pair of sclerotized jaws (j) in the anterior pharynx then one to two sulci (phs), a short esophagus (e), stomach (st), intestine (i), and an anus (a) opening dorso-medially in segment 10. The esophagus, stomach, and intestine are lined with a ciliated epithelium and entry and exit of food is controlled by esophageal and anal sphincter muscles, respectively.

Vascular System The vascular system consists of dorsal (adbv, pdbv) and ventral (vbv) longitudinal vessels that are connected by four paired lateral branches (lsbv) in the head and a single pair of branches in segments 1, 7, and 10, respectively. The dorsal vessel surrounds the stomach and intestine in segments 3 to 7, forming a gastric sinus (gs) in direct contact with the base of the gut epithelium. Blood containing hemoglobin and amebocytes is circulated through the system by

dorsal vessel wall peristaltic contractions in segments 1 to 3, which forces the fluid anteriorly (Figure 22.4).

Neural System Moore (1895) described in detail the central nervous system (Figure 22.5) of Bdellodrilus illuminatus (Moore, 1894), which consists of a dorsal cephalic ganglionic (cg) mass (brain) in the segment behind the peristomium with a pair of circumesophageal connective nerve cords arising laterally that pass ventrally around the pharynx. The nerve cords (vnc) then come together, running closely parallel to each other mid-ventrally and deflecting around the spermathecal duct and bursa before ending in segment 11. Two pairs of adjacent ganglia are located in each segment on the nerve cords, with those in segments 9 to 11 condensed into nearly a single mass. The details are largely the same for European Branchiobdella species except that the brain appears to be in, rather than behind, the peristomium.

Secretory Glands Branchiobdellidans attach themselves to the substrate by secretions from adhesive glands (aag) located laterally along the head and opening on the ventral side of the peristomial ventral lip (aa), and from glands in segments 9 and 10 (pag) opening onto the posterior disc (pa) (Figures 22.3 and 22.5). Histochemical characterization of the secretions show that the

Chapter | 22  Clitellata: Branchiobdellida

FIGURE 22.5  Lateral view of generalized branchiobdellidan to show nervous (black), excretory (green), adhesive (fawn), female (purple) and male (blue) reproductive systems. Aag, Anterior attachment gland; b, bursa; cg, cerebral ganglia; ga, glandular atrium; ma, muscular atrium; n, nephridia; np, nephridial pore; o, ovum; od, ovary duct; pag, posterior attachment gland; pg, prostate gland; s, spermatozoa; sg, segmental ganglia; spb, spermathecal bulb; spd, spermathecal duct; vd, vasa deferentia; vnc, ventral nerve cord. Gelder, in preparation; with permission. Copyright © 2015 Dr Stuart R. Gelder. Published by Elsevier Inc. All rights reserved.

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FIGURE 22.6  Schematic longitudinal section through the bursa to show a generalized eversible penis in a retracted position. B, bursa; E, epidermis; gp, genital pore; ma, muscular atrium; p, penis (dotted); ps, penial sheath. Gelder, in preparation; with permission. Copyright © 2015 Dr Stuart R. Gelder. Published by Elsevier Inc. All rights reserved.

glands are composed of two types of pyriform secretory cells (Gelder and Rowe, 1988), although only the adhesive gland cell ducts open onto the attachment surfaces (Weigl, 1994).

Excretory System In branchiobdellidans, nephridia (n) are reduced to two pairs: an anterior pair located in segments 2 to 4 discharging urine via the nephridrial pore (np) on the dorsal surface of segment 3, and a posterior pair in segment 8 opening separately onto the segment’s surface. The anterior pair is asymmetrically arranged with the nephridial tube complex on one side in segments 2 and 3 and the opposite in segments 3 and 4 (Figure 22.5).

Reproductive System Reproductive systems consist of female organs that include a pair of ovaries in segment 7 and a spermatheca or sperm receptacle in segment 5, and male organs in segments 5 and 6. Ovigenesis occurs in the coelom of segment 7 and mature ova are released to the exterior through a pair of short oviducts (od) in the segment’s body wall. A spermatheca, when present, consists of a glandular bulb (spb) connected to a muscular duct (spd) that opens medially onto the ventral surface of the segment. A pair of testes is located in both segment 5 and 6 (only one pair in segment 5 is present in Branchiobdella), and spermiogenesis (s) occurs in the coelom of those segments. In a review of spermatozoa ultrustructure from 25 branchiobdellidan species representing each subfamily, Cardini and Ferraguti (2004) found shared characters previously thought to be indicative of either oligochaetes or leeches. In addition, the morphological diversity of branchiobdellidan spermatozoa was greater than that found in either the leeches or oligochaetes. Differences within Branchiobdellida are largely intergeneric, whereas intrageneric variation was primarily organelle dimensions (Ferraguti, 2000). Two pairs of vasa efferentia (in segments 5 and 6) each merges into a vas deferens (vd). The two vasa

FIGURE 22.7  Schematic longitudinal section through the bursa to show a generalized protrusible penis in retracted a position; labels as in Figure 22.6. Gelder, in preparation; with permission. Copyright © 2015 Dr Stuart R. Gelder. Published by Elsevier Inc. All rights reserved.

deferentia enter the single median glandular atrium (ga) in segment 6. When present, a prostate gland (pg) forms a protuberance or diverticulum from the glandular atrium. The glandular atrium connects with a muscular atrium (ma) and in turn a penis, which is encompassed by a bursa (b). Most penes (p) are surrounded by a penial sheath (ps), and are either eversible (Figure 22.6) or protrusible (Figure 22.7), although a few species have a penis that shows varying degrees of reduction. The bursal atrium opens medially on the ventral surface of segment 6 through a genital pore (gp). The terms, “glandular atrium” (=spermiducal gland) and “muscular atrium” (=ejaculatory duct) are the traditional names used in clitellate descriptions. As such, for consistency these terms have been reintroduced to the branchiobdellidans rather than continuing use of the archaic terms shown in parentheses above and found in Holt (1986).

Physiology Berry and Holt (1959) experimentally showed that Xironodrilus formosus Ellis, 1919 and Xironogiton instabilis (Moore, 1894) had similar tolerances to low oxygen tensions, although survival across a range of high temperatures (>22  °C) differed. Under laboratory conditions, as water temperature falls below 7  °C, Cambarincola ­fallax Hoffman, 1963, C. mesochoreus, X. instabilis, and

SECTION | V  Phylum Annelida

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X. victoriensis have difficulty moving and attaching to the substrate and will die after several hours if maintained at 300 spp. The common name for the Pisauridae is nursery-web spiders, but the semi-aquatic species are often called fishing spiders or raft spiders. There are no unique morphological synapomorphies for the Pisauridae (Santos, 2007), and membership has been volatile, with many genera originally

SECTION | VI  Phylum Arthropoda

FIGURE 25.2  A female Pirata (Lycosidae) carrying an egg sac. Photograph by Steve Marshall.

included now moved to other families. In a recent cladistic analysis, Santos (2007) tested hypotheses about composition, monophyly, and behavioral evolution in the family using Ancylometes (Ctenidae) and Trechalea (Trechaleidae) to root the tree. He concluded that capture of prey on the water’s surface evolved independently in Ancylometes, Trechaleidae, and three times within the Pisauridae (in Nilus, Dolomedes and Tinus + Thaumasia); however, Santos also suggested that semi-aquatic foraging may have been basal within the Pisauridae and was repeatedly lost. Lack of information about the behavior of many pisaurids makes testing this hypothesis difficult. Among the semi-aquatic genera, the cosmopolitan Dolomedes is the best studied genus. There is some ecological information for Nilus (=Thalassius), which is found in the Old World tropics, but almost nothing has been published about the ecology of the mostly New World Tropics genera Tinus and Thaumasia (Santos, 2007). As for species of Pirata (Lycosidae), Dolomedes species vary in how closely tied they are to water. Dolomedes triton (Walckenaer) is very aquatic in its habits, but Dolomedes tenebrosus Hentz wander considerable distances from water (Carico, 1973). Graham et al. (2003) used floating and terrestrial pitfall traps to determine the distribution of D. triton on and around a pond in Alberta. Of all specimens, 95% were on the pond, and the remaining 5% were 2000 spp. Almost all crab spiders are sit-andwait predators that hunt in flowers, foliage, or leaf litter. Many species have been collected from phytotelms such as bromeliads and pitcher plants. Misumenops nepenthicola (Pocock) is associated with Nepenthes pitcher plants in Southeast Asia. It has been observed crawling into the water of a pitcher plant, carrying a bubble of air on its abdomen, to capture mosquito larvae (http://www.bbc.co.uk/ nature/life/Crab_spider#p0038stg). It is not clear whether M. nepenthicola is unique among phytotelmata-associated crab spiders in foraging below water. Chua and Lim (2012) determined the effects of M. nepenthicola and another crab spider typically found on Nepenthes, Thomisus nepenthiphilus (Fage), on survivorship of dipteran larvae in the pitchers. They found that declines in larval abundance occurred only in the presence of M. nepenthicola, and that T. nepenthiphilus was unable to maintain body weight when only aquatic prey were available.

This family of 16 genera and 119 species is found in South and Central America. It was originally considered part of the Pisauridae, and at least some species are similar in behavior to semi-aquatic pisaurids. Trechaleids are mostly active at night. Juveniles tend to occupy the riparian zones of streams, whereas the adults are found on the rocks in streams. da Silva et al. (2005) observed predatory behavior of Trechaleoides (previously Trechalea) biocellata (MelloLeitão) in a stream in Brazil. Like pisaurids, these spiders keep their hind legs on solid objects and front legs resting on the water’s surface, lunging out to grasp insects caught in the surface (Figure 25.4). The courtship behavior of Paratrechalea is unusual in that males wrap and present prey to females prior to copulation. This giving of a “nuptial gift” is known only for a few other species of spiders, all within the Pisauridae (the nonaquatic P. mirabilis, Pisaura lama Bösenberg and Strand and Perenethis fascigera [Bösenberg and Strand, 1906]). Costa-Schmidt et al. (2008) observed the mating behavior of Paratrechalea azul Carico and Poecilotheria ornata in Brazil. Behavior of the two is almost identical. A male interested in mating first spins a small sperm-web, deposits a drop of sperm, and charges his palps. He then captures a prey item (usually an aquatic insect) and wraps it in silk. With the package in his chelicerae he walks about apparently randomly until he encounters silk lines deposited by a female. Once the female is perceived, the male taps the female with his forelegs and then leans far toward her with his legs anchored behind his prothorax, and presents his nuptial gift. Brum et al. (2011) determined that chemicals produced by the male and incorporated into the silk induce the female to grasp the gift with her chelicerae. Once the female has grasped the package, the pair touch palps, the female rotates her abdomen to one side to expose her genital opening, and the male mounts the female while maintaining

FIGURE 25.4  A female Trechalea tirimbina Silva and Lapinski (Trechaleidae) hunting at the water’s edge in Costa Rica. Photograph by Witold Lapinski.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

contact with the gift with his legs III. After inserting one palp and injecting some sperm, the male inserts his other palp on the opposite side (presumably filling both the left and right spermatheca of the female). This is often repeated several times on alternate sides, with the male returning to face the female and biting/releasing the nuptial gift each time. Typically the mating ended once the female began to behave aggressively toward the male, but the authors observed no post-copulatory cannibalism. The female usually retained the nuptial gift but in a few cases the male departed with it. Costa-Schmidt et al. (2008) suggested that giving of nuptial gifts may be a homology between Trechaleidae and Pisauridae; however, this hypothesis does not fit the phylogeny of Santos (2007) parsimoniously. In at least one Trechalea species, the female carries her hatchlings on her abdomen until they become capable of fending for themselves.

ACARI: PARASITIFORMES The Parasitiformes makes up one of the two major lineages of arachnids collectively called “mites” (see initial section for discussion of acarine diphyly). About 25% of all mite species belong to the Parasitiformes, which is also known as Anactinotrichida in reference to the non-birefringent nature of their setae when under polarized light (the lineage of mites with birefringent setae being the Acariformes, or Actinotrichida). The name Parasitiformes reflects the inclusion of ticks (order Ixodida) within this group. There are three

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other orders in the Parasitiformes: Opilioacarida, Holothyrida, and Mesostigmata. Almost all Parasitiformes, and all freshwater species, are in the Mesostigmata, which includes ∼80 families and >12,000 spp. (Krantz and Walter, 2009). The vast majority of parasitiform mites are terrestrial, living in soil, on plants, or as symbionts of larger-bodied terrestrial animals. Only a few members of the Mesostigmata live in freshwater habitats. Therefore, we focus on the morphology of this group of parasitiformans below.

Morphology of Mesostigmata Like all other mites, Mesostigmata have two tagmata, an anterior gnathosoma bearing the palps, chelicerae and mouth opening, and a posterior idiosoma bearing the legs and containing the reproductive, digestive, and excretory systems (Krantz and Walter, 2009). Mesostigmata lack eyes. Their chelicerae are 3-segmented with the distal two segments forming a pincer-like chela in most free-living and all freshwater species (Figure 25.5). In males of some taxa, the chelicerae are modified to transfer sperm. A small bifurcating protuberance called the tritosternum is present ventrally on the idiosoma, where it joins with the gnathosoma (Figure 25.5). The anal opening is small as befits fluid-feeders and is located ventrally toward the end of the idiosoma. Variously developed ventral and dorsal shields are present, depending on the taxon. Males are usually more thoroughly sclerotized than females, as the idiosomas of the latter have to expand

FIGURE 25.5  Ventral view of a mesostigmatan mite, showing three-segmented chelate chelicerae (left) and bifurcate tritosternum (right). Photographs by H.C. Proctor.

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FIGURE 25.7 Lateral view of a semi-aquatic Cheiroseius (Blattisociidae), with inset showing broad peritreme with mictrotrichiae. SEMs by David Walter. FIGURE 25.6  Lateral view of a terrestrial mesostigmatan mite, with inset showing stigma and peritreme SEMs by David Walter.

in order to accommodate eggs. A single pair of respiratory openings called stigmata are present dorsolaterally near the insertions of the posterior legs (the mid-body placement of these openings gives the order Mesostigmata its name). Each stigma is associated with a peritreme, a shallow channel running anteriorly and posteriorly along the side of the mesostigmatan (Figure 25.6). The peritremes of most mesostigmatans are circular in cross-section, with only a narrow slit running along the external surface to connect the peritrematal duct to the outside atmosphere (as if one had taken a long thin strip from a tube). In the semi-aquatic species of Blattisociidae, a cross-section of the peritreme appears to be semi-circular; so the surface area available for gas exchange in these mites is much larger than in fully terrestrial mesostigmatans (Hinton, 1971). A lining of microtrichiae presumably helps to keep the layer of air attached to the blattisociid’s peritreme (Figure 25.7, inset). Legs have six free segments (coxa, trochanter, femur, genu, tibia, tarsus). The tarsus usually bears paired claws and a median pulvillus that is well-developed in some freshwater species (e.g., Cheiroseius). Ontogeny in mesostigmatans typically involves the following stages: egg, six-legged larva, eightlegged deutonymph, and adult.

Biology of Freshwater Mesostigmata Freshwater species of Mesostigmata have been reported from the Blattisociidae (Cheiroseius, Platyseius, and Cheiroseiulus spp.) and Dithinozerconidae (Caminella peraphora Krantz and Ainscough). Although the overall diversity of freshwater mesostigmatans is low (probably fewer than 30

known species), they occupy a moderate diversity of habitats. Cheiroseius spp. have been collected from saturated soil, phytotelmata (e.g., tree holes) and, for many Platyseius spp., in vegetated margins of large and small wetlands and mosses in streams (Lindquist, 2003; Krantz and Walter, 2009). Cheiruseiulus reniformis Evans and Baker is known from phytotelmata in epiphytes (Lindquist, 2003). C. peraphora occurs in wet moss in mountain streams in Oregon. Platyseius italicus (Berlese) can be found in great numbers in sewage filter-beds (Learner and Chawner, 2003), which perhaps can be granted status as “flowingwater” habitat. One species in the Phytoseiidae, Macroseius biscutatus Chant, Denmark and Baker, has been collected from water-filled Sarracenia pitcher plants in the southeastern United States (Muma and Denmark, 1967). Other than a rough idea of habitat, virtually nothing is known of the biology of freshwater Mesostigmata. Locomotion is by crawling; no species are known to swim. Diets of freshwater mesostigmatans have not been well investigated, although some Cheiroseius and Platyseius are known to eat mosquito eggs (Smith, 1983). It is likely that many species feed on nematodes and/or soft-bodied microarthropods, as is common in free-living terrestrial Mesostigmata. Muma and Denmark (1967) reared the pitcher-plant mite M. biscutatus through its life cycle by feeding them nematodes. Development was poor if the diet included only the soft-bodied mites Sarraceniopus hughesi (Hunter and Hunter) (Astigmatina: Histiostomatidae). Adult female Cheiroseius and Platyseius sometimes disperse by phoresy on insects, in particular flies of the families Tipulidae, Ceratopogonidae, and Culicidae (Lindquist, 2003; Krantz and Walter, 2009). Caminella peraphora females often bear a ring of amorphous material around their opisthosomas, connected to a

Chapter | 25  Subphylum Chelicerata, Class Arachnida

sac of similar material resting on their dorsums, the product of an unusual sperm-transfer system (Walter and Proctor, 2013). Prior to sperm transfer, the male mounts the female and faces the opposite way. The female produces a mass of “ring” material from under her genital shield. The male presses a spermatophore directly from his genital opening into the amorphous material near her anal opening. The spermatophore is enveloped in the material and somehow a hollow tube forms in the material leading from the spermatophore to the female’s genital shield. The male returns to his original position and uses his legs to manipulate the still-extruding ring material along the sides of the female up onto her dorsum. The material eventually hardens to form a double-chambered cell on the female’s back. Embedding of the spermatophore in the ring material may be an adaptation to prevent water damage to the ejaculate.

ACARIFORMES: FRESHWATER SARCOPTIFORMES Acariform mites living in fresh water can easily be distinguished from parasitiforms by the presence of genital papillae (also known as genital acetabula when flat and plate-like) (Figure 25.8) and the absence of stigmata and peritremes (Figures 25.6–25.7).

Oribatida Excluding Astigmatina The Oribatida includes the cohort Astigmatina; but because astigmatans differ strongly from other oribatids morphologically, developmentally, and ecologically, we discuss

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them separately in a subsequent section. The information in this section refers to those non-astigmatan oribatids that are obligatory inhabitants of fresh water. This includes representatives from 10 families (Behan-Pelletier and Eamer, 2007; see Table 25.1). Because their exoskeleton is relatively resistant to decomposition, terrestrial oribatids are often found in aquatic samples after having tumbled into the water and drowned, and so one should not assume that an oribatid found in a freshwater sample is actually aquatic.

Morphology With few exceptions (none of them being species that inhabit freshwaters), adult oribatid mites are well armored, with the only soft integument being located in regions of articulation. This heavy sclerotization gives them one of their common names: “beetle mites.” In contrast, juvenile oribatids are often mostly soft-bodied and can look quite different from their adult counterparts (e.g., Zetomimus, Figure 25.9). They reach a maximum of about 1.5 mm in length. Aquatic oribatids lack paired eyes but some have a lightly sclerotized median area on the anterior prodorsum that is sensitive to light (the lenticulus, Figure 25.10). Gas exchange in aquatic species may be through unsclerotized integument or sclerotized integument punctured with many fine pores (porose areas) (Norton and Behan-Pelletier, 2009). When under water, adult Hydrozetes (Hydrozetidae) often have a silvery appearance due to a fine layer of air trapped by a plastron-like arrangement of micropapillae. This layer can be maintained through a huge range of air pressures (−98 KPa to +550 KPa, Behan-Pelletier and

FIGURE 25.8  Ventral view of a freshwater acariform mite (Prostigmata: Halacaridae) showing (a) absence of the stigmata and peritremes characteristic of the Mesostigmata, and (b) numerous genital papillae and their porous structure. SEMs by H.C. Proctor.

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FIGURE 25.9  Zetomimus francisi (Habeeb) (Zetomimidae) showing how the poorly sclerotized and highly setose nymph (left) contrasts with the armoured, smooth-bodied adult (right). Scale bars = 100 μm. SEMs by David Walter.

FIGURE 25.10  Hydrozetes spp. (Hydrozetidae) have a light sensitive area on the prodorsum called the lenticulus (arrow). Scale bar = 100 μm. SEM by David Walter.

Eamer, 2007). The unsclerotized juvenile Hydrozetes lack this physical gill, and presumably exchange gases through their thin integument (Krantz and Baker, 1982). Most but not all oribatids have a distinctive pair of mechanosensory setae called trichobothria on the gnathosoma adjacent to its articulation with the idiosoma. The bases of the trichobothria (the bothridia) are deep and cup-shaped (Figure 25.11). Some freshwater oribatids have the trichobothria reduced or absent. All aquatic oribatids are particulate feeders, and have very large anal openings relative to fluid-feeding mites in order to allow defecation of large fecal pellets (Figure 25.12). They have chelate, toothed chelicerae for biting off chunks of food. Adult and tritonymphal oribatids have three pairs of genital papillae immediately flanking the genital opening. Proto- and deutonymphs have one and two pairs, respectively, located in the provisional genital area. Both sexes have a tube-like

FIGURE 25.11  Prodorsum of an Ameronothrus sp. (Ameronothridae) with arrows indicating the deep bases (bothridia) of the club-shaped trichobothrial setae. SEM by H.C. Proctor.

FIGURE 25.12  Ventral view of a trhypochthoniid mite showing the genital opening (g) and large anal opening (a). SEM by H.C. Proctor.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

structure that can be retracted within the body or extended through the genital valves, and that is used for spermatophore deposition (by males) or oviposition (by females). Oribatids go through six developmental stages: egg, larva, protonymph, deutonymph, tritonymph, and adult.

Diversity and Biology of Freshwater Oribatida Members of 15 genera in 10 families of Oribatida are truly freshwater species (where “truly” means that members of the species can feed, locomote, and mate when on or under water) (Behan-Pelletier and Eamer, 2007; Table 25.1). The proportion of a given family’s diversity that is freshwater varies, as does whether they are fully subaquatic or semi-aquatic. All 20 or so species in the Hydrozetidae are subaquatic in ponds, lakes (Krantz and Baker, 1982), and streams (H.C. Proctor, personal observation). Numerous freshwater inhabitants are scattered through the families Trhypochthoniidae and Ameronothridae. A number of other taxa are found in aquatic systems but appear equally at home in damp moss and other sodden vegetable matter. These “amphibians” include Limnozetidae, some Trhypochthoniidae, and two genera of Malaconothridae. Zetomimidae are also associated with shallow waters, but anecdotal literature suggests that they may live on the water’s surface rather than beneath it. Many species in the Camisiidae and Mycobatidae inhabit soggy areas in moss and litter next to water bodies, but they are rarely completely immersed. At least three species of Oribatida in the Ameronothridae inhabit rock pools that evaporate completely. When this happens, the mites remain in the arid silt for days or weeks until the next rains arrive (Norton et al., 1996). Two species of Mucronothrus in the family Trhypochthoniidae are benthic in cold running water (Norton et al., 1988). Most other trhypochthoniids are terrestrial, with a few species being amphibious in water and in sodden littoral vegetation. For these oribatids, it seems reasonable to suggest a mode of invasion from terrestrial to amphibious to obligatorily aquatic. Mucronothrus nasalis (Willmann) has been found in streams, springs, and the bottoms of very cold lakes throughout the world. As M. nasalis has no way of traveling long distances over land or sea, this distribution suggests that its ancestor invaded water before the break-up of Pangaea 200 million years ago (Norton et al., 1988). In a survey of springs in the Bavarian and Italian Alps, Schatz and Gerecke (1996) found that M. nasalis occupied only permanent springs, where it made up about 45% of all oribatids collected. Many of the oribatids that they found were clearly accidental specimens from nearby terrestrial and arboreal habitats; however, Platynothrus peltifer (Koch) (Camisiidae) was also extremely common in their samples (34% of individuals). They concluded that it, too, was a true subaquatic denizen of springs. Unlike M. nasalis, P. peltifer occasionally occurred in intermittent springs.

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Like their land-locked relatives, aquatic oribatids feed primarily on fungus and decaying plant matter. Hydrozetes are often found inside stems of aquatic plants and on damaged and deteriorating leaves, but it is not known whether their feeding causes the damage or if they simply exploit decaying wounds. Norton et al. (1988) examined the gut contents of the widely distributed spring-dwelling species M. nasalis. Adults and immature mites from Canadian, French, Australian, and New Zealand populations contained a mixture of filamentous algae, diatoms, hyphae, decayed vascular plants, and the occasional bit of arthropod cuticle, suggesting that these mites graze on the periphyton growing on benthic substrates. An interesting exception to the typical vegetable diet is shown by a species of freshwater ameronothrid that includes bdelloid rotifers in its meals (Norton et al., 1996). These mites have no obvious adaptations for capturing moving prey, and it is likely that the hapless rotifers are engulfed along with detritus as the mites graze. No known species of oribatid has legs modified for swimming. Aquatic oribatids of the family Hydrozetidae do not look much different from terrestrial oribatids and move at the same sedate walking pace as they crawl among water weeds, but Hydrozetes makes use of air in a novel way. Newell (1945) noted that H. lacustris (Michael) and H. petrunkevitchi Newell often contain bubbles of gas in their midguts; and that by releasing their grip on the substrate, they would float up to the water’s surface much more rapidly than they could crawl—rather like underwater ballooning. Newell noted that agitating mites would induce the formation of these bubbles, suggesting that this may be a form of rapid escape response. Levitation also occurred whenever ambient light conditions fell low enough. Newell suggested that levitation would allow animals that had been dislodged from their normal home on plants in the photic zone to escape starvation and death by the anoxia that would occur at the dark bottom of a pond. Almost all sexually reproducing oribatids transfer sperm via spermatophores deposited on a substrate, typically with little to no contact between the sexes (Walter and Proctor, 2013). The spermatophores of terrestrial oribatid mites appear to have some osmoregulatory ability (Alberti et al., 1991), suggesting that this may also be a pre-adaptation for invasion of aquatic systems in oribatids. However, the relatively high proportion of parthenogenetic versus sexual taxa of oribatids in aquatic systems may indicate that freestanding spermatophores suffer physiological stress when submerged (Behan-Pelletier and Eamer, 2007).

Acariformes: Freshwater Sarcoptiformes— Astigmatina The Astigmatina, or Astigmata, are a cohort within the Oribatida. Morphologically and developmentally they appear to be paedogenetic versions of oribatids, in that they

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FIGURE 25.14  Lateral view of an freshwater histiostomatid mite with the inset image showing the serrate chelicerae characteristic of the Histiostomatidae. SEM by H.C. Proctor.

FIGURE 25.13  Ventral view of a female Schwiebea (Acaridae), with the inset image showing the genital area bearing two pairs of genital papillae. Photograph by H.C. Proctor.

are soft-bodied like oribatid nymphs and reach reproductive maturity with only two rather than three pairs of genital papillae (OConnor, 2009) (Figure 25.13). However, they also have some very non–oribatid-like features, including the presence of a highly modified phoretic deutonymphal stage in most free-living species, direct transfer of sperm via intromission, and absence of the prodorsal trichobothrial setae so characteristic of other oribatids.

Morphology Aquatic astigmatans are teardrop-shaped or slightly flattened, poorly sclerotized, and often pale in color. They reach a maximum of about 1 mm in length. Like other oribatids, they have a large anal opening associated with their consumption of particulate food (Figure 25.13). They lack obvious respiratory structures (hence their name) and presumably exchange gases directly through their cuticle. Most aquatic astigmatans have robust two-segmented chelicerae, but members of the family Histiostomatidae have serrate chelicerae (Figure 25.14) and whip-like palps used for sweeping up and separating small particles from the substrate. They lack eyes and obvious light-sensing structures, and also lack the pair of sensory trichobothria characteristic of non-astigmatan oribatids. As mentioned above, at maturity, astigmatans have only two pairs of genital papillae (Figure 25.13). In the Algophagidae, which includes some freshwater species, the papillae have been either modified or lost, and their ion-exchange function is achieved by large plate-like structures associated with the leg bases (Fashing and Marcuson, 1996). Some freshwater astigmatans have their hind legs modified for swimming (see below).

FIGURE 25.15  Deutonymphal astigmatan mite (Acaridae: Histiogaster) with arrow indicating the sucker plate used to phoretically attach to a transport host. SEM by H.C. Proctor.

Males have a sclerotized intromittent organ (the aedeagus) used for transferring sperm to the female’s porelike secondary genital opening, located posterodorsally on the idiosoma. The female’s primary genital opening, through which eggs are laid, is located ventrally on the idiosoma between or just behind the hindleg coxal bases (Figure 25.13). Astigmatans lack the tubular ovipositor characteristic of other female oribatids. Their life cycle includes a maximum of six stages: egg, larva, protonymph, phoretic deutonymph (not always expressed), tritonymph, and adult. The phoretic deutonymphs are also called hypopodes (or “hypopi,” singular hypopus). They are strongly heteromorphic to the other stages, being dorsoventrally flattened, often well sclerotized, lacking mouthparts, and having setae near the anal opening modified as suckers to hold on to the phoretic host (typically an insect) (Figure 25.15).

Chapter | 25  Subphylum Chelicerata, Class Arachnida

Diversity and Biology of Freshwater Astigmatina Although their unsclerotized integument would seem well suited for aquatic gas exchange, there are few families of astigmatans with freshwater species: Histiostomatidae, Algophagidae, and Acaridae. The Histiostomatidae and Algophagidae are essentially all aquatic or semi-aquatic, with even the most terrestrial of species requiring a thin film of water to wade through. Some algophagids are marine rather than freshwater (OConnor, 2009). The Acaridae are mostly terrestrial (including several important pests of stored products), but the family includes a number of freshwater genera. Members of the Histiostomatidae, Acaridae, and Algophagidae are common inhabitants of phytotelmata (OConnor, 1994). Histiostomatids and acarids have also been found as associates of freshwater fish and leeches (Walter and Proctor, 2013). Most aquatic astigmatans have also maintained diets similar to those of their terrestrial progenitors, feeding on fungus, decaying plants, and bacteria. Fashing (1994, 1998) compared the feeding methods of three treeholeinhabiting species. Naiadacarus arboricola Fashing (Acaridae) uses its powerful chelicerae to bite off chunks of decaying leaves and ingests them whole. Algophagus pennsylvanicus Fashing and Wiseman (Algophagidae) uses its more gracile mouthparts to scrape fungus from leaf surfaces. Hormosianoetus mallotae (Fashing) (Histiostomatidae) has serrate chelicerae that filter suspended bacteria and bits of detritus that it stirs up using its whiplike palps. Judging by their gut contents, hyadesiids and other species of freshwater and marine algophagids are algivores (OConnor, 1994). There are numerous records of aquatic acarids and histiostomatids, and even oribatids, collected from the bodies of fish, both wild and farmed, and one report of these mites inhabiting the integument of leeches (Olmeda et al., 2011; Walter and Proctor, 2013). Although this may appear to represent parasitism, in the case of the leech and in many of the fish, the “host” animals were diseased, and it appears more likely that the mites were feeding on bacteria and fungus associated with necrotic flesh. Among the Astigmata, only a few species in the genera Zwickia and Creutzeria (Histiostomatidae) have exchanged walking for swimming (Fashing, 2004). In a Creutzeria species from Australian pitcher plants, nymphs, males, and females have highly modified legs bearing long whip-like setae at their tips. The male bears a remarkable likeness to a planktonic copepod nauplius. However, this crustacean-like morphology is the result of a compromise between sex and swimming. The male uses his strong forelegs to grasp the female while mating, and the second pair of legs are used both for swimming and for holding onto a substrate when the first pair are connubially occupied.

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ACARIFORMES: PROSTIGMATA: OVERVIEW AND FRESHWATER TAXA EXCLUDING HYDRACHNIDIAE The Prostigmata includes approximately 25,800 named species (Zhang et al., 2011) and is the most morphologically and ecologically diverse suborder of mites. Its name derives from the anterior placement of the stigmatal openings to the respiratory system, which are present in many but not all prostigmatans. Although all are fluid-feeders (in contrast to the particle-feeding Sarcoptiformes), the Prostigmata includes predators, herbivores, microbivores, and parasites of vertebrates and invertebrates. Prostigmatans range in morphology from very soft to highly sclerotized, dull to brilliantly colored, and include the tiniest (80-μm gall mites) and the largest (1-cm giant velvet mites) of the Acariformes (Walter and Proctor, 2013). Although the majority of prostigmatans are terrestrial, there are thousands of species that live in fresh water, including the most species-rich group of aquatic mites, the true water mites (Hydrachnidiae). Below we cover freshwater prostigmatans other than the Hydrachnidiae, which are discussed in their own, subsequent section.

Superfamily Halacaroidea: Families Halacaridae and Pezidae The Halacaroidea includes about 1100 mostly marine species (Bartsch, 1996). There are no terrestrial halacaroids. Almost all halacaroids are within the family Halacaridae. Pezidae consists of only two named species. Halacaroids are small (most less than 400 m, but with a few species up to 2 mm in length), dorso-ventrally flattened, diamond-shaped mites with the front two pairs of legs projecting forward and the latter two backwards, giving them a splay-legged stance (Figure 25.8). Pezids have their legs arrayed more radially, however (Krantz and Walter, 2009). No halacaroids have legs modified for swimming. Most have one to two pairs of eyes and one to many dorsal and ventral platelets. Like true water mites, halacaroids have flattened porous genital papillae that likely act as organs of ionic regulation (Figure 25.8). Almost all freshwater halacarids have a life cycle including a larva, protonymph, deutonymph, and adult; however, Astacopsiphagus parasiticus Viets and some marine halacarids have a tritonymphal stage as well (Bartsch, 2007). Approximately 60 species of halacaroids live in nonmarine aquatic habitats (Bartsch, 2008). In some cases the move to fresh water may have been passive. Bartsch (1996) hypothesized that many lake-dwelling halacaroid species originated from brackish-water mites whose habitats became land-locked and slowly transformed into fresh water. Other lineages of halacaroids may have invaded standing water by gradually moving up estuaries and adapting to

614

the lower salinity in a leisurely way. Variation in tolerance to fresh water can be seen among estuarine species. Isobactrus uniscutatus (Viets) may occur at all salinities, whereas other species such as Thalassarachna basteri (Johnston) are restricted to high salinities (Walter and Proctor, 2013). The presumed terrestrial ancestor of the Halacaroidea was a member of the superfamily Bdelloidea (discussed in Harvey, 1990). This taxon is composed entirely of freeliving predators that feed on eggs, larvae, and adults of small arthropods (Krantz and Walter, 2009). In striking contrast, halacaroids act as everything from fungivores to predators to parasites. However, this dietary breadth is best known for marine taxa. Freshwater species appear to be algivorous (based on color of gut contents) or, for some crayfish-associated species, possibly parasitic. Astacopsiphagus parasiticus Viets dwells in the gills of the freshwater crayfish Euastacus spinifer (Heller) in Australia (Bartsch, 1996). Another halacarid, Limnohalacarus wackeri (Walter), has been collected from the gill chambers of Astacus spp. in Europe. It has also been found in surface- and groundwater, implying that it may be an accidental inhabitant of crayfish gills. On firmer ground is Harvey’s (1990) identification of Peza daps Harvey (Pezidae) as a parasite of the gill chambers of the Australian crayfish Engaeus fultoni Smith and Schuster. In addition to its habitat, P. daps lacks eyes, suggesting a cloistered and potentially parasitic life. On the other hand, Engaeus species are burrowing crayfish that dig tunnels up to a metre in depth at the margins of swamps and streams; so it is possible that P. daps is a nonparasitic mite normally inhabiting interstitial areas that is accidentally “breathed in” by E. fultoni. The exact diet has not been determined for any presumed parasitic species of halacarid, freshwater or marine, nor has the effect of halacaroid parasitism on

SECTION | VI  Phylum Arthropoda

hosts been investigated. Marine halacaroids transfer sperm indirectly via spermatophores deposited on a substrate, and presumably freshwater species do as well.

Superfamily Raphignathoidea: Families Homocaligidae and Stigmaeidae The vast majority of species in the Raphignathoidea are terrestrial, and many of them are characteristic of very dry habitats (Krantz and Walter, 2009). In contrast, all Homocaligidae (eight named species in two genera, Homocaligus and Annerossella) and a few in the Stigmaeidae live in freshwater habitats. Like other raphignathoids, members of these two closely related families have non-chelate, knifelike mouthparts and transmit sperm directly with a small sclerotized aedeagus. No species has legs modified for swimming. Their life cycle typically involves four active instars: larva, protonymph, deutonymph, and adult (Krantz and Walter, 2009). Homocaligids are small (∼300–500 μm long), red or orange, well-armored hemispherical mites (Figure 25.16). The dorsum is completely covered by a single helmet-like shield with a complex reticulated pattern. They have a pair of bulbous single lensed eyes behind each of which is an internal tube (in males) or large sac (in females) that may be associated with gas exchange (Gonzalez, 1978; Krantz and Walter, 2009). Homocaligids have been observed walking on the surface of standing water and on the leaves of water plants, including invasive aquatic weeds such as water hyacinth, Eichhornia crassipes (Mart.) Solms. Gonzalez (1978) noted that Annerossella knorri Gonzalez laid its eggs on the leaves of water lettuce (Pistia stratiotes L.), but there was no evidence that these mites fed on the plants.

FIGURE 25.16  The family Homocaligidae consists of two genera of small, spherical, highly sclerotized semiaquatic mites. Annerosella (left), Homocaligus (right). SEMs by David Walter.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

The Stigmaeidae is a family of about 500 species (Zhang et al., 2011) of mostly predatory mites associated with soil, plants, and, to a much lesser extent, fresh water. Stigmaeids are small (less than 500 μm), usually have several dorsal and ventral plates, and are often red or orange in color (F­igure 25.17). Most species have either one or two pairs of eye lenses. Members of the genus Eustigmaeus (previously Ledermuellaria) are unusual in feeding on mosses rather than being predators of small invertebrates (­Flechtmann, 1985), and some species inhabit water-saturated mosses associated with standing water (H.C. Proctor, personal observation). Other genera of stigmaeids with members that occupy standing water habitats are Caligohomus and Cheylostigmaeus (Flechtmann, 1985; Fan and Walter, 2004). Caligohomus durus Fan and Walter from wet soil was observed to feed on nematodes in the laboratory (Fan and Walter, 2004), suggesting that other aquatic stigmaeids may be predators of small invertebrates.

Cohort Parasitengonina—Overview The Parasitengonina includes about 80 families and over 11,000 described species of terrestrial and aquatic mites (Zhang et al., 2011). They include many relatively large and brightly colored (typically red or orange) terrestrial and aquatic mites. One unifying characteristic of this morphologically and behaviorally diverse group is their very complicated life cycle (as exemplified by that of water mites) (Figure 25.21), which includes four inactive and three active stages. The mite passes through a very brief, inactive prelarval stage that is typically spent within the egg, though in some cases the prelarva pops out of the egg as it grows and swells. With few exceptions, the active post-hatching

615

larva is a parasite, while the active deutonymph and adult are predators. Depending on the species, the larva spends from days to months on the host and may be moved a considerable distance from its site of hatching. After engorging on host fluids, the swollen larva drops off and walks or swims to a safe site, where it transforms to an inactive protonymph within the exoskeleton of the larva. This inactive, pharate stage is termed a calyptostase. Terrestrial species usually seek a humid site or bury themselves in soil, whereas water mites attach to aquatic vegetation or seek shelter in the tissues of freshwater sponges and bivalves (Hevers, 1980). The deutonymph develops within the skin of the protonymph, and hence is doubly enclosed by cuticle. The deutonymph that emerges is a very different creature from the larva. Morphologically, these two stages bear no resemblance to each other in the structure of palps, legs, or body (with the exception of Calyptostomatidae, see below). This radical transformation is similar to the structural rearrangement that occurs in insect pupae, which has led some water mite biologists to call the life cycle of parasitengones “essentially holometabolous” (Smith et al., 2010). As well as looking different from the larva, the deutonymph behaves differently. It is an active predator of the eggs, larvae, and pupae of insects, soft-skinned mites, or (in water mites) small crustaceans (see section on “Water Mites”). The deutonymph eventually enters a second calyptostase, the tritonymph. In this case the transformation is less extreme, and the eclosed adult is usually morphologically similar to the deutonymph. However, in water mites the adults are often more highly sclerotized than the deutonymphs, and in sexually dimorphic species the female resembles the deutonymph more closely than the often bizarrely modified male. Adult parasitengones are mostly predatory and have diets similar to their deutonymphal stages, although they may feed on larger prey.

Non-hydrachnidian Parasitengones: Families Calyptostomatidae, Johnstonianidae, and Stygothrombidiidae

FIGURE 25.17  Eustigmaeus frigida (Habeeb), a stigmaeid mite that feeds on freshwater mosses. SEM by David Walter.

Although the vast majority of freshwater Parasitengonina are Hydrachnidiae (“true” water mites), many families of non-hydrachnidian parasitengones include taxa associated with very wet areas (Wohltmann, 2001). In particular, members of the Calyptostomatidae (6 spp.) and Johnstonianidae (53 spp.) are commonly found in sodden moss and other nearshore vegetation, whereas all members of the Stygothrombidiidae (16 spp.) live subaquatically in the substrates of streams and rivers (species counts from Zhang et al., 2011). Like almost all non-hydrachnidian parasitengones, members of these families are orange-to-red soft-bodied mites. They locomote via crawling rather than swimming. Unlike in many taxa of water mites, there is no strong sexual dimorphism in these three families, and for

616

all studied species sperm transfer is indirect via spermatophores deposited on a substrate in the absence of direct contact with the female, rather than by males placing sperm in the female’s genital opening, as occurs in many water mites (Walter and Proctor, 2013). Calyptostomatids can fully retract their gnathosoma within the anterior of their idiosoma, which gives them an odd blunt-nosed appearance (Figure 25.18). Unlike other parasitengones, larvae of this group look quite similar to adults. They parasitize adults of aquatic Diptera, particularly crane flies (Tipulomorpha) Deutonymphs and adults prey on larval tipulids (Wohltmann, 2001). Johnstonianids cannot retract their mouthparts and are relatively unremarkable looking parasitengones with no obvious morphological adaptations for a semi-aquatic

SECTION | VI  Phylum Arthropoda

life, except perhaps for lacking the dense pelage of setae found on terrestrial members of their superfamily (Trombiculoidea) (Figure 25.19). Like calyptostomatids, johnstonianid larvae commonly parasitize crane flies (Tipulomorpha) and, in some cases, larger-bodied parasitengone mites sharing the same semi-aquatic habitat (Wohltmann, 2001) (Figure 25.19). Postlarval stages feed on a diversity of small arthropods including true water mites and other johnstonianids. Johnstoniana rapax Wendt and Eggers is unusual among parasitengones in that the larva can act as either as a predator or as a parasite (Wohltmann, 2001). The family Stygothrombidiidae is sufficiently distinctive as to warrant placement in its own subcohort, the Stygothrombiae (Krantz and Walter, 2009). Stygothrombidiids

FIGURE 25.18  Calyptostoma (Calyptostomatidae) is unusual among the Parasitengonina both in being able to completely retract its gnathosoma, giving it an odd blunt-nosed appearance, and in having larvae (middle image) that resemble post-larval stages. SEM by David Walter, photographs by Joanna Makol.

FIGURE 25.19  Members of the Johnstonianidae (Parasitengonina) live in waterlogged shoreline vegetation. Deutonymphs (left) and adults are predatory, and larvae are parasitic/predatory on arthropods, including other parasitengones such as these unfortunate Microtrombidium individuals (right). Photographs by Andreas Wohltmann.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

617

(a)

(b)

(c)

(d)

FIGURE 25.20  SEM images of a Stygothrombium (Parasitengona: Stygothrombidiidae) individual from Alberta, Canada. (a) lateral view of whole animal; (b) prodorsum showing trichobothrial bases (arrows); (c) glandularium consisting of sensory hair and gland opening; (d) tarsal claws and median empodium (arrow). SEMs by H.C. Proctor.

are morphologically unusual mites, being elongate and wormlike (Figure 25.20), a body form that has independently evolved in several mite taxa that live in interstitial habitats, and have very sparse body setation. Although they have the two defining characteristics of water mites that are lacking in terrestrial Parasitengonina (two setae on the larval palpal genu, combinations of skin glands and trigger hairs called glandularia) (Krantz and Walter, 2009), stygothrombidiids appear to retain prodorsal trichobothria and have well-developed empodial claws in postlarval stages (Figure 25.20), characteristics that have been lost in the Hydrachnidiae sensu stricto. Larvae are parasitic on stoneflies (Plecoptera). The diet of deutonymphal and adult stygothrombidiids is unknown, but they are very likely to be predators.

WATER MITES (PROSTIGMATA: PARASITENGONINA: HYDRACHNIDIAE) Introduction to Water Mites The Hydrachnidiae include about 6000 named species in 57 families, and occupy all possible freshwater habitats on

all continents except Antarctica (Di Sabatino et al., 2008; Krantz and Walter, 2009). There are even a few species that live in marine littoral waters. Water mites are among the most abundant and diverse benthic arthropods in many habitats. One square metre areas of substrate from littoral weed beds in eutrophic lakes may contain as many as 2000 deutonymphs and adults representing up to 75 species in 25 or more genera. Comparable samples from an equivalent area of substrate in rocky riffles of streams often yield over 5000 individuals of more than 50 species in over 30 genera (including both benthic and hyporheic forms). Water mites are associated with some of the dominant insect groups in freshwater ecosystems, especially nematocerous Diptera, and typically interact intimately with these insects at all stages of their life ­histories. In developing this section on arachnids, we have made numerous additions and refinements to the information included in previous editions of this chapter (Smith et al., 2010), as well as some revisions. We have tried to avoid unnecessary use of specialized acarological terms. Those wishing to consult a more comprehensive account of mite structure and function are referred to the appropriate sections of Cook (1974) and Krantz and Walter (2009).

618

General Relationships Hydrachnidiae, along with the enigmatic interstitial Stygothrombidioidea and terrestrial Calyptostomatoidea, Trombidioidea, and Erythraeoidea, belong to a remarkably diverse natural group of actinedid acariform mites, the Parasitengonina, which are characterized by a complex life cycle involving parasitic, calyptostatic, and predaceous stages (Figure 25.21). Larval water mites can be distinguished morphologically from those of other parasitengones by having two setae, rather than one, on the genu of the palp. Deutonymphal and adult water mites differ from all other Acari in having a series of paired glandularia on the idiosoma, as described below.

Origin and Phylogeny Hypotheses about the origin of water mites usually assume a terrestrial parasitengone ancestor (Davids and Belier, 1979; Mitchell, 1957a). An alternative hypothesis (Wiggins et al., 1980; Smith and Oliver, 1986; Krantz and Walter, 2009) suggests that the primitive parasitengone stock may have been water mites resembling certain extant Hydryphantoidea. According to this postulate, water mites diverged

FIGURE 25.21  Life-cycle of a parasitengone mite (Parasitengonina: Hydrachnidiae: Torrenticola sp.). (1) Eggs; (2) larva searching for host; (3) larva parasitic on host; (4) engorged larva detached from host; (5) calyptostatic protonymph; (6) predatory deutonymph; (7) calyptostatic tritonymph; (8) predatory adult. Illustration by Reinhard Gerecke.

SECTION | VI  Phylum Arthropoda

from terrestrial ancestors with direct development, perhaps resembling extant Anystoidea, while evolving the basic parasitengone life history pattern as a set of adaptations for exploiting spatially and temporally intermittent aquatic habitats. Terrestrial parasitengones subsequently radiated into various habitats, including more xeric ones. The fossil record provides little insight into the evolutionary history of water mites. The only reported fossils that undoubtedly are water mites (Cook, 1957; Poinar, 1985) are larval specimens from Tertiary deposits representing highly derived taxa. However, distributional and host association data indicate to us that the group originated no later than the Triassic or early Jurassic period (Smith and Cook, 1999; Krantz and Walter, 2009). Morphological and behavioral data suggest that extant water mites are monophyletic (Mitchell, 1957a; Barr, 1972; Cook, 1974; Smith and Oliver, 1986), and that the major phyletic lineages of water mites, and possibly all Parasitengonina, were derived from hydryphantoid-like ancestors (Mitchell, 1957a; Wainstein, 1980). At the time of writing, no molecular phylogenetic studies have explicitly addressed the monophyly or sister-group relationships of water mites within the Parasitengonina. Water mite evolution since at least the Triassic appears to have involved repeated episodes of rapid diversification, resulting in novel behavioral and morphological traits associated with exploitation of new host groups by larvae and invasion of new aquatic habitats by deutonymphs and adults. As discussed below, members of Hydryphantoidea and other early derivative superfamilies such as Eylaoidea and Hydrovolzioidea retain essentially terrestrial larvae that locate their hosts on the surface film. Those of more derived clades comprising the superfamilies Hydrachnoidea, Lebertioidea, Hygrobatoidea, and Arrenuroidea have fully aquatic larvae that locate hosts in the water column or on the substrate (Smith and Oliver, 1986; Smith and Cook, 1999; Krantz and Walter, 2009). Evolution of subaquatic larvae probably occurred independently in the Hydrachnoidea and in the ancestors of the clade comprising the other three superfamilies. Larvae of most groups of the Lebertioidea, Hygrobatoidea, and Arrenuroidea parasitize nematocerous Diptera, especially Chironomidae, and exhibit high levels of taxonomic and ecological diversity. The complex life history of these mites has provided them with virtually unlimited opportunities for experimenting with new host associations and habitats and members of numerous clades with diverse origins have developed similar suites of morphological and behavioral adaptations. The high levels of diversity coupled with extensive morphological homoplasy make resolution of phylogenetic relationships among and within these superfamilies very challenging in the absence of molecular data. This problem continues to be compounded by inadequate knowledge of the fauna,

Chapter | 25  Subphylum Chelicerata, Class Arachnida

especially from springs, streams, and interstitial habitats in most regions of the world.

Diversity, Classification, and Biogeography Nearly 6000 species of water mites are currently recognized worldwide, representing more than 300 genera and subgenera in over 100 families and subfamilies (Viets, 1987; di Sabatino et al., 2008; Krantz and Walter, 2009; Smith et al., 2010). Acarologists usually consider water mites to be a taxon of intermediate rank between superfamily and suborder. The group rivals several orders of aquatic insects in diversity and is probably comparable to them in age. The families are conservatively grouped into seven superfamilies. Of these, the Hydrovolzioidea, Hydrachnoidea, and Eylaoidea probably represent natural monophyletic groupings. The Hydryphantoidea, Lebertioidea, Hygrobatoidea, and Arrenuroidea are all either paraphyletic or polyphyletic assemblages in need of revision. A comprehensive overview of water mite genera to harmonize information on morphological and behavioral traits of all active instars with new molecular data is essential to improve understanding of water mite phylogeny and to produce a more natural and informative higher classification. Water mites occur throughout the world, except Antarctica (Cook, 1974). All superfamilies, except Hydrovolzioidea, as well as many early derivative families and subfamilies, are richly represented in all zoogeographic regions. Knowledge of global distribution patterns of water mite taxa has been substantially improved by recent studies of the fauna of austral regions, especially those by Kurt Viets and David Cook. The basic patterns can be explained as the result of vicariance due to plate tectonics (Smith and Cook, 1999). Dispersal (along with hosts) between nearby land masses (e.g., Southeast Asia and Australia) or within continents (e.g., Africa) significantly altered these patterns as more recently derived groups progressively joined, and perhaps displaced, ancient ones. Smith and Cook (1999) identified four major sets of water mite lineages by assessing modern geographic distributions, phylogenies, levels of endemism, and potential for dispersal (see Table 25.2). Pangean clades exhibit substantial endemism on all continents and apparently represent ancient lineages that were widely distributed in Pangea during mid-Jurassic times. Laurasian or Gondwanan clades have broad northern or southern distributions, respectively, and probably represent derived lineages that originated in Laurasia (north) or Gondwanaland (south), during late Jurassic or Cretaceous times. Recent clades have relatively narrow distributions restricted to one region or two contiguous regions in either the Northern or Southern Hemisphere, and likely originated after the break-up of Laurasia or Gondwanaland, respectively.

619

These authors inferred the existence of three more-orless distinct regional faunas in Pangea during early and mid-Jurassic times:  Pangean Tropical Fauna distributed rather uniformly around the equator in areas that would later become tropical Laurasia and Gondwanaland, including representatives of at least 11 extant families (13 subfamilies); l Pangean North Temperate Fauna distributed widely in the Northern Hemisphere in most of the areas that would later become temperate Laurasia, including representatives of at least 28 extant families (43 subfamilies); and l  Pangean South Temperate Fauna distributed widely in the Southern Hemisphere in the areas that would later become temperate Gondwanaland, including representatives of at least 21 extant families (31 subfamilies). l

According to this assessment, there may have been some interchange of taxa between northern and southern temperate areas of Pangea along highland corridors, but considerable endemism existed in cooler areas away from the equator in both hemispheres at the time of rifting. The gradual separation of Laurasia and Gondwanaland resulted in further divergence of endemic clades in temperate latitudes and initiated differentiation of distinctive clades in tropical areas of both supercontinents. The northern and southern clades that resulted from the separation of Laurasia and Gondwanaland diverged and diversified as the two supercontinents became further fragmented during Cretaceous and Tertiary times. In the Northern Hemisphere, regional vicariance was established as the opening of the North Atlantic Ocean progressively split eastern North America from western Eurasia. The final separation of North American and Eurasian populations in cool temperate areas on either side of the widening Atlantic probably occurred sometime after the establishment of connections in the Beringian region initiated a new wave of dispersal. Two major distinctive faunas apparently inhabited Laurasia at the time of the separation of eastern North America from western Eurasia: Laurasian Temperate Fauna initially widely distributed throughout most of North America and Eurasia, including representatives of at least 34 extant families (55 subfamilies). A small group of these taxa (represented by five extant families and seven subfamilies) may have been endemic to either extreme western North America or eastern Eurasia prior to establishment of the filter bridge, and later corridor, through Beringia. l  Laurasian Tropical Fauna distributed in southern Laurasia, in the areas that later became tropical southeastern Eurasia and tropical North America. The component of this fauna inhabiting southeastern Laurasia included representatives of at least 21 extant families (29 subfamilies). Tropical fauna apparently nearly disappeared from l

Pangean

 Hydrodromidae

Unknown Terrestrial

Pangean

Laurasian

Pangean

 Protziinae

 Pseudohydryphantinae Pangean

Pangean

 Hydryphantinae

 Tartarothyadinae

 Wandesiinae

Pangean

 Thermacaridae

 Apheviderulicidae

Laurasian

Gondwanan

 Zelandothyadinae

Eylaoidea

Gondwanan

 Australiothyadinae

 Zelandothyadidae

Gondwanan

North or South American

 Santiagocarinae

 Teratothyadidae

Unknown

South American

 Clathrosperchoninae

Terrestrial

Unknown

Unknown

Terrestrial

Unknown

Unknown

Unknown

Aquatic

Terrestrial

 Rhynchohydracarinae South American

 Rhynchohydracaridae

Terrestrial

Pangean or Laurasian

 Euthyadinae

Terrestrial

Laurasian

Unknown

Terrestrial

Unknown

Type

 Cyclothyadinae

 Hydryphantidae

Gondwanan

Biogeographic Origin

 Ctenothyadidae

Hydryphantoidea

Taxa

Thorax, abdomen

Thorax

Unknown

Thorax

Unknown

Attachment Site

Coleoptera

Unknown

Unknown

Anura (Amphibia)

Unknown

Unknown

Unknown

Unknown

Plecoptera

Unknown

Unknown

Abdomen

Unknown

Unknown

Body dorsum

Unknown

Unknown

Unknown

Unknown

Thorax

Unknown

Unknown

Plecoptera, Diptera, Thorax, abdomen Trichoptera

Hemiptera, Odonata, Diptera

Collembola, Diptera, Trichoptera

Unknown

Diptera

Unknown

Order of Hosts

Larvae

Springs

Interstitial

Interstitial

Hot springs

Riffles

Riffles

Riffles, interstitial

Riffles, interstitial

Springs, interstitial

Springs, riffles

Pools, lakes

Riffles

Lakes, temporary pools

Springs, riffles, temporary pools

Springs, interstitial

Diptera larvae

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Diptera eggs

Insect eggs

Unknown

Insect eggs

Unknown

Prey

Crawling

Unknown

Crawling or walking Unknown

Crawling or walking Unknown

Crawling

Crawling

Crawling

Crawling

Crawling

Walking

Walking, crawling

Swimming

Walking

Swimming

Walking, crawling

Crawling

Swimming

Crawling

Mode of Locomotion

Deutonymphs and Adults

Riffles, pools, lakes

Mosses

Habitat Type

Ecology

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Dissociated

Dissociated

Unknown

Dissociated

Unknown

Spermatophore Transfer Mode

Adults

TABLE 25.2  Summary of Some Biogeographic and Ecological Characteristics of Broadly Distributed Genera of Water Mites (Prostigmata: Hydrachnidiae)

620 SECTION | VI  Phylum Arthropoda

Pangean

Pangean

Pangean

Laurasian

 Oxidae

 Rutripalpidae

Gondwanan

Laurasian

 Teutoniidae

Laurasian

 Sperchontinae

 Stygatoniidae

Gondwanan

 Apeltosperchontinae

 Sperchontidae

Laurasian

Pangean

 Nilotoniinae

 Lebertiidae

Pangean

 Anisitsiellinae

 Anisitsiellidae

Lebertioidea

 Hydrachnidae

Pangean

Laurasian

 Hydrovolziidae

Hydrachnoidea

Laurasian

Laurasian

 Acherontacaridae

Hydrovolzioidea

 Piersigiidae

 Rhyncholimnocharinae Gondwanan

 Limnocharinae

 Limnocharidae

 Eylaidae

Aquatic

Unknown

Aquatic

Unknown

Unknown

Aquatic

Aquatic

Aquatic

Aquatic

Terrestrial

Aquatic

Terrestrial

Aquatic

Terrestrial

Terrestrial

Unknown

Unknown

Thorax, abdomen

Thorax

Thorax, abdomen

Thorax, abdomen

Thorax, abdomen

Unknown

Abdomen

Abdomen

Thorax, abdomen

Thorax, abdomen

Diptera

Unknown

Abdomen

Unknown

Diptera, Trichoptera Thorax, abdomen

Unknown

Unknown

Diptera

Diptera

Diptera

Coleoptera, Hemiptera

Hemiptera, Diptera

Unknown

Coleoptera

Coleoptera

Hemiptera, Odonata

Coleoptera, Hemiptera

Springs, pools

Interstitial

Springs, riffles, pools, lakes

Riffles

Springs, pools

Pools, lakes

Springs, riffles, pools, lakes

Springs, riffles, pools, interstitial

Lakes, temporary pools

Springs, riffles

Interstitial

Springs, interstitial, temporary pools

Riffles, interstitial

Riffles, pools, lakes

Crawling, swimming

Walking

Crawling

Crawling

Crawling

Swimming

Walking, swimming

Crawling, swimming

Swimming

Crawling

Crawling

Crawling

Crawling

Crawling, swimming

Pools, lakes, tempo- Swimming rary pools

Diptera larvae

Unknown

Diptera larvae

Unknown

Unknown

Diptera larvae

Diptera larvae

Unknown

Insect eggs

Unknown

Unknown

Ostracoda

Unknown

Diptera larvae

Ostracoda, Cladocera

Continued

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Dissociated

Unknown

Dissociated

Unknown

Unknown

Unknown

Unknown

Unknown

Dissociated; paired, indirect; paired, direct

Chapter | 25  Subphylum Chelicerata, Class Arachnida 621

Laurasian

 Torrenticolinae

Pangean

Pangean

Laurasian

 Frontipodopsidae

 Hygrobatidae

 Lethaxonidae

 Limnesiidae

South American

Gondwanan

 Notoaturinae

 Ferradasiidae

Pangean

 Axonopsinae

Laurasian

Laurasian

 Aturinae

 Feltriidae

Pangean

 Albiinae

 Aturidae

 Astacocrotonidae

Australian

Laurasian

 Testudacarinae

Hygrobatoidea

Pangean or Laurasian

Biogeographic Origin

 Neoatractidinae

 Torrenticolidae

Taxa

Unknown

Aquatic

Unknown

Unknown

Aquatic

Unknown

Aquatic

Aquatic

Aquatic

Unknown

Aquatic

Aquatic

Unknown

Type

Unknown

Diptera, Trichoptera

Unknown

Unknown

Diptera

Unknown

Diptera

Diptera

Trichoptera

Unknown

Diptera

Diptera

Unknown

Order of Hosts

Larvae

Unknown

Abdomen

Unknown

Unknown

Abdomen

Unknown

Abdomen

Abdomen

Abdomen

Unknown

Thorax

Thorax

Unknown

Attachment Site

Interstitial

Springs, riffles, interstitial, pools, lakes

Interstitial

Riffles

Springs, riffles, interstitial

Riffles, interstitial

Springs, riffles, interstitial, pools, lakes

Springs, riffles, interstitial, pools

Pools, lakes

Crayfish gills

Riffles, interstitial, pools, lakes

Unknown

Diptera larvae

Unknown

Diptera larvae

Diptera larvae

Unknown

Unknown

Diptera larvae

Diptera larvae

Unknown

Prey

Walking

Crawling, walking, swimming

Unknown

Diptera larvae, Ostracoda, Cladocera

Walking, swimming Unknown

Crawling

Crawling

Crawling

Walking, swimming

Crawling

Swimming

Crawling

Crawling

Crawling

Crawling

Mode of Locomotion

Deutonymphs and Adults

Riffles, interstitial

Riffles

Habitat Type

Ecology

Unknown

Dissociated

Unknown

Unknown

Paired, direct

Unknown

Dissociated; paired, direct

Paired, direct

Unknown

Unknown

Unknown

Unknown

Unknown

Spermatophore Transfer Mode

Adults

TABLE 25.2  Summary of Some Biogeographic and Ecological Characteristics of Broadly Distributed Genera of Water Mites (Prostigmata: Hydrachnidiae)— cont’d

622 SECTION | VI  Phylum Arthropoda

Laurasian

Pangean

South American

North or South American

South American

Eurasian

North American

North or South American

Eurasian

South American

 Kawamuracarinae

 Limnesiinae

 Mixolimnesiinae

 Neomamersinae

 Neotorrenticolinae

 Nicalimnesiinae

 Psammolimnesiinae

 Protolimnesiinae

 Stygolimnesiinae

 Tyrrellinae

Pangean or Gondwanan

 Omartacarinae

Laurasian

Pangean

Laurasian

Laurasian

Laurasian

 Foreliinae

 Huitfeldtiinae

 Hydrochoreutinae

 Najadicolinae

 Pioninae

 Pionidae

Laurasian

 Maharashtracarinae

 Omartacaridae

North American

 Epallagopodinae

Aquatic

Aquatic

Aquatic

Aquatic

Aquatic

Unknown

Unknown

Aquatic

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Aquatic

Unknown

Unknown

Diptera

Diptera

Diptera

Diptera

Diptera

Unknown

Unknown

Diptera

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Diptera

Unknown

Unknown

Abdomen

Abdomen

Abdomen

Abdomen

Abdomen

Unknown

Unknown

Abdomen

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Thorax

Unknown

Unknown Unknown

Unknown

Crawling

Swimming

Swimming

Crawling, swimming

Walking

Walking

Crawling

Walking

Walking

Walking

Walking

Crawling

Walking

Walking

Copepoda, Cladocera, Diptera larvae

Mollusc tissue

Cladocera

Cladocera

Diptera larvae

Unknown

Unknown

Diptera eggs and larvae

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Walking, swimming Copepoda, Cladocera, insect eggs and larvae

Walking

Crawling

Springs, riffles, Crawling, pools, lakes, tempo- swimming rary pools

Lakes (mollusc parasite)

Pools, lakes

Pools, lakes (profundal)

Springs, interstitial, pools, lakes

Interstitial

Interstitial

Springs, margins of pools and lakes

Interstitial

Interstitial

Interstitial

Interstitial

Riffles

Interstitial

Interstitial

Riffles, interstitial, pools, lakes

Interstitial

Springs

Continued

Paired, direct

Unknown

Unknown

Unknown

Paired, direct

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Chapter | 25  Subphylum Chelicerata, Class Arachnida 623

Gondwanan

Eurasian

 Pollicipalpinae

 Encentridophorinae

North American

North American

Pangean

 Amoenacaridae

 Arenohydracaridae

 Arrenuridae

 Athienemanniidae

Laurasian

 Acalyptonotidae

Arrenuroidea

Pangean

Pangean

 Unionicolinae

 Wettinidae

Pangean

 Pionatacinae

 Unionicolidae

Pangean

Gondwanan

 Schminkeinae

 Pontarachnidae

Pangean

Biogeographic Origin

 Tiphyinae

Taxa

Aquatic

Unknown

Unknown

Aquatic

Aquatic

Unknown

Unknown

Aquatic

Aquatic

Unknown

Unknown

Aquatic

Type

Unknown

Unknown

Abdomen

Attachment Site

Diptera, Odonata

Unknown

Unknown

Diptera

Diptera

Unknown

Unknown

Thorax, abdomen

Unknown

Unknown

Thorax

Abdomen

Unknown

Unknown

Diptera, Trichoptera Abdomen, legs

Diptera, Trichoptera Abdomen

Unknown

Unknown

Diptera

Order of Hosts

Larvae Mode of Locomotion

Deutonymphs and Adults

Springs, riffles, interstitial, pools, lakes, temporary pools

Interstitial

Pools

Unknown

Ostracoda, Cladocera, Copepoda, Diptera larvae

Prey

Swimming

Swimming

Swimming

Crawling, swimming

Swimming

Unknown

Unknown

Walking, swimming Ostracoda, Cladocera, Copepoda, insect larvae

Walking

Swimming

Unknown

Unknown

Unknown

Unknown

Copepoda, Cladocera, Diptera larvae, mollusc tissue

Diptera larvae, Copepoda

Walking, swimming Unknown

Crawling

Springs, pools, lakes Walking

Pools, lakes

Pools, lakes

Pools, lakes

Pools, lakes, mollusc parasites

Pools, lakes

Pools, marine littoral

Interstitial

Pools, lakes, tempo- Swimming rary pools

Habitat Type

Ecology

Paired, direct

Unknown

Unknown

Unknown

Dissociated

Unknown

Unknown

Dissociated; paired, indirect; paired, direct

Dissociated; paired, indirect; paired, direct

Unknown

Unknown

Paired, direct

Spermatophore Transfer Mode

Adults

TABLE 25.2  Summary of Some Biogeographic and Ecological Characteristics of Broadly Distributed Genera of Water Mites (Prostigmata: Hydrachnidiae)— cont’d

624 SECTION | VI  Phylum Arthropoda

Australian

North American

 Notomudamellinae

 Stygameracarinae

Eurasian

Pangean

North American

Laurasian

Pangean

Pangean

 Kantacaridae

 Krendowskiidae

 Laversiidae

 Mideidae

 Momoniidae

 Nudomideopsidae

Eurasian

South American

 Plaumanniinae

 Nipponacaridae

Pangean

  Mideopsinae +   Mideopsellinae

Laurasian

Gondwanan

 Guineaxonopsinae

 Neoacaridae

Australian

 Gretacarinae

 Mideopsidae

Laurasian

 Hungarohydracaridae

Laurasian

 Uchidastygacarinae

Gondwanan

Laurasian

 Morimotacarinae

 Harpagopalpidae

Laurasian

 Chappuisidinae

 Chappuisididae

Laurasian

Pangean

 Athienemanniinae

 Bogatiidae

Eurasian or African

 Africasiinae

Unknown

Aquatic

Unknown

Aquatic

Unknown

Unknown

Aquatic

Aquatic

Aquatic

Aquatic

Aquatic

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Aquatic

Unknown

Unknown

Diptera

Unknown

Diptera

Unknown

Unknown

Diptera

Trichoptera

Diptera

Diptera

Diptera

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Diptera

Unknown

Unknown

Thorax

Unknown

Thorax

Unknown

Unknown

Thorax

Thorax, abdomen

Abdomen

Thorax

Abdomen

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Thorax

Unknown

Interstitial

Riffles, interstitial, lakes

Pools

Springs, riffles, interstitial, pools, lakes

Interstitial

Pools

Springs, interstitial

Riffles, interstitial, pools, lakes

Pools, lakes

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Diptera larvae

Unknown

Unknown

Swimming

Diptera larvae

Unknown

Unknown

Unknown

Unknown

Diptera larvae

Walking or crawling Unknown

Walking

Swimming

Walking, crawling, swimming

Walking or crawling Unknown

Swimming

Walking

Walking, swimming Diptera larvae

Swimming

Unknown

Unknown

Walking or crawling Unknown

Walking

Walking or crawling Unknown

Walking

Walking

Walking

Walking

Walking

Walking

Walking

Swimming

Springs, riffles, lakes Walking (profundal)

Pools, lakes

Interstitial

Interstitial

Riffles

Interstitial

Interstitial

Interstitial

Interstitial

Interstitial

Interstitial

Springs, riffles, interstitial

Pools

Unknown

Paired, direct

Unknown

Unknown

Unknown

Unknown

Paired, direct

Unknown

Paired, direct

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Chapter | 25  Subphylum Chelicerata, Class Arachnida 625

SECTION | VI  Phylum Arthropoda

626

southwestern Laurasia during the Cretaceous, persisting only in Mexico and on some of the larger Caribbean islands, probably because the area that later became continental North America moved entirely north of the Tropic of Cancer. During mid-Tertiary times, tropical fauna of southwestern Laurasia included representatives of at least 16 extant families (22 subfamilies). In the Southern Hemisphere, rifting was more complex and resulted in the establishment of five continent-sized fragments, each with its own complement of Gondwanan clades. Here, two distinct faunas in Gondwanaland are inferred to have existed before rifting began to separate Africa and India from the supercontinent. Gondwanan Tropical Fauna distributed throughout equatorial areas that later became northern Africa and South America, including representatives of at least 14 extant families (16 subfamilies); l Gondwanan Temperate Fauna initially distributed widely in temperate areas that later became Africa and India, New Zealand, South America, Australia, and Antarctica, including representatives of at least 26 extant families (and 44 subfamilies). l

Following the separation of Africa and India, elements of this fauna apparently differentiated significantly in the land masses attached to the eastern and western parts of Antarctica. In the Northern Hemisphere, movement of water mites between eastern North America and western Eurasia ceased early in the Tertiary. Later, significant faunal interchange across Beringia began in the Paleocene or Eocene and continued as successive waves of dispersal throughout the Tertiary and Pleistocene. Climatic cooling and drying that began in the Miocene and culminated in the extensive glaciations of the Pleistocene displaced the temperate fauna southward in both North America and Eurasia. This series of displacements resulted in highly disjunct “Tertiary relict” distributions for many clades that had previously been more or less continuously spread throughout temperate Laurasia. Cold-adapted clades differentiated and replaced Tertiary lineages in Arctic and Boreal latitudes. Africa docked with southern Eurasia during the Miocene, initiating extensive faunal interchange between the two areas that continues to the present. South America approached southern North America during the Miocene, establishing a filter bridge that permitted limited dispersal, and finally docked in the Pliocene, leading to interchange that has significantly altered the faunas of both continents. In the Southern Hemisphere, tropical taxa persisted in equatorial areas as Africa drifted northward, but were largely replaced in northern parts of the continent by groups dispersing from southern Eurasia during the late Tertiary. Some of these temperate clades eventually reached southern Africa by

traversing mountainous areas. Elements of the old temperate Gondwanan fauna have survived only in extreme South Africa. The original temperate Gondwanan fauna of India was completely replaced as India drifted north across the equator and ultimately docked with southern Eurasia. The modern fauna of India is composed entirely of groups that have dispersed from Eurasia since the Miocene. In South America, ancient tropical groups persisted and diversified in northern equatorial areas throughout the Tertiary. The fauna of this part of the continent has been greatly altered as a result of dispersals from North America since the Pliocene. The old temperate Gondwanan fauna has survived only in southern Argentina and Chile, and some groups were able to disperse northward in the high Andes during late Tertiary and Quaternary times. Old temperate Gondwanan clades remain dominant in New Zealand, but in Australia, they are now confined largely to the southeastern mountains. Temperate Gondwanan fauna was eliminated from most of Australia as it moved northward and the climate became progressively warmer and drier. The fauna of northern parts of Australia is now composed largely of groups that have dispersed from tropical Eurasia during the late Tertiary and Quaternary.

External Morphology and Internal Anatomy Early water mite workers developed an idiosynchratic set of morphological terms for external structures of adult water mites. Taxonomic studies of larval water mites generated an additional set of special terms for the exoskeleton of this strongly heteromorphic instar; however, it has become evident that the relatively plesiotypic body plan of larvae permits much of the standard acariform terminology of Grandjean (see Krantz and Walter, 2009) to be used with confidence.

Morphology of Larvae Like larvae of all mites, those of the Hydrachnidiae have only three pairs of legs and hence are easily differentiated from the eight-legged nymphal and adult stages. The short gnathosoma of the water mite larva bears a pair of stocky pedipalps (or just “palps”) that have five free segments (trochanter, femur, genu, tibia, tarsus) (Figure 25.22). The tarsus of the palp is relatively long and cylindrical in some early derivative genera, but is typically reduced to a dome- or button-shaped pad in most derived groups. A thick, curved seta is present dorsally at the end of the tibia, the homologue of the tibial claw that characterizes the palps of most Anystoidea, terrestrial Parasitengonina, and related groups. The paired chelicerae, each consisting of a cylindrical basal segment and a movable terminal claw, lie between the palps. The idiosoma of larval water mites varies in degree of sclerotization. In the Hydryphantoidea, it is mainly

Chapter | 25  Subphylum Chelicerata, Class Arachnida

(a)

627

(b)

(c)

FIGURE 25.22  Labeled anatomy of water mite larva, Sperchonopsis ecphyma Prasad and Cook (Sperchontidae), excluding legs: (a) dorsal view, (b) ventral view excluding gnathosoma, (c) ventral view of gnathosoma.

unsclerotized (Figure 25.23). The dorsal integument bears a median eye-spot, two pairs of lens-like lateral eyes, four pairs of lyrifissures and the numerous pairs of setae. Ventrally, the integument bears the three pairs of coxal plates, paired urstigmata (which are precursors to genital acetabula =  genital papillae) laterally between coxal plates I and II,

the excretory pore, one pair of lyrifissures, and numerous pairs of setae (Figure 25.23). The size, shape, and position of these dorsal and ventral structures, and the degree of fusion of the sclerites associated with them, provide useful taxonomic characters. The legs are inserted laterally on the coxal plates, and in early derivative groups have six movable

SECTION | VI  Phylum Arthropoda

628

D

E

FIGURE 25.23  Dorsal plates of water mite larvae: (a) Hydrodroma despiciens (Müller) (Hydrodromidae); (b) Eylais major Lanciani (Eylaidae).

segments: trochanter, basifemur, telofemur, genu, tibia, and tarsus. The segments have taxonomically specific complements of setae and solenidia (chemosensory setae). Each leg tarsus bears a pair of claws and a claw-like empodium. Larvae of Hydryphantoidea and Eylaoidea retain six movable leg segments, but those of all other groups have the basifemoral and telofemoral segments fused (Figure 25.24). Larval Hydryphantoidea also have the largest complement of setae on the leg segments. Larvae of each of the more derived groups exhibit characteristic leg chaetotaxies that reflect reductions from the presumed plesiotypical complement. Certain ventral setae on the genua, tibiae, and tarsi are elongate and plumose in larvae adapted for swimming. Coxal plates are often fused (Figure 25.25).

Morphology of Deutonymphs and Adults In deutonymphal and adult water mites, the gnathosoma consists of the gnathosomal base (or “capitulum”), chelicerae, and palps (Figure 25.26). The capitulum is plesiotypically a simple, short channel derived from extensions of the palpal coxae, leading to the esophagus. A protrusible tube of integument connecting the capitulum to the idiosoma has developed independently in several distantly related genera (e.g., Rhyncholimnochares) (Figure 25.27). The paired

palps are used both for sensing and for capturing prey. In the plesiomorphic condition, the palps have five movable segments, namely the trochanter, femur, genu, tibia, and tarsus, that are essentially cylindrical. The tibia bears a thick, blade-like, dorsal seta distally in many ancient genera (e.g., Hydryphantes, Tartarothyas, Pseudohydryphantes) (Figure 25.28). As in larvae, this seta is the homologue of the tibial claw of terrestrial relatives, and it often makes the palps appear to be chelate. In other groups, other setae along with various denticles and tubercles may be elaborated to enhance the raptorial function of the palps. Segmentation of the palps is reduced by fusion in a few genera. A modification that has developed independently in various groups of Arrenuroidea is the so-called uncate condition. In these genera, the tibia is expanded and produced ventrally to oppose the tarsus (Figure 25.28), permitting the mites to grasp and hold slender appendages of prey organisms. The paired chelicerae lie in longitudinal grooves between the palps on the dorsal surface of the capitulum. Plesiotypically they consist of a cylindrical basal segment bearing a movable terminal claw (Figure 25.29). This cheliceral structure is designed for tearing the integument of prey organisms, and is retained in nearly all derivative groups. Hydrachnidae are unique in having unsegmented and stiletto-like chelicerae (Figure 25.29), which they use to

Chapter | 25  Subphylum Chelicerata, Class Arachnida

629

D

E

F

FIGURE 25.24  Legs of larval Sperchonopsis ecphyma Prasad and Cook (Sperchontidae).

pierce insect eggs. The chelicerae are separate in all groups except Limnocharidae and Eylaidae where they are fused medially. The most comprehensive comparative studies of water mite mouthparts were published by Motaş (1928) and Mitchell (1962). The idiosoma, or body proper, is highly variable in shape among water mites (see next section), but in early derivative taxa is round or ovoid, slightly flattened dorsoventrally, and mostly unsclerotized, as in certain extant members of ancient genera such as Hydryphantes, Tartarothyas and Pseudohydryphantes. In these taxa the dorsal integument

bears an unpaired median eye, paired lateral eyes that are usually enclosed in capsules, paired preocular and postocular setae, and longitudinal series of paired glandularia, muscle attachment sites, and lyrifissures (five). Ventrally the integument bears the paired coxal plates (often fused into anterior and posterior groups on each side), the genital field (comprising the gonopore, three pairs of acetabula [­ =genital papillae] (Figure 25.30), and paired genital valves), five pairs of ventroglandularia, and the excretory pore. As in larvae, these idiosomal structures provide a wealth of useful taxonomic characters.

SECTION | VI  Phylum Arthropoda

630

D

E

FIGURE 25.25  Dorsum and venter of larval Torrenticola sp. (Torrenticolidae).

FIGURE 25.26  Labeled ventral view of adult female Limnesia sp. (Limnesiidae).

Chapter | 25  Subphylum Chelicerata, Class Arachnida

631

The legs are inserted laterally on the coxae, plesiotypically articulate on a vertical major axis, and have six movable segments. The segments are essentially cylindrical and have variable complements of setae. Although chaetotaxy of the legs provides a variety of taxonomic characters in deutonymphs and adults, the expression and position of individual setae are highly variable within taxa. Consequently, the rigorous analysis of chaetotactic patterns that proves so useful in the case of larvae is not practicable for later instars. The leg tarsi typically bear paired claws terminally, but this number is often reduced. This soft-bodied condition is retained in early derivative genera adapted for walking on the substrate or swimming in shallow habitats such as seepage pools or temporary pools. Walking forms have relatively short, stocky leg segments and setae, whereas swimmers have longer segments bearing fringes of slender swimming setae. As pointed out by Mitchell (1957a,b, 1958, 1960), the soft integument of these mites permits the highly precise local control of body shape FIGURE 25.27  Limnocharid water mites showing extendible gnathosoma. Rhyncholimnochares kittatinniana Habeeb (top) with mouthparts retracted and extended, and Limnochares americana Lundblad (bottom) with mouthparts extended and grasping the head capsule of a chironomid larva. Photographs by Bruce Smith.

D

D

E

E FIGURE 25.28  Palps of Trichothyas sp. (Hydryphanidae) and Arrenurus sp. (Arrenuridae).

D F

E

FIGURE 25.29  Chelicerae of Hydrachna sp. (Hydrachnidae) and Hygrobates sp. (Hygrobatidae).

FIGURE 25.30  Genital fields of Albertathyas montanus Smith and Cook (Hydryphantidae), Atractides sp. (Hygrobatidae) and Najadicola ingens Koenike (Pionidae) showing diversity of genital acetabula number and arrangement.

SECTION | VI  Phylum Arthropoda

632

and internal pressure that is needed to produce the walking and swimming movements of the legs.

D

E

F

G

Evolution of Adult Exoskeleton The major lineages of water mites apparently differentiated from ancestral stock that resembled extant s­ oft-bodied Hydryphantoidea with a slightly flattened, oblate form (Mitchell, 1957a,b; Smith and Oliver, 1986; Krantz and Walter, 2009), similar to that in extant Calyptostomatidae (Figure 25.19). Adaptation to habitats such as seepage areas and springs or substrata in streams required a change in locomotor habits from walking or swimming. Water mites associated with moss mats and wet litter habitats have a hydrophilic integument that draws a film of water over the dorsal surface, creating sufficient downward force to press the body to the substrate and prevent efficient walking (Mitchell, 1960). Those living in streams evolved a wedge-shaped body designed for negotiating confined spaces under rocks and within gravel. Locomotion in both of these types of habitats necessitated evolution of a crawling gait. This change involved a shift in orientation of the major axis of the legs, shortening and thickening of leg segments and setae, enlargement of tarsal claws, and development of stronger, more massive muscles to move the legs. Expansion of coxal plates and sclerites occurred to provide rigid exoskeletal support for these muscles. Fusion of these sclerotized areas led to repeated evolution of complete dorsal and ventral shields in many groups of Hydryphantoidea, Lebertioidea, Hygrobatoidea, and Arrenuroidea (Figures 25.31(a)–(d) and 25.32(a)–(d)). Different lineages have exploited interstitial habitats in subterranean waters and the hyporheic zone of springs and streams. These mites tended to lose eyes and integumental pigmentation, and required further streamlining of the body to facilitate locomotion in interstitial spaces. Originally soft-bodied forms adopted a vermiform shape, whereas sclerotized mites became extremely compressed laterally or dorsoventrally. The crawling mode of locomotion became secondarily modified in several groups of well-sclerotized interstitial genera (e.g., Neomamersa, Frontipodopsis, Chappuisides) to permit rapid and agile running in hyporheic and ground water habitats. Repeated habitat diversification within the major clades of Hydryphantoidea, Lebertioidea, Hygrobatoidea, and Arrenuroidea resulted in several parallel and convergent trends in body sclerotization. Members of different clades developed superficially similar sclerite arrangements in adapting to lotic or interstitial habitats (e.g., species of Diamphidaxona and certain Axonopsinae); others underwent homoplastic reduction or loss of sclerites in secondarily invading lentic habitats (e.g., species of Forelia and Piona). Finally, some groups retained extensive sclerotization while becoming adapted for swimming in standing

FIGURE 25.31 (a) Arrenurus fissicornis Marshall (Arrenuruidae), male. (b) Arrenurus (Megaluracarus) pseudocylindratus Marshall (Arrenuridae), male. (c) Kongsbergia sp. (Aturidae), female. (d) Aturus sp. (Aturidae), male.

water (e.g., certain species of Axonopsis and Mideopsis). Consequently, modern water mite assemblages are highly heterogeneous phylogenetically and morphologically. Evolutionary trends involving other readily observed and taxonomically useful characteristics, such as position and number of genital acetabula, positions of glandularia and lyrifissures, number and arrangement of setae on the body plates, and modification of claws on the tarsi of the legs, are still not adequately explained. For example, although the genital acetabula are plesiotypically borne in the gonopore in most Hydryphantoidea and Lebertioidea, they are either fused with the genital flaps or incorporated into acetabular plates flanking the gonopore in most taxa of other superfamilies (Figure 25.30). In addition, the number of acetabula has proliferated from the ancestral complement of three pairs independently in each of the six largest superfamilies. Alberti (1979) proposed that acetabula in water mites function as osmoregulatory chloride epithelia, and Barr (1982) concluded that analysis of morphological data for early derivative taxa supports this hypothesis. However, until the function of acetabula is clearly understood on the basis of well-designed experiments, the significance of the trend to increasing number of acetabula will remain speculative. The often striking and elaborate color patterns of adult water mites present an intriguing enigma. Members of most ancient clades (e.g., Hydrachnoidea, Eylaoidea, Hydryphantoidea) have colorless integument but are red, like terrestrial parasitengones, due to the presence of

Chapter | 25  Subphylum Chelicerata, Class Arachnida

633

D

E

F

G

FIGURE 25.32 (a) Koenikea wolcotti Viets (Unionicolidae), female. (b) Uchidastygacarus acadiensis Smith (Chappuisididae), female. (c) Paramideopsis susanae Smith (Nudomideopsidae), female. (d) Volsellacarus sabulonus Cook (Neoacaridae), female, pedipalp.

pigment granules that appear to be distributed throughout the body. Members of other superfamilies, except for taxa in interstitial habitats, exhibit highly distinctive patterns resulting from symmetrically arranged concentrations of pigment granules of various colors. In soft-bodied taxa the pigments are located beneath the integument, but in sclerotized groups they are incorporated into the plates. Members of well-sclerotized taxa of Lebertioidea, Hygrobatoidea, and Arrenuroidea may be various shades of red, orange, yellow, green, or blue (Figure 25.33), and the dorsal and ventral shields frequently exhibit intricate patterns combining several contrasting colors. The dorsal patterns of these mites can readily be interpreted as disruptive camouflage, but the adaptive value of their bright colors is more difficult to understand and may be a combination of aposematism and photoprotection (Proctor and Garga, 2004).

Internal Anatomy The organ systems of water mites occupy the haemocoel, bathed by haemolymph which is circulated by movements of the body musculature (Schmidt, 1935; Bader, 1938). As is typical for arthropods, there is no closed circulatory system.

Digestive System Digestion of food materials begins pre-orally, and only fluids are ingested. Food is drawn into the mouth (or buccal cavity) by the muscular pharynx, then passed through the tubular esophagus to the lobed midgut (Bader, 1938), where digestion and absorption occur. Undigested material accumulates as insoluble particles in a posterodorsal lobe of the midgut. All lobes of the midgut end blindly, and there is no connection between the gut and the excretory pore. Excretory System This system consists of a large, thin-walled excretory tubule, apparently derived from the primitive hindgut that lies dorsal to, and in close contact with, the midgut. Waste products are absorbed from haemolymph and stored in the excretory tubule as insoluble, whitish or yellowish crystals of guanine and possibly other chemicals. When filled with crystalline material, the excretory tubule may be visible through the dorsal integument as a yellowish-white “T”or “Y”- shaped structure. The excretory tubule connects ventrally with the excretory pore, and is evacuated periodically by pressure generated through movements of the body muscles.

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(a)

(b)

(c)

(d)

FIGURE 25.33  Arrenurus species of different shapes and colors. (a) A. (Megaluracarus) birgei female; (b) A. (Megaluracarus) rotundus male; (c) A. (Megaluracarus) megalurus male; (d) A. (Arrenurus) bleptopetiolatus male. Compilation of images from Bruce Smith.

Osmoregulation is apparently accomplished by the urstigmata in larvae, and by the genital acetabula in deutonymphs and adults (Alberti, 1979; Barr, 1982). Both of these structures have porous caps that apparently provide an ample surface area of chloride epithelium for maintaining internal ionic balance. Respiratory System Many groups of water mites retain paired stigmata located between the bases of the chelicerae in all active instars. Plesiotypically, the stigmata lead to tracheal trunks, which anastomose repeatedly into tracheolar tubules extending to all parts of the body. However, they appear to be nonfunctional in deutonymphs and adults, and respiration occurs by diffusion through the integument. In adults of many large species inhabiting standing water, a network of closed tubes of tracheolar dimensions lying beneath the integument transports gases to and from the tracheae that lead to internal organs (Mitchell, 1972). Heavily sclerotized mites have the body plates well supplied with regularly arranged pores that permit diffusion of gases between the tracheolar loops and surrounding water through areas of thin integument. In some species, there are regions of tracheal anastomosis lateral to the brain (Wiles, 1984). Neural System The so-called brain, a fused, undifferentiated central ganglionic mass, surrounds the esophagus. Nerve trunks dorsal to

the esophagus lead to the anterior sense organs and mouthparts, and ventral trunks lead to the legs and genital region. The lateral eyes appear to be the primary light-sensing structures in water mites. There are typically two pairs of lateral eyes, with those on each side located close together and often enclosed in lens-like capsules that lie above, within, or beneath the integument, depending on the taxon. In postlarval instars of Eylaoidea, the eyes are borne on a median sclerite. Many Hydryphantoidea also have an unencapsulated median eyespot between the lateral eyes, but this structure is absent in most members of the other superfamilies. Muscles attached to the inner parts of the lateral eyes can move these structures, and eye oscillation is readily seen in living mites subjected to changes in incident light, but the function of eye movements in water mites is not clear. The eyes seem to function as ocelli, permitting the mites to detect the intensity and direction, and, at least in some taxa, the wavelength of incident light, but not to resolve images. Experiments with species of Unionicola have demonstrated that some species respond differentially to various wavelengths and that the response of the mites can be influenced by chemicals produced and released by their mollusc hosts (Roberts et al., 1978). Retinal development, albeit rudimentary, has been reported for a variety of taxa (Lang, 1905). All water mites have specialized tactile organs, called glandularia, distributed in pairs on both the dorsum and venter of the body. Each glandularium consists of a small, goblet-shaped sac, or gland, with a tiny external opening, and an associated seta. The gland contains a milky, viscous

Chapter | 25  Subphylum Chelicerata, Class Arachnida

fluid that is ejected abruptly when the seta is stimulated. This substance quickly stiffens to a sticky gel after it comes in contact with water, and apparently evolved as a deterrent to attackers. In males of Arrenurus, however, material secreted by the glandularia seems to function also in cementing the body of the female in place during copulation (Proctor and Wilkinson, 2001). The five pairs of lyrifissures that are arranged in series on the dorsum and venter of the body are thought to be proprioceptors (Krantz and Walter, 2009). In addition, the distal segments of the palps and legs are well supplied with a variety of specialized setiform structures, apparently derived from outgrowths of the integument, which function as chemoreceptors (Baker, 1996). The most conspicuous of these organs are the solenidia that adorn the dorsal surfaces of the genua, tibiae, and tarsi of the legs, and the tarsi of the palps. Reproductive System Males have paired testes and vasa deferentia that lead to an elaborate ejaculatory complex positioned immediately internal to the gonopore, and consisting of a series of membranous chambers attached to a sclerotized framework (Barr, 1972). The ejaculatory complex functions as a syringe-like organ for compacting masses of spermatozoa, assembling spermatophores, and expelling them from the genital tract through the gonopore. Females have paired ovaries that are more or less fused, and paired oviducts that also fuse to form a single duct leading to the genital chamber within the gonopore. Paired spermathecae, for storing spermatozoa picked up in spermatophores, flank the genital chamber and are connected to it by short ducts.

Life History The basic life history pattern of water mites is shown in Figure 25.21 and has been studied in a wide range of taxa (see Smith and Oliver 1986 for a summary of relevant literature). The first well-documented life-history studies were of Palearctic species, especially members of the genus Arrenurus, investigated by Münchberg (1935a,b). Subsequently, Mitchell (1959) dealt intensively with a number of Nearctic species of Arrenurus, and Böttger (1972a,b) described life cycles of representative Palearctic species of Hydrachna, Limnochares, Eylais, Limnesia, Unionicola, Piona, and Arrenurus in considerable detail. A meticulously comprehensive account of the life history of Hydrodroma despiciens (Müller) (family Hydrodromidae) was published by Meyer (1985). Many species of water mites in temperate latitudes typically live for just over 1 year, most of which is spent in the deutonymphal and adult stages. Information is lacking on those living in tropical regions. Limnocharids, and various species living in temperate zone temporary waters, are exceptional in that adults will normally live several

635

years in the laboratory and presumably also in nature (B.P. Smith, personal observation). Most species are univoltine, with long-lived female adults producing multiple clutches of eggs. Iteroparity may reduce risks associated with the parasitic larval stage, as larvae of most species can survive only about 1 week without a host. Species in ephemeral habitats (e.g., Arrenurus spp. in temporary pools) have a short (ca. 2-week) concentrated period of oviposition, and the adult either dies shortly after or spends most of the year in a reproductive diapause. In temperate latitudes, hosts are usually seasonally limited, and parasitism occurs mainly in late spring and early summer; if tropical regions have well defined wet and dry seasons, parasitism of hosts should also reflect seasonality and host availability. Larvae of most species of Hygrobatoidea, Lebertioidea, and Arrenuroidea spend no more than several days on their hosts, and engorgement results in modest growth, typically a two- to 16-fold increase in weight and volume. Larvae of Eylaioidea, Hydrachnoidea, and odonate-associated Arrenurus spp. have longer associations and proportionately much greater larval engorgement. In the temperate zone, deutonymphal water mites typically feed and grow throughout the summer, and adults appear in late summer or early fall and mate almost immediately. Inseminated females overwinter and lay fertilized eggs the following spring. Reproductive diapause is more likely to be facultative in lower latitudes, and can be absent altogether in tropical populations. Seasonal patterns in abundance of deutonymphs and adults may still occur in the tropics, especially when there are well-defined wet and dry seasons. Several derivative life-history strategies occur among water mites, representing variations on this basic pattern (Smith and Oliver, 1986; Smith, 1988, 1998, 1999). One of these involves elimination of larval feeding and host association altogether, thus avoiding risk of not encountering a host but foregoing host-mediated dispersal (Smith, 1988; Bohonak et al., 2004). Loss of parasitism has evolved many times in unrelated genera in a wide diversity of families. Comparison of closely related species pairs with and without larval parasitism shows that loss of parasitism correlates with accelerated maturity and metamorphosis to adult at a smaller body size (Smith, 1998; Bohonak et al., 2004). Removal of the seasonal limitations of host availability in species with nonparasitic larvae appears to favor a shift to a multivoltine life history, loss of reproductive diapause, and production of smaller clutches of eggs over a longer period of time. Several clades exhibit a third strategy of greatly extending the duration of attachment to the host, in some cases for several months. Species in these groups exhibit typically undergo remarkable growth during the larvae stage, increasing in volume by 600 times or more their original size (Münchburg, 1960; Davids, 1973). Growth of these mites during the deutonymphal stage is much more

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modest, and this period often lasts only a few days or weeks. Deutonymphs in a few species with massive larval growth do not need to feed at all (Münchburg, 1960). Extended larval association with hosts is exhibited by all species of Hydrachnoidea and Eylaioidea (excluding Limnochares), and also by atypical members of a few other taxa, such as Hydryphantes tenuabilis Marshall (Lanciani, 1971a) and certain species of Arrenurus apparently belonging to the circumtropical subgenus Brevicaudaturus (Münchburg, 1960). Nearly all known examples of extended association involve water mite species parasitic on aquatic Hemiptera and Coleoptera, presumably reflecting the longevity, large size, and mainly aquatic existence of these hosts. The only reported exceptions are the species of Arrenurus (Brevicaudaturus) mentioned above that parasitize large libellulid dragonflies. This type of life history allows at least some species to adopt multivoltine life cycles (Davids, 1973). Variation in these major life history patterns can be seen even among congeneric species (Lanciani, 1971b; Cook et al., 1989).

Eggs Eggs are typically laid in clusters in a gelatinous matrix attached to plants, wood, or stones, but females of some species, such as Midea expansa Marshall, may scatter them individually on the substrate. Females of the genus Hydrachna use a unique elongate ovipositor to lay eggs individually in stems of aquatic plants, and those of certain species of Unionicola use stiff modified setae on the genital plates to prise open the tissues of the sponges or mussels in which they lay

SECTION | VI  Phylum Arthropoda

their eggs. The most comprehensive morphological study of water mite eggs was published by Sokolow (1977). Detailed accounts of oviposition behavior have been published by Ellis-Adam and Davids (1970) and Davids (1973). Lifetime egg production varies greatly among species. For example, females of Eylais discreta Koenike may produce individual clutches of 1000–2000 eggs, with one female recorded as producing over 13,000 eggs in a 3-month period (Davids, 1973). At the other extreme, species of Todothyas, Hygrobates, Piona, and Arrenurus that forego larval feeding typically produce two to five eggs per clutch (Smith, 1988). Most water mite species typically produce about a dozen to several hundred eggs in a clutch. There is a clear trade-off between clutch size and egg size, probably related to risks to the larval instar and the proportion of total growth that occurs during this stage. An analysis of this trade-off among 27 species of Arrenurus revealed two strategies defined by egg size, clutch size, and female size, the one with few, large eggs represented by species parasitic on nematoceran flies and that with many, small eggs those species parasitic on Odonata (Cook et al., 1989). Female water mites appear to possess adaptations for selecting appropriate oviposition sites, based on convergence in oviposition site choice among individuals. Large, communal clutches of eggs have been observed for several species. Patches of eggs of Eylais euryhalina Smith in western Canada frequently cover areas of more than 100 cm2 on plant or rock substrata. Massive rafts of Eylais larvae observed in Alaska implies similar mass-oviposition behavior (Figure 25.34). Females of Arrenurus pseudosuperior Cook produce communal egg masses entirely covering

FIGURE 25.34  Successively closer views of a raft of larval Eylais on East Mackey Lake, Alaska. Compilation of images from David Wartinbee.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

submerged oak leaves in central Canadian lakes. The stimulus to lay eggs communally may stem from a shortage of oviposition sites in nature, or it may be that the first clutches chemically stimulate oviposition by subsequent females. There may be a selective advantage to mass oviposition if eggs are distasteful or if their sheer numbers exceed what local predators can consume. Within the egg membrane, water mites pass through a transitory prelarval instar (Meyer, 1985) and then usually develop rapidly and directly into larvae. Just prior to eclosion, fully formed larvae can be observed moving within the egg membrane 1–3 weeks after oviposition. Arrested larval development has been reported in certain species of Unionicola, Teutonia, Utaxatax, and Laversia, whose ­larvae do not emerge until at least 6 months after eggs are laid ­(Mitchell, 1955).

Larvae The vast majority of water mite species have larvae that are parasitic on winged adult insects (reviewed by Smith (1976), Smith and Oliver (1986) and summarized in Table 25.2), with the host providing nutrition as well as a means of dispersal. Engorgement on the host is modest in many clades, but feeding is apparently essential for development, as there are no records of purely phoretic associations. Mitchell (1969a) calculated the probability of success for larval water mites as the product of the probabilities of discovering a host, attaching at an appropriate site on the host, completing engorgement on the host, and detaching from the host in a suitable habitat. The risk of failure can be high. Collins (1975) estimated that more than 75% of larval Wandesia (Partnuniella) thermalis Viets fail to locate a host in a system in which the distribution of hosts is extremely clustered and unpredictable (Collins et al., 1976). By comparing numbers of eggs found on the substrate in a lake to numbers of parasitic mites found on hosts, Meyer (1985) estimated that nearly 99% of larval H. despiciens die before locating a host. Searching for Hosts Above or Below the Water’s Surface In many species, larvae can survive only about a week without a host, and the likelihood of successfully parasitizing a host declines after even a few days. Insect hosts provide water mites with both the source of nutrition necessary for larval growth and their primary dispersal mechanism. Host associations of larval water mites were reviewed by Smith and Oliver (1986) and are summarized in Table 25.2. After hatching, larvae of Hydrovolzia and most genera of Eylaoidea and Hydryphantoidea rise to the water’s surface search for potential hosts on the aerial half of the surface film, presumably the plesiotypical terrestrial behavior. In contrast, larvae of Acherontacarus, Rhyncholimnochares,

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Wandesia, Hydrachna, Lebertioidea, Hygrobatoidea and Arrenuroidea search for hosts by swimming in the water column or crawling on the substrate under water. This behavior arose independently several times, within the superfamilies Hydrovolzioidea, Eylaioidea, and Hydryphantoidea, in ancestral Hydrachnoidea, and again in ancestor(s) of the other three superfamilies. Surface-dwelling larvae are essentially terrestrial, and typically can locate hosts only when they reach or pass through the surface film. Larvae of Limnochares americana Lundblad are unusual in being able to search for hosts on emergent plants up to 50 cm from the water surface. Opportunities to contact hosts occur irregularly in space and time, and are strongly influenced by the structure of a host population. For example, parasitic larvae of Todothyas barbigera (Viets) are found only on parous female mosquitoes, and those of L. americana Lundblad are found predominantly on territorial male dragonflies (Smith, 1999). Some water mites with terrestrial larvae reduce the risks of failure by using a relatively broad range of host organisms and producing large numbers of eggs. For example, larvae of Hydrovolzia and certain genera of Hydryphantidae exploit a wide range of hosts including insects that are only casual visitors to the habitat of the mites. This strategy presumably results in considerable wastage, as larvae attaching to hosts that do not return to suitable mite habitats will die. Larvae of most Eylaoidea (Eylais, Piersigia, Neolimnochares, some Limnochares) and various Hydryphantidae have strong specificity for particular groups of insect hosts that live on the surface film (e.g., Hydrometridae, Gerridae), regularly visit it to replenish air supplies (e.g., aquatic Hemiptera and Coleoptera), or pass through it just before ecdysis and regularly revisit it after emergence (e.g., Odonata, Trichoptera, Diptera). This strategy improves the probability that a larva finding a host will be returned to an appropriate habitat, but limits the availability of potential hosts. These mites apparently compensate for the high risk of failure in finding hosts by producing many hundreds of eggs per female. Species with terrestrial larvae are generally limited to relatively shallow-water microhabitats, given that these larvae must swim or crawl to the surface in order to search for potential hosts. Enclosure experiments by one of us (B.P. Smith) indicated that the likelihood of water striders of the genus Gerris being parasitized by larval Limnochares aquatica L. is strongly dependent on water depth. Differences in rates of infestation are substantial, even between insects caged at 0.5 m deep water and those caged at 1.5 m. In contrast, larvae that search for hosts subaquatically can actively seek them in the water column or on the substrate. Larvae of Hydrachnidae exploit virtually the same range of hosts as Eylaidae, but locate hosts beneath the surface film. Members of certain Anisitsiellidae and Teutoniidae (Lebertioidea), as well as Krendowskiidae and Arrenuridae, seek

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their hosts above the substrate, but exhibit preparasitic attendance of hosts during the host’s penultimate instar prior to ecdysis. Their larvae parasitize tanypodine Chironomidae, Culicidae, Ceratopogonidae, Chaoboridae, and Odonata, all groups that have the final aquatic instar active in the water column below the surface film. Mites locating their hosts subaquatically apparently exploit them more efficiently than do those with terrestrial larvae, as their females tend to lay smaller, though still substantial, numbers of eggs. For example, females of Hydrachna conjecta (Koenike) lay only about 10% as many eggs as those of E. discreta Koenike, yet these species cooccur in nature and parasitize the same species of corixids (Davids, 1973). Larvae of Rhyncholimnochares and Wandesia are also aquatic and apparently independently evolved adaptations to locate their hosts in the benthic substrate. Larvae of ­Rhyncholimnochares are subelytral parasites of adult elmids that remain in water. Those of Wandesia ­pre-parasitically attend nymphal stoneflies before transferring to the emerging adults and becoming parasites. Interestingly, larval stygothrombidiids exhibit similar behavior in exploiting the same hosts. Larvae of the remaining families of Lebertioidea, Hygrobatoidea, and Arrenuroidea locate ­ final instar larvae or pupae of their hosts in cases or burrows in the substrate. Their hosts, various Chironomidae or ­Trichoptera, construct cases or retreats where they remain during prepupal and pupal stages. Larvae in all of these clades attend the host during the preparasitic phase, clinging to the integument of the pupa until it rises to the surface. They typically then transfer to the imago as it emerges, embed their chelicerae, and enter the parasitic phase. Individual females in these groups lay relatively small numbers of eggs (often fewer than 10), indicating both that the larvae efficiently find hosts and that the hosts that they select have a high probability of returning to a suitable habitat. The development of fully aquatic larvae was a crucial preadaptation for ancestors of the various groups of Lebertioidea, Hygrobatoidea, and Arrenuroidea that became specialized to exploit nematocerous Diptera as hosts. The origin and evolution of modern genera and families of these mites apparently followed the explosive radiation of the Nematocera, and especially Chironomidae, during the Jurassic and early Cretaceous. Refinement of their adaptations for parasitizing these flies permitted the so-called higher water mites to co-diversify with them, and to exploit their potential for dispersal and invasion of new habitats. Variations among Taxa in Attachment Sites on Host Larvae of many groups of water mites exhibit strong selectivity for attaching to particular sites on the body of the host (Smith and Oliver, 1986; Martin, 2004), as shown in Table 25.2. It is not clear why these apparent taxon-specific preferences for certain parts of the host’s body exist. Presumably,

SECTION | VI  Phylum Arthropoda

some of the preference reflects selective pressures to not interfere with the host’s ability to disperse. Larvae of Hydrovolzia and certain genera of Hydryphantidae that parasitize Hemiptera, Plecoptera and Trichoptera (e.g., Protzia, Wandesia), seem to attach rather indiscriminately to various parts of the host’s body. Those of all other groups are more selective. Larval Eylais are essentially terrestrial organisms and select attachment sites bathed by the host’s air supplies, under the elytra of aquatic bugs and beetles (Figure 25.35(a)). The fully aquatic larvae of Hydrachna are able to utilize a much wider range of sites on the same hosts (Figure 25.35(b)). Larval hydryphantoid mites that parasitize Diptera tend to attach to thoracic sites, exhibiting what might be plesiotypical behavior. Larvae of many early derivative groups of Lebertioidea, and most Arrenuroidea parasitizing Diptera, also prefer thoracic sites. These larvae tend to be relatively large, and only a few individuals can share a single host. When more than one of these larvae attach to the same part of the host’s thorax, they are usually symmetrically arranged on either side of the body. Larvae of some species of Sperchon, Lebertia, Oxus, and Torrenticola can use the anterior abdominal segments of hosts when thoracic sites are already occupied. In virtually all Hygrobatoidea (except Limnesiidae, which may not be closely related to other taxa in the superfamily) and a few families of Arrenuroidea (e.g., Mideidae, Momoniidae), larvae attach exclusively to abdominal sites on their hosts, usually nematocerous flies but in some cases caddisflies. This likely apotypical behavior correlates with a marked trend to smaller size, allowing several larvae to share each attachment site on a host. These small larvae occupying abdominal attachment sites can exploit even the smallest species of Chironomidae as hosts without preventing them from flying and dispersing (Figure 25.35(d)). The relatively small larvae of various species of Arrenurus (sensu stricto) show preferences for either thoracic or abdominal sites on odonate hosts (Figure 25.35(c)). Duration of the parasitic phase varies considerably. Larvae of certain species in early derivative genera such as Limnochares and Hydryphantes that feed on hemimetabolous insects retain their mobility and can transfer from one instar to the next of the same host individual. Larval L. aquatica engorge gradually by feeding on successive host instars, but larvae of H. tenuabilis exhibit true preparasitic attendance and delay feeding until the host reaches adulthood. This strategy of tracking hemimetabolous hosts through several molts may reflect the plesiotypical condition in water mites. Larval Hydrachna and Eylais remain attached to the same adult bug or beetle throughout the parasitic phase, for several weeks or more. Larvae of species in these families that are adapted to temporary pools often remain on hosts throughout the dry phase of the habitat, and engorge only after a period of arrested

Chapter | 25  Subphylum Chelicerata, Class Arachnida

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(a)

(b)

(c)

(d)

FIGURE 25.35  (a) Larvae of Eylais sp. (Eylaidae) attached to the subelytral abdominal integument of adult water boatmen (Heteroptera: Corixidae) illustrating larval size before and after engorgement. (b) Larvae of Hydrachna sp. (Hydrachnidae) attached to the thorax and elytra of a giant water bug (Heteroptera: Belostomatidae). (c) Larvae of Arrenurus sp. (Arrenuridae) attached to the abdominal segments of a damselfly (Odonata: Zygoptera). (d) Larvae of Feltria sp. (Feltriidae) and Aturus sp. (Aturidae) attached to the abdominal segments of a chironomid midge (Diptera: Chironomidae).

development that may last as long as 10 months. Larvae of species parasitizing nematocerous Diptera or other aerial insects typically engorge rapidly and mature within several days. Fully fed larvae then enter the quiescent (calyptostatic) protonymph stage. Larvae of Hydrachna, Eylais, and certain Hydryphantidae grow substantially, increasing several hundred–fold in size in some cases, while feeding on their relatively long-lived hosts. Larvae of many relatively recently evolved groups associated with short-lived insects, especially nematocerous Diptera, still engorge on fluids from the host but increase more modestly in size in the process. The dominant strategic role of the larval instar in the life history of most Hydryphantoidea, Lebertioidea, Hygrobatoidea, and Arrenuroidea appears to be dispersal rather than growth. Species that have the parasitic larval stage suppressed illustrate that development of large eggs can obviate the need for larval feeding. However, the relatively rarity of this phenomenon attests to the crucial role of larval dispersal in more derived water mite species. Most feeding and growth occur during the deutonymph and adult stages in these mites. The larva is suppressed as an active instar in a few unrelated species in widely divergent genera (e.g., Todothyas, Lebertia, Limnesia, Hygrobates, Piona, Axonopsis, and Arrenurus), apparently permitting the mites to accelerate development and maximize exploitation of optimal

conditions (Smith and Oliver, 1986; Smith, 1988). In these cases, larvae either remain within egg membranes or emerge for a brief period of activity before entering the protonymph stage. Species known to forego a parasitic phase appear to be isolated and ephemeral evolutionary experiments (Smith and Oliver, 1986; Smith, 1988). There are relatively high levels of morphological divergence and lower levels of heterozygosity within populations of apparently recently diverged species lacking parasitism when compared with closely related species that retain parasitic larvae (Bohonak, 1999; Bohonak et al., 2004). Although elimination of parasitism may confer a short-term advantage in larval survival and potential for population growth, given the absence of diverse lineages of nonparasitic water mites, it seems that host-mediated dispersal is essential for long-term persistence of a lineage. Seeking and Locating Hosts After emerging, larvae of most species with a parasitic phase immediately begin host seeking. The period of survival for larvae in the preparasitic phase ranges from 4 days to 6 weeks (Smith, 1988) but is about 1 week for most species. Larval Arrenurus remain active and exhibit preparasitic attendance over a period of 2 weeks in the laboratory but have the greatest chance of locating and infecting a host during the first few days. Larvae older than 1 week are rarely able to successfully attach to hosts.

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Some authors have suggested that water mite larvae find hosts by accidental contact, but close observation of both terrestrial and aquatic larvae reveals noticeable changes in larval behavior when near hosts. Terrestrial larvae seem to use visual, tactile, and chemical cues to locate hosts at the surface film. Those of several Hydryphantidae have hind legs modified for jumping and leap toward objects positioned above them, an obvious adaptation for encountering hosts on the surface film. Larval L. aquatica exhibit questing behavior when within several millimetres of water striders that involves hesitation and reaching upward with the first pair of legs. Details of the sensory mechanisms used by aquatic larvae to locate hosts remain unclear, but there is evidence that both chemical and tactile cues are important (Böttger, 1972b). Larvae of Hydrachna often extend the gnathosoma when approaching a host before actual contact is made. Larval Arrenurus are attracted to mosquito pupae and slow their swimming just prior to contact (Smith and McIver, 1984b). These larvae swim in tight circles, as if attempting to orient to some cue, before making a direct approach (B.P. Smith, personal observation). Recognizing Hosts Some water mite species with aerial larvae have remarkably broad host spectra encompassing several orders of insects, whereas species with preparasitic attendance are normally limited to species within a subfamily or family of insects. Commonly, one or two host species carry the majority of parasitic larvae of a given mite species in a particular habitat. Although co-occurrence in space and time set the absolute boundaries for host utilization, larval mites also actively select among potential host species. In laboratory experiments, larval mites of diverse genera such as Hydrachna, Eylais, Limnochares, and Arrenurus exhibit preferences when exposed simultaneously to hosts of various species (Smith and McIver, 1984b). In addition, there is evidence that larval mites favor certain individuals within a host population while rejecting others (Smith and McIver, 1984b). In some instances, hosts react strongly to the presence of mite larvae with avoidance behavior or vigorous grooming activity (Smith and McIver, 1984b; Forbes and Baker, 1990), and there may be differences in intensity of parasite avoidance among host species and individuals. Strong bias for hosts of a specific sex or age class is evident in many water mite species with terrestrial larvae, apparently resulting less from choice than from differences in the likelihood of mites encountering males and females of particular ages as determined by host behavior (Smith, 1999). Robb and Forbes (2006) found that in nature, newly emerged females of the damselfly Lestes disjunctus were more heavily parasitized than males. This bias could not be explained by host body size or times of emergence, but could be attributed to the difference between the sexes of

SECTION | VI  Phylum Arthropoda

the host in the duration of the last preadult instar and thus its exposure to searching mite larvae. Among mosquitoes, dragonflies, and some other host groups, the likelihood of an adult returning to water is strongly influenced by its sex (females having to return to oviposit), and this should result in selective pressure for larval mites to recognize the sex of potential hosts. Lanciani (1988) observed sex-biased selection of hosts by larval Arrenurus subaquatically searching for mosquito pupae. He found that the mites exhibited a small but significant preference for female pupae of Anopheles crucians, given equivalent exposure to both sexes of host pupae and adjusting for any sex-related differences in body size. Selecting a Feeding Site on the Host After encountering a potential host, terrestrial larvae locate an appropriate attachment site and begin to feed. Species with larvae that exhibit preparasitic attendance must transfer from the penultimate instar of the host to the imago during ecdysis. Larval Arrenurus cling passively to the preimaginal host instar, becoming active only when it begins to molt. Interestingly, many larvae of species parasitizing mosquitoes appear to have difficulty in locating the split in the pupal exuvia, and some that do find it are unable to penetrate the surface film quickly enough to reach the adult mosquito as it emerges. As a result, 30–50% of preparasitic larvae fail to attach to the adult host (Smith and McIver, 1984b). Larvae of Arrenurus species parasitic on odonates are carried out of the water on the penultimate instar of the host, and almost all of them successfully move from the larval exuvia onto the adult odonate (Mitchell, 1969b). Larvae of species of Atractides, Hygrobates, and Unionicola pierce the pupal cuticle of their chironomid hosts and embed their chelicerae in the pharate adult. This behavior virtually ensures successful transfer to the adult, because the larvae are passively pulled through the pupal exuvia as the host emerges (Böttger, 1972b). Little is known about the mechanisms by which mite larvae select feeding sites on hosts, and virtually all available information pertains to a few species of the genus Arrenurus. Site selection of larval Arrenurus on odonates appears to be largely a function of timing. Progress of the host’s ecdysis at the point when mite larvae become active determines where the mites first come in contact with the host, and they typically attach close to that point. Larvae experimentally removed from the host and placed elsewhere on the body readily attached there. In contrast, there appears to be little correlation between the process of host ecdysis and site selection in larval Arrenurus parasitizing mosquitoes. For example, larval Arrenurus kenki almost always attach between the head and thorax when infesting adult Aedes excrucians but to the abdomen when attacking Aedes cinereus, without reference to the amount of time taken by the host for ecdysis.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

Attaching to Host and Engorgement Larval mites hold onto the host with their palps and use them to produce the leverage needed to pierce the cuticle of the host with their chelicerae. Larvae of some species of Arrenurus secrete a substance that solidifies and apparently helps to cement the mite to the host (Åbro, 1984), but this has not been observed in other cases. After attachment, larvae of many genera form a feeding tube, or stylostome, in the tissues of the host (Smith, 2003). Gross structure of stylostomes appears to be consistent among species in a genus, but differs greatly between genera. Stylostomes formed by species of Eylaoidea, Hydrachnoidea, and some Hydryphantoidea are multibranched and potentially open-ended structures (Davids, 1973), whereas those of species of Hygrobatoidea and Arrenuroidea are simple, blind tubes (Smith, 2003). Stylostomes have not been reported for species of Lebertioidea or certain species of Hydryphantoidea. Details of stylostome morphology appear to be constant for a given species of mite regardless of host but differ among mite species, leading to the conclusion that their basic structure is determined by the larval mites rather than their hosts. Stylostomes are acellular, consisting of acid mucopolysaccharides, and have no perforations other than a terminal pore, when present. The mechanism of stylostome function is not well understood. Davids (1973) argued that the stylostomes of larval Hydrachna are formed by coagulation of host haemolymph in reaction to mite saliva, and that the mites imbibe haemolymph from the host. Stylostomes of Arrenurus larvae are typically surrounded by a fluid-filled abscess, and it has been suggested that the mites feed upon liquefied tissues in much the same way as larval chiggers (Åbro, 1984). It is possible that the different forms of stylostomes formed by larvae in these remotely related clades are not homologous. Some host species mount an effective immune response to stylostomes. For example, adults of the dragonfly Sympetrum internum are nearly always successful at blocking engorgement by larval Arrenurus planus, although members of the co-occurring species Sympetrum obtrusum are highly susceptible to parasitism by these mites (Forbes et al., 2002). Partial immunity has been reported for various damselfly species, related to differences in the host’s age, sex, condition, and date of emergence (Robb and Forbes, 2005, 2006). Immune responses to parasitic larval water mites have also been observed in water boatmen, mosquitoes, and chironomids, and these defenses are presumably widespread among host taxa. Duration of the period of larval engorgement varies considerably among water mite taxa. Larvae of L. aquatica and H. tenuabilis require 6–13 days to engorge (Lanciani, 1971a; Böttger, 1972a; Smith, 1989), but those of various species of Hydrachna and Eylais may spend from 2 weeks to 10 months on the host (Davids, 1973; Smith, 1988;

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­ iggins et al., 1980). Larvae of some species of Eylais W can complete engorgement within 2 weeks at the time of year when female hosts are producing eggs, but apparently undergo diapause in a partially engorged state at other times of the year (Smith, 1988). Presumably, growth rate of ­larvae in these groups is affected by the physiological condition of the host. Larvae associated with short-lived insects, especially nematocerous Diptera, require shorter times for engorgement. Those of pionid species inhabiting temporary pools, such as Piona napio and Tiphys americanus, can engorge in as little as 24 h on their chironomid hosts. ­Similarly, ­larvae of Unionicola foili spend an average of 3 days attached to chironomids (Edwards and Dimock, 1995), and those of Arrenurus feeding on mosquitoes take approximately 6 days for full engorgement (Mullen, 1974). Some species of Arrenurus (e.g., Arrenurus papillator in Spain, ­Arrenurus madaraszi in Pakistan Reisen and ­Mullen, 1978) are atypical of the genus, as they also appear to diapause on the host in a fully engorged condition, extending the association through inhospitable periods of the year. Larval growth during engorgement is also highly variable. Larvae parasitic on nematocerous flies increase only modestly in size. For example, larvae of Limnesia maculata and Unionicola crassipes expand to 3 to 4.3 times their original volume on their chironomid host, and those of Arrenurus species feeding on mosquitoes increase in volume about 16-fold (Münchberg, 1938; Böttger, 1972b). Larvae of species of the subgenus Arrenurus parasitic on odonates typically augment their volume 5 to 140 times, with an extreme outlier in the subgenus Brevicaudaturus at 1300 times (Münchberg, 1960). Larval Hydrachna and Eylais grow substantially, increasing up to 600-fold in size in some cases (Davids, 1973; Figure 25.35(a)). Larvae parasitizing long-lived hosts for extended periods risk premature forcible detachment when the host molts. Cast skins of nepid and belastomatid bugs often have larval and protonymphal Hydrachna still attached. Larvae of L. aquatica retain their mobility and can transfer from one host instar to the next to continue engorgement. Detaching from Host Following engorgement, mite larvae must detach from the host and return to an aquatic habitat suitable for postlarval development. Larvae of various taxa may use host behavior, along with mechanical, visual, and chemical stimuli, as cues to initiate detachment (Böttger, 1972a,b). Environmental cues such as moisture may be sufficient to induce detachment in some species, but physiological cues from the host are essential in others (Smith and Laughland, 1990). Larvae of the Eurasian species Arrenurus cuspidator parasitic on the damselfly Coenagrion puella require two simultaneous cues to stimulate detachment, high humidity, and either mating or oviposition behavior by the host (Rolff

SECTION | VI  Phylum Arthropoda

642

and M ­ artens, 1997). Edwards and Smith (2003) found that under experimental conditions, larvae of U. foili were more likely to detach into water than would be expected by chance. Although there was no sex bias in the likelihood of success, detachment from females was positively correlated with oviposition behavior by the host. Presumably, the mite larvae can detect hormones or some other physiological correlate with reproductive behavior in the female host. Engorged larval mites risk detaching into unsuitable habitats. Fully fed larvae with limited mobility falling on dry ground quickly perish, and those introduced into unsuitable aquatic habitats may fail to survive or reproduce. At least some species of Arrenurus parasitic on odonates are regularly introduced into lakes that apparently cannot support their reproduction (Mitchell, 1969b). Adult Cenocorixa bifida water boatmen living in inland saline lakes of central ­British Columbia that are too salty for water mites often bear larval Eylais and Hydrachna that were picked up in other, less saline water bodies. Fully fed larvae are capable of only limited movement, and those successfully re-entering water typically seek out plant material or detritus, attach themselves by their chelicerae, and become quiescent. Engorged larvae of Piona and Arrenurus may attempt to swim or crawl for up to 2 days before transforming to protonymphs. Phenology of Larvae In permanent water habitats at north temperate latitudes, most species of water mites overwinter primarily as inseminated females and oviposit in late spring. Consequently, aquatic insect species whose adults emerge in early spring are rarely parasitized by water mites, even within groups that are otherwise suitable hosts. Interestingly, some of these early emerging species show high susceptibility to parasitism under laboratory conditions. For example, although only a small proportion of Aedes communis and Aedes punctor mosquitoes carry larval Arrenurus angustilimbatus and A. kenki in nature, they prove to be more susceptible to parasitism in choice experiments than the species typically utilized as hosts by these mites in the field (Smith and McIver, 1984a,b). A few mite species that apparently overwinter as eggs or ­larvae are able to exploit hosts that emerge in early spring. For example, l­arvae of Huitfeldtia rectipes Thor are found regularly on early season species of Chironomus emerging from oligotrophic lakes in eastern Canada throughout the ice-free period beginning in mid-April. Available evidence suggests only loose synchrony between life cycles of mites and their host species, even when host specificity is strong (Smith and McIver, 1984a,c). In the case of Coquillettidia ­perturbans mosquitoes in Florida parasitized by larval Arrenurus ­danbyensis and Arrenurus delawarensis, peak infestation did not closely correlate with peak abundance of hosts, and there was no close seasonal correspondence between population

densities of the two mite species, although both are dependent on the same host (Lanciani and McLaughlin, 1989). Comparable studies from the tropics are largely lacking: as mentioned earlier, seasonal peaks in occurrence of particular life history stages occurs in regions with well-defined wet and dry seasons.

Protonymph (Nymphochrysalis) Engorged larvae of all groups of water mites enter a quiescent protonymph stage during which larval tissues are resorbed and reorganized and the deutonymph develops. After a few days, fully formed deutonymphs emerge from the larval skins and become active. Mites that parasitize large, long-lived insects, including Hydrachna, most Eylaoidea, and certain Hydryphantidae, are unusual in passing through the protonymphal stage while still attached to the host, within the larval integument. This trait is correlated with extreme growth during the parasitic phase, which renders engorged larvae incapable of locomotion. Along with other adaptations, extended attachment to the host allows many of these mites to exploit periodically temporary habitats. Members of A. planus Marshall and Arrenurus ventropetiolatus Lavers survive for up to 10 months of the year in the dry basins of vernal temporary pools in eastern and western North America, respectively, as diapausing protonymphs and pharate deutonymphs. These mites require a period of desiccation, and probably cold temperatures, to release them from diapause (Wiggins et al., 1980).

Deutonymph The deutonymphal instar is typically active and predaceous, and resembles the adult but is sexually immature. Deutonymphs typically exhibit relatively undeveloped idiosomal sclerites and chaetotaxy compared to adults. They have only a rudimentary (“provisional”) genital field bearing an incomplete complement of acetabula. They feed voraciously, often preying upon immature instars of the same taxa of insects that they parasitized as larvae (see Table 25.2). Deutonymphs and adults of Hygrobates (Lurchibates) spp. of Southeast Asia are quite exceptional, being ectoparasitic on newts, and deutonymphs of at least one of these species are parasites within the mouths of newts (­Goldschmidt and Fu, 2011). At the other extreme, a small number of species forego deutonymphal feeding altogether (e.g., Arrenurus (Brevicaudaturus) spp. Münchburg, 1960). The deutonymphal stage varies in duration from a few days or weeks in early derivative groups such as Hydrachnidae, Eylaoidea, and certain Hydryphantidae, to several months in many groups of Lebertioidea, Hygrobatoidea, and Arrenuroidea. Deutonymphs of some hygrobatoid families are especially long-lived. Those of Pionidae typically live for many months through summer and the following winter, and

Chapter | 25  Subphylum Chelicerata, Class Arachnida

this trait has preadapted certain species to exploit annually temporary pools. In these pionids, deutonymphs are able to endure the dry phase of the cycle in damp retreats in the substrate (Smith, 1976; Wiggins et al., 1980). The deutonymph is the primary growth instar in most groups of water mites, and body size increases dramatically during this stage. On attaining adult size, deutonymphs prepare to transform to tritonymphs by embedding their chelicerae in plant tissue or soft detritus and becoming inactive.

Tritonymph (Imagochrysalis) During this stage, further structural reorganization occurs to produce the adult. This final metamorphosis is rapid, and adults typically emerge within a few days. Mature, fully-fed deutonymphs anchor themselves to living or decaying plant material using the legs and mouthparts as they approach transformation to the tritonymph stage and become quiescent. Tritonymphs of several species of Pionidae inhabiting vernal temporary pools are regularly found attached to aquatic mosses in clusters, with each mite attached at a leaf axil. Conspicuous clumps of up to 500 tritonymphs comprising several species Tiphys and Piona can be observed in moss mats at the edges of these pools as the ice is melting in early spring. The significance of this clustering is not known. The tritonymphal instar remains enclosed by the deutonymphal exoskeleton, and the pharate adult may often be seen forming within it. Mites of various species of Arrenurus typically spend 4–5 days in the tritonymph stage under laboratory conditions at 20 °C. Water mites appear to be particularly vulnerable to poor water quality during the tritonymph stage, perhaps because the metamorphosing adults must exchange gases via diffusion across three levels of cuticle. For example, when dead and decaying prey are present in laboratory cultures, tritonymphs of Arrenurus may take up to 17 days to complete development, and mortality rates of this instar may exceed 50%, despite minimal mortality among deutonymphs and adults living in the same container.

Adults Eclosion and Sclerotization of Exoskeleton Adults emerge from the deutonymphal integument with soft, pliable, and colorless sclerites. The teneral adults immediately become active, crawling or swimming about while the process of sclerotization of the body is completed and distinctive color patterns become evident. Shortly after emergence, both males and females mature sexually and begin to mate. In many taxa, males tend to emerge and become mature a few days earlier than conspecific females, and are ready to mate as soon as females become active in the habitat. Adults of certain species of Limnochares (­Böttger, 1972a), and possibly some Arrenurus, may undergo additional (“supernumary”) molts as adults.

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Mating: Indirect Spermatophore Transfer Water mites exhibit a greater variety of spermatophore transfer methods than any other group of arachnids (­Proctor, 1991a), and can be grouped into categories based on the degree of interaction between males and females during mating (Proctor, 1992a). Complete dissociation involving no chemical communication or physical contact between the sexes has been observed in species of a wide variety of early derivative genera (Proctor, 1992a). In these species, males deposit stalked spermatophores on the substrate for subsequent retrieval by conspecific females, and those of some species have been observed to deposit large numbers when isolated in small containers of water, indicating that chemical cues about female presence are not necessary to induce spermatophore deposition. After finding the spermatophores, perhaps using pheromonal cues, females pick them up in the gonopore. Incomplete dissociation requiring chemical or physical cues from females to stimulate males to deposit spermatophores occurs in several unrelated genera (­Proctor, 1992a). Dissociation during spermatophore transfer is widespread in prostigmatan mites and probably occurs in most groups of water mites in which males and females are morphologically very similar. A more interactive type of mating, pairing of males and females for indirect spermatophore transfer, has been observed in several species. In these cases, males deposit spermatophores on the substrate, then actively assist females to find them and pick them up. Males of some Eylais lead receptive females in a circular dance, stopping repeatedly to deposit spermatophores at the same location on the circumference of the route. Halfway through each revolution of the dance, the female pauses with the male over his mass of spermatophores and picks up one or more of them in her gonopore (Lanciani, 1972). Males of N. papillator Marshall apparently exploit the hunting behavior of females to attract females to spermatophores (Proctor, 1991b, 1992b; Proctor and Wilkinson, 2001). Upon encountering a female, a male vibrates his forelegs rapidly, simulating the stimuli produced by copepod prey, until she responds with characteristic clutching behavior. The male then turns away, deposits a complex array of interconnected spermatophores, and uses his hind legs to fan water over the spermatophores toward the female, presumably directing pheromones toward her. A receptive female then ceases aggression, approaches the spermatophore net, and begins to collect sperm packets. Males of some species of Unionicola also use leg-trembling to generate vibrational cues (Proctor, 1992b; Proctor and Wilkinson, 2001). Mating: Direct Spermatophore Transfer Pairing with direct spermatophore transfer has evolved independently in many water mite lineages. Active transfer

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of spermatophores to females is accomplished either by modified appendages or idiosomal sclerites of males, or by direct gonopore-to-gonopore contact. Proctor (1992a) considered both of these modes as examples of copulation. In Unionicola intermedia, a species that inhabits the mantle cavity of freshwater mussels, males pick up their own spermatophores and carry them between the genua or tibiae of their hind legs while seeking receptive females. On finding a female, the male crawls beneath her and places his spermatophores in her gonopore (Hevers, 1978). Males of many groups of Hygrobatoidea produce unstalked spermatophores that they carry between modified claws on the tarsi of their flexed third pair of legs before transferring them directly into the gonopore of the female. This type of mating has been reported in various species of Feltria, Forelia, Tiphys, Piona, and Brachypoda (see Proctor, 1992a). In each case, the male uses specially modified appendages, usually the fourth pair of legs, to hold the female in a characteristic posture so that he can transfer spermatophores to her gonopore by simply extending his third pair of legs. A variety of complex modes of direct spermatophore transfer occur in Arrenurus. Males of the various species have the idiosoma modified posteriorly to form characteristically shaped “cauda” (Figure 25.33). A male captures a female and positions her so that her gonopore rests over the back edge of his cauda. Secretions from glandularia of the male temporarily cement the pair together, and the male then deposits one or more stalked spermatophores on the substrate. In some species, the male then rocks his body down and forward to bring the gonopore of the female in contact with the spermatophore (Lundblad, 1929). In others, the male’s cauda is further modified with the addition of a petiole, a small spoon- or cup-shaped appendage, to scoop up sperm from deposited spermatophores into a mass that he then inserts into the gonopore of the female (Proctor and Smith, 1994). Proctor and Wilkinson (2001) suggested that the petiole may enable males of the subgenus Arrenurus to circumvent female choice. Variations in copulatory behavior occur throughout the many species of Arrenurus, along with the multitude of different modifications in caudal morphology that characterize the males of various species. Finally, direct transfer of spermatophores from gonopore to gonopore during copulation occurs in certain species of Eylais (Lanciani, 1972) and in the arrenuroid genera Midea (Lundblad, 1929) and Nudomideopsis (I.M. Smith, personal observation). In some of these cases, males either have the gonopore protruding from the venter on a cone-shaped projection (Eylais) or are equipped with wing-like appendages flanking the gonopore (Midea) to facilitate transfer. Direct spermatophore transfer has evolved many times in the Hydrachnidiae. As defined by Proctor (1991a), copulation has developed independently many dozens of times within the water mites, including at least three times within the family Pionidae (Smith, 1976). Direct transfer from

SECTION | VI  Phylum Arthropoda

gonopore to gonopore certainly developed independently in Eylais and Arrenuroidea. Sex pheromones appear to mediate pairing in at least some species of water mites. Males of Limnesia undulata increase spermatophore deposition when females are present, and produce comparable numbers when placed in containers from which females have recently been removed. The vigorous fanning of water over spermatophore nets toward females by males of N. papillator strongly suggests chemical communication (Proctor, 1991b). Females of various species of Arrenurus produce male-arrestant/attractant pheromones that can be extracted from water. Unlike insect pheromones, which are often species-specific, the pheromones of Arrenurus species frequently elicit response in males of several closely related and co-occurring species (Smith and Florentino, 2004). There is also evidence that males of some species of Arrenurus produce pheromones attractive to females (Baker, 1996). Preliminary evidence suggests that the first male to mate with a female has almost total sperm precedence over subsequent mates. Given that sperm is regularly stored for many months over winter, there must be substantial selective pressure for behavioral and phenological adaptations to gain early access to females. Guarding of tritonymphs (presumably females) by young males has been observed in A. planus. Males do not seem to discriminate between virginal and previously mated females. Males of N. papillator attempt to mate with all of the females that they encounter. They eventually produce significantly greater numbers of spermatophores for virginal females, but this appears to be the result of avoidance behavior by inseminated females rather than discrimination by males (Proctor, 1991b). Males of Arrenurus manubriator choose larger females over smaller ones, perhaps selecting for greater egg production capacity (B.P. Smith and L.M. Bovenzi, unpublished data). Males in many groups mate and die within a few days of emerging. Those of some groups, such as Arrenurus, may live for several months. Males of some genera of Foreliinae and Tiphyinae are so modified morphologically that they their movement is limited to awkward swimming or crawling over the substrate using the anterior pairs of legs. They are restricted to crawling slowly over the substrate using the anterior pairs of legs. Males appear to be much less common than females in many genera of water mites (Meyer and Schwoerbel, 1981). This may in part be due to the shorter lifespan of males. Biased sex ratios in the field can also result from phenological protandry, which has been demonstrated in at least some water mite species (Proctor, 1989, 1992b), such that collecting early in the season can result in male-biased catches and collecting later in female-biased ones; however, bias can also occur at oviposition. Females of A. manubriator produce strongly sexbiased clutches (B.P. Smith, unpublished data). The bias for males or females appears to be consistent among successive

Chapter | 25  Subphylum Chelicerata, Class Arachnida

clutches of eggs for a given female, and appears to be a trait largely inherited from the maternal parent (B.P. Smith, unpublished data). The mechanism for this is unclear, as water mites are diploid (and hence cannot determine sex as do haplodiploid species), with no recognizable sex chromosomes (Sokolow, 1954). Mated females may live for many months, continuing to feed while they produce several clutches of eggs. In many temperate-zone groups, mating occurs in late summer and fertilization is delayed while females overwinter. These females lay their eggs in the following spring or early summer. In contrast, females of species inhabiting temporary pools must lay their eggs within a few days of mating to ensure that their offspring reach a life history stage capable of avoiding or withstanding the dry period before water disappears from the habitat (Wiggins et al., 1980; Smith and McIver, 1984c).

Habitats and Assemblages Most water mite species and many monophyletic groups representing genus or family level taxa are restricted to one or a few similar types of habitats. The strong correlation between certain clades and habitats suggests that physiology and/or requirements for locating hosts, prey, mates, and oviposition sites tend to constrain adaptive radiation. Nevertheless, over their evolutionary history, water mites have successfully invaded a great diversity of freshwater habitats. The typical water mite assemblages found in the major types of freshwater habitats in North America are outlined in detail by Smith et al. (2010), and below we include an abbreviated, more globally applicable summary (Table 25.2).

Springs (Including Seepage Areas) A diversity of water mites live in moss and wet detritus associated with rheocrenes and helocrenes. Hydryphantoidea are particularly well represented. Members of ancient clades that may well have originated in this type of habitat are soft-bodied or partly to fully sclerotized. Adults of derivative groups that apparently invaded spring and seepage habitats from flowing water more recently have entire dorsal and ventral shields. Most spring inhabiting species are cold-adapted stenophiles, but some Wandesia (subgenus Partnuniella) and all Thermacarus live only in hot springs.

Riffle Habitats Diverse assemblages of mites live on the substrate in rapidly flowing areas of streams and rivers; samples from a single location may contain up to 50 species in 30 genera. Members of a few early derivative taxa are soft-bodied (e.g., Protzia), but adults of most riffle-associated groups are both strongly flattened and well sclerotized (e.g., Aturus, Figure

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25.31(d)). Most species from these habitats are cold- or cool-adapted stenophiles.

Interstitial Habitats A surprising diversity of mostly small-bodied water mites live in sand and gravel deposits to depths of 1 m or more, mainly in the hyporheic zone of streams and rheocrenes. Deutonymphs and adults crawl through the spaces between particles, often with considerable speed and agility. Members of most early derivative taxa are soft-bodied (e.g., Rhyncholimnochares), and some are also vermiform (e.g., Wandesia). In adults of more recently derived groups, the dorsum and venter are partly covered by various plates or small shields, or are well sclerotized with either entire dorsal and ventral shields or greatly expanded coxal plates (e.g., Uchidastygacarus (Figure 25.32(b)). Mites adapted to interstitial habitats have reduced eyes and lack pigmentation of the integument.

Stenothermic Pools Unlike mites that live in interstitial habitats, those that live in silty substrata in spring-fed pools, fen pools, and depositional areas of streams show little evidence of morphological adaptation to these habitats. Members of ancient taxa are soft-bodied, whereas adults of early derivative clades exhibit various degrees of sclerotization ranging from slight enlargement of dorsal and ventral plates to development of entire dorsal and ventral shields. Adults of certain taxa that invaded pools relatively recently from lotic and hyporheic habitats have become secondarily adapted for swimming (e.g., Axonopsis, Mideopsis). Certain groups inhabiting depositional areas of streams (e.g., Teutonia, Oxus, ­Wettina) include the fastest and most agile swimmers among the water mites. Most inhabitants of pools are cool-adapted stenophiles.

Lakes, Permanent Ponds, Marshes, Swamps, and Bogs Members of ancient and early derivative genera found in large standing water bodies are generally soft-bodied (e.g., Hydrodroma, Limnesia, Hygrobates). Adults of many genera in more recently evolved clades have extensive dorsal and ventral shields, in at least some cases because the group has secondarily invaded standing water from lotic or hyporheic habitats (e.g., Axonopsis, Mideopsis). Most species are strong swimmers and have fringes of long, slender setae on the genua and tibiae of the second, third, and fourth pairs of legs (Figure 25.31(a) and (b)). These swimming setae help to propel the mites as they move about at the surface of the substrate or among plants. Some species of Unionicola and Piona have unusually long swimming setae and are essentially pelagic or planktonic (Riessen, 1982). A few species

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that lack swimming setae either walk or crawl actively on plants and detritus, likely reflecting recent running-water ancestry (e.g., Sperchon, Torrenticola). Members of Tyrrellia are unusual in that they inhabit wet litter at the edges of permanent bodies of water, including lakes, where they crawl about at the surface film. The number of congeners that co-occur in lakes can be high. For example, more than 25 species in each of Arrenurus and Piona often occur together in a single small lake. The high levels of sympatry are remarkable and enigmatic. Most likely, the clustered distributions of larval mites on hosts results in density dependent mortality, and competition between deutonymphs and adults of the various species is probably not significant in natural communities. Most lentic taxa tolerate a broad range of temperatures, which they encounter in weed beds and silty substrata in the littoral and sublittoral zones. However, a few are coldadapted stenophiles restricted to either arctic-alpine glacial lakes or the profundal zone of oligotrophic lakes. Certain species of some genera (e.g., Limnochares, Hydrodroma, Limnesia, Piona, and Arrenurus) are highly tolerant of extreme thermal and chemical regimes and are able to inhabit desert sloughs, alkaline lakes, or acid bogs.

Temporary Pools Members of ancient genera that may have originated in temporary-pool habitat range from soft-bodied walkers and swimmers (e.g., Eylais) to mites with unique arrangements of integumental plates that either crawl (e.g., Piersigia) or swim awkwardly (e.g., Hydrachna, Hydryphantes). Species of derivative groups that apparently invaded temporary pools secondarily tend to be relatively good swimmers (e.g., Tiphys, Piona, Arrenurus). Water mites inhabiting temporary pools are adapted either to avoid (e.g., species of Hydrachna and Eylais) or to endure the dry phase of the annual cycle (Smith, 1976; Wiggins et al., 1980).

Phytotelmata Phytotelmata are small, often temporary, pools of water held in the leaves and stems of plants. A few species of water mites from the families Anisitsiellidae, Arrenuridae, and Hydryphantidae live in water-filled tree holes and leaf axils (Smith and Harvey, 1989; Walter and Proctor, 2013). The best known of these is Arrenurus kitchingi Smith and Harvey, 1989 (Arrenuridae), which inhabits tree holes in the Australian rainforest.

Ecology The parasitic larvae and predatory deutonymphs and adults of water mites have direct and almost certainly significant effects on the size and structure of insect populations in many

SECTION | VI  Phylum Arthropoda

habitats (Lanciani, 1983; Smith, 1983, 1988; Proctor and Pritchard, 1989). Unfortunately, their impact has rarely been measured accurately because of the routine neglect of mites in ecological studies of freshwater communities (Proctor, 2007). Many freshwater biologists are unfamiliar with mites and tend either to disregard them as too difficult to identify, and either ignore them or lump them together in a meaningless way when conducting community studies. Failure to include water mites must routinely result in flawed analyses of the structure and dynamics of freshwater communities.

Impact as Parasites Effects on Individual Hosts The impact of larval water mites on host insects was reviewed by Lanciani (1983) and Smith (1988). Numerous laboratory studies have demonstrated reduced survival or longevity of various insect hosts parasitized by larval water mites, including mosquitoes, ceratopogonid midges, chironomid midges, juvenile water striders, backswimmers, and damselflies (Lanciani, 1982, 1986a; Smith, 1989; Forbes and Baker, 1990; Leung and Forbes, 1997; Weiberg and Edwards, 1997). Lanciani (1983) concluded that the ratio of larval mite weight to host weight is a good indicator of the probable impact of parasitism on the host. Studies of parasite-induced mortality in field populations have not been conclusive (Smith, 1999). There is evidence of occasional crashes of host populations related to parasitism by mites (Smith, 1988). Water mite parasitism reduces egg production of insect hosts. The most pronounced effects have been reported for Hemiptera infested with larval Hydrachna and Eylais (Davids, 1973), but parasitized nematocerous flies also show lowered fecundity (Smith and McIver, 1984c). Development of juvenile instars is retarded in backswimmers (Notonectidae) parasitized by larvae of Hydrachna and in water striders (Gerridae) attacked by L. aquatica (Smith, 1989). Parasitism by larval water mites has been shown to delay maturation in adult damselflies. Mite parasitism influences frequency of foraging, intensity of territorial behavior, and likelihood of mating in males of the damselfly Enallagma ebrium (Hagen) (Forbes, 1991). In contrast, Rolff et al. (2000) found that parasitism by water mites had no effect on mating success in male damselflies but did result in reduced fat storage by the hosts. Mite parasitism also apparently reduces the capacity of the damselfly Nehalennia speciosa (Charpentier) to fly and disperse (Reinhardt, 1996). Reilly and McCarthy (1991) reported that the corixid Cymatia bonsdorfi (Sahlberg) increases feeding activity when parasitized by larvae of H. conjecta. The presence of engorged water mite larvae on the bodies of adult hosts would seem to have the potential to physically interfere with copulation (Figure 25.36); however, we know of no studies that have tested this.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

FIGURE 25.36  Water mite larvae (Limnocharidae) on adult water striders (Gerridae) in Australia. Photograph by John Prats.

Some hosts exhibit strong avoidance behavior in the presence of larval water mites, involving grooming behavior to dislodge them. Larval Ischnura verticalis (Say) typically respond to fish by reducing movement, but in the presence of both fish and preparasitic larvae of A. pseudosuperior they engage in mite-avoidance behavior despite the increased risk of detection by predators (Baker and Smith, 1997). Variation in Mite Load on Hosts The distribution of parasitic larval water mites typically follows a clustered pattern within a host population, with most carrying few or no mites and a few host individuals heavily infested (Smith and McIver, 1984a; Smith, 1988; but see Rolff, 2001). We have seen examples of over 1000 larvae of Arrenurus parasitizing a dragonfly (Erythemus simplicicolis), over 350 larvae of L. aquatica on a water strider (Gerris comatus), over 50 larvae of Arrenurus on a ceratopogonid midge (Bezzia sp.), and over 40 larvae of A. danbyensis on a mosquito (C. perturbans). Individual chironomid midges emerging from mesotrophic lakes in the Great Lakes Basin often carry 20 or more mite larvae representing as many as seven different genera. Under natural conditions, clustering may reflect differences in the temporal and spatial distributions of host-seeking larval mites and their potential hosts. However, clustering also occurs in laboratory experiments involving synchronous exposure of insects to mites, where duration of exposure and spatial heterogeneity are controlled. This suggests either differential susceptibility to parasitism among host individuals or facilitation of additional loading on hosts that have become parasitized. Leung et al. (2001) found that although unstressed and nutritionally stressed damselfly larvae were just as likely to be colonized by larval mites, the stressed hosts were less successful in removing mites by grooming. This suggests that clustered distributions of mite larvae may in part reflect variation in host condition. Indeed, intensity of infestation by larval Arrenurus appears to be negatively correlated with

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body size and condition (i.e., ratio of mass to body size) of their damselfly hosts in at least some cases (Leung and Forbes, 1997). This may be a result of more vigorous avoidance behavior by larger and more robust potential hosts (Smith and McIver, 1984b; Forbes and Baker, 1990). Larval Arrenurus have greater intensity of parasitism on damselflies with asymmetry in forewing length, but it is not clear whether increased infestation is a response to or a cause of this asymmetry (Walter and Proctor, 2013). Behavioral subtleties can increase vulnerability of hosts of certain sex or age classes to colonizing larval mites, with a resulting differential in parasite loads (Smith, 1988, 1999). Observed differences in parasite loads among sex and age classes in natural populations can prove useful for inferring the behavior of the host and parasite and the age structure of the host population (Smith, 1999). Effect on Other Mites Sharing the Same Host Clustering of parasitic larvae on certain host individuals results in intraspecific competition between mites on those hosts that can affect the mites’ own growth and survival (Lanciani, 1984, 1986b). Clustering may result in densitydependent control of mite populations through intraspecific competition at the infrapopulation level (i.e., on individual hosts), and keeps these populations below carrying capacity, effectively precluding intra- and interspecific competition at the component population level (i.e., on all hosts taken collectively). Parasitism of a host individual by larvae of more than one species of water mite occurs more commonly than would be expected by chance (Stechmann, 1980; Smith and McIver, 1984a). In addition, many species of water mites exploit a variety of related host species. As a result, parasitic associations often involve complex assemblages with considerable overlap in exploitation of host resources, including use of similar attachment sites. Assuming that larval success is the limiting factor determining population size in water mite species, there is probably little competition for food resources among deutonymphs and adults. This may help to explain the remarkably high levels of species richness in many water mite communities.

Impact as Predators The feeding habits of deutonymphal and adult water mites were reviewed by Böttger (1970) and Proctor and Pritchard (1989), and an overview is presented in Table 25.2. Deutonymphs and adults of free-living species are voracious predators on a wide range of small aquatic organisms. In laboratory settings, some species will occasionally scavenge on recently killed organisms. Species of Hydrachna, Todothyas, Hydryphantes, and Hydrodroma are specialized predators of arthropod eggs (Davids, 1973; Mullen, 1975; Meyer, 1985). Many members of Eylais and Piersigia and

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the large genus Arrenurus appear to depend on ostracod crustaceans as prey (although some Eylais and Arrenurus feed on cladocerans); many Neumania ambush copepods; pelagic and planktonic species of Unionicola and Piona are efficient predators of cladoceran zooplankton; and species of many genera of Lebertioidea, Hygrobatoidea, and Arrenuroidea prey upon larval Chironomidae (Böttger, 1970; Proctor and Pritchard, 1989, 1990; Proctor, 1991b). Limnocharid mites have an interesting stalking method of hunting chironomids in which they gradually approach a midge larvae in its tube, extend the gnathosoma, slowly press the tip of the gnathosoma against the back of the head capsule of the larva, and then rapidly drive the chelicerae to pierce the prey’s head capsule (Figure 25.27). After external digestion, the liquefied prey contents are imbibed through the opening in the head capsule. Some species of Limnesia and Piona appear to be more opportunistic in their choice of prey, and frequently even attack other mites in crowded containers (Elton, 1922). Parasitism of freshwater molluscs during the deutonymphal and adult stages has evolved at least twice in the superfamily Hygrobatoidea. For insectfeeding mites, deutonymphs and adults frequently prey upon the immature instars of the same species that they parasitize as larvae (Davids, 1973; Proctor and Pritchard, 1989). Many species of Unionicola inhabit the mantle cavity of mussels and snails (Mitchell, 1955), and the pionid Najadicola ingens has similar habits (Simmons and Smith, 1984). Mollusc-associated Unionicola species are often host specific, with those associated with the gills of the host exhibiting the highest levels of specificity (Edwards and Vidrine, 2006). Some water mites exhibit high feeding rates in experimental studies that, if reflective of natural rates, are high enough to influence the size and structure of prey populations in nature (Mullen, 1975; Paterson, 1970; Lanciani, 1979; Ten Winkel et al., 1989; Matveev et al., 1989). ­Gliwicz and Biesiadka (1975) reported that individuals of Piona limnetica consume 10–20 cladocerans per day in a neotropical lake, and estimated that they could reduce the average standing crop of prey by 50% each week. Members of another species of Piona each killed 28–168 cladocerans per day, representing 53% and 40% of the mortality in Daphnia laevis and Diaphanosoma birgei, respectively (Matveev et al., 1989). Adults of Hygrobates nigromaculatus Lebert, with reported population densities as high as 1000/m2, apparently consumed 14,500 larvae of the chironomid Cladotanytarsus mancus (Walker) per square metre in a European lake, accounting for a 50% mortality rate in the prey population (Ten Winkel et al., 1989). Tactile cues, including contact chemoreception, appear to be of primary importance to predatory water mites (­Böttger, 1970; Davids et al., 1981). Hunting mites often ignore stationary potential prey even at close range, but immediately attack and kill individuals of the same prey

SECTION | VI  Phylum Arthropoda

species after contact is made. Adults of species of Piona typically swim in wide arcs until colliding with potential prey. Upon contact, the mites attack aggressively, and, if initially unsuccessful, swim in tight spirals for several seconds until the prey is encountered again (B.P. Smith, personal observation). Several species of Neumania and Unionicola orient toward vibrations in the water as a cue to locate potential prey (Proctor and Pritchard, 1990). Adults of H. despiciens seem to respond to chemicals in the gelatinous coating of the insect eggs on which they feed. Adults of a number of species show behavioral responses to chemical cues from prey organisms, and Baker (1996) demonstrated how the mites detect these stimuli. Adults of both free living and parasitic species of Unionicola react to host extracts, suggesting that they use chemical cues to initiate regional searching (Roberts et al., 1978; Dimock and Davids, 1985; Proctor and Pritchard, 1989).

Importance as Prey Water mites are typically underrepresented in the stomach contents of predatory freshwater vertebrates (Pieczyński, 1976; Eriksson et al., 1980). Nevertheless, deutonymphs and adults at least occasionally form a significant part of the diet of fish and turtles (Marshall, 1940), and we have seen examples of stomach contents from individual brook trout and coregonids that consisted exclusively of several hundred adults of one species of Piona and Hygrobates, respectively. It appears that fish can develop narrowly focused search images for water mites in exceptional cases. Adults of many species of water mites eject viscous, sticky fluid from the glandularia when handled roughly, apparently as a deterrent to predation. Experiments have shown that red-colored water mites are distasteful to fish, which quickly learn to reject them as potential prey (Elton, 1922; Kerfoot, 1982; Proctor and Garga, 2004). Kerfoot (1982) concluded that red pigmentation in water mites is aposematic coloration, but the situation in nature is more complex (Proctor and Garga, 2004). Most red water mites live in temporary pools and seepage areas where fish are absent. Terrestrial anystoid and parasitengone mites, and members of many early basal water mite clades, are predominantly red or orange, suggesting that suffusion of the body with red carotenoid pigments is plesiotypical for the entire lineage. This type of pigmentation may have initially evolved in these mites as protection from ultraviolet wavelengths in sunlight (Proctor and Garga, 2004). It is noteworthy that, at least in temperate latitudes, most species of the derivative water mite superfamilies Lebertioidea, Hygrobatoidea and Arrenuroidea, which predominate in permanent water habitats, are highly colored but not bright red. Many of the relatively small number of species in these groups that are bright red inhabit vernal temporary pools or springs. The preponderance of red species in these habitats, and their

Chapter | 25  Subphylum Chelicerata, Class Arachnida

comparative scarcity in larger bodies of water, has yet to be adequately explained. Few studies have examined invertebrate predation on water mites. Elton (1922) reported an instance of predation by an adult dytiscid beetle on a species of Hydrachna. Two author groups have reported positive associations between fish and the mite Piona carnea, and have suggested both that fish prefer insects to mites as prey and that predaceous insects, including hemipterans, larval odonates, and larval chaoborid midges, feed on the mites (Eriksson et al., 1980; Punčochář and Hrbáček, 1991). Higher activity by male N. papillator appears to make them more susceptible than females to predation by damselfly larvae (Proctor, 1992c).

Potential as Indicators of Environmental Quality Many species of water mites are are limited to narrow ranges of physical and chemical regimes, as well as to particular biotic attributes of particular water bodies. This is clearly demonstrated by studies involving multivariate analysis of environmental conditions and occurrence of water mites (Smit and Van der Hammen, 1992; Van der Hammen and Smit, 1996; Goldschmidt, 2004). Consequently, water mites should be exceptionally sensitive indicators of habitat conditions and the impact of environmental changes on freshwater communities. Preliminary studies of physicochemical and pollution ecology of the relatively well-known fauna of Europe have demonstrated that water mites are excellent indicators of habitat quality (Pieczyński, 1976; ­Kowalik and Biesiadka, 1981; Cicolani and di Sabatino, 1991). Gerecke and Schwoerbel (1991) analyzed changes in water mite assemblages in the Danube River between 1959 and 1984, and concluded that they were highly correlated with changes in levels of organic pollution. One of the species that they studied, Hygrobates fluviatilis (Ström), appears relatively resistant to pollution and increased in dominance during the period of declining water quality. Growns (2001) found that abundance and diversity of water mites were diminished by pollution in streams in southeastern Australia, and was able to use species assemblages to discriminate between polluted and unpolluted sites. The results of these studies, along with our own observations in sampling a wide variety of habitats in North America and elsewhere, lead us to conclude that water mite diversity is dramatically reduced in habitats that have been degraded by pollution or physical disturbance. There have been few laboratory studies of the tolerances of water mites to environmental variables. Deutonymphs of A. manubriator are more susceptible than other life history stages to iron in solution, possibly because of greater physiological activity of this instar (Rousch et al., 1997). Edwards (2004) showed that survival of U. foili decreased when the mites were exposed to pH values of less than 5.3.

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Progress toward the establishment of essential baseline documentation on water mite assemblages in North America, however, continues to be hampered by lack of sufficient ecological information at the species level, by inadequate sampling and extraction procedures, and by a tendency by freshwater ecologists to ignore or lump water mites into uninformative categories (Proctor, 2007). Application of potentially useful information on water mites in assessing and monitoring the state of freshwater habitats would benefit from a clearer understanding of the individual biotic and abiotic requirements of species. This can be achieved by improving reporting of the group by ecologists, and by developing more appropriate collecting, observing, and rearing techniques, including the use of mesh sizes fine enough to capture mites.

COLLECTING, REARING, AND PREPARATION FOR STUDY These subjects were treated comprehensively by Barr (1973), and we discuss here only those aspects needing further emphasis or refinement. Our emphasis is on Hydrachnidiae, as very little effort has been put into collecting other groups of aquatic mites. However, most general methods of freshwater sampling can also be applied to aquatic mite fauna, provided that the mesh size is appropriate for the body size of the mites (∼250 μm). For some habitats, such as springs, specific collecting methods are required.

Collecting and Extracting Techniques Most field work on water mites has focused on qualitative sampling to obtain specimens for systematic or life history studies. Modifications of these procedures for rigorous and cost-effective quantitative sampling are needed, including those exploiting behavioral traits to extract mites from detritus and silt. Although water mites are frequently found in samples collected for quantitative analysis in ecological studies (Proctor, 2007), they are often underrepresented in number and diversity. Collecting devices such as dredges and grabs are designed to capture organisms that are larger or more sedentary than mites. Due to their small size and ability to cling tenaciously to substrata, mites in lotic habitats can often resist casual efforts to dislodge them or can simply pass through nets with coarse mesh. By running or swimming, mites in hyporheic or lentic habitats can avoid capture by traps that are designed to instantaneously sample small areas of superficial substrata. Mite specimens are frequently overlooked when preserved samples are sorted, because they match the size and shape of substrate particles or pass through coarse-meshed sieves during sample preparation. Some of the methods below could be used or modified for use in quantitative sampling, but it is important to consider whether your sample is likely to accurately represent

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true abundance. In many cases, it is better to view such samples as appropriate only for comparison with others collected using the same method.

SECTION | VI  Phylum Arthropoda

simply digging down through the gravel and sand beneath the riffle zone and removing all large stones to allow the current to flush out the pockets of interstitial silty detritus and the organisms that live there. By digging out an area of substrate approximately 1 m2 to a depth of 0.5 m, we Deutonymphs and Adults are often able to recover good numbers of many hyporheic It is easiest to discuss sampling methods by breaking the species. This technique can often be used to sample the process into two phases: collecting and concentrating a hyporheic zone in habitats where the standard Karamansample, then extracting the mites from that sample. Methods Chappuis method is not practical. for collecting a sample are dependent on the structure of the In lentic habitats or depositional zones of lotic waters, habitat: water flow, depth, substrate composition, and rela- the net should be actively and repeatedly swept back and tive density and resistance of vegetation, etc. Once a sample forth, bouncing the net on the surface of the substrate and is taken, the size and behavior of the organisms will best sweeping through any subaquatic vegetation. In habitats determine the optimal methods for separating them from where there is a thick, soft flocculant layer on the substrate, nontarget materials in a sample, and similar methods may consider how deeply to dig: if deeper layers are anoxic, then be used with samples collected from different habitat types. most organisms will be restricted to the top few millimeThe most common combination is to use some type of ters of substrate. In such conditions, going deeper increases net for concentrating mites into a sample, and a large white your effort and the volume of debris in the sample without tray filled with water for sorting mites out of that sample. It significantly increasing your catch. This is a common misis often also useful to add an additional step before extract- take made by people new to collecting water mites. ing mites: pass the sample through a series of sieves to In very shallow habitats, such as seepage areas, small exclude organisms and particles larger and smaller than the springs, fens, and the edges of pools and ponds, the subtarget mites. We find that a pair of upper and lower sieves strate should be gathered and stirred by hand. The net can with mesh sizes of 1.4 mm and 250 μm, respectively, yields be worked beneath the edge of the wet moss, leaf litter, or good results for Hydrachnidiae; however, some large-­ detritus that is to be sampled so that mats of wet substrate bodied mites in lentic waters may be stopped by even 2-mm can be separated and rinsed over the opening. Where there mesh, whereas some halacarids and hyporheic species of is sufficient surface water, successive handfuls of loose water mites may pass through 200-μm sieves. material can be picked or scooped up with the aid of a small trowel, thoroughly picked apart, and washed in the Collecting a Concentrated Sample net. In seepage areas, spring edges, fens, and bog margin In most habitats and for virtually all types of substrata, con- habitats where there is an extensive carpet of wet moss and centrating mites in a sample involves using a strong net with associated plants, mites can also be collected by methodia wide opening (25–40 cm diameter) and fine mesh size cally treading down the plants while dragging the net along (200–250 μm). In habitats with dense submerged vegeta- to scoop up the detritus, silt, and organisms that become tion, leaves, or woody debris, coarse-meshed metal screen- temporarily dislodged as water is forced up through the ing (e.g., hardware cloth, with ∼6 × 6-mm mesh) can be bent disturbed layer of vegetation. This should not be done in over the front of the net to exclude larger debris while the any ecologically sensitive areas, however. In either case, sample is being taken. Some models of net have a flat lower when a mixture of fine silt, plant fragments, and organisms edge (D-frame net), which makes it easier to contact the begins to fill the bottom of the net, it should be transferred substrate without leaving gaps. Once the desired habitat or to a container along with a small volume of water. Repeat microhabitat has been selected, the substrate must be dis- this process until the container is approximately half full of material. The container is then nearly filled with clean turbed and the debris passed through the net. In lotic erosional zones, the net can be held against the water and the contents stirred gently but thoroughly to allow substrate while the collector actively stirs the substrate heavy inorganic material such as gravel and sand to settle to upstream with their foot, or with a spade when necessary. the bottom. The contents of the container are then poured It is best to dig deeply (15 cm or more when possible) and carefully through a set of sieves so that gravel and sand to thoroughly stir up the substrate. In stream riffles with a remain in the container. This operation should be repeated heterogeneous substrate of rocks, gravel, sand, and detri- until all loose organic material is in the sieves. The coarse tus, vigorous and persistent agitation is usually necessary material in the upper sieve can then be thoroughly stirred to dislodge the mites that cling tenaciously to rocks or that and rinsed with clean water. The fine silt and small organburrow in the silt under larger stones. Where well developed isms accumulating in the lower sieve should be regularly interstitial habitats occur in a stream bed, this approach can transferred to a large (250–1000 ml) container with enough be extended to yield rich collections of mites including both water to just cover the surface of the silt to be transported surface and hyporheic species. This is accomplished by back to the laboratory or sorting area.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

Quantitative samples of hyporheic arthropods including large and diverse collections of water mites have been obtained in desert streams in Arizona by driving polyvinyl chloride (PVC) tubing into the substrate to various depths and using a mechanical pump to evacuate the resulting wells periodically. This method was also used in a New Zealand study (Scarsbrook and Halliday, 2002), as well as freezecoring and colonization pots. Water mites were the most common taxonomic group among the samples, although relative proportions differed significantly depending on sampling method, with mites being relatively more common in freeze-core samples. In a study on meiobenthos from various streams, the classical Karaman-Chappuis method (digging and sieving in dry stream banks) turned out to be more effective in terms of number of specimens compared to pumping animals directly from the hyporheos. Although animals from Karaman-Chappuis diggings were generalists adapted to a diversity of flowing-water conditions, mites from pumping samples were typically hyporheic specialists. Drift nets, as those used for stream-dwelling insects, also capture lotic water mites if the mesh is fine enough (200–250 μm) (Martin, 1999). At least in temperate zones, these mites apparently drift primarily in daylight irrespective of the presence of fish (Martin, 1999). Most sampling for water mites has taken place in shallow water conditions, and sampling in deep water requires special methods. Planktonic water mites can be collected using plankton tow nets, Schindler traps, and other devices commonly used for zooplankton collection. Unless the mites are in relatively high densities, the most successful method is to use a large (e.g., 0.5-m diameter) plankton net, hauled through a reasonably large volume of water (e.g., 15-m vertical haul). Sampling substrates at depth is difficult: dredges and grabs capture only a small volume of material and dig deeply into the substrate, whereas most mites will be found on the top of the flocculent layer of the substrate. A thorough but more expensive way to sample mites on the substrate at depth would be to employ scubaequipped divers wielding nets. Under certain circumstances, underwater traps are an effective method for collecting water mites in lentic habitats. Although they are useful in the littoral zone, they can also be deployed in the benthic sublittoral and profundal zones, or suspended at various depths in the planktonic zone. Pure activity traps consist of funnel pointing into a closed container, and simply depend on the mites entering the trap during normal locomotion. The effectiveness of these traps can be increased by orienting the trap downward when mites are expected to be moving upward, etc. The same trap design baited with light (e.g., Barr, 1979) can collect several orders of magnitude more specimens than unbaited traps. Subaquatic light traps are not new, but earlier models were heavy and cumbersome, involving lead-acid batteries and waterproofed light bulbs. Chemo-luminescent light

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sticks and inexpensive compact submersible flashlights make these traps much more practical (e.g., Barr, 1979). Light traps are highly biased to certain taxa. For example, Oxidae are typically underrepresented in light trap samples when compared to numbers collected in D-frame net samples, even though these taxa are highly mobile (B.P. Smith, personal observation). In contrast, Hydrodroma and freeliving Unionicola species are typically overrepresented (Barr, 1979; B.P. Smith, personal observation). Also, water clarity, obstructions in the water, wave action, and water temperature can influence the effectiveness of the traps. On the positive side, light traps can make it easier to sample in difficult situations, e.g., at depths greater than 1.5 m, where substrate has a thick soft layer, among obstructions such as submerged branches, or where there is risk to the collector because of snakes or crocodilians. Although the efficacy of net-sampling can be highly biased by the collector’s experience, light traps are less affected. A very simple light trap can be constructed from a 2-L transparent plastic bottle of the sort in which carbonated beverages are sold. If the top is cut off just where it reaches the widest point, it can be inverted and used as a funnel. This end can be held in place with small pieces of waterproof tape. Covering the outside of the trap (except for the funnel) with aluminum foil will block stray light and greatly increase the numbers of organisms caught. Light can be provided by a chemo-luminescent glowstick, which provides a reasonable intensity of light for about 3 h (e.g., Barr, 1979). As an indication of its efficiency under ideal conditions, a single glowstick trap caught ca. 8000 Hydrodroma in 3 h in a highly productive Canadian lake (B.P. Smith, personal observation). Such traps can be constructed and operated very inexpensively, so the risk of trap loss is trivial, but a significant amount of waste is generated with singleuse glowsticks. Waterproof flashlights or battery-operated glowsticks are also potential light sources. For repeated use, an economical trap can be constructed from a white plastic tube, e.g., 10-cm diameter PVC tubing used for drains and sewage, or a white polyethylene 3- to 4-L bucket. The external surface can be painted black to minimize stray light. A transparent funnel is attached at one end, and, if using tubing, an end cap is used to seal the other end. A small waterproof flashlight can be attached to a hole drilled through the end cap (or bottom of the bucket). When combined with rechargeable NiMH batteries, a reuseable PVC tubing flashlight trap can currently be made in North America for less than US $50 in materials (B.P. Smith, personal experience). A very simple kind of trapping by light has been used in spring areas in the European waterproof flashlights Alps (P. Martin, personal observation). A light source, powered by a car battery, was suspended from a tripod at night over a helocrene spring. Mites are attracted by the light, and Hydryphantidae in particular are strongly attracted. This

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method may be especially useful where sites are sensitive to mechanical damage. Extraction The most common method for extracting mites from a concentrated sample involves using a white tray partially filled with clean water. A large tray (e.g., 30 × 40 cm) is most convenient for sorting mites from net samples. A bolus of collected materials is gently placed in the center of the tray, minimizing disturbance of the water. As mites crawl or swim free of the debris, they can be picked up with a pipette or captured with forceps. Separating the mites from the sample at the collecting site may be practical when collecting larger species in good weather and bright sunlight. However, it is often more effective and efficient to separate mites from substrate under controlled conditions in the laboratory. If specimens are to be transported alive, then a portable insulated carrier (provided with ice, chiller packs, or an electronic cooling system) is very useful. Although it is more laborious, sorting samples under a stereo-microscope is much more thorough, and may be advantageous when collecting small and less conspicuous species. Many swimming species are strongly positively phototropic and can be recovered efficiently from trays kept in bright light. However, many crawling mites and a considerable number of swimming species are less attracted, or even repelled, by light and become active only at low levels of illumination. The effectiveness and rate of recovering mites from all types of habitats can be substantially increased by reducing ambient lighting to low levels indoors or waiting until after sunset outdoors, and then periodically scanning the trays with a flashlight beam. An experienced collector can then spot the mites as they move across a contrasting background provided by either the darkly colored silt or the white bottom of the tray. Mites remain active and continue to emerge from silt in the trays for up to 72 h. For this reason we suggest that, whenever possible, mites should be extracted from samples in situations that permit periodic observation of undisturbed trays over a period of at least 4–6 h with controlled lighting. Under these conditions, mites tend to congregate around the edges of the trays, especially in the corners, where they can be easily spotted and picked out. Although attraction to or repulsion from light can be used to separate water mites from debris, mites and other aquatic arthropods can also be extracted from samples of substrate by a temperature gradient. This method has yielded very good results for sphagnum samples, and could probably be adapted for use with any type of substrate inhabited by mites. Salt solutions can also be used to separate mites from debris. For example, Radwell and Brown (2008) separated meiofauna from core samples, first by swirling and decanting the sample in stream water and then in a saturated solution of calcium chloride, inducing the mites to detach from the substrate and

float on the surface. Differences in specific gravity can also be used to separate mites from other organisms in fresh water, e.g., from light-trap samples. Water mites have a specific gravity around 1.25, whereas zooplankton are much closer to neutral buoyancy. If a sample is swirled in a bucket of water, as the water slows, the mites will settle near the center while zooplankton continue swirling. At this point, a pipette can be used to remove the mites from the center. We have also had some success extracting crawling water mites from damp macrophyte samples using a Tullgren funnel (light bulb suspended over a funnel) (H.C. Proctor, personal observation), a method more commonly used for soil and litter extraction. It is not always possible to live-sort samples, for example, if many samples need to be collected simultaneously, or if the researchers doing the collecting are not the same as those doing the sorting (which is typical in many biomonitoring programs). In these circumstances, samples are usually preserved in the field and sorted later in the laboratory. Proctor (2001) compared the thoroughness and efficiency of three methods of mite extraction from substrates: livepicking, exhaustive sorting of preserved samples using a dissecting microscope, and kerosene-flotation of preserved samples followed by sorting with a dissecting microscope. This flotation method relies upon the affinity between arthropod cuticle and kerosene. Kerosene is added to an ethanolpreserved sample, which is gently shaken and then allowed to rest. Mites and other arthropods adhere to the kerosene, which is less dense than the water–alcohol mixture and rises to the top, where it and the accumulated arthropods can be removed using a large-bore pipette. Proctor (2001) found that live-picked samples yielded significantly fewer individuals and species than other methods, and were biased toward large species of water mites. Oribatid and halacarid mites were not recovered using live-picking. Live-picking and kerosene-float methods provided similar numbers of mites per minute of sampling effort, whereas microscopepicking had a lower efficiency than kerosene-floatation. A combination of live-picking and kerosene-flotation can be used for thorough surveys of stream acarofauna.

Larvae There is no method developed specifically for collecting free-living (not yet parasitic) water mite larvae, reflecting the lack of effort to do so in past studies. Free-living larvae as well as quiescent protonymphs and tritonymphs are often incidentally collected from aquatic habitats along with deutonymphs and adults. This is especially common in interstitial habitats (P. Martin, personal observation) and when sorting through mossy substrates for crawling mites (B.P. Smith, personal observation). Parasitic larvae can be recovered from insect hosts. Aquatic hosts are usually captured using a dip net, whereas aerial adults can be collected in emergence traps, netted while swarming (especially over

Chapter | 25  Subphylum Chelicerata, Class Arachnida

water at dusk), swept from vegetation, or attracted to lights at night. The “aerial” larvae of most Eylaioidea, Hydryphantoidea, and Hydrovolzoidea are so called because they move to the water’s surface and associated vegetation shortly after having hatched. Sometimes these can occur in exceptionally large numbers, as in Eylais spp. in western North America (Figure 25.34 from Alaska; B.P. Smith, personal observation). In such cases, larvae can easily be collected directly, e.g., by using a fine paint brush dipped in alcohol. Terrestrial larvae of Limnochares can be by placing thin dowels vertically in the soft substrate of a pond, simulating emergent vegetation, with the top of each dowel covered with a piece of masking tape, sticky surface out. Aquatic larvae (Hydrachnoidea, Lebertioidea, Hygrobatoidea, Arrenuroidea) are frequently observed in underwater light traps. Although the main use of such traps is for collecting adult and deutonymphal mites, these traps may be reasonably effective for sampling larvae as well. A promising approach to sampling larval mites in nature is to use sentinel hosts, set out in cages. This has proved useful for collecting larval Arrenurus and Limnochares on caged damselfly larvae and water striders, respectively (B.P. Smith and R. Baker, and B.P. Smith, unpublished).

Rearing The heteromorphic life cycle of water mites, which typically includes a parasitic larval stage, makes it challenging to recognize conspecifics among instars, and to culture mites in the laboratory. For most taxa, rearing through a full generation requires co-maintenance of live hosts for the larvae and prey for the predatory deutonymphs and adults. Numerous species have been kept in the laboratory over one or more of their life-history transformations, but the full transformation series of species with parasitic larvae has been completed in captivity for only a handful of species (e.g., Ullrich, 1978; Hevers, 1980). For the minority of water mite species that lack parasitic larvae (Smith, 1998), culturing is somewhat easier, as one needs only appropriate prey for the predatory stages. Raising unengorged larvae from field-collected females is comparatively easy and is widely used for making taxonomic associations between larval and adult stages. Techniques for encouraging oviposition have previously been described (e.g., Prasad and Cook, 1972; Smith, 1976; Gerecke and Di Sabatino, 2007). In temperate regions of the northern hemisphere, gravid females are usually most abundant in the field in spring and early summer; however, some species lay eggs in late summer or fall and may overwinter as eggs (e.g., Rhyncholimnochares kittatinianna, Smith). The probability of oviposition generally decreases with the time since collection, so it is best to collect females around the expected time of egg laying in the field and to

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immediately isolate individual females in containers so as to be able to associate egg masses with the correct female. In some cases, females kept for weeks to months will eventually lay eggs. With some taxa (e.g., Arrenurus spp.), providing food for the adults increases the chances of eggs being laid. For taxonomic purposes, it is essential to hold individual females separate; however, some species lay eggs in communal masses, and both the probability of oviposition and the number of eggs are often greater when females are kept together. Thus, if one is certain that females are conspecific, holding them in the same container may improve the likelihood of getting larvae. If females fail to oviposit, it may be that they are still virginal, in which case success may be achieved by maintaining them with conspecific males; however, at least in temperate areas, fertilized females of many taxa must overwinter prior to oviposition. Generally females prefer to oviposit on substrates other than smooth glass or plastic, and small pebbles, fragments of moss, wood, or slips of paper should be provided. The addition of organic materials can be problematic in that decomposition can result in low oxygen levels and high egg mortality. As an alternative, the mites can be housed in tissue-culture well-plates in which the floors of the wells are roughened to provide structure. Almost all Hydrachna spp. lay eggs in emergent vegetation; however, most will accept polystyrene foam (e.g., cups sold for hot beverages) as a proxy, avoiding the need to maintain plants. Some species seem to prefer to oviposit below the surface of a substrate, probably indicating photophobic behavior, and may not oviposit unless a relatively deep and penetrable substrate is provided. Containers for rearing larvae should be covered to limit evaporation and, for taxa with “aerial” larvae, to keep the larvae from escaping. Parafilm with a few small perforations is a good option for covering containers. An air space should be left between the cover and the surface of the water to allow for gas exchange. Maintain containers at temperatures and light conditions near levels experienced in the natural habitat of the mites. Most mites from cold habitats must be refrigerated, although some stream-living species may survive several days at room temperature during the summer (P. Martin, personal observation). The rearing containers should be checked periodically for the presence of egg masses. For taxonomic purposes, females should be preserved immediately after they have laid their eggs to ensure that good-quality specimens are available for confirming species identifications. Larvae should be either preserved in ethanol or slide-mounted very soon after hatching; otherwise opaque waste products (guanine crystals) may build up in their bodies and interfere with observation of morphological features when mounted. In some groups, it may be advantageous to expand larvae so that sclerites and setae on the body are well separated. To do so, first kill the larvae with hot water, then let them sit for 1 to 2 days in water prior to clearing and slide mounting.

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Providing a suitable host for unengorged larval mites can be difficult, depending upon the stage of the host that is parasitized, duration of the larval association, and degree of host specificity. Larvae that will attach to both juvenile and adult stages of the host are relatively amenable to engorging in a laboratory situation. For example, Smith (1989) exposed various field-collected water strider species (Heteroptera: Gerridae) to newly hatched larval L. aquatica, which attached, engorged, and detached successfully under laboratory conditions. In contrast, species of Hygrobatoidea, Lebertioidea, and Arrenuroidea are difficult to rear through the larval stage because the larvae form a preparasitic association with the pre-imaginal stage of the host and attach to the adult host at eclosion, resulting in a narrow window of time in which hosts can be colonized. Edwards and his collaborators (Edwards and Dimock, 1995; Edwards, 2004) collected freshwater mussels (hosts for many species of Unionicola) from the field and either maintained them for several weeks in the laboratory or dissected the developing mite eggs from the mussel tissue and kept them in artificial lake water until hatching. The newly hatched larvae then successfully attended pupae of laboratory-reared Chironomus and completed engorgement on the adult midges. Although rearing of larvae through the parasitic stage is more feasible for lentic species, Ullrich (1978) investigated complete life cycles of several streamdwelling species of water mites and succeeded in inducing parasitism on various insects. For his experiments he used vials with standing water, and succeeded in infesting juvenile Chironomidae, Simuliidae, and Trichoptera collected from the field. Parasitized adult hosts were maintained alive in the laboratory until larval mites finished engorgement, with simuliids fed on glucose and human blood. Engorged larvae can be collected from field-caught parasitized hosts and raised to their later active stages. Larval exuviae can be slide-mounted for taxonomic use. Host insects may be maintained over water until the larval mites naturally detach. Alternatively, killing the host and letting it sit in water may trigger detachment (Mullen, 1974). As a last resort, engorged larvae can be gently dislodged from their hosts and, if uninjured and sufficiently engorged, may survive to transform into deutonymphs and, with luck, into adults, which can then be identified to species. The ease of maintaining the post-larval stage of water mites depends on the taxon. Limnocharidae and various Hydryphantidae can often live without food for 1 year or more, and although best kept under refrigeration, will even survive long periods at room temperature (B.P. Smith, personal observation; Martin, 2004). Most taxa, however, need some food if kept for more than a week, especially if they are to be used for ecological or behavioral studies. Inducing transformation of deutonymphs to adults typically also requires feeding. Benthic species from cool habitats (springs, headwater streams) typically need less food than

SECTION | VI  Phylum Arthropoda

actively swimming mites from warmer lentic habitats, provided that they are maintained at their normal temperatures. Although it is often possible to find acceptable prey, it is often not sufficient for growth and successful transformation from deutonymph to adult. This has been accomplished for relatively few species, but is generally easiest with mites that are specialist predators of culturable prey, as one or a few species of prey can be provided from laboratory colonies. One of us (B.P. Smith) has had great success using the ostracod Cypridopsis vidua (Müller) as an easily cultured food source for raising at least a dozen North American Arrenurus spp. Small individuals of Daphnia species are acceptable prey for other Arrenurus species, Piona spp., and for non-mussel-associated Unionicola; however, most Daphnia are too large, and smaller cladocerans such as Ceriodaphnia are a better choice. Some specialist egg-predators are also candidates for raising to adult. At least one species of Hydryphantes accepted dragonfly eggs and matured to adults in the laboratory, and several species of Hydrachna were adequately fed on corixid eggs. In both cases, eggs serving as prey were obtained from field-caught gravid insects. Hydrodroma species consume chironomid eggs and could potentially be co-cultured along with Chironomus; however, the thickness of the egg jelly-coat can interfere with the mites’ ability to reach the eggs with their palps. Many water mites will feed on chironomid larvae, and hatchling Chironomus would likely be suitable prey. Limnochares can consume quite large tube-building chironomids. The maintenance of continuous laboratory cultures for water mites with parasitic larvae has been accomplished for very few species. The best success has been with Arrenurus spp. that parasitize mosquitoes. One of us (B.P. Smith) has raised species of two subgenera of Arrenurus (Megaluracarus and Truncaturus) through a number of generations, using Anopheles spp. as hosts for members of the first subgenus and Aedes or Culex for the second, and C. vidua as prey for postlarval stages. Populations or species that forego larval parasitic associations are ideally suited for culturing as the need for a host is eliminated. Most mite species from the temperate zone exhibit a reproductive diapause in which adults will remain active, both mating and feeding, but females will not oviposit until “overwintered” at around 5 °C for several months. As a final note, laboratory cultures of water mites should be periodically cleaned of remnants of prey and dead mites, as this organic debris can result in buildup of colonies of bacteria and other microorganisms that can entangle mites or produce anoxic conditions.

Preservation and Preparation for Study For morphological study, deutonymphal and adult mites are typically preserved in modified Koenike’s solution (or GAW), consisting of 5 parts glycerin, 4 parts water, and 1 part glacial acetic acid, by volume (Barr, 1973) so that

Chapter | 25  Subphylum Chelicerata, Class Arachnida

they can be easily cleared, dissected, and slide-mounted for identification and study. Specimens preserved in alcohol or other hygroscopic agents often become so distorted and brittle that subsequent preparation is difficult or impossible. Deutonymphs and adults must be cleared in either acetic corrosive or 10% KOH, dissected, and mounted in glycerin jelly, following the techniques described by Barr (1973) and Cook (1974). One of us (H.C. Proctor) has had success in slide-mounting post-larval water mites preserved in ethanol by first clearing them overnight in 85% lactic acid after piercing the soft integument in a few places, massaging out the body contents, and then mounting the exoskeleton in PVA (available commercially from BioQuip, USA). Reared larvae are often slide-mounted directly, but can be preserved in 70% alcohol for subsequent mounting. Parasitic larvae should be preserved with the host in 70% alcohol, although larvae removed from hosts that have been pinned and dried often can be slide-mounted successfully. Before larvae are removed from hosts, the attachment sites should be noted and recorded. Larvae should be mounted whole in Hoyer’s medium or PVA, which also acts as an efficient clearing agent for these small specimens (Barr, 1973).

DEDICATION The authors dedicate this collaborative effort to the memory of our mentor and friend Rodger Mitchell, who passed away in June 2013. Rodger’s curiosity and originality, intellectual rigor, and always insightful interpretations of complex problems helped set the stage for modern research on mite structure, function, and ecology. His enthusiasm for sharing his passionate interest in mites with colleagues and students ensured that we all benefited from his unique perspectives. We will miss him.

ACKNOWLEDGMENTS We thank Michelle MacKenzie at Agriculture and Agri-food Canada for her invaluable assistance in preparing the illustrations and scanning electron micrographs of water mites. Peter Martin from the Christian-Albrechts-Universität zu Kiel provided helpful comments on some early draft sections.

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Baker, G.T., 1996. Chemoreception in four species of water mites (Acari: Hydrachnida): behavioural and morphological evidence. Exp. Appl. Acarol. 20, 445–455. Baker, R.L., Smith, B.P., 1997. Conflict between antipredator and antiparasite behaviour in larval damselflies. Oecologia 109, 622–628. Barr, D.W., 1972. Life Sciences Contributions. The Ejaculatory Complex in Water Mites (Acari: Parasitengona): Morphology and Potential Value for Systematics, vol. 81. Royal Ontario Museum. 87 pp. Barr, D.W., 1973. Methods for the Collection, Preservation, and Study of Water Mites (Acari: Parasitengona). Life Sciences Miscellaneous Publication. Royal Ontario Museum. 28 pp. Barr, D.W., 1979. Water mites (Acari, Parasitengona) sampled with chemoluminescent bait in underwater traps. Int. J. Acarol. 5, 187–194. Barr, D.W., 1982. Comparative morphology of the genital acetabula of aquatic mites (Acari, Prostigmata): Hydrachnoidea, Eylaoidea, Hydryphantoidea, Lebertioidea. J. Nat. Hist. 16, 147–160. Bartsch, I., 1996. Halacarids (Halacaroidea, Acari) in freshwater. Multiple invasions from the Paleozoic onwards? J. Nat. Hist. 30, 67–99. Bartsch, I., 2007. The freshwater mite Porolohmannella violacea (Kramer, 1879) (Acari: Halacaridae), description of juveniles and females and notes on development and distribution. Bonn. Zool. Beiträge 55, 47–59. Bartsch, I., 2008. Global diversity of halacarid mites (Halacaridae: Acari: Arachnida) in freshwater. Hydrobiologia 595, 317–322. Behan-Pelletier, M., Eamer, B., 2007. Aquatic Oribatida: adaptations, constraints, distribution and ecology. In: Morales-Malacara, J.B., ­Behan-Pelletier, V., Ueckermann, E., Pérez, T.M., Estrada-Venegas, E.G., Badii, M. (Eds.), Acarology XI: Proceedings of the International Congress. Instituto de Biololgía; Facultad de Ciencias; ­Universidad Nacional Autónoma de México; Sociedad Latinoamericana de ­Acarología, México, pp. 71–82. 2002. Bohonak, A.J., 1999. Effect of insect-mediated dispersal on the genetic structure of postglacial water mite populations. Heredity 82, 451–461. Bohonak, A.J., Smith, B.P., Thornton, M., 2004. Distributional, morphological and genetic consequences of dispersal for temporary pool water mites (Acari: Arrenuridae: Arrenurus). Freshw. Biol. 49, 170–180. Böttger, K., 1970. Die Ernährungsweise der Wassermilben (Hydrachnellae, Acari). Int. Rev. Gesamten Hydrobiol. 55, 895–912. Böttger, K., 1972a. Vergleichend biologisch-ökologische Studien zum Entwicklungszyklus der Süßwassermilben (Hydrachnellae, Acari). I. Der Entwicklungszyklus von Hydrachna globosa und Limnochares aquatica. Int. Rev. Gesamten Hydrobiol. 57, 109–152. Böttger, K., 1972b. Vergleichend biologisch-ökologische Studien zum Entwicklungszyklus der Süßwassermilben (Hydrachnellae, Acari). II. Der Entwicklungszyklus von Limnesia maculata und Unionicola crassipes. Int. Rev. Gesamten Hydrobiol. 57, 263–319. Brum, P.E.D., Costa-Schmidt, L.E., de Araújo, A.M., 2011. It is a matter of taste: chemical signals mediate nuptial gift acceptance in a neotropical spider. Behav. Ecol. 23, 442–447. Carico, J.E., 1973. The nearctic species of the genus Dolomedes (Araneae, Pisauridae). Bull. Mus. Comp. Zool. 7, 435–488. Chua, T.J.L., Lim, M.L.M., 2012. Cross-habitat predation in Nepenthes gracilis: the red crab spider Misumenops nepenthicola influences abundance of pitcher dipteran larvae. J. Tropical Ecol. 28, 97–104. Cicolani, B., di Sabatino, A., 1991. Sensitivity of water mites to water pollution. Mod. Acarol. 1, 465–474. Collins, N.C., 1975. Tactics of host exploitation by a thermophilic water mite. Misc. Publ. Entomol. Soc. Am. 9, 359–370.

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Fashing, N.J., 1998. Functional morphology as an aid in determining trophic behaviour: the placement of astigmatic mites in food webs of water-filled tree-hole communities. Exp. Appl. Acarol. 22, 435–453. Fashing, N.J., 2004. Mate-guarding in the genus Zwickia (Astigmata: Histiostomatidae), a natatorial inhabitant of the fluid-filled pitchers of Nepenthes (Nepentheaceae). Phytophaga 14, 149–154. Fashing, N.J., Marcuson, K.S., 1996. Fine structure of the axillary organs of Fusohericia lawrencei Baker and Crossley (Astigmata: Algophagidae). In: Mitchell, R., Horn, D.J., Needham, G.R., Welbourn, W.C. (Eds.), Acarology IX. Proceedings, vol. 1. Ohio Biological Survey, Columbus, OH, pp. 381–384. Flechtmann, C.H.W., 1985. Eustigmaeus bryonemus sp. n., a moss-feeding mite from Brasil (Acari, Prostigmata: Stigmaeidae). Rev. Bras. Zool. 2, 387–391. Forbes, M.R., 1991. Ectoparasites and mating success in male Enallagma ebrium damselflies (Odonata: Coenagrionidae). Oikos 60, 336–342. Forbes, M.R., Baker, R.L., 1990. Susceptibility to parasitism: experiments with the damselfly Enallagma ebrium (Odonata: Coenagrionidae) and larval water mites, Arrenurus spp. (Acari: Arrenuridae). Oikos 58, 61–66. Forbes, M.R., Muma, K.E., Smith, B.P., 2002. Diffuse coevolution: constraints on a generalist parasite favor use of a dead-end host. Ecography 25, 345–351. Futami, K., Sonye, G., Akweywa, P., Kaneko, S., Minakawa, N., 2008. Diving behavior in Anopheles gambiae (Diptera: Culicidae): avoidance of a predacious wolf spider (Araneae: Lycosidae) in relation to life stage and water depth. J. Med. Entomol. 45, 1050–1056. Gerecke, R., Di Sabatino, A., 2007. Water mites (Hydrachnidia and Halacaridae) in spring habitats: a taxonomical and ecological perspective. In: Bertuzzi, M., Cantonati, E., Spitale, D. (Eds.), The Spring Habitat: Biota and Sampling Methods, pp. 193–216. Trento. Gerecke, R., Schwoerbel, J., 1991. Water quality and water mites (Acari: Actinedida) in the upper Danube region, 1959–1984. Mod. Acarol. 1, 483–491. Gettmann, W.W., 1978. Untersuchungen zum Nahrungsspektrum von Wolfsspinnen (Lycosidae) der Gattung Pirata. Mittl. Dtsch. Ges. für Allg. Angew. Entomol. 1, 63–66. Gliwicz, Z.M., Biesiadka, E., 1975. Pelagic water mites (Hydracarina) and their effect on the plankton community in a neotropical man-made lake. Arch. Hydrobiol. 76, 65–88. Goldschmidt, T., 2004. Environmental parameters determining water mite assemblages in Costa Rica. Exp. Appl. Acarol. 34, 171–197. Goldschmidt, T., Fu, V.W.K., 2011. Description of Hygrobates aloisii sp. nov. from Hong Kong, a new species of Hygrobates (Lurchibates) subgen. nov. (Acari, Hydrachnidia, Hygrobatidae), with data on the parasite–host relationship to the Hong Kong Newt Paramesotriton hongkongensis (Amphibia, Caudata, Salamandridae). Zool. Anz. 250, 19–31. Gonzalez, R.H., 1978. Annerossella knorri, a new species of mite on water lettuce from Thailand (Acari: Homocaligidae). Int. J. Acarol. 4, 221–225. Graham, A.K., Buddle, C.M., Spence, J.R., 2003. Habitat affinities of spiders living near a freshwater pond. J. Arachnol. 31, 78–89. Greenwood, M.J., McIntosh, A.R., 2008. Flooding impacts on responses of a riparian consumer to cross-ecosystem subsidies. Ecology 89, 1489–1496. Growns, J.E., 2001. Aquatic mites as bioindicators, with an Australian example. In: Halliday, R.B., Walter, D.E., Proctor, H.C., Norton, R.A., Colloff, M.J. (Eds.), Acarology: Proceedings of the 10th International Congress. CSIRO Publishing, Melbourne, pp. 136–142.

Chapter | 25  Subphylum Chelicerata, Class Arachnida

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SECTION | VI  Phylum Arthropoda

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Chapter | 25  Subphylum Chelicerata, Class Arachnida

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Sokolow, I.I., 1977. The protective envelopes in the eggs of Hydrachnellae. Zool. Anz. 198, 36–42. Stechmann, D.-H., 1980. Zum Wirtskreis syntopischer Arrenurus-Arten (Hydrachnellae, Acari) mit parasitischer Entwicklung an Nematocera (Diptera). Z. für Parasitenkd. 62, 267–283. Suter, R.B., Stratton, G., Miller, P.R., 2003. Water surface locomotion by spiders: distinct gaits in diverse families. J. Arachnol. 31, 428–432. da Silva, E.L.C., Picanço, J.B., Lise, A.A., 2005. Notes on the predatory behavior and habitat of Trechalea biocellata (Araneae, Lycosoidea, Trechaleidae). Biociências 13, 85–88. Ten Winkel, E.H., Davids, C., de Nobel, J.G., 1989. Food and feeding strategies of water mites of the genus Hygrobates and the impact of their predation on the larval population of the chironomid Cladotanytarsus mancus (Walker) in Lake Maarsseveen. Neth. J. Zool. 39, 246–263. Ullrich, F., 1978. Biologisch-ökologische Studien an den Larven rheophiler Wassermilben (Hydrachnellae, Acari), Schlitzer Produktionsbiologische Studien (29). Arch. Hydrobiol. Suppl. 54 (2), 189–255. Van der Hammen, H., Smit, H., 1996. The water mites (Acari: Hydrachnidia) of streams in the Netherlands: distribution and ecological aspects on a regional scale. Neth. J. Aquatic Ecol. 30, 175–185. Viets, K.O., 1987. Die Milben des Sußwassers (Hydrachnellae und Halacaridae [part.], Acari). 2. Katalog, vol. 8. Sonderbande Naturwissenschaftlichen Vereins Hamburg. 1012 pp. Wainstein, B.A., 1980. Opredelitel lichinok vodjanych kleshchei. (Key to Larval Water Mites). Institut Biologii Vnutrennikh Vod., Akademiia Nauk SSSR. 238 pp.

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Walter, D.E., Proctor, H.C., 2013. Mites: Ecology, Evolution and Behaviour – Life at a Microscale. Springer, Dordrecht. 494 pp. Weiberg, M., Edwards, D.D., 1997. Survival and reproductive output of Chironomus tentans (Diptera: Chironomidae) in response to parasitism by larval Unionicola foili (Acari: Unionicolidae). J. Parasitol. 83, 173–175. Wiggins, G.B., Mackay, R.J., Smith, I.M., 1980. Evolutionary and ecological strategies of animals in annual temporary pools. Arch. Hydrobiol. Suppl. 58, 97–206. Wiles, P.R., 1984. Watermite respiratory systems. Acarologia 25, 27–31. Wohltmann, A., 2001. Closely related species of Parasitengonae (Acari: Prostigmata) inhabiting the same areas: features facilitating cohabitation. In: Halliday, R.B., Walter, D.E., Proctor, H.C., Norton, R.A., Colloff, M.J. (Eds.), Acarology: Proceedings of the 10th International Congress. CSIRO Publishing, Melbourne, pp. 121–135. Zhang, Z.-Q., Fan, Q.-H., Pesic, V., Smit, H., Bochkov, A.V., Khaustov, A.A., Baker, A., Wohltmann, A., Wen, T., Amrine, J.W., Beron, P., Lin, J., Gabrys, G., Husband, R., 2011. Order Trombidiformes Reuter, 1909. In: Zhang, Z.-Q. (Ed.), Animal Biodiversity: An Outline of Higher-Level Classification and Survey of Taxonomic Richness. ­Zootaxa 3148. Magnolia Press, pp. 129–138. Zimmermann, M., Spence, J.R., 1989. Prey Use of the Fishing Spider Dolomedes triton (Pisauridae: Araneae) an Important Predator of the Neuston Community. Zimmermann, M., Spence, J.R., 1998. Phenology and life-cycle regulation of the fishing spider Dolomedes triton Walckenaer (Araneae, Pisauridae) in central Alberta. Can. J. Zool. 76, 295–309.

Chapter 26

Subphylum Myriapoda, Class Diplopoda Jean-Jacques Geoffroy Département Ecologie et Gestion de la Biodiversité, Muséum National d’Histoire Naturelle, UMR 7204 CESCO CNRS-MNHN-UPMC, Brunoy, France

Chapter Outline Introduction to the Subphylum 661 Evolution, Classification, and Phylogenetic Relationships661 Biogeography and Diversity 662 General Biology 662 Millipede Anatomy 662 External Morphology 662 Internal Anatomy 662 Millipede Physiology 663 Respiration663 Osmoregulation and Water Balance 663 Nervous System and Neurosecretion 663 Defensive Glands and Toxic Substances664

INTRODUCTION TO THE SUBPHYLUM The subphylum Myriapoda comprises four classes among terrestrial arthropods. Two of them, Pauropoda and Symphyla, are represented by minute micro-myriapods living in soils (Sheller, 2008, 2011; Szucsich and Sheller, 2011). The class Chilopoda (the centipedes) is comprised of five orders of venomous predators, very active and well-adapted for running and hunting living prey (Minelli, 2011). None of them are aquatic animals, except maybe some strongly halophilous-adapted species living in the marine littoral zone that can survive up to several hours total immersion in water during high tide. Despite the ability of a very few species to tolerate long periods of submersion in the intertidal zone, no myriapods are truly adapted for life in the sea (Mauriès, 1982; Barber, 2009; Golovatch and Kime, 2009). The class Diplopoda (the millipedes) is the most abundant and diverse group within the Myriapoda, comprising 15 orders and numerous families (Hoffman, 1980; Shelley, 2003). Most millipedes are saprophagous detritivores and mull formers in soils, converting vegetable debris into humus and playing a vital role in the cycling of matter, energy, and nutrients (Figure 26.1). They are well represented

Ecology and Behavior of Freshwater Millipedes 664 Physiological Problems Faced by Millipedes in Freshwater664 Millipedes of Tropical Islands and Amazonian Floodplains665 Millipedes of Swamps and Rivers 666 Aquatic Australian Millipedes 666 Millipedes in Subterranean Habitats of Europe 666 Collecting, Rearing, and Preparation for Identification 667 Sampling Methods 667 Extraction667 Fixing and Mounting 667 Culturing668 References668

in any kind of terrestrial ecosystem, as well as in subterranean habitats (Hopkin and Read, 1992; Shear, 1999). Millipedes range in size from very tiny (∼5 mm in Poly­ xenida) to the longest terrestrial invertebrates (Figure 26.2), some Spirostreptida being up to 35 cm long. Diplopods are also sources of various compounds useful for medicine and possibly industry, since they produce a large array of toxic substances that they use not only for selfdefense but probably also for inter-individual communication among populations (Geoffroy and Mauriès, 2014). Almost all the ∼15,000 described species of millipedes are strictly terrestrial, and the rare presence of diplopods in freshwater is generally occasional or accidental. No aquatic or filterfeeding millipedes are known in surface habitats. However, certain Amazonian species can survive months of floods by breathing air bubbles that are trapped on their bodies.

Evolution, Classification, and Phylogenetic Relationships The phylogenetic relationships of the various clades among the Myriapoda are not completely known, and active

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00026-7 Copyright © 2015 Jean-Jacques Geoffroy. Published by Elsevier Inc. All rights reserved.

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can survive in several kinds of extremely wet habitats or water in swamps, during overflow in tropical and temperate floodplains, and in subterranean aquatic environments.

GENERAL BIOLOGY Some aspects of the biology of millipedes are briefly presented below and more detailed information related to reproduction, development, molting, and life histories is available in Hopkin and Read (1992) and Minelli (in press). FIGURE 26.1  Cylindroiuluspunctatus (Diplopoda, Julida, Julidae; Leach, 1815). A typical julid millipede from a European temperate ecosystem. Photograph by Régine Vignes-Lebbe and Geofffroy, J.-J.

Millipede Anatomy External Morphology

FIGURE 26.2  Anadenobolus sp. (Diplopoda, Spirobolida) in a tropical forest of Brazilian Pantanal. Photograph by Zillikens.

research programs are conducted on the subject. Molecular studies are being used together with morpho-anatomical approaches to show relationships within the subphylum, and the systematics are progressively supported by genomic studies. The present understanding of myriapod phylogeny, and more precisely diplopod phylogeny, strongly support the monophyly of the Myriapoda and the phylogenetic relationships among the Arthropoda or between clades in the Myriapoda (Koch, 2003; Edgecombe, 2004, 2010; Gai et al., 2006; Regier et al., 2010; Giribet and Edgecombe, 2012) and particularly the Diplopoda (Enghoff, 2001; ­Sierwald et al., 2003; Sierwald and Bond, 2007).

Biogeography and Diversity Freshwater diplopods are very rare and distributed in a very few cases in aquatic habitats. Their ecological role in freshwater is not understood, and they cannot be considered as important components in the freshwater food web. Most “freshwater millipede species” are distributed in rather limited or regional localities, and almost nothing is known about the dispersal of freshwater diplopods occurring in particular aquatic habitats. Semi-aquatic millipedes

Millipedes are comprised of three body sections: head, body, and telson (Figure 26.3). The head bears two pairs of mouthparts, the mandibles and the gnathochilarium, which is formed from the maxillae. This very particular structure forms the “floor” of the buccal cavity, is made of several plates, and bears sensory organs. The head also bears sensory structures, including one pair of antennae, Tömösvary organs, and ocelli in an ocular field. The body is long and cylindrical, and sometimes exhibits prominent lateral projections. Behind the head is the apodous collum, followed by three “thoracic” segments bearing one pair of legs each. The following rings are diplosegments that bear two pairs of legs. In adult males, some legs are deeply modified in gonopods and associated with sperm transfer and copulation. In adult females, the vulvae open behind the second pair of legs. Body rings are comprised of an anterior prozonite and a posterior metazonite on which open paired defense glands. The telson consists of a pre-anal ring with a pygydium developed into a projection, anal valves that open during defecation, and a ventral scale. Between the posterior leg-bearing rings and the telson, are several apodous rings and a proliferation zone where future segments are initiated and develop. The cuticle, deeply calcified, consists of extensively modified dorsal tergite, ventral sternite, and lateral pleurites. Walking legs comprise seven to eight articles: coxa, trochanter, prefemur, femur, postfemur, tibia, tarsus, and claw. Walking in millipedes is rather complicated. Each leg takes a step with a propulsive stage followed by a recovery stage. The legs on the right side move in phase with those on the left but each leg is out of phase with immediately anterior and posterior legs. This causes the very spectacular metachronal wave that is so characteristic of millipedes.

Internal Anatomy The digestive tract is a tube comprising foregut, midgut, and hindgut, and is linked to associated organs such as the salivary glands, maxillary glands (nephridial organs), fat

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FIGURE 26.3  Nanogona polydesmoides (Diplopoda, Chordeumatida, Craspedosomatidae; Leach, 1814). Anterior part, lateral view of a female. Drawing by Michelle Bertoncini and Geoffroy, J.-J.

body, and Malphigian tubules involved in osmoregulation and excretion. The reproductive organs open anteriorly on the ventral side of the third body ring: paired ovaries and vulvae, tubular testes, and bilobed penis. The ventral nerve chord consists of paired segmented ganglia. It is swollen in the head to form the brain and associated with neurosecretory activity.

Millipede Physiology Respiration During the respiration process, living organisms take oxygen from the environment and release carbon dioxide. Chemical breakdown of organic substances takes place in tissues and cells, while energy is released and carbon dioxide is produced. In millipedes, exchange of oxygen and carbon dioxide between tissues and atmosphere takes place via tracheae. Fundamentally, diplopods are tracheaetes living in air. The tracheal system opens via cuticular holes called spira­cles. A pair of spiracles opens on each sternite, anteriorly and laterally to the coxae of the leg. In some groups, they are protected by a cuticular lattice or are closeable; both structures help control water loss from the moist linings of the tracheae. The spiracle opens in an atrium from which numerous tracheae ramify among the tissues. Oxygen uptake in millipedes shows quite different rates, depending on the group or species, the natural environment or experimental conditions, and possible adaptations of the respiratory metabolism to ambient parameters change.

Osmoregulation and Water Balance The water content of millipedes is related both to the species and the particular physiological state of the individual. Millipedes, especially large ones, are highly susceptible to water loss, and species differ in their susceptibility to desic­ cation. Obvious differences between species are strongly related to environmental factors, but millipedes have evolved various physiological and behavioral compensatory mechanisms to reduce water loss and avoid dehydration in osmotically unfavorable situations. For example, they may rapidly walk to a wet area, roll up to minimize cuticular or respiratory water losses, increase internal osmotic pressure, and/or add water from the environment via metabolism or transport. The very low cuticular permeability and success in water conservation of some species allow these welladapted taxa to live in very dry habitats, including deserts. But most obviously, millipedes avoid dry areas, are most active at night, and reduce the surface area of the body exposed to the atmosphere by rolling up into a sphere, coiling up to form a spiral, or aggregating in large numbers to reduce the collective surface area exposed to the atmosphere.

Nervous System and Neurosecretion Millipedes have a strong sense of direction, and they receive environmental cues via several sensory systems: mechanoreceptors, gustatory receptors, olfactory receptors, and chemoreceptors. Linked with the nervous system, these simple sensory structures are present in several complex sense organs such as eyes (or ocelli gathered in an ocular field),

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Tömösváry organs (or post-antennal temporal organs), trichobotria, sensilla, and antennae. The antennae are covered in a variety of sensory structures linked with gustatory and olfactory receptors, thermo- and hygro receptors, and detection of pheromones before mating. All these functional structures are particularly active in the apical section of the antennae that bears characteristic cone-shaped sensilla. Many activities of millipedes are under secretory control, such as reproductive cycle, molting, and metabolic processes. Several neurohemal organs have been described in the brain of diplopods; these are located in the periesophageal ring and subesophageal nerve mass in julids. The Gabe’s organ or cerebral glands, a primitive structure, contains no intrinsic secretory cells and could be involved in the controlling of molting. l The paraesophageal bodies are located lateroventrally to the brain. l Connective bodies are cephalic neurohemal organs that may control ovocyte growth. l  The periesophageal blood sinus formations are associated with the paraesophageal bodies. l  The paired paracommissural plates lie near the transverse commissure of the circumesophageal connectives. l The periesophageal gland, specific of Penicillata, is considered to be an ecdysial gland. l

Defensive Glands and Toxic Substances These slow-moving diplopods cannot avoid predator attack, and have, therefore, evolved physical and chemical defensive measures. A physical protection is offered by the heavy calcified exoskeleton, and rolling- or coilingup behavior. Paired repugnatory defense glands are present in most millipedes, and ozopores open laterally on each diplosegment. The secretion is not really dangerous for humans but may cause considerable discomfort, pain, or burns when it comes into contact with sensitive skin or eyes. Diplopod substances may be used in traditional medicine, poisoning arrowheads, or even as inhibitors against tumoral cells and other clinical treatments. Products secreted by millipedes also elicit self-anointing or even social anointing in many groups of vertebrates such as birds, monkeys, and lemurs. Several main types of defensive glands and toxins are recognized in millipedes (Eisner et al., 1978; Geoffroy and Mauriès, 2014). They are quite effective against a wide range of potential predators, even if predation in the field has rarely been observed. The most effective predators are certainly among arachnids and insects, such as large spiders, opilionids, true bugs, and above all, ants. The structure of defensive glands present in most diplopod orders is simple. A spherical sac contains secretory cells that open laterally in a small pore. The secretions in Spirobolida, Spirostreptida, and Julida are benzoquinones and

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hydroquinones. Polyzoniida produce nitrogen-containing terpenoids called “polyzonimine,” which are irritants to insects. In the Callipodida, defensive secretions are typically phenols. The Polydesmida is the only order able to secrete cyanide. The secretion, manufactured by the millipede from simple precursors, is emitted from two compartments of the glands. The inner compartment stores mandelonitrile; whereas in the outer small compartment, an enzyme catalyzes breakdown of mandelonitrile into hydrogen cyanide and benzaldehyde. The secretion is remarkable for its highly repellant defensive properties and may also protect against fungal attack. In Glomerida, the pill millipedes, defensive glands open mid-dorsally and the secretion is issued as a sticky droplet containing specific quinazolinones called “glomerin” and “homoglomerin,” acting to discourage feeding and as sedatives and toxins to predators such as wolf-spiders.

ECOLOGY AND BEHAVIOR OF FRESHWATER MILLIPEDES Physiological Problems Faced by Millipedes in Freshwater As a first look and rough answer, we could consider that there are no “truly aquatic” millipedes anywhere in the world because all millipedes lack a gill or other structure to extract oxygen from water, unlike almost all other aquatic organisms. Their respiratory system is specially adapted for extracting oxygen from air. The exchange of oxygen and carbon dioxide between the cells of these arthropods and the atmosphere takes place via tracheae. The tracheal respiratory system opens via cuticular spiracles located on sternites (Blower, 1985; Hopkin and Read, 1992). Being air-breathing, most millipedes living in flooded or waterlogged soils usually come to the surface, move to upper litter layers, or even climb trees to avoid drowning. The excretory system of millipedes is also not adapted for life in water. Terrestrial millipedes primarily excrete nitrogenous wastes as ammonia (which has to be excreted rapidly as gas) and uric acid (which is stored before excretion), with the latter requiring more energy but less water to produce and excrete. They have not evolved to release nitrogenous excretory products directly into water as liquid ammonia. Nevertheless, some millipedes can survive under water for surprising lengths of time. Most of them belong to the order Polydesmida (Figure 26.4), like the pantropical Oxidus gracilis (C.L. Koch, 1847). These millipedes may rely on an air bubble that gets trapped around their spiracles, as in some diving beetles. Once the oxygen in that air bubble is depleted, however, the diplopod must replenish it with more air, or drown. Such a special situation among the

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FIGURE 26.4  Oxidus gracilis (Diplopoda, Polydesmida, Paradoxosomatidae; C.L. Koch, 1847). A pantropical widespread millipede. Photo Lemaire, J.-M.

FIGURE 26.6  Diplopoda, Polydesmida, Pyrgodesmidae. Habitus of a minute pyrgodesmid millipede from Vanuatu. Photograph by Deharveng, L.

FIGURE 26.5  Polydesmus angustus (Diplopoda, Polydesmida, Polydesmidae; Latzel, 1884). A common saprophagus polydesmid millipede from Western Europe, shown here walking on oak litter. Photograph by Régine Vignes-Lebbe and Geofffroy, J.-J.

terrestrial millipedes was reported about polydesmid millipedes (Figure 26.5) under stones in French streams and tentatively explained a century ago as a form of branchial respiratory system in millipedes. Observations of the diplopods making pumping movements of the rectal area were interpreted as a respiratory surface formed by the rectal wall (Causard, 1903). Several cases of such aquatic or semiaquatic diplopods have been documented in different localities, ecosystems, and biogeographical regions in the world. Examples are reported for species in Polyzoniida, Julida, Chordeumatida, and above all, Polydesmida.

Millipedes of Tropical Islands and Amazonian Floodplains The widespread pantropical polydesmid Aporodesminus wallacei Silvestri, 1904 (Pyrgodesmidae) is distributed in several islands (St. Helena, Hawaii, Tahiti) and in urban freshwater creeks near Sydney, Australia (see below). It belongs to a group of minute polydesmid millipedes (Figure 26.6), some of them well adapted to survive long submersions. Adults and subadults have been sampled underwater, and the species is considered to be semi-aquatic millipedes, with similar habits to the related minute pyrgodesmid Cryptocorypha ornata (Attems, 1938). The aquatic habit in these taxa is supported by the well-adapted structure of the mouthparts and the presence of a cerotegument enabling plastron

respiration (Adis et al., 1998). Morphological adaptations of the mouthparts and spiracles have been reported in diplopods strongly suspected of entering freshwater bodies and feeding on fine-grained organic particles in the water (Enghoff, 1985; Burrows et al., 1994). In most of these species, submersion tolerance or resistance lasting weeks or even months is possible because of plastron respiration using very special structures and cuticular secretions, including a cerotegument that covers the spiracles (Messner and Adis, 1992, 1994, 1997; Messner et al., 1996). This is particularly well documented in another pyrgodesmid diplopod, Myrmecodesmus adisi (Hoffman, 1985), which was reported to have survived more than a 6-month long flood period under a submerged tree trunk in an Amazonian inundation forest (Hoffman, 1985; Adis, 1986; Messner and Adis, 1988; Adis and Messner, 1997). All tergites are covered with this thick cerotegument layer, extending to the coxal region of the sternite. The spiracles are, therefore, entirely or partly covered by the externally hydrophilous secretion that facilitates long-term submersion. Air is held in the hydrophobic cavity below the cerotegument and around the spiracles. Then, plastron respiration adds oxygen from the surrounding water to the trapped reserve of air; bubbles of atmospheric air are also captured in the water and added to the plastron (Adis, 1992; Messner and Adis, 1992, 1994, 1997). However, all field observations and laboratory experiments are related to adults or subadults, and development of juveniles probably remains restricted to very moist terrestrial habitats and banks of water bodies (Adis et al., 1998). In contrast, other species, such as Myrmecodesmus duodecimlobatus (Golovatch, 1996), lacking either morphological or physiological adaptations, rely on vertical migrations along tree trunks to survive flood periods (Adis et al., 1996; Adis, 1997; Minelli and Golovatch, 2001; Adis and Junk, 2002). Although almost nothing is known about the small chelodesmid Pandirodesmus disparipes Silvestri, 1932 from

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Guyana, it is suspected of being semi-aquatic. It is good climber, swimmer, and glider, and several morphological features of the legs, tubiform spiracles, sternites, and hydrophobe setae strongly suggest a special ecology closely associated with freshwater, despite the absence of cerotegument and microtrichia in the spiracles (Golovatch and Kime, 2009).

Millipedes of Swamps and Rivers Several reports of “aquatic” Polydesmida (mainly Paradoxosomatidae) were recently noted for streams in North America (Shelley, personal communication). Numerous specimens, thousands of them, were observed in the middle of a stream, covering water algae; all were alive and well as far as it could be determined. These polydesmid millipedes are sightless, and their mass migrations seem not to be deterred by crossing a stream. They cannot be considered as a truly aquatic species of millipedes, but they certainly seem to show capacity to survive in water. Paradoxosomatid species observed in the Hudson Valley were under wet soggy leaf litter all along a very small headwater stream. They are not “aquatic” but living and well adapted to damp areas close to the streams. Similar situations have been observed in Australia, and they are thought to have occurred in swamps, floodplains, and river banks in Central Europe during major floods last century (Tajovsky, 1999). European millipedes can survive in floodplains using a riskstrategy, due to a combination of high reproductive rates, dispersal, and re-immigration following catastrophic events (Adis and Junk, 2002; Golovatch and Kime, 2009).

Aquatic Australian Millipedes Populations of two millipede species were discovered under stones submerged in a creek on the Macquarie University campus in the northern suburb of Sydney. One was a Polyzoniida belonging to the family Siphonotidae, and the second was a Polydesmida belonging to the family Pyrgodesmidae. The latter is closely related to pyrgodesmid species described in oceanic islands and Amazonian inundation forests. This was the first published report of “aquatic millipedes” in Australia. They became targets in the center of a hot debate about protection and conservation of invertebrate populations threatened by construction works of a dam (Burrows et al., 1994; Black, 1997). There was obvious risk of damage to the clay walls of the creek, where the millipedes were suspected to lay their eggs. Animals were present in numbers in and near the water from autumn to late spring, but moved to other habitats during the summer, probably burrowing into damp soil or the creek bed. They were observed leaving the water in large numbers during early winter to mate and lay eggs in cracks in the banks. They apparently

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breathe cutaneously, and both adult and juvenile stadia have been found totally submerged; long-term submergence is considered a normal part of their behavior (­Burrows et al., 1994; Adis et al., 1998). The specimens were later identified as Aporodesminus wallacei Silvestri, 1904 (Pyrgodesmidae), and a well-documented re-description of the species was published (Adis et al., 1998). A. wallacei has also been reported from St. Helena (southern Atlantic Ocean), Tahiti, and the Hawaiian Islands. It is not known to be native anywhere in its range but belongs to the pyrgodesmid millipede fauna of Australia (Mesibov, 2012). The modified mouthparts in A. wallacei exhibit hypertrophied, hair-shaped teeth of pectinate lamellae and reduced masticating parts (Adis et al., 1998: Figs 5-8), and the species show highly adapted features of the tegument allowing long-term resistance to water submersion (see discussion above on the cerotegument). Having both mouthparts adapted for food uptake underwater and a cerotegument for plastron respiration, A. wallacei is one of the best semiaquatic millipede examples we have, with the others distributed in deep caves.

Millipedes in Subterranean Habitats of Europe Caves and subterranean habitats more generally are shelters for many different lineages of myriapods, particularly diplopods. Most of the cave-dwelling and strongly adapted troglobionts differ from their epigean relatives in showing striking morphological and physiological features (Shear, 1969; Mauriès, 1994, 2004; Culver and Shear, 2012). Among these adaptations to cave life, mouthparts are commonly convergently modified in Julidae, Blaniulidae, Polydesmidae, etc. in different geographic areas (Enghoff, 1985). The cave environment is sometimes considered equivalent ecologically to an island, and their insularity combined with general habitat conditions in subterranean environments appear to be highly significant components of millipede diversity and adaptation (Enghoff, 1993). Therefore, cave-dwelling millipedes are obviously adapted to hygrophily and most certainly comprise the best candidates to possess aquatic or at least semiaquatic ways of life among terrestrial myriapods. Serradium semiaquaticum Enghoff et al., 1997 (Polydesmida, Polydesmidae) has been described from several northern Italian caves (Figure 26.7) and is a close, derived relative and sister species of Serradium hirsutipes Verhoeff, 1941, which lives in the same caves but shows a wider distribution (Enghoff et al., 1997). The species is utterly remarkable in exhibiting peculiar semiaquatic and feeding habits and modified anatomical features, especially mouthparts and spiracles (Caoduro, 1995; Enghoff et al., 1997; Adis et al., 1997, 1998). S. semiaquaticum appears to be

Chapter | 26  Subphylum Myriapoda, Class Diplopoda

667

COLLECTING, REARING, AND PREPARATION FOR IDENTIFICATION Sampling Methods

FIGURE 26.7  Serradium semiaquaticum (Diplopoda, Polydesmida, Polydesmidae; Enghoff et al., 1997). A cavernicolous millipede from northern Italian caves, which is remarkable because of its semiaquatic habits. Photograph by Luca Cavallari.

the single truly semiaquatic millipede occurring in a cold subterranean environment, at least for subadults and adults (Adis et al., 1997; Golovatch and Kime, 2009). This troglobitic species is remarkable in showing a combination of morphological, ecophysiological, and ecoethological adaptative traits that facilitate its amphibious mode of life. These include the following:  The modified, broom-like hypertrophied pectinate lamellae of the mandibular gnathal lobes and reduced masticating parts, which allow uptake of fine organic particles, clay, and limestone material from moist surfaces along the banks of water bodies and from underwater and the bottom of cave rivulets. l  A hydrophilic surface of the cuticle, which facilitates entry to the water. l Hydrophobic microtrichia of the spiracles, which allow plastron respiration under freshwater and small bubbles of atmospheric air in the water to be captured if water currents are strong enough. l An ion-catching chloride epithelia in the intersegmental membranes, which allows additional uptake of ions and of dissolved oxygen from the water. l

This semiaquatic troglobiont is a stenothermal species that enters cold subterranean water bodies voluntarily as part of its natural semiaquatic behavior, which contrasts with the behavior of its relative S. hirsutipes. Taken together, the listed adaptations enable specimens to enter subterranean water bodies periodically during the day or and to remain submerged for up to four weeks in laboratory conditions; the former demonstrates the species’ tolerance and the latter its submersion resistance (Adis et al., 1997). In Papua, New Guinea, the hygrophilous montane troglobite Selminosoma chapmani Hoffman, 1978 (Polydesmida, Paradoxosomitadae) is also highly adapted both to ecological conditions in caves in general and to total immersion in water in particular (Hoffman, 1977; Messner et al., 1996). Moreover, several other cases are strongly suspected from China or Southwest Asia where millipedes have been observed entering water without apparent difficulties (Deharveng and Bedos, 2000).

Diplopods are collected qualitatively and quantitatively. Qualitative collections are made to determine which kinds of millipedes are present in a studied ecosystem, leading to the best possible specific richness. Quantitative samples are used to evaluate the components of a community, the proportion of each species population in the environment, and the seasonal and temporal dynamics of the population. Qualitative methods for sampling myriapods include hand collecting, pitfall traps, and field sieving. The type and size of millipede being collected will determine the mesh size of the sieve selected. Traps can be set dry or contain a bait or a killing-preservative liquid such as ethanol or ethylene glycol. Bait can be rotting fruit or meat. Heavily scented cheese is particularly efficient for millipedes in caves or in peculiar MSS habitat (mesovoid shallow substratum; or originally milieu souterrain superficiel in French). Quantitative sampling methods involve the use of a device to remove core samples of measurable sizes from the soil or sediment, on the basis of need and the desired biotype to be investigated. Techniques applied to any soil macroarthropod group are suitable for millipedes, but qualitative techniques will be better for freshwater investigations. Millipedes are not particularly fragile animals; however, sampling and extraction techniques should be done as carefully as possible, and you should be careful to avoid the risk that the millipede will die by desiccation.

Extraction The process of extraction involves removing diplopods from samples collected in various habitats. The main principle of millipede extraction is based on the fact that diplopods are naturally negatively phototropic and positively geotropic. One of the oldest and simplest methods of extracting di­­­­ plopods from a wide range of samples is the Berlese-­Tullgren funnel method, or a modification thereof, or a Kempson extractor. The Berlese collector consists of a funnel, a holder to maintain it upright, a sieve inside the funnel, with a mesh size of ∼2 mm, a container with killing-­preservative liquid (ethanol 70% added to 25% ethylene glycol or hypersaturated saltwater), and a top. The sample will simply dry under the environmental conditions or a bulb can be added at the top in order to speed up the process and make the light and temperature gradient harder.

Fixing and Mounting Specimens should be stored in vials in 75–80% ethanol for their later study, or preserved alive in boxes with wet dead

668

organic matter or sediment. The diplopods should be fixed for at least 2–3 days in clean ethanol before transfer to glycerin after dissection for mounting on microscope slides. Gonopods of adult males are generally cleared in 80% lactic acid and temporarily mounted in 60% lactic acid for optical microscopy. Other body parts can be temporarily mounted in a 1:1 glycerol to water mixture.

Culturing Some soil - or freshwater millipedes can be maintained alive in the laboratory in plastic boxes with convenient food and a separate compartment with water and natural sediment. This rearing method allows subadults to molt and reach the definitive mature stadium, or gravid females to lay eggs. It becomes, therefore, possible to associate young juveniles with adult forms.

REFERENCES Adis, J., 1986. An “aquatic” millipede from central Amazonian inundation forest. Oecologia 68, 347–349. Adis, J., 1992. Überlebensstrategein terrestrischer Invertebraten in überschwemmungswäldern Zentralamazoniens. Verhandlungen Des. Naturwiss. Vereins Hamburg 33, 21–114. Adis, J., 1997. Terrestrial invertebrates: survival strategies, group spectrum, dominance and activity patterns. In: Junk, W. (Ed.), The Central Amazon floodplain. Ecology of a Pulsing System. Ecological Studies, vol. 26. Springer Verlag, Berlin, pp. 299–317. Adis, J., Caoduro, G., Messner, B., Enghoff, H., 1997. On the semiaquatic behaviour of a new troglobitic millipede from northern Italy (Diplopoda, Polydesmida: Polydesmidae). Entomol. Scandinavia Suppl. 51, 301–306. Adis, J., Golovatch, S.I., Hamann, S., 1996. Survival strategy of the terricolous millipede Cutervodesmus adisi Golovatch (Fuhrmannodesmidae, Polydesmida) in a blackwater inundation forest of Central Amazonia (Brazil) in response to the floodpulse. Mémoires Du. Muséum Natl. d’Histoire Nat. 169, 523–532. Adis, J., Golovatch, S.I., Hoffman, R.L., Hales, D.F., Burrows, F.J., 1998. Morphological adaptations of the semiaquatic millipede Aporodesminus wallacei Silvestri, 1904 with notes on the taxonomy, distribution, habitats and ecology of this and a related species (Pyrgodesmidae Polydesmida Diplopoda). Trop. Zool. 11, 371–387. Adis, J., Junk, W., 2002. Terrestrial invertebrates inhabiting lowland river floodplains of Central Amazonia and Central Europe: a review. Freshw. Biol. 47, 711–731. Adis, J., Messner, B., 1997. Adaptations to life under water: tiger beetles and millipedes. In: Junk, W.J. (Ed.), The Central Amazon Floodplain. Ecology of a Pulsing System. Ecological Studies, vol. 126. Springer Verlag, Berlin, pp. 318–330. Barber, A.D., 2009. Littoral myriapods: a review. Soil. Org. 81 (3), 735–760. Black, D., 1997. Diversity and biogeography of Australian millipedes. Memoirs Mus. Vic. 56 (2), 557–561. Blower, J.G., 1985. Millipedes. Linnean Society Synopses of the British Fauna (New Series), vol. 35, E.J. Brill/Dr W. Backhuys, London. 242 pp.

SECTION | VI  Phylum Arthropoda

Burrows, F.J., Hales, D.F., Beattie, A.J., 1994. Aquatic millipedes in ­Australia: a biological enigma and a conservation saga. Aust. Zool. 29 (3/4), 213–216. Causard, M., 1903. Recherches sur la respiration branchiale chez les myriapodes diplopodes. Bull. Sci. la Fr. la Belg. 37, 461–479. Caoduro, G., 1995. First observations of the behaviour in captivity of two troglobitic species from Monte Lessini, Verona: Italaphaenops dimaioi Ghidini (Coleoptera, Carabidae) and Serradium hirsutipes Verhoeff (large form: Strasser, 1981) (Diplopoda, Polydesmidae). Mémoires Biospéologie 22, 7–12. Culver, D.C., Shear, W.A., 2012. Myriapoda. In: White, W.B., Culver, D.C. (Eds.), Encyclopedia of Caves, second ed. Elsevier, Academic Press, Amsterdam, New York, Oxford, pp. 538–542. Deharveng, L., Bedos, A., 2000. The cave fauna of Southeast Asia: origin, evolution and ecology. In: Wilkens, H.D., Culver, D.C., Humphreys, W. (Eds.), Ecosystems of the World. Subterranean Ecosystems, vol. 30. Elsevier, Oxford, pp. 603–632. Edgecombe, G.D., 2004. Morphological data, extant Myriapoda, and the myriapod stem-group. Contrib. Zool. 73 (3), 207–252. Edgecombe, G.D., 2010. Arthropod phylogeny: an overview from the perspectives of morphology, molecular data and the fossil record. Arthropod Struct. Dev. 39, 74–87. Eisner, T., Alsop, D., Hicks, K., Meinwald, J., 1978. Defensive secretions of millipedes. In: Bettini, S. (Ed.), Arthropod Venoms. Hanbook of Pharmacology, vol. 48. Springer-Verlag, Berlin, pp. 41–72. Enghoff, H., 1985. Modified mouthparts in hydrophilous cave millipedes. Bijdr. tot Dierkd. 55, 67–77. Enghoff, H., 1993. Millipedes, caves and islands. Mémoires Biospéologie 20, 77–80. Enghoff, H., 2001. Millipede phylogeny: how much do we know and what is it good for? Fragm. Faun. (Warszawa) 43 (Suppl. 2000), 1–17. Enghoff, H., Caoduro, G., Adis, J., Messner, B., 1997. A new cavernicolous, semiaquatic species of Serradium (Diplopoda, Polydesmidae) and its terrestrial, sympatric congener. With notes on the genus Serradium. Zool. Scr. 26 (3), 279–290. Gai, H.Y., Song, D.X., Sun, H.Y., Zhou, K.Y., 2006. Myriapod monophyly and relationships among myriapod classes based on Nearly Complete 28S and 18S rDNA sequences. Zool. Sci. 23 (12), 1101–1108. Giribet, G., Edgecombe, G.D., 2012. Reevaluating the arthropod tree of life. Annu. Rev. Entomol. 57, 167–186. Geoffroy, J.-J., Mauriès, J.-P., 2014. Myriapodes Diplopodes. In: Goyffon, M., Rollard, C. (Eds.), La Fonction Venimeuse., Lavoisier, Paris. Golovatch, S.I., Kime, R.D., 2009. Millipede (Diplopoda) distributions: a review. Soil. Org. 81 (3), 565–597. Hoffman, R.L., 1977. Diplopoda from papuan caves (Zoological results of the British speleological expedition to Papua-New Guinea, 1975, 4). Int. J. Speleol. 9, 281–307. Hoffman, R.L., 1980. Classification of the Diplopoda. Muséum d’Histoire Naturelle, Genève. 237 pp. Hoffman, R.L., 1985. A new millipede of the genus Gonographis from an inundation forest near Manaus (Pyrgodesmidae). Amazoniana 9 (2), 234–246. Hopkin, S.P., Read, H.J., 1992. The biology of Millipedes. Oxford University Press, Oxford and New York. 233 pp. Koch, M., 2003. Monophyly of the Myriapoda? Reliability of current arguments. Afr. Invertebr. 44 (1), 137–153.

Chapter | 26  Subphylum Myriapoda, Class Diplopoda

Mauriès, J.-P., 1982. Dolichoiulus tongiorgii (Strasser), diplopode halophile nouveau pour la faune de France. Remarques sur la classification des Pachyiulini (Myriapoda, Diplopoda, Iulida). Bull. Du. Muséum Natl. d’Histoire Nat. 4eme Série, Sect. A 4 (3–4), 433–444. Mauriès, J.-P., 1994. Diplopoda. In: Juberthie, C., Decu, V. (Eds.), Encyclopaedia Biospeologica, Tome 1. Société Internationale de Biospéologie, Moulis-Bucarest, pp. 255–262. Mauriès, J.-P., 2004. Myriapoda (Centipedes and millipedes). In: Gunn, J. (Ed.), Encyclopaedia of Cave and Karst Science. Fitzroy Dearborn, New York, London, pp. 534–536. Mesibov, R., 2012. The first native pyrgodesmidae (Diplopoda, polydesmida) from Australia. Zookeys 217, 63–85. Messner, B., Adis, J., 1988. Die Plastronstrukturen der bisher einzigen submers lebenden Diplopodenart Gonographis adisi Hoffman, 1985 (Pyrgodesmidae, Diplopoda). Zool. Jahrbücher Abt. für Anat. 117, 277–290. Messner, B., Adis, J., 1992. Die Plastronatmung bei aquatischen und flutresistenten terrestrischen Arthropoden (Acari, Diplopoda und Insecta). Mittl. Dtsch. Ges. für Allg. Angew. Entomol. 8, 325–327. Messner, B., Adis, J., 1994. Funktionsmorphologishe Untersuchungen an den Plastronstrukturen der Arthropoden. Verhandlungen Westdtsch. Entomol. 1993, 51–56. Messner, B., Adis, J., 1997. Über die Vielfalt der Plastronatmung – Vorschlag zur Neuefassung des Begriffes “Plastron”. Verhandlungen Westdtsch. Entomol. 1996, 89–92. Messner, B., Adis, J., Zulka, P., 1996. Stigmale Plastronstrukturen, die einigen Diplopoden-Arten eine submerse Lebensweise in kaltem und fliessendem Wasser ermöglichen. Rev. Suisse Zool. 103 (3), 613–622. Minelli, A. (Ed.), 2011. Treatise on Zoology – Anatomy, Taxonomy, Biology. The Myriapoda, vol. 1. Brill, Leiden, Boston, p. 530±VIII. Minelli, A. (Ed.), (in press). Treatise on Zoology – Anatomy, Taxonomy, Biology. The Myriapoda, vol. 2. Brill, Leiden, Boston, 500 pp.

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Minelli, A., Golovatch, S.I., 2001. Myriapoda. In: Levin, S.A. (Ed.), Encyclopedia of Biodiversity, vol. 4. Academic Press, San Diego, pp. 291–303. Regier, J.C., Schultz, J.W., Zwick, A., Hussey, A., Ball, B., Wetzer, R., Martin, J.W., Cunningham, C.W., 2010. Arthropod relationship ­ revealed by phylogenomic analysis of nuclear protein-coding sequences. Nat. Lett. 463, 1079–1083. Shear, W.A., 1969. A synopsis of the cave millipedes of the United States, with an illustrated key to genera. Psyche 76, 126–143. Shear, W.A., 1999. Millipeds. These “thousand-legged” arthropods are little known but appear to hold many secrets for scientists. Am. Sci. 87, 232–240. Sheller, U., 2008. A reclassification of the Pauropoda. Int. J. Myriapodol. 1, 1–38. Sheller, U., 2011. Pauropoda. In: Minelli, A. (Ed.), Treatise on Zoology – Anatomy, Taxonomy, Biology. The Myriapoda, vol. 1. Brill, Leiden, Boston, pp. 467–508. Shelley, R.M., 2003. A revised, annotated, family-level classification of the Diplopoda [For 2002]. Arthropoda Sel. 11 (3), 187–207. Sierwald, P., Bond, J.E., 2007. Current status of the myriapod class Diplopoda (millipedes): taxonomic diversity and phylogeny. Annu. Rev. Entomol. 52, 401–420. Sierwald, P., Shear, W.A., Shelley, R.M., Bond, J.E., 2003. Millipede phylogeny revisited in the light of the enigmatic order Siphoniulida. J. Zool. Syst. Evol. Res. 41, 87–99. Szucsich, N., Sheller, U., 2011. Symphyla. In: Minelli, A. (Ed.), Treatise on Zoology – Anatomy, Taxonomy, Biology. The Myriapoda, vol. 1. Brill, Leiden, Boston, pp. 445–466. Tajovsky, K., 1999. Impact of inundations on terrestrial arthropod assemblages in southern Moravia floodplain forests, the Czech Republic. Ekológia Bratisl. 18 (Suppl. 1), 177–184.

Chapter 27

Introduction to “Crustacea” James H. Thorp Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, Lawrence, KS, USA

D. Christopher Rogers Kansas Biological Survey, and Biodiversity Institute, University of Kansas, Lawrence, KS, USA

Alan P. Covich Odum School of Ecology, University of Georgia, Athens, GA, USA

Chapter Outline Introduction671 General Systematics and Phylogenetic Relationships 671 Distribution and Diversity 672 General Biology 673 External Anatomy 673 Exoskeleton673 Appendages673 Internal Anatomy and Physiology 675 Digestive System 675 Circulation and Respiration 675 Fluid and Solute Balance 675 Neural System and Receptors 676 Chemoreception677 Reproduction and Development 678 Reproduction678 Development678 Growth678

INTRODUCTION General Systematics and Phylogenetic Relationships Recent phylogenetic studies demonstrate that the traditional “Crustacea” is most likely paraphyletic and is, in reality, comprised of several important clades including Hexapoda (see Chapter 24). This larger group is treated as the subphylum Tetraconata (sometimes known as Pancrustacea). In this chapter, we will address only those clades in the traditional “Crustacea.” Crustaceans adapted to freshwater habitats very early in their history (Abele, 1982). Indeed, the known crustacean

General Ecology and Behavior 679 Habitat Selection 679 Physiological Constraints 680 Salinity680 Temperature680 Hardness and pH 681 Feeding Behavior 681 Predators and Parasites 682 Collecting, Culturing, and Specimen Preparation 682 Collecting682 Culturing682 Specimen Preparation 682 References683

evolutionary record extends back to the lower Cambrian with fossils associated with a broad range of aquatic habitats, especially marine (Schram, 1982; Babcock et al., 1998). The early freshwater species were able to avoid a diverse array of marine predators and to benefit from access to inland food resources by developing physiological adaptations for variable salinities and temperatures in rivers and lakes. The wide range of sizes, shapes, and behaviors among species living in inland waters today indicates how this group continues to evolve in response to changing environmental conditions and complex biotic interactions (Augusto et al., 2009). Although only about 15% of the extant crustacean species currently occur in inland waters

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00027-9 Copyright © 2015 Elsevier Inc. All rights reserved.

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(Table 27.1; and Bowman and Abele, 1982; Crandall and Buhay, 2008), crustaceans are extremely important to ecosystem processes in freshwaters (Hutchinson, 1967, 1993). They are also among the most vulnerable species, and a large number of freshwater crustaceans have gone extinct, primarily from loss of habitat (Dudgeon, 2006; Strayer, 2006; Väinölä et al., 2008; Yeo et al., 2008). The “traditional Crustacea” are divided into six classes (Martin and Davis, 2001; Ahyong et al., 2011): Branchiopoda, Cephalocarida, Remipedia, Maxillopoda, Ostracoda, and Malacostraca. The Branchiopoda are represented in fresh and marine habitats all over the world, and their monophyly is well supported (see review by Olesen, 2007).

TABLE 27.1  Major Taxonomic Groups of Crustaceans within Inland Waters of the World along with Common Names in the United States Taxonomic Group

Common Names

Subphylum Crustacea   Class Branchiopoda

Water fleas, fairy shrimp, etc.

  Class Maxillopoda   Subclass Copepoda

(None or copepods)

  Subclass Branchiura

Fish lice

  Subclass Cirripedia

Barnacles

  Class Ostracoda

Mussel or seed shrimps

  Class Malacostraca   Subclass Eumalacostraca   Order Mysida

Opossum shrimps

  Order Stygiomysida

Opossum shrimps

  Order Thermosbaenacea

(Hot-springs shrimps)

  Order Cumacea

(None or cumaceans)

  Order Tanaidacea

(None or tanaids)

  Order Isopoda

(Aquatic sowbugs)

  Order Amphipoda

Scuds, sideswimmers

  Order Bathynellacea

(None or bathynellaceans)

Remipedia and Cephalocarida are both marine and their monophyly is strongly indicated because of their morphological uniformity and by the few molecular studies conducted (Richter et al., 2009). The phylogenetic status of both Maxillopoda and Ostracoda (Figure 27.1) are still under debate. Several morphological studies have found unifying characters establishing their monophyly as separate classes or with the Ostracoda as part of the Maxillopoda; however, no molecular analysis has supported either theory. Instead, the Maxillopoda appears to be split into several clades, and the Ostracoda may be paraphyletic as well (Richter et al., 2009). The Malacostraca appears to be monophyletic based upon a variety of evidence; however, internal relationships remain disputed (Richter et al., 2009). Some studies directly contradict each other, as in the Peracarida (isopods, amphipods, tanaidaceans, and mysids), which may or may not be polyphyletic (Richter and Scholtz, 2001).

Distribution and Diversity Crustaceans are more diverse in anatomic structure than any group of arthropods, and thrive in a wide array of habitats. Of the 125,530 species identified from freshwaters (as of 2008), approximately 12,000 species are crustaceans (Balian et al., 2008), making them third in diversity to insects (75,874) and vertebrates (18,235). In fact, they may be second in abundance because a higher percentage of extant vertebrates than crustaceans may have been identified. Crustaceans occur in inland waters from the arctic (e.g., fairy shrimp Branchinecta paludosa (Müller, 1877); Lindholm et al., 2012) to Antarctica (e.g., branchiopods, copepods, and ostracods; Pugh et al., 2002), from 5000 m in the Himalayas (most branchiopods, amphipods, and isopods; Mann, 1968) to the sea, and from surface waters to

  Order Decapoda   Suborder Dendrobrachiata Dendrobrachitae shrimp   Suborder Pleocyemata   Superfamily Caridea

Shrimp

  Superfamily Astacidea

Crayfish

  Suborder Brachyura

Crabs

Common names listed in brackets indicate that either no common name is used or one has been recently suggested.

FIGURE 27.1  Ostracod (seed or mussel shrimp) shown against a United States penny. Photograph by M.A. Hill in Thorp & Rogers (2011).

Chapter | 27  Introduction to “Crustacea”

groundwaters. In many aquatic ecosystems, they rival or surpass the insect fauna in numbers of individuals and species when the entire aquatic system (infauna, epifauna, and plankton) is considered. They span almost the entire range of osmotic conditions from fresh to saline and hypersaline waters, although faunas are typically more limited in very soft water streams and lakes.

GENERAL BIOLOGY External Anatomy Crustaceans are more anatomically diverse compared to other Arthropoda (Schram, 1986) and to most of phyla in general. For example, once you have examined any insect or spider, you will have a relatively good picture of the general body form of other insects and arachnids. The crustaceans, however, have evolved a wide diversity of body forms by fusing various segments or developing highly specialized body segments and appendages (Figure 27.2). Crustaceans are basically metameric, protostomate coelomates with a hard exoskeleton and are often divided into three regions, or tagmata: the head (cephalon), thorax, and abdomen. Sometimes, the first two of these regions are combined as a cephalothorax (Figure 27.3). The jointed appendages are biramous and may be present in all three body regions. Mandibles, two

673

pairs of maxillae, and two pairs of antennae are nearly always present.

Exoskeleton The name “Crustacea” is derived from the Latin word for shell and refers to the exoskeleton, which is not a true “shell” as is found among the Mollusca. Crustaceans have a durable, exoskeleton composed of chitin, usually hardened with calcium carbonate. It varies in rigidity among taxa and life-history stages. This exoskeleton consists of a thin, stiffened proteinaceous epicuticle and a thick, multilayered procuticle strengthened in most groups by calcium carbonate. The skeleton attains its maximum thickness and rigidity in the decapods (e.g., crayfish and crabs). Projecting inward are chitinized struts, called apodemes, which serve as sites for internal attachment of the muscles, and lend partial protection to some organs.

Appendages Crustacean appendages are variously modified among taxa for locomotion (walking, swimming), feeding, grooming, respiration, sensory reception, reproduction, and defense. Consequently, the primitive, generally biramous appendages (terminal exopod and endopod) are often modified with additional lateral and medial projections. Two extreme forms are

(a)

(b)

(c)

(d)

FIGURE 27.2  (a) Dorsal view of the Red Swamp Crayfish Procambarus clarkii; (b) Spider Cave Crayfish, Troglocambarus maclanei; (c) a common amphipod Gammarus; and (d) the Eastern Tubebuilding Scud, Apocorophium. Photographs (a-b) courtesy of Gunter Schuster and c-d courtesy of Matt Hill (in Thorp and Rogers, 2011).

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SECTION | VI  Phylum Arthropoda

FIGURE 27.3  Lateral view of a generalized shrimp (Decapoda), with the carapace covering the first two major tagma (head and thorax). From Figure 22.1 in Hobbs and Lodge (2010).

FIGURE 27.4  Representative crustacean appendages: (a) a phyllopod appendage of Anostraca; (b) biramous appendage of Anaspidacea (superorder Syncarida); and (c) uniramous stenopod appendage of the decapod Stenopodidea. Redrawn from McLaughlin (1982).

recognized among adults (Figure 27.4): the lamellar phyllopod appendage (as found among branchiopods) and the branched (the limb has a basal gill), segmented walking leg, or stenopod (typical of crayfish and crabs). The cephalic region contains six basic paired appendages: (1) compound eyes; (2) first antennae, which are biramous in the malacostracans;

(3) second antennae; (4) mandibles; (5) first maxillae; and (6) second maxillae. The second two pairs generally have a sensory function (aiding some taxa in food location and filtering), whereas the last three pairs normally function in food acquisition, handling, or processing. The number of appendages on the thorax and abdomen vary greatly among large

Chapter | 27  Introduction to “Crustacea”

675

(a)

(b)

FIGURE 27.6  Structure of decapod gill branches, as shown in transverse (upper) and lateral (lower) views: (a) dendrobranchiate; (b) phyllobranchiates; and (c) trichobranchiate. From McLaughlin (1982).

mandibles that assist in cutting and shredding this organic matter. Food is typically ground into small, digestible particles by the gastric mill (the posterior end of the cardiac stomach) of the muscular foregut (Schmitz and Scherrey, 1983). Enzymes (primarily from the hepatopancreas) empty into the short midgut to digest and assimilate food. Peristalsis conveys waste through the long hindgut and out the terminal anus.

Circulation and Respiration

FIGURE 27.5  Scanning electron micrographs of crayfish chelae: (a) distal portion of a chela (300×); and (b) plumose sensilla (feathery hairs) on the outer edge of a crayfish chela (6000×). Images by J. L. Borash (1997).

taxonomic groups. Malacostracans (such as decapods and amphipods) generally possess five to eight pairs of thoracic appendages (sometimes called thoracopods or pereiopods) and six pairs of abdominal appendages (pleopods and terminal uropods). Primary abdominal appendages are absent in all non-malacostracans except Anostraca.

Internal Anatomy and Physiology Digestive System Crustaceans predominantly use their antennae, maxillae, or thoracopods to filter, capture, or collect food. Crayfish and crab chelae (“claws”) and the gnathopods of amphipods and isopods can be used to grasp larger food items, and they also serve in defense. The numerous sensilla (sensory setae) on various crustacean appendages may aid in chemically locating food (Figure 27.5). This food is then passed through a ventral mouth into a tripartite alimentary tract. Malacostracans have sharp

Crustaceans have an open circulatory system consisting of a dorsal, neurogenic heart, one or more arteries (in malacostracans only), and several sinuses for returning blood to the heart. Many non-calanoid copepods (and marine ­barnacles) lack a heart altogether. In those cases, fluids are ­circulated as a result of general body movements. The heart is in the thorax or cephalothorax but extends into the abdomen in taxa with abdominal gills (such as most malacostracans). The blood contains low concentrations of a viscous, ­copper-based respiratory pigment (hemocyanin), which is dissolved in the hemolymph. Respiration takes place entirely across the body integument in non-malacostracans, but malacostracans typically use thoracic and abdominal gills. Individual species differ in their rates of oxygen consumption and their ability to regulate their respiration as dissolved oxygen concentrations decrease (Väinölä, 1990). Most freshwater decapods have trichobranchiate gills composed of a central axis (with afferent and efferent blood vessels) bearing numerous filamentous branches (Figure 27.6).

Fluid and Solute Balance The principal excretory organs are paired maxillary and antennal glands (also called green glands) (Figure 27.7).

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FIGURE 27.7  Structure of the antennal gland of a crayfish. From Mantel and Farmer (1983).

The antennal gland’s labyrinth is involved in reabsorption of glucose, amino acids, and divalent ions from tubule fluids. Maxillary glands predominate within lower crustaceans, whereas the slightly more complex antennal glands characterize most malacostracans. More than 90% of the nitrogenous wastes are voided as ammonia. Water influx and salt loss taking place primarily across the gills require the release of copious quantities of urine that is isosmotic to the hemolymph (a few taxa produce urine that is hyposmotic). Freshwater branchiopods and some copepods maintain a dilute hemolymph equal to about 200 mOsm (20% of seawater), while the blood of crayfish is more saline (about 350 mOsm) (Mantel and Farmer, 1983). Hyporegulating crustaceans living in inland salt lakes have the opposite situation. The brine shrimp Artemia franciscana Kellogg, 1906 tolerates salinities ranging from 10% seawater to crystallizing brine. It is hypertonic to the medium in dilute waters but hypotonic in concentrated solutions.

Neural System and Receptors The crustacean neural system is comprised of a tripartite brain and paired ventral, ganglionated nerve cords linked by commissures in a ladder-like arrangement. The protocerebrum of the brain normally innervates the eyes, sinus gland, frontal organs, and head muscles. The deutocerebrum controls the antennules, whereas the tritocerebrum innervates the antennae and a portion of the alimentary tract. Crustaceans are generally sensitive to light, chemicals, temperature, touch, gravity, pressure, and sound. Photoreception Crustaceans have evolved photoreceptors to detect shapes, movement, and slight differences in light intensities,

FIGURE 27.8  Stalked eyes shown in the fiddler crab Uca. Photograph by M.A. Hill in Thorp & Rogers (2011).

especially in deep, thermally stratified waters in some taxa. The eyes of adult malacostracans and branchiopods are compound and frequently mounted on a stalk (= peduncle), as in decapods and anostracans (Figure 27.8). However, the adult eye of ostracods and copepods is a simple cluster of inverse pigment cup ocelli that also comprises the naupliar (larval) eye. The peak sensitivity of the crayfish eye is at 520–525 nm (Kennedy and Bruno, 1961). Crayfish, at least, can detect polarized light and are very sensitive to orange light and much less sensitive to blue light (Muller, 1973). Moreover, some crustaceans, including marine mantis shrimp, are sensitive to ultraviolet wavelengths, with maximum absorption at 345 nm (Cronin et al., 1994). Visual as well as chemical cues are important to many crustaceans, including decapods. For example, the giant Australian crayfish Cherax destructor Clark, 1936, or yabbie, can visually recognize the “faces” of fight opponents and commit them to

Chapter | 27  Introduction to “Crustacea”

memory for at least 24 h and probably much longer (Van der Velden et al., 2008). Subordinate Neohelice granulatus (Dana, 1851) have better memory retention than dominant individuals of this freshwater-to-marine crab (Kaczer et al., 2007).

Chemoreception Many crustaceans have well developed chemosensory systems to help regulate daily and seasonal patterns of movement during periods of stable flow regimes, to select foods or mates, and to avoid predators (Ache, 1982; Atema, 1988; Zimmer-Faust, 1989). Some exhibit specific pheromonal responses (Dahl et al., 1970, 1998; Dunham, 1978; Breithaupt and Thiel, 2011), especially marine species. Studies of freshwater species have found pheromones in some species (Yen et al., 2011), none in others (e.g., Itagaki and Thorp, 1981; Crease and Hebert, 2006), and questionable in the remainder (Caskey et al., 2009). Sex pheromones seem more likely to be present when the female crustacean has a very temporally constrained reproductive period, especially when tied to the molt process (Thorp and Itagaki, 1982). Although the effects of stream flow and lake current hydrodynamics mediate diffuse chemicals effectiveness, the signal’s persistent unidirectional nature may provide a basis for orientation and communication. Crustaceans can sometimes detect potential predators by chemical means. The amphipod Gammarus lacustris Sars, 1863 responds to fish predator alarm signals from northern pike (Esox lucius L.) but not to dragonfly larvae (Aeshna eremita Kirby, 1896), apparently because fish are more effective predators (Wudkevich et al., 1997). Similarly, the isopod Lirceus fontinalis Rafinesque, 1820 reduces activity in the presence of fish mucous alarm substances from predatory green sunfish (Lepomis cyanellus Rafinesque, 1819) but not from grazing fish (Short and Holomuzki, 1992). The importance of chemical communication in minimizing risk of fish predation among decapod prey by changing their behavior and morphology is also well documented among tropical species, such as the shrimp Xiphocaris elongata (Guérin-Méneville, 1855) (Covich et al., 2009) and among atyid shrimp species living in cave streams (Jugovic et al., 2010). Crayfish as benthic consumers are important in nutrient cycling, grazing, and predation. Their predation can alter gastropod prey morphology (Krist, 2002) and affect the life histories of some prey species that have evolved avoidance strategies (Crowl and Covich, 1990; Covich, 2010). Chemical defenses in some autotrophs, such as aquatic mosses (e.g., Fontinalis spp.), can limit crayfish grazing and thereby affect not only plant community composition but also the subsequent spatial refuges for amphipods (e.g., Crangonyx gracilis [S.I. Smith, 1871]) and isopods (Asellus aquaticus L., 1758) (Parker et al., 2007). In other cases, herbivory on chemically defended plants can provide

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a deterrent to predation such as among the amphipod Hyalella (Rowell and Blinn, 2003). Chemical communication is also important in complex life cycles of crustacean hosts and their parasites. Some parasite-infected Gammarus pulex (L., 1758) are chemically attracted to fish predators, that uninfected individuals actively avoid after detecting chemical cues from these fish (Baldauf et al., 2007; Perrot-Minnot et al., 2007). Reversal of chemically mediated predator-avoidance behaviors greatly increases the likelihood of the infected amphipod being consumed by the final host. Gammarus minus (Say, 1818) respond differently to chemicals released from injured conspecific individuals relative to chemical cues released from other species (Wisenden et al., 1999). Finally, chemoreception is effective in two-dimensional orientation within the stream flow boundary layer. Chemical cues may be extremely important for organisms migrating up or downstream, with organism size relative to the boundary layer depth being an important ratio (Dodds, 1990). Mechanoreception The ability to orient into currents (positive rheotaxis) is a common trait in many freshwater crustaceans. These physical, hydrodynamic cues are often associated with chemical cues that stimulate directed movement. Crustaceans have many types of mechanical and chemical sensory setae (Bush and Laverack, 1982). These receptor setae are mostly positioned near the antennae and mandibles, with a few receptors scattered across the body; in this way, potentially vital information can be obtained from several directions simultaneously. The functional anatomy of decapod mechanoreceptors has been extensively explored. The cleaning behavior of freshwater crustaceans has not been as thoroughly studied as in marine taxa nor have there been many studies on how current speed changes behavioral responses. Copepod behavioral studies have emphasized mechanoreceptors for predator avoidance (see review in Chapter 29), but relatively little is known for other groups. Thermoreception Metabolic rates and need for dissolved oxygen increase as temperatures rise. Unfortunately, dissolved oxygen concentrations diminish as temperatures increase and lowoxygen stress can cause mortality. Low-oxygen stress can be avoided to some extent by moving into shallow near shore areas or more turbulent water, but these may have more predators or physical stress, respectively. Deeper lake waters are also cooler, but some may lack sufficient oxygen because of high decomposition rates. Consequently, temperature sensitivity is important attribute for crustaceans, as it allows them to locate optimal microhabitats for growth and reproduction. Many species can orient in thermal gradients and combine information on

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temperature with simultaneous inputs from photo- and chemoreceptors (Ache, 1982). Freshwater crustaceans have distinct thermal preferences (Cheper, 1980; Oberlin and Blinn, 1997), and considerable information is available for crayfish (see Chapter 32). As mentioned previously, the interactions between thermal cues and light intensity are often associated with vertical migrations of zooplankton. Evidence is growing that zooplankton may use chemical cues to orient themselves and to avoid predators (see Chapters 28 and 29).

Reproduction and Development Reproduction Crustaceans are primarily dioecious and reproduce sexually, but hermaphroditism, parthenogenesis, and gonochory occur sporadically among ostracods and branchiopods and in a few marine malacostracans. Paired gonads lie above or lateral to the midgut. In mysid shrimp (Figure 27.9), the ovaries are partially fused and linked by a cellular bridge. Paired, or rarely single, reproductive ducts open ventrally through simple gonopores (paired or single), elevated papillae, or elaborate modified limbs called gonopods (anostracans and most malacostracans). The location of the malacostracan gonopore opens on the sternite or coxae of the sixth thoracic somite in females and at the eighth somite in males, whereas in other groups its position varies. Internal fertilization is the general rule in crustaceans, and females may be polyandrous (mating more than once in the same breeding season). Fertilization may take place immediately, or the spermatozoa may be stored for months, as in many crayfish. Following the initial copulation, a second male may attempt to remove the spermatophores of the first male partner (e.g., in Austropotamobius italicus (Faxon, 1914)) (Galeotti et al., 2007). Females of most species protect their embryos either by retaining them in a brood pouch (e.g., branchiopods) or

FIGURE 27.9  Opossum shrimp, Neomysis. Photograph courtesy of John Pfieffer at EcoAnalyst, Inc.

external ovisac (e.g., copepods) or by gluing them to certain appendages (decapods employ their abdominal pleopods). Mysids, isopods, and amphipods safeguard their young in a ventral brooding shelf, the marsupium, until they reach an advanced stage. Ovigerous amphipods are readily recognizable because of their extended brood pouch and its yellowish to gray coloration. Stream-dwelling isopods apparently release their juveniles from the brood pouch when at risk from fish predation but not when exposed to lower risk of predation by salamanders (Sparkes, 1996).

Development Crustacean embryogeny includes modified spiral, total cleavage, and gastrulation by invagination. All taxa possess an initial nauplius stage in their development, but these larvae may be free swimming (e.g., copepods and most branchiopods) or enclosed in an egg (e.g., malacostracans and cladocerans). Direct development (i.e., without any external larval stages) characterizes taxa such as cladocerans (Figure 27.10), peracardians, and some freshwater decapods. The young appear morphologically similar to adults. In contrast, indirect development with a free nauplius followed by either distinct metamorphosis stages or gradual development to an adult is typical of most ostracods, copepods, decapods (except crayfish), and Branchiopoda (other than cladocerans).

Growth Larval, juvenile, or adult growth requires periodic shedding of the older, smaller exoskeleton in a process termed molting or ecdysis. Rapid body expansion occurs immediately after the old exoskeleton is shed and before the new one hardens. The degree of expansion varies significantly among species, ranging from 8% to 9% in some mysids to 22% in many decapods to as high as 83% in certain cladocera (Hartnoll, 1982). Some taxa have indeterminate growth (e.g., the cladoceran

FIGURE 27.10  Water flea (Daphnia). Photograph by M.A. Hill in Thorp & Rogers (2011).

Chapter | 27  Introduction to “Crustacea”

Daphnia and apparently most crayfish species), whereas others reach a finite body size in a fixed number of molts (e.g., ostracods, copepods, and some isopods). Hormones released by several organs control molting. Molt-inhibiting hormone production by the cephalic X-organ and sinus gland complex decline and the Y-organs secrete the molt-stimulating hormone ecdysone. The actual molt requires as little as a few minutes to as long as several hours. This molt period can be very hazardous because the animal is vulnerable to predators (and cannibalism) and it may die from physiological stress or eventually succumb if it cannot extract its body from the old exoskeleton. The newly molted individual is also vulnerable to rapid colonization by epibionts and ectoparasites. Some species of ciliated protozoans and sessile rotifers are well adapted to quickly recolonize the clean surfaces of newly molted crustaceans (Cook et al., 1998; Roberts and Chubb, 1998). For these reasons, some taxa seek shelter prior to molting. The larger and older an individual is, the longer for the molting process to complete, and the greater the vulnerability to cannibalism or predation. Various environmental factors influence crustacean susceptibility to predators after molting. For example, medium and large crayfish experience lower predation risks from salamanders when living in waters rich in calcium carbonate (tufa and travertine) because lime deposits on their bodies apparently make them less vulnerable (Ruff and Maier, 2000). Each habitat may vary in how environmental complexity influences rates of molting and survivorship, although indeterminate growth typically has limitations imposed by physiological stress, parasite infections, epibiont loads, and predatory pressures.

GENERAL ECOLOGY AND BEHAVIOR Habitat Selection Crustaceans are major players in freshwater planktonic and benthic habitats of lentic and lotic systems and are common groundwater inhabitants in hyporheic waters adjacent to rivers and especially hypogean streams of caves. Copepods, cladocerans, and occasionally mysids comprise the eukaryotic holoplankton of lentic systems in addition to rotifers. Decapod crabs, shrimp, and crayfish are often the largest mobile invertebrates of freshwater benthic habitats, although the smaller ostracods, amphipods, harpacticoid copepods, chydorid cladocerans, and occasional isopods are more abundant than these larger taxa. Small streams support many amphipods and some isopods along with crayfish, shrimp, copepods, and ostracods in temperate latitudes. In tropical streams, large populations of shrimp and crabs replace the crayfish found in temperate habitats. As rivers increase in size, crustacean zooplankton become ecologically important especially in lateral slackwater areas (Casper and Thorp, 2007). With the increase in large

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freshwater fish capable of eating crayfish, shrimp, and crabs, the numbers of these crustaceans often decrease except in more protected shallow slackwaters, or the crustaceans escape some predators by adopting more nocturnal habits. More permanent lakes also typically support abundant populations of large fish, so benthic crustaceans are generally confined to protective vegetated areas of the littoral zone. However, diverse populations may still exist, including the speciose assemblage of amphipods in ancient lakes such as Baikal (Väinölä et al., 2008). Hyporheic and other groundwater habitats contain syncarid and bathynellacean crustaceans along with taxa more common in other habitats such as copepods, ostracods, and isopods. While the biomass of all invertebrates within hypogean streams is low relative to that in surface (epigean) streams, isopods, amphipods, and decapods can be important components of the fauna. Many rare and geographically isolated species of crustaceans are found in karst caverns and other hypogean habitats, some species of which are both blind and unpigmented. Branchiopods, cladocerans, and copepods typically constitute the greatest biomass of invertebrates within ephemeral wetlands (Brendonck et al., 2008; Rogers, 2009). They are adapted to these environments because of their abilities to grow from dormant eggs very rapidly after rains fill the pool, reproduce, and then produce resistant diapausing eggs embryonic stage when the pool dries. In contrast, most peracarids (e.g., isopods and amphipods) have no adaptation for avoiding desiccation. Thus, their chances for passive dispersal (e.g., being carried to new habitats by wind or migratory animals) are low in comparison with branchiopods, copepods, and ostracods (Hairston and Caceres, 1996; Bohonak, 1999). Large branchiopods, such as Artemia salina (L., 1758), occur in great numbers within saline pools and lakes (e.g., Williams, 1998), where they are among only a handful of metazoans able to tolerate the hyperosmotic environment. Planktonic crustaceans exploit multiple, vertical and sometimes lateral habitats within lakes and some rivers (Jack et al., 2006). Freshwater zooplankton exploit thermal gradients by migrating daily and seasonally in both vertical and horizontal patterns to obtain preferred conditions. The adaptive value of this migration may be a combination of: (1) optimizing growth by seeking deeper cool waters where metabolic costs are low; (2) rising into upper waters to feed on algae (some also consume bacterioplankton); and (3) avoiding exposure to visual predators and harmful UV radiation in shallow waters during the day. The relative importance of these variables can change seasonally and shift the exact timing of migration to maximize net energy gain under different conditions. When algal food are scarce, zooplankton may ascend early from deep, cool waters and begin grazing 1–2 h before sunset, thereby gaining access to limited food supplies before competitors, but at a higher metabolic cost and risk of predation (Enright, 1979; ­Gliwicz, 1986). Arctic crustacean zooplankton are exposed

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to continuous daylight during their growing season and do not vertically migrate on a daily basis. However, phototactic reactions and photosensitivity are maintained at approximately the same values as temperate species (Buchanan and Haney, 1980). Evaluation of the adaptive significance of variable migratory responses to different light intensities in the presence of predators remains controversial.

Physiological Constraints Salinity Many crustacean species have adapted to living solely in freshwater habitats, whereas others spend only portions of their lives in these highly variable habitats (Mantel and Farmer, 1983; Vernberg and Vernberg, 1983; Wollheim and Lovvorn, 1995; Bauer, 2011a, b). Species richness in inland waters is highest in fresh water, but some groups have rapidly evolved and colonized hypersaline waters resulting in accelerated rated molecular evolution (Hebert et al., 2002; Jellison et al., 2008). Branchiurans (e.g., fish lice such as Argulus; Figure 27.11) are distributed worldwide in both marine systems and freshwater habitats; some species can tolerate a rapid shift from salt to freshwater and may even switch hosts from marine to non-marine fishes. Among the Peracarida, some species are confined to coastal inland waters and apparently are not completely adapted to many of the relatively unbuffered, dilute waters found farther inland (Sutcliffe, 1971, 1974; Dormaar and Corey, 1978).

FIGURE 27.11  Ventral view of the fish lice Argulus. Photograph courtesy of John Pfieffer at EcoAnalyst, Inc.

Amphipod species differ in their ability to osmoregulate in variable salinities. Some Hyalella are found in saline lakes with salinities up to 25 g/l (Hammer et al., 1990). Other amphipod species are sensitive to direct or indirect effects of increased salinity (Wollheim and Lovvorn, 1996). Salt pollution from various sources can increase hypo-osmotic stress and create adverse effects on amphipod metabolism (Koop and Grieshaber, 2000). Although the current volume is primarily limited to freshwater environments, some crustaceans (especially crabs) found in freshwater streams and some lakes can tolerate estuarine conditions or even full-strength seawater (see Chapter 32). For example, adult mitten crabs (Eriocheir sinensis Milne-Edwards, 1853) migrate to estuaries to breed and then release planktonic larvae (which require full-strength seawater to develop). The juveniles eventually migrate back into freshwaters to grow and live sometimes hundreds of kilometers upstream. The Harris mud crab (Rhithropanopeus harrisii (Gould, 1841)) is moving inland from the Gulf of Mexico into Texas and Oklahoma and is adapting to freshwater habitats of higher ionic content (Boyle et al., 2010; Patton et al., 2010). A number of “marine” crabs (e.g., blue, xanthid, and saber crabs) are increasingly found far upstream in completely freshwaters, although most are more common in estuaries and costal marine habitats.

Temperature Temperature is an important metabolism, growth rate, and survival regulator, and crustaceans have evolved a number of means of either avoiding or tolerating thermal shifts (see review in Lagerspetz and Vainio, 2006). The upper lethal temperatures limiting survival vary greatly among species. Some hot spring crustaceans have the highest known temperature tolerance among aquatic metazoa. The ostracod Potamocypris lives on algal-coated substrates in habitats ranging from 30 to 54 °C (Wickstrom and Castenholz, 1973), and the malacostracan Thermosbaena mirabilis Monod, 1924 survives in waters of 45–48  °C. The bathynellacean Thermobathynella adami Carpart, 1951 lives in 55  °C water (Capart, 1951). Some isopods are found in warm springs of the southwestern United States. The endangered Thermosphaeroma thermophilum (Richardson, 1897) and T. subequalum Cole and Bane, 1978 thrive in 32–35 °C pools (Cole and Bane, 1978; Shuster, 1981). Some species, such as the amphipod Hyallela azteca (Saussure, 1858), have populations with variable thermal tolerances, including those that can reproduce at temperatures ranging from 12 to 40 °C; the densest assemblages were grouped in the 20–25 °C, but others colonized areas of the geyser ponds of Hunter Hot Springs in Oregon, United States (Strong, 1972).

Chapter | 27  Introduction to “Crustacea”

In general, water temperatures affect individual crustacean growth rates directly and indirectly. Water temperatures variations often result in seasonal movements by different aged individuals. For example, field observations demonstrate that 20 °C is a threshold for reproductive resting stage induction and termination of some Hyalella. This genus moves from warmer, shallow littoral zones into colder, deeper water as individuals mature (Panov and McQueen, 1998). If water depths decrease and water temperatures increase in response to both successional and climatic warming, benthic taxa, such as Hyalella, will likely increase their growth rates and breed earlier in shallow lakes, ponds, and intermittent streams. These shifts in water temperatures and habitat fragmentation could affect intra- and interspecific competition and gene flow, especially during prolonged droughts (Hogg and ­Williams, 1996; Covich et al., 1997; Hogg et al., 1998). Other amphipod species require colder, relatively constant waters and are often associated with cold springs. For example, Gammarus minus (Say, 1818) is rarely found in waters warmer than 15 °C and is absent in waters above 20 °C (Culver et al., 1995; Culver and Pipan, 2008).

Hardness and pH Widespread acidification of lakes and streams has drawn attention to the importance of pH tolerances of many organisms, especially crustaceans (Dangles et al., 2004). Because carapace calcification immediately following molting is sensitive to low pH, changes in crustacean distributions may reflect regional levels of acidification and natural differences in calcium concentrations (Nero and Schindler, 1983; Hammer et al., 1990; Meyran, 1998). Both lethal and sublethal effects of increased acidity alter ion regulation, especially by juveniles in soft-water lakes. Although many studies have emphasized lentic species, evidence suggests that lotic species may have a narrower low pH tolerance range. Crustaceans generally are good indicators of changes in pH values in well-defined habitats. A regional study in 32 central Pennsylvania springs documented the limitation of Gammarus minus to waters with a pH of 6 and above, with most populations found in waters near pH 7, with minimal temporal variation (Glazier et al., 1992). Although crayfish are generally thought to require habitats with calcium concentrations in excess of 2 mg/l, some species may survive by extracting sufficient calcium from their food (Zehmer et al., 2002). Calcium uptake in adult Orconectes virilis (Hagen, 1870) is impaired below pH 5, with survival greatly influenced by age and molt stage. The tendency of crayfish to consume their shed exuviae to gain the calcium carbonate increases in soft compared to hard waters (J.H. Thorp, personal observation).

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Feeding Behavior Although opportunistic omnivory is the most common feeding mode in crustaceans in general, some species are primarily algivores (some zooplankton), others are detritivores, and some are mostly predators and scavengers (e.g., benthic decapods, certain copepods, and others). Ontogenetic changes in feeding patterns are common. Filter-feeding by benthic species occurs in some tropical atyid shrimp in Puerto Rican streams (Atya lanipes Holthuis 1963) where they are preyed upon by a large predatory shrimp, Macrobrachium carcinus (L., 1758) (Crowl and Covich, 1994). Predation is a common feeding mode in crustaceans, but it is often a component of a general omnivory habit. Intraspecific predation—or cannibalism—is common among benthic crustaceans, such as decapods, amphipods, and isopods, but is mostly confined to attacks on newly molted individuals and young or hurt individuals (e.g., Dick, 1995; Jormalainen and Shuster, 1997). Detritivores may often consume a mass of somewhat unrecognizable organic material but then selectively assimilate the most nutritious elements, including fungi, bacteria, and small metazoans (e.g., Findlay et al., 1986; Mancinelli et al., 2013). Crayfish will consume vascular plants, although usually not as a primary food source. However, they can wreak havoc on macrophyte stands by disturbing the plants while eating relatively small amounts (Lodge et al., 1994). This reflects the general but not exclusive rule that high cellulose plants are generally considered of low quality food and difficult to digest (Newman, 1991) and may also contain tannins and other chemicals that inhibit herbivory (e.g., Parker et al., 2007). However, a stable isotope, food web study in karst springs has demonstrated that some percarid crustaceans can feed extensively on mosses and vascular macrophytes (Carroll and Thorp, in review). Moreover, some amphipods (Monk, 1977) and mysids (Friesen et al., 1986) have digestive enzymes that hydrolyze cellulose in vitro. In this case, these cellulases were apparently confined to degrading small particles rather than whole cell walls. Microbial breakdown of ingested cellulose appears to be the most common mechanism used by crustaceans, and this would require a complex microorganism community. Given that crustaceans molt periodically and would need to reestablish this microbial community frequently, the cellulase evolution would be an important adaptation. Parasitism is generally uncommon among crustaceans as a group. Exceptions include: (1) specialized copepods and branchiurans that attack fish (Yamaguti, 1963; ­Margolis and Kabata, 1988); (2) bopyrid isopods, such as Probopyrus pandalicola (Packard, 1879), which infest freshwater shrimp (Beck, 1980; Collart, 1990; Roman Contreras, 1996); and (3) and the amphipod Pachyschesis in Lake Baikal, which parasitizes large nektobenthic amphipods (Takhteev, 2000).

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anostracans species are parasitized by microsporidians (Daborn, 1976).

COLLECTING, CULTURING, AND SPECIMEN PREPARATION Collecting

FIGURE 27.12  Murray River crayfish. (Euastacus armatus (Von Martens, 1866)). Photograph by J.H. Thorp.

Predators and Parasites Crustacean predators are primarily fish and/or other crustaceans in most habitats, and these predators plus birds in wetlands and mostly shallow areas of lakes and ponds. Aquatic mammals (e.g., otters, water shrews, and yapoks), terrestrial mammals (e.g., raccoons and crabeating macaques), and some snakes eat decapods. Intraclass through interspecific predation and scavenging is common in many crustaceans, although the former may only be triggered by failure to respond adequately to normal agonistic interactions. During edysis, crustaceans are especially vulnerable to intra- and interspecific predators, which causes many to seek shelter before the molt occurs and until the exoskeleton hardens. Some crustaceans have evolved escape, armor, and/or defensive mechanisms to escape predation from aquatic predators (Figure 27.12), although this is often futile when encountering large fish, wading birds, and other large aquatic and terrestrial vertebrates. Larger crabs and wellarmored decapods can partially outgrow most predation threats, except those from really large aquatic or periodically amphibious vertebrates. Decapods are often nocturnal and avoid visual predators by remaining inactive and undercover during the day. Many benthic amphipods and isopods seek dark microhabitats, such as beneath leaf litter. This normal avoidance behavior can be affected by endoparasites. Infected hosts increase their diurnal activity and their predation vulnerability, thereby providing a greater chance for the parasite to reach its predatory host (Maynard et al., 1998; Médoc et al., 2006). How photoreception, activity cycles, and other behaviors of some crustacean species are altered by specific parasites remain an active area of research in physiology, ecology, and parasitology (­Wellnitz et al., 2003; Bollache et al., 2008; Hernandez and Sukhdeo, 2008). Many anostracans are important intermediate hosts for avian cestode parasites (Sánchez et al., 2013), and several

Tips on collecting crustaceans are provided in Chapters 28–32, but, in general, there are four broad approaches. First, zooplankton can be collected with a fine mesh plankton net (typically >100 μm), or a plankton trap sampler (e.g., Schindler-Patalas box sampler or various vertical water samplers) can be used for more quantitative sampling. Plankton can be obtained by throwing the net out into the water body and then pulling it back, or by using a hose, high velocity pump, and mesh bucket to bring water and animals to investigators in a boat. Second, a benthic grab (e.g., Ponar or Ekman grab samplers) is used to obtain a known amount of material from a soft bottom habitat, and the collected material is then sieved to reveal the captured crustaceans (especially ostracods but sometimes amphipods if the grab is used in vegetated areas). Third, a net can be used to catch crayfish, shrimp, and crabs hiding under rocks, in vegetation, or under a river bank. Nets are also useful for collecting large branchiopods from ephemeral pools. Fourth, a baited trap is effective for collecting crabs and crayfish (the latter with minnow trap baited with canned dog food, chicken wings, or some other meat). Once collected, crustaceans should usually be preserved in ∼75% ethyl alcohol, which should then be replaced the next day for better preservation. Formalin can also be used if the sample is not destined for genetic analysis.

Culturing Crustaceans vary considerably in the ease by which they can be cultured in the laboratory. Many can be raised in batch cultures fairly easily if supplied with the proper food and water conditions. Decapods, especially crabs and crayfish, are more prone to cannibalism and either need separate quarters or at least plenty of individual refuges. Biological supply houses can provide stock cultures of some common species along with instructions for their care and sometimes food. Fairy shrimp and other large branchiopods can also be obtained by hatching their resistant “eggs” after obtaining them from dry ephemeral pools. See the individual crustacean chapters in this book for more detailed information on culturing crustaceans and Chapter 3 for general guidelines for collecting and labeling aquatic invertebrates.

Specimen Preparation See Chapters 28–32 for instructions for preparing specimens in different crustacean orders for preservation.

Chapter | 27  Introduction to “Crustacea”

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Chapter | 27  Introduction to “Crustacea”

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Yen, J., Sehn, J.K., Catton, K., Kramer, A., Sarnelle, O., 2011. Pheromone trail following in three dimensions by the freshwater copepod ­Hesperodiaptomus shoshone. J. Plankton Res. 33, 907–916. Yeo, D.C.J., Ng, P.K.L., Cumberlidge, N., Magalhaes, C., Daniels, S.R., Campos, M.R., 2008. Global diversity of crabs (Crustacea: Decapoda: Brachyura) in freshwater. Hydrobiologia 595, 275–286. Zehmer, J.K., Mahon, S.A., Capelli, G.M., 2002. Calcium as a limiting factor in the distribution of the amphipod Gammarus pseudolimnaeus. Am. Midl. Nat. 148, 350–362. Zimmer-Faust, R.K., 1989. The relationship between chemoreception and foraging behavior in crustaceans. Limnol. Oceanogr. 34, 1367–1374.

Chapter 28

Class Branchiopoda Carla E. Cáceres School of Integrative Biology, University of Illinois at Urbana-Champaign, Urbana, IL, USA

D. Christopher Rogers Kansas Biological Survey and Biodiversity Institute, University of Kansas, Lawrence, KS, USA

Chapter Outline Introduction687 General Phylogenetic Relationships 687 Fossil Record 688 Distribution and Diversity 689 General Biology 690 Morphology690 Anostraca690 Notostraca690 Laevicaudata691 Diplostraca693 Physiology694 Reproduction and Development 696

INTRODUCTION The arthropod group Branchiopoda comprises about 2180 species of small (mostly 0.5-mm to 2-cm) crustaceans with flattened, leaf-like limbs (Dumont and Negrea, 2002; ­Brendonck et al., 2008; Rogers, 2009, 2013; Ahyong et al., 2011). This heterogeneous group of crustaceans includes fairy shrimp (Anostraca), tadpole shrimp (Notostraca), smooth clam shrimp (Laevicaudata), spiny clam shrimp (Spinicaudata), tropical clam shrimp (Cyclestherida), and the various species of “water-fleas” (Cladocera) (Table 28.1). They occur in most aquatic habitats and occupy a central position in aquatic communities, both as important herbivores eating algae and bacteria and as important prey items for vertebrates and invertebrates. Their fossil remains open a window into past climate patterns and lake ecology (Kurek et al., 2011).

General Phylogenetic Relationships There is considerable variation in body form among branchiopods, and multiple hypotheses regarding the evolutionary relationships of these taxa have been proposed.

Life History and Ecology 697 Behavior697 Foraging698 Predators and Parasites 699 Population Regulation 699 Collecting, Culturing, and Specimen Preparation 700 Field Collection 700 Sample Preparation 700 Culture Methods 701 Acknowledgments701 References701

Morphological analyses have traditionally used legs, antennae, and mouth parts for branchiopod classification (Daday, 1910; Dumont and Silva-Briano, 1998; Negrea et al., 1999; Richter, 2004; Richter et al., 2007). Morphological similarities resulted in four orders of Branchiopoda being generally recognized until the mid-1980s: Cladocera, Notostraca, Anostraca, and Conchostraca. Fryer (1987a,b) questioned this classification scheme (echoing concerns of Linder, 1945), and proposed that the branchiopods be classified into eight extant (Anostraca, Spinicaudata, Laevicaudata, Ctenopoda, Anomopoda, Onychopoda, Haplopoda, Notostraca) and two extinct (Lipostraca, Kazacharthra) orders. A combination of molecular (e.g., Braband et al., 2002; Swain and Taylor, 2003; deWaard et al., 2006; Stenderup et al., 2006; Richter et al., 2007; Fritsch et al., 2013) and morphological (Negrea et al., 1999; Olesen, 2000, 2007; Richter, 2004; Richter et al., 2007) analyses have elucidated many of the evolutionary relationships of these morphologically diverse taxa. The most recent phylogenies provide strong support for the notion that the Branchiopoda is monophyletic, as are the Anostraca, Notostraca,

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00028-0 Copyright © 2015 Elsevier Inc. All rights reserved.

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TABLE 28.1  The Families and Present Number of Extant Genera of the Class Branchiopoda Order

Suborder

Anostraca

Family

Number of Genera

Artemiidae

1

Branchinectidae

2

Branchipodidae

6

Chirocephalidae

9

Parartemiidae

1

Streptocephalidae

1

Tanymastigitidae

2

Thamnocephalidae

5

Notostraca

Triopsidae

2

Laevicaudata

Lynceidae

3

Cyzcidae

3

Leptestheriidae

3

Limnadiidae

8

Diplostraca

Spinicaudata

Cyclestheridia Cyclestheriidae Cladocera

1

Daphniidae

8

Moinidae

2

Bosminidae

2

Ilyocryptidae

1

Macrothricidae

10

Neothricidae

2

Acantholeberidae

1

Ophryoxidae

2

Chydoridae

37

Dumontidae

1

Sididae

7

Holopediidae

1

Podonidae

7

Polyphemidae

1

Cercopagidae

2

Leptodoridae

1

Data compiled from: Smirnov (1974, 1992, 1996), Belk (1982), Martin and Belk (1988), Fryer (1988, 1991), Korovchinsky (1992), DeMelo and Hebert (1994), Alonso (1996), Dumont and Silva-Briano (1998), Rivier (1998), Santos-Flores and Dodson (2003), Rogers (2009, 2013), and Rogers et al. (2012)

Laevicaudata, and the Diplostraca (Olesen, 2007, 2009). The Diplostraca is composed of three monophyletic suborders: Spinicaudata, Cyclestherida, and Cladocera (­Olesen, 2007, 2009). The exact affiliations of the Laevicaudata

(as an independent order or as a suborder of the Diplostraca) are still debated (Rogers, 2009; Pessacq et al., 2011; Olesen and Richter, 2013). The term “Conchostraca” has had no taxonomic meaning since the 1980s and should be entirely abandoned.

Fossil Record Kerfoot and Lynch (1987), Schram (1986), and Walossek (1995) provide summaries on the earliest fossils of branchiopods, but revisions to their timelines continue as new discoveries are made. There are branchiopod-like fossils from the Burgess Shale, but these animals are not considered branchiopods (Gould, 1989). Walossek (1995) described a possible branchiopod ancestor, Rehbachiella, from Upper Cambrian sediments of Sweden. Harvey et al. (2012) also reported extensive deposits of branchiopod-type mandibles (analogous to modern anostracans) in Middle to Upper Cambrian marine strata from western Canada. By the Early Devonian (ca. 410 mya), when jawed fish also make their appearance, the Notostraca and the Spinicaudata are established (Tasch, 1969). The fossil taxa Lepidocaris, an anostracan-like animal, and Castricollis, a branchiopod with characteristics of both notostracans and spinicaudatans, are reported from Devonian deposits in Scotland (Walossek, 1995; Fayers and Trewin, 2002). These animals are from sediments that indicate marine and estuarine habitats. Anderson et al. (2004) described what is currently the earliest known cladoceran from Early Devonian sediments. Recent findings by Womack et al. (2012) have expanded the cladoceran record by providing the first record from the Carboniferous. Previously Smirnov (1970) described some possible Daphnia-like and chydorid-like fossils from the Late Permian (ca. 240 mya). Smirnov (1992) also described good specimens of Daphnia-like fossils from the Jurassic (ca. 225 mya). Deposits from the Carboniferous through Cretaceous contain a diverse array of taxa. By the Carboniferous (ca. 286–360 mya), tadpole and clam shrimp species are diverse and occur in many habitats (marine, freshwater, streams) (Tasch, 1969). The Spinicaudata greatly diversified during the Permian and Mesozoic into many families, including those recognized today. The fossil group Kazacharthra, which strongly resemble notostracans, appeared and flourished during the Jurassic (Tasch, 1969). The first recognizable fossil anostracans appear in the Early Cretaceous (Trussova, 1971). Laevicaudata first appear in the Late Cretaceous (Tasch, 1969). Starting with the Oligocene (ca. 34 mya), there are many good fossils of cladocerans, especially of ephippia (the specialized egg case that covers dormant eggs in some cladocera) from lake deposits. Similarly, the current distribution of cladoceran species in North America and Europe suggests that the major bout of speciation was associated with

Chapter | 28  Class Branchiopoda

glaciation, which produced modern species (Brooks, 1957). Fossils of modern anostracan genera appear in the Miocene (Belk and Schram, 2001). In addition to true fossils, cladoceran microfossils have long been used to reconstruct processes ranging from changes in lake productivity to regional climate patterns (Kerfoot, 1974; Bos et al., 1999; Jeppesen et al., 2001; Kurek et al., 2011). Microfossils can also be used to record the recent establishment of invasive species (Sprules et al., 1990), and to document changes in the cladoceran assemblage through time (Hann et al., 1994; Mergeay et al., 2004; Davidson et al., 2007). Even the most sophisticated paleoecological methods cannot directly test the causes of the patterns that they describe. Beginning in the 1980s, increasing attention was devoted to the viable diapausing eggs deposited in sediments (e.g., De Stasio, 1989; Wolf and Carvalho, 1989). The discovery in the 1990s that the eggs of rotifers, copepods, and cladocerans can remain viable in sediment egg banks for decades, if not centuries (Marcus et al., 1994; Hairston et al., 1995; Weider et al., 1997; Cáceres, 1998), brought with it the recognition that these eggs could be used as a tool to experimentally address the processes driving the patterns that they recorded. Kerfoot et al. (1999) coined the term “resurrection ecology” to explain how viable eggs “resurrected” from the sediment egg banks can be used to test questions regarding past mechanisms. Egg banks accumulate dormant progeny from many different years, preserving a record of genotypes and species that were dominant at different times in the past (Hairston, 1996). In the deep water of stratified lakes, the vast majority (>90%) of diapausing eggs fail to receive appropriate hatching cues within the first year and are subsequently buried (Cáceres and Tessier, 2004). When these eggs are hatched and reared in the laboratory, they represent a living link to past populations and communities, thereby providing a way in which to test the ecological and evolutionary changes that have occurred in the past (Weider et al., 1997; Hairston et al., 1999a,b; Cousyn et al., 2001; Decaestecker et al., 2007, 2013; Steiner et al., 2007). Studies of egg bank genetics in Daphnia have indicated that it is possible to extract DNA from individual eggs that may be centuries old, and changes in population genetic structure can be observed in the time span of a few decades (Weider et al., 1997; Reid et al., 2000; Limburg and Weider, 2002).

Distribution and Diversity Branchiopod crustaceans can be found in most freshwater habitat types around the world. There are more than 2000 species described, and new taxa are still being discovered. There are 353 recognized anostracan species (Rogers, 2013), and representatives can be found on every continent and every major island system except New Zealand (Brendonck et al., 2008; Rogers, 2009). Anostracans are

689

generally found in temperate, dry tropical, subpolar, and polar regions, with very few wet tropical species. Roughly half of the anostracan species are narrow range endemics, known from less than 10 localities each (Belk and Brtek, 1995, 1997; Rogers, 2013). The Notostraca comprise 15 species, and the Laevicaudata 36 species, across all major island systems and every continent except Antarctica (Brendonck et al., 2008; ­Rogers, 2009). Both groups range from the Arctic to the tropics and temperate regions of the southern hemisphere. The only two genera of tadpole shrimp, Triops and Lepidurus, do not co-occur as adults due to the former being active in summer and the latter being active mostly in winter and spring (Rogers, 2009). Spinicaudata are most common in temperate regions, although many are found in dry tropics, whereas Cyclestherida predominately occurs in tropical areas, occasionally extending into subtropical areas (Brendonck et al., 2008; Rogers, 2009). There are nearly 180 described Spinicaudata species (Ahyong et al., 2011), although many are probably not valid (Rogers et al., 2012). Similarly, there is one described Cyclestherida species, although there are probably more (Schwentner et al., 2013). The Cladocera may be found from pole to pole, in nearly any part of the world (Adamowicz et al., 2002; ­Bekker et al., 2012; Crease et al., 2012). There are currently an estimated 1070 described species of cladocera (­Adamowicz and Purvis, 2005; Ahyong et al., 2011; Kotov et al., 2013). Our understanding of the distribution and evolutionary relationships of cladocerans depends on how well we can distinguish species. Many taxa were originally assumed to be cosmopolitan, but molecular work often demonstrates strong population subdivision (De Meester et al., 2002; Ishida and Taylor, 2007a,b). Thus, the cladocerans of different continents are often different species (even though many have had, or now have, the same specific names) (Schwenk et al., 2000; Xu et al., 2009; Millette et al., 2011). Some species of chydorid, macrothricid, and sidid cladocerans are termed “meiobenthos” (Frey, 1988) because they live on and in various surfaces, particularly aquatic plants (macrophytes), coarse plant detritus (decaying plant material), and organic bottom sediments. Other species are primarily or totally “planktonic” (inhabiting the open water zone), where their facility for swimming makes them quite independent of surfaces. We know much more about pelagic cladoceran species richness and distribution than we know about the littoral (or meiobenthic) species. The large branchiopods and many cladocerans typically occur in seasonally astatic wetlands and saline lakes, and many rock pool specialists occur in Australia and Africa ­ ogers, (Kerfoot and Lynch, 1987; Brendonck et al., 2008; R 2009; Zofkova and Timms, 2009; Jocque et al., 2010; Timms, 2012). Twenty or so species of anostracans, notostracans, and spinicaudatans are listed as rare, threatened, or endangered at either state or federal levels in the USA,

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690

Brazil, Australia and Europe, and many species are on the Red List of the International Union of Conservation of Nature and Natural Resources. Typical large branchiopod habitat exists on open, level ground, which is often the most desirable for agricultural, urban, or industrial development. The greatest threat to ephemeral wetland invertebrates is the elimination or modification of their habitat.

GENERAL BIOLOGY Morphology Martin (1992) provides a masterful and detailed description of branchiopod morphology, with many drawings and photographs. The four extant orders (Table 28.1) are not closely related and are morphologically diverse (Fryer, 1987a; ­Olesen, 2007). The general morphology of each order is considered separately below.

Anostraca The body structures of anostracans (Figures 28.1–28.3(a)) have been described in great detail in many publications (e.g., Linder, 1941). The order Anostraca (fairy shrimp) have 10, 11, 17, or 19 pairs of thoracic legs along their rather delicate, transparent, and elongate body. Anostracans are typically 1–5 cm long; however Dendrocephalus alachua (Dexter, 1953) and Branchinella nana Timms, 2002 are

usually around 6 mm in length, whereas Branchinella occidentalis (Dakin, 1914) will grow to 10 cm, and Branchinecta gigas Lynch, 1937 and Branchinecta raptor may reach 18 cm (Dakin, 1914; Dexter, 1953; Rogers et al., 2006). The head is free and bears a pair of stalked compound eyes, a pair of small, divided first antennae, and a larger pair of second antennae that are not used in swimming. The female second antennae are undivided, and typically are unadorned. However, the male second antennae typically have two antennomeres, sometimes bearing additional appendages (Figures 28.4 and 28.5); these are used to amplex the female prior to mating. Sometimes cephalic appendages are also part of the premating display (Rogers, 2002). Posterior to the second antennae are mandibles, and the first and second maxillae. The thorax lacks a carapace, and bears the locomotor appendages and gonopods. Most anostracans have 11 pairs of thoracopods, however the arctic Polyartemiella has 17 and Polyartemia has 19, and most females of the ­Australian genus Parartemia have 10. Posterior to the legs are two fused thoracic segments which have the genital structures, comprising modified limbs. The gonads extend into abdominal and thoracic segments. Males have a pair of ventral gonopods (Rogers et al., 2007). Females possess a medial brood pouch. Within the brood pouch are lateral oviducal pouches, a single median ovisac, and shell glands. There are no abdominal limbs, but rather a pair of terminal lobes (cercopods) on the last abdominal segment (telson).

Notostraca

FIGURE 28.1  Eubranchipus bundyi Forbes, 1871 (female above, male below).

FIGURE 28.2  Linderiella occidentalis (Dodds, 1915) (pair in amplexus).

The Notostraca, or tadpole shrimp, have a pair of dorsal compound eyes, and possess a broad and flat carapace that covers the head and thorax (Figures 28.3(b) and 28.6). Adults are 10–58 mm long and have 35–70 pairs of trunk legs partially hidden by their yellow, green, brown, or black carapace. Immature notostracans have a transparent carapace that is partially folded along the dorsal midline and encloses the body and legs. Long filaments (endites) protruding from either side of the carapace are attached to the first pair of thoracic legs. Endites of the second pair of legs are less filamentous, and the succeeding legs lack filaments (Fryer, 1988). The first pair of legs is used in swimming; the succeeding pairs are used for swimming as well as crawling, digging, and manipulating food (Fryer, 1988; Rogers, 2001). The notostracan thorax is defined by the first 11 body segments. Posterior to the 11th segment is the abdomen, however the limbs continue past the genital segments and there are many more limbs than segments. The limbs gradually decrease in size posteriorly. The body ends in a pair of long thin cercopods; and in Lepidurus, a posterior extension of the telson (supra-anal plate). Eggs are carried in brood pouches in the specialized 11th limb pair. The anterior operculum of the brood pouch is derived from the exopodite, and the bottom arises from

Chapter | 28  Class Branchiopoda

691

(a)

(b)

(c)

(d)

(e)

(f)

FIGURE 28.3 (a) Branchinecta raptor (Rogers et al., 2006). Photograph by S. Quinney; (b) Lepidurus bilobatus (Packard, 1883). Photograph by M. Hill; (c) Daphnia pulicaria. Photograph by John Crawford; (d), Cyzicus sp. Photograph by D. Murrow; (e) Lynceus brachyurus Muller, 17776. Photograph by M. Hill; and (f) Cercopagis pengoi (Ostroumov, 1891). Photograph by K. Schnake.

the subapical lobe (Fryer, 1988). In males, the eleventh pair of thoracic legs resembles the 10th and 12th pairs.

Laevicaudata The Laevicaudata (smooth clam shrimp) (Figures 28.3(e), 28.7, and 28.8) share the large, bivalved carapace and the modified male first thoracopods with the Spinicaudata (Figure 28.9) and Cyclestherida (Figure 28.10). However, these characters are convergent between the Laevicaudata and the Diplostraca, and have different origins (Pessacq et al., 2011). The laevicaudatan carapace begins in the early instars as a shield-like carapace, as in the Notostraca, and then splits

to form two separate valves, hinged together in a groove (Tasch, 1969; Olesen, 2000; Dumont and Negrea, 2002). The Laevicaudatan carapace does not develop growth lines (there may be one unspecified exception, reported by Fryer, 1987a), and does not form an umbone (Linder, 1945; Martin et al., 1986; Fryer, 1987a). The Laevicaudatan head is massive compared to the body and may be larger than the body (Linder, 1945; Tasch, 1969; Martin et al., 1986; Fryer, 1987a; Olesen, 2000; Dumont and Negrea, 2002; Richter, 2004). Contrary to Tasch (1969), Pennak (1978), Smith (2001) and Dumont and Negrea (2002), the head as well as the body are encompassed by the carapace. However, the head can be projected outside the carapace and withdrawn

692

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FIGURE 28.6  Lepidurus couesii (Packard, 1875), dorsal view.

FIGURE 28.4  Branchinella ondonaguae (Barnard, 1924), male head, leftside, anterior view. FIGURE 28.7  Lynceus brachyurus (Müller, 1785), dorsoposterior view.

FIGURE 28.8  L. brachyurus left lateral view, left carapace valve removed.

FIGURE 28.5  Streptocephalus moorei Belk, 1973, male head, leftside, anterior view.

again, unlike the Spinicaudata and Cyclestherida. Laevicaudata typically range in size from 2.5 mm (Paralimnetis) to 12 mm (Lynceus). The antennae are small and are not used for swimming. The thoracopods are used for locomotion and feeding. The

segmentation and number of the thoracomeres (10–12) in the Laevicaudata is similar to the Anostraca (Walossek, 1993). Internally, the Laevicaudata have a heart with three ostia, and the vas deferens terminates in the abdomen (Dumont and Negrea, 2002). The sexes are separate, with males having a shorter, blunter rostrum than females, and males having the first one or two thoracopods modified as claspers to amplex the female prior to mating (Martin and Belk, 1988). The female has flanges on the dorsal surface and/or on the carapace inside surface for holding eggs (Martin et al., 1986).

Chapter | 28  Class Branchiopoda

693

FIGURE 28.11  Limnadia lenticularis (Linnaeus, 1761), left lateral view.

FIGURE 28.9  Leptestheria compleximanus (Packard, 1877) right lateral view.

FIGURE 28.12  Ceriodaphnia cf quadrangula (Mueller, 1785) left ­lateral view.

FIGURE 28.10  Cyclestheria hislopi Baird, 1859, left lateral view. Redrawn from Sars, 1886

Diplostraca The Diplostraca contains three suborders: Spinicaudata (Figures 28.9, 28.11, and 28.3(d)), Cyclestherida (Figure 28.10), and Cladocera (Figures 28.12–28.16, 28.3(c), 28.3(f)). In the Diplostracan suborders, the carapace begins as a folded structure, and continues to grow without developing a true hinge (Tasch, 1969; Martin et al., 1986; Walossek, 1993; Olesen and Grygier, 2003). Spinicaudata and Cyclestherida The carapaces of Spinicaudata (spiny clam shrimp) (Figures 28.3(d), 28.9, and 28.11) and Cyclestherida (tropical clam shrimp) (Figure 28.10) closely resemble a clamshell, even having growth lines and umbo. The head has an angular or rounded

FIGURE 28.13  Kurzia longirostris (Daday, 1898) right lateral view.

rostrum, a pair of compound eyes (close-set and partially to completely fused), simple first antennae, and elongated, biramal second antennae that are used for swimming (Linder, 1945; Tasch, 1969; Fryer, 1987a; Dumont and Negrea, 2002; Olesen and Grygier, 2003; Richter, 2004). The head is entirely encompassed by the carapace and cannot be projected outside, although the limbs and abdomen may.

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694

FIGURE 28.14  Macrothrix rosea (Jurine, 1820) left lateral view.

FIGURE 28.15  Leptodora kindti (Focke, 1844) left lateral view.

The body segments number at least 32 (Walossek, 1993); the heart has four ostia; and the vas deferens terminate in the 11th thoracomere (Dumont and Negrea, 2002). Spinicaudata and Cyclestherida swim with their legs down or forward. Adults are 2–16 mm long and have 10–36 pairs of thoracic limbs. Cladocera Most cladocerans are small (80% (Doggett and Porter, 1996; Gries and Güde, 1999; Bittner et al., 2002; Cáceres et al., 2006; Johnson et al., 2006a; Wolinska et al., 2006). However, there is also evidence that many parasites are Daphnia specialists (Green, 1974; Stirnadel and Ebert, 1997; Little and Ebert, 1999) and both intra- and interspecific variation in host resistance has been documented (Little, 2002; Little et al., 2006; Duffy et al., 2008). Given the frequency and high prevalence of infections, and the variation in susceptibility to disease both within and among species, parasites can influence the ecological and evolutionary dynamics of their cladoceran hosts (Ebert, 2005). Parasitism likely interacts with predation and competition in complex ways (Hall et al., 2006). For example, parasitized zooplankton are often at a greater risk of predation (Johnson et al., 2006b); and in some shallow lakes, selective predation may prevent epidemics from occurring (Ebert et al., 1997; Hall et al., 2010). Some parasites also castrate their hosts (­Johnson et al., 2006a; Duncan and Little, 2007), which likely alters competitive outcomes. Daphnia–­parasite interactions have also been used as a model system to investigate general questions of host–parasite coevolution (Ebert, 1994; Decaestecker et al., 2007).

Population Regulation Cladoceran populations, especially of planktonic species, are characterized by hundred-fold annual variations in population size (Rudstam et al., 1993). The peak population generally occurs during an algae bloom. Cladoceran populations are typically at low points during cold weather or when edible algae are scarce. Littoral species show similar population dynamics (Whiteside et al., 1978). Competition for food among cladocerans is well documented (Threlkeld, 1988; DeMott, 1989; Cáceres, 1998). In some cases, this competition is intense enough to result in marked reduction in abundance of other cladocerans and potentially to result in competitive exclusion. Until the work of Hall (1964), it was assumed that the only significant factor driving cladoceran population dynamics was the availability of edible algae. However, Hall showed that although the increase in Daphnia numbers was due to an algal bloom, the decrease was due to predation rather than starvation. Predation can affect dynamics either directly,

700

via consumption (Lampert et al., 1986), or indirectly, via nonconsumptive effects (Pangle et al., 2007) such as diel vertical migration. The population regulatory roles of parasitism and interactions between food limitation and predation are receiving increasing attention (see Lampert, 2011 for review). Planktonic cladocerans such as Daphnia are of great importance to the ecology of lakes. They affect the amount of algae, via grazing and nutrient regeneration, and provide food for juvenile fish (Carpenter, 1985; Sommer, 1989; Kitchell, 1992; Lampert and Sommer, 1997; Lampert, 2011). Hrbácek (1958) demonstrated the importance of certain fish in excluding cladocerans from communities. These ideas were extended to the effects of selective grazing of cladocerans on algae to produce a model of a dynamic food chain, in which the number and abundance of species in a community is the result of the interaction of competition, selective predation, and nutrients. Anostracans, clam shrimp, notostracans, and many cladocerans of temporary habitats typically have one generation each time their habitat fills (Belk and Cole, 1975; Loring et al., 1988; Rogers, 2009). The primary strategy is to make as many resistant eggs as possible in the shortest amount of time, thereby developing and maintaining an egg bank. A fraction of the eggs in the egg bank then hatch when the habitat fills again, generally leaving some eggs unhatched in case the habitat dries before the animals have reached sexual maturity and reproduced (Rogers, 2009). These astatic habitat species typically show rapid growth due to a combination of high temperatures and physiological capabilities specialized for rapid growth (Loring et al., 1988). Population regulation can also be accomplished by physiologic factors (e.g., salinity; Broch, 1988).

COLLECTING, CULTURING, AND SPECIMEN PREPARATION Field Collection Branchiopods can be collected using a net pulled through the water, a sieve through which water is poured, or, for the larger species, a dip net. Several species are also easily collected by collecting mud from a dried habitat, adding water, and rearing the animals that hatch from the resistant eggs—the so called “Sars’ Method” (Rogers and Fugate, 2001; Van Damme and Dumont, 2010). When using a net, smaller species obviously require smaller mesh sizes. Nets are most useful for towing in open water. Weedy habitats can be sampled by a plankton net attached to a handle, and with a wide-mesh (1-cm or so) cover over the opening of the net to exclude large plants. Weedy or muddy habitats can be sampled by collecting water (but not the mud) in a bucket and pouring it through a series of coarse (to exclude weeds and detritus) to fine nets or sieves (Downing, 1984).

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Estimations of population dynamics or productivity require quantitative samples using special gear (Bernardi, 1984; Downing, 1984), a careful experimental design, and statistical analysis of samples (McCauley, 1984). You can make inexpensive filters from a plastic bottle. Cut the bottom from the bottle, remove the center from the cap at the opposite end, and glue fine mesh over the cap hole. When the cap is screwed back onto the bottle, water can be poured into the inverted bottle, and branchiopods will be caught by the mesh. The captured animals can then be washed into a jar of water previously filtered through the mesh.

Sample Preparation Preserve samples with ethanol (Black and Dodson, 2003). Wash the animals off the screen of a small sieve with a wash bottle filled with 95% ethanol, or transfer the sample into a larger volume of 95% ethanol. This will kill the animals rapidly enough so there will be little distortion and will preserve the sample indefinitely. It will be necessary to keep the samples tightly capped and to add 95% ethanol from time to time. Although messy, the addition of a few drops of glycerine will ensure that the sample never dries up. Do not use formalin to preserve zooplankton, because of its shortand long-term toxicity. Large samples will need the ethanol replaced after 24 h and again after 3 days, as the water volume released from the specimen’s bodies may dilute the preservative to the point at which decomposition will occur. Animals trapped in a thin film of water can usually be identified with a dissecting microscope, but it may be necessary to mount them on slides. The best mounting medium is one that is more or less permanent, and that is soluble in water and alcohol solutions so that specimens can be added directly to the medium without tedious dehydration. Hoyer’s solution can be made by dissolving 30 g of ground Gum Arabic in 150 mL of hot distilled water. Add 200 g of chloral hydrate and 20 g of glycerine. Divide the Hoyer’s into two batches. Let one batch sit open in a warm place until it is as thick as honey, and keep the second batch in a closed jar so it remains thin. You can always add more water to thin either batch. Keep at least some of the thick and thin Hoyer’s in screw-top eye dropper bottles for easy use, and be careful to keep the Hoyer’s off the threads of the bottles. To make a slide, use the eye dropper to put a small streak of dilute Hoyer’s (about a quarter of a drop) toward one end of the slide. Arrange the specimens in this streak. It is wise to place four or so of what you think are the same kind of animal on a slide to allow for comparison and viewing of difficult characters. Delicate specimens can be protected from compression by putting a few pieces of broken cover slips around the specimens. Let the Hoyer’s streak dry on a slide warmer or in a warm place (below 100 °C). When

Chapter | 28  Class Branchiopoda

the streak is dry, add a drop or two of the thick Hoyer’s and gently lower a cover glass over the Hoyer’s. Put the slide back into a warm place to dry again. Label the slide with the date and location of the collection. It is important to use thickened Hoyer’s in the last step; thin Hoyer’s will shrink as it dries, and will produce large bubbles in the final preparation. If you use thickened Hoyer’s, small bubbles produced when you lower the coverslip will disappear as the medium dries. These slides are semi-permanent. If the climate is humid, the Hoyer’s will thin and cover glasses will slip off the slide. If the climate is arid, the Hoyer’s will desiccate enough so that the cover glasses pop off the slide. These problems may be avoided by storing the slides horizontally in a climate that has moderate humidity, or you can ring the slide with clear nail polish. An alternative mounting method, especially useful for bosminids, chydorids, and macrothricids, is to use polyvinyl lactophenol stained with lignin pink. Temporary slides can be made using glycerol or glycerine jelly. Best of all for museum specimens is possibly Canada Balsam, although this requires that the specimens be dehydrated through an alcohol series.

Culture Methods To maximize survival, bring the animals into the laboratory and let them slowly equilibrate to the laboratory temperature. Most cold-water anostracans and notostracans must be kept at temperatures similar to their original collection site or they will quickly die. Some cladocera tend to be captured by the surface film, especially if they are initially in cold water with saturated oxygen, because bubbles form on the carapace as the water warms. In extreme cases of mortality, keep the surviving animals in a narrow-mouth bottle filled to the top and plugged with cotton pushed below the water’s surface. Animals will live in filtered pond water for a few days at least, especially if there are fewer than 50 animals per liter. Animals can be transferred to new water using filters and/or pipettes. Cladoceran cultures are temperamental, sometimes persisting for years, and sometimes dying overnight, so regular daily attention is essential. It is also best to maintain multiple cultures. A reliable high-quality water source or artificial media is required. The most common culture medium for Artemia is artificial seawater, although more specialized media may give better viability (Bowen et al., 1985). Many cladocerans can be grown successfully in “COMBO” (Kilham et al., 1998). Do not allow any container of formalin to be opened in the room with cladocera, because they are very sensitive to this compound. The recommended diet varies among the four orders. Anostracans and clam shrimp eat commercial tropical fish food. Notostracans can be fed anostracans or dried cladoceran fish food (Rogers, 2001). Feed cladocerans mixed algae from a fish aquarium or single-species algal cultures

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purchased from biological supply houses or grown in the laboratory. Grow the algae in 2-L clear bottles. For fast growth, aerate the cultures and use bright light, but make sure water temperatures remain moderate. Alternative and less-desirable foods include bacteria from a manure suspension or a yeast suspension. Feed the cultures a food suspension. The optimal culture appears faintly murky but not opaque. As the water in each jar clears, add more food. Even an overnight bout of clear water will cause some clones to starve. It is also possible to overfeed clones, but starvation seems to be a much more common occurrence. Continue adding algae suspension until the jar is full. Change culture jars when full or after excess food has collected on the bottom and sides. Cladocerans tend to grow and reproduce well at room temperature.

ACKNOWLEDGMENTS We are very grateful to Brenda Hann for her careful and detail oriented review of the first draft. This chapter is based on those from earlier additions, originally written by Stanley I. Dodson and David G. Frey. Earlier versions were improved by many reviewers, especially James Thorp, Alan Tessier, Brenda Hann, Denton Belk, Carlos SantosFlores, Henri Dumont, Professor Smirnov, Jon Marcot, Tara Stewart, and John Havel.

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Chapter | 28  Class Branchiopoda

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Chapter 29

Class Maxillopoda Eduardo Suárez-Morales El Colegio de la Frontera Sur, Chetumal, Mexico

Chapter Outline Introduction710 Subclass Copepoda 710 Subclass Branchiura 711 Subclass Thecostraca 711 General Biology of Copepoda 712 External Anatomy 712 Internal Anatomy 717 Digestive System 717 Circulatory System 717 Excretory System 717 Neural System 718 Reproductive System 718 Reproduction and Life History 718 Mating720 General Ecology of Copepoda 720 Adaptations, Behavior, and Feeding 720 Adaptations720 Behavior722 Feeding722 Population Regulation 723 Regulation by Temperature 723 Regulation by pH 724 Regulation by Food Density and Quality 724 Regulation by Heavy Metals 725 Larval Mortality 725 Predator Control 725 Impact of Parasites 725 Diversity, Distributional Patterns, and Conservation 726 Diversity726 Distributional Patterns 727 Conservation728 Importance for Humans 732 Biological Control of Mosquito Larvae 732 Intermediate Hosts of Pathogenic Agents 733 Copepods as Food, Biological Indicators, and Source of Chitin 733

General Biology and Ecology of Branchiura 734 External Anatomy 734 Internal Anatomy 737 Reproduction and Life History 739 Mating739 Life History 739 Diversity and Distribution 740 Host, Infection, and Control 741 Infection741 Control742 General Biology and Ecology of Cirripedia 742 External Anatomy 742 Internal Anatomy 743 Reproduction and Life History 744 Life History 745 Ecology, Distribution, Impacts, and Control 746 Distribution747 Impacts and Control 747 Collecting, Culturing, and Specimen Preparation 748 Copepoda748 Collecting Copepods 748 Culturing Copepods 748 Specimen Preparation of Copepods 748 Argulidae749 Collecting and Culturing Argulids 749 Specimen Preparation of Argulids 750 Cirripedia750 Collecting Barnacles 750 Culturing Barnacles 750 Specimen Preparation of Barnacles 751 Acknowledgement751 References751

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00029-2 Copyright © 2015 Elsevier Inc. All rights reserved.

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INTRODUCTION The crustacean class Maxillopoda (Dahl, 1856) is represented by a diverse array of taxa sharing a combination of characteristics, including (1) short bodies with no more than 11 trunk segments of which 6 are thoracic somites; (2) a reduced abdomen lacking appendages; (3) a naupliar eye with three cups; (4) well-developed mouthparts including a mandibular palp in adults; and (5) filtering-adapted maxillules and maxillae. Nevertheless, because of the diversity of forms contained in this group, it is not possible to single out a key distinguishing character to define it. Results from molecular analysis of different crustacean taxa suggest that the Maxillopoda is a paraphyletic group (Regier et al., 2005). This controversial assemblage of taxa (Martin and Davies, 2001) currently includes the following subclasses: Copepoda, Branchiura, Thecostraca, Pentastomida, Mystacocarida, and Tantulocarida. Only the first three of these subclasses have freshwater representatives. Pentastomids are an intriguing group of 130 species that are parasitic of vertebrates and are presumably linked to the branchiurans (Lavrov et al., 2004); however, they are also considered to be representatives of stem-arthropods because their lack of a naupliar larval stage. Mystacocarids are marine meiobenthic maxillopodans dwelling in interstitial habitats of sandy beaches. The group contains 13 nominal species belonging to two valid genera, Derocheilocaris Pennak and Zinn, 1943 (8 species) and Ctenocheilocharis Renaud-Mornant, 1976 (5 species), both contained in a single family. All known mystacocarids have been recorded from the Atlantic Basin only. They have been linked, on morphological grounds, to the Copepoda, but molecular data suggest that they are closer to the Remipedia and Cephalocarida. Tantulocarids are small, ectoparasitic forms infesting other marine crustaceans (Boxshall and Lincoln, 1983); the anchialine tantulocarid Stygotantulus stocki Boxshall and Huys, 1989 is the smallest crustacean known, with a length 20 Subfamilies, including the following:    Subfamily Cyprinotinae Bronshtein, 1947    Subfamily Eucypridinae Bronshtein, 1947    Subfamily Megalocypridinae Rome, 1965    Subfamily Isocypridinae Hartman and Puri, 1974    Subfamily Mytilocypridinae de Deckker, 1974    Subfamily Callistocypridinae Shornikov, 1980    Subfamily Cypridopsinae Kaufmann, 1900 Family Notodromadidae Kaufmann, 1900    Subfamily Notodromadinae Kaufmann, 1900    Subfamily Indiacypridinae Hartmann and Puri, 1974    Subfamily Oncocypridinae De Deckker, 1979    Subfamily Cyproidinae Hartmann, 1963 Family Ilyocyprididae Kaufmann, 1900 Superfamily Cytheroidea Baird, 1850 Family Cytheridae Baird, 1850 Family Hemicytheridae Puri, 1953 Family Kliellidae Schafer, 1945 Family Limnocytheridae Klie, 1938    Subfamily Limnocytherinae Klie, 1938    Subfamily Timiriaseviinae Mandelstam, 1960 Family Cytherideidae Sars, 1925 Family Loxoconchidae Sars, 1925 Family Leptocytheridae Sars, 1925 Family Xestoleberidae Sars, 1866 Family Entocytheridae Hoff, 1942 Superfamily Darwinuloidea Brady and Robertson, 1885 Family Darwinulidae Brady and Robertson, 1885 Modified from Martens and Horne, 2009, Table 1.

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Cyprididae, Notodromadidae, and Ilyocyprididae. The remaining superfamilies include the Cytheroidea, composed predominantly of marine forms with some freshwater lineages (notably Limnocytheridae), and Darwinuloidea, which is entirely nonmarine. Both Cypridoidea and Darwinuloidea also have some semi-terrestrial representatives. The semi-terrestrial Superfamily Terrestricytheroidea, however, is usually found in marine-influenced environments such as salt marshes and is not considered further in this chapter. The bases of ostracode systematics are the appendage morphology, the characteristics of the reproductive organs, and the carapace morphology. Although dissection of appendages and soft tissue is recommended to securely identify ostracodes to the species level, this is clearly not possible with fossil material. The carapace or separated valves may be the only material available. However, carapace shape and structural or ornamental features of the valves can be used with confidence to identify most taxa to the species level, whereas adductor muscle scar patterns are characteristic of superfamilies. For this reason, it is important, whenever possible, to record and preserve both “soft parts” (e.g., appendages) and “hard parts” (calcified valves) in any taxonomic analysis of ostracodes. By developing well-documented collections of species carapaces and appendages, it will be possible to link such studies to a variety of environmental and biogeographical applications. Up-to-date regional taxonomic keys with illustrations are available for some parts of the world, notably in the work of Meisch (2000), Smith and Delorme (2010) and ­Karanovic (2012). Many monographs exist for particular taxa, and these are referenced in the regional keys.

An Overview of Phylogenetic Relationships At the family level and below, research results that combine descriptive taxonomy (using either living or fossil material) with genetic analysis are still relatively rare but do exist (e.g., Martens et al., 2005). However, at higher levels of classification, several studies have been conducted with differing results. In a summary review by Koenemann et al. (2010), the authors described the current consensus that arthropods are a monophyletic group composed of five clades, with ostracodes included within the Pancrustacea clade (Hexapoda + Crustacea). The Pancrustacea clade itself is composed of several clades, but where the ostracodes fit within these groups has been the subject of considerable research. There has not been much agreement regarding monophyly of the Ostracoda [see Jenner (2010) for a discussion of monophyly vs polyphyly in this group]. Horne et al. (2005) found weak support for ostracode monophyly in an analysis of morphologic characters. As recently as 2010, DNA sequence data from nuclear ribosomal coding genes, nuclear ribosomal 18S rDNA, and

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mitochondrial regions have been the primary source of information for arthropod phylogenetic studies, including Crustacea (Koenemann et al., 2010) and Pancrustacea (Regier et al., 2005, 2008). The phylogenies that Regier et al. (2005) reconstructed with DNA sequence data from three nuclear protein coding genes show that Ostracoda are at the base of Pancrustacea and might be polyphyletic. Regier et al. (2008) confirmed with data from 62 nuclear coding genes that Ostracoda together with Branchiura are a sister group to all other pancrustaceans, but could not conclude whether Myodocopa and Podocopa are indeed different phylogenetic clades. In the same year, Tinn and Oakley (2008) used 18S rDNA data available for ostracode families and noted that fossil estimates and genetic estimates of ostracode lineages are incongruent. They concluded that lineage-specific rate differences may indicate erratic rates of molecular evolution and that the ostracode lineage may not be monophyletic. In a broader study of the arthropod group, Koenemann et al. (2010) constructed phylogenetic trees from 18S rDNA data as well as mitochondrial 16S rDNA and cytochrome C oxidase subunit 1 (COI). The ostracodes did not demonstrate a clear monophyletic relationship in this study. However, advances in technology have provided new opportunities to analyze the crustacean phylogenetic problem in more detailed and far-reaching ways. By using transcriptome (only coding regions) sequence data generated by next-generation sequencing technology from nine ostracode taxa together with available data from expressed sequence tags (ESTs), mitochondria and rDNA of ostracods and other crustaceans, Oakley et al. (2012) demonstrated strong support for ostracode monophyly. This study by Oakley et al. (2012) is the most recent comprehensive analysis on ostracode phylogenetics, and places the Ostracoda in the Oligostraca clade of Pancrustacea. At lower taxonomic levels, DNA sequence data have been used successfully to resolve phylogenetic relationships between closely related species or genera, for example, for ostracods from the ancient lakes Baikal and Tanganyika (Schön and Martens, 2012). For this purpose, mainly ribosomal nuclear regions such as ITS or the mitochondrial COI gene have been sequenced. Molecular data can also be used to test specific phylogeographic hypotheses (Schön, 2007; Brandao et al., 2010; Schön et al., 2010) or to provide relative age estimates (Schön and Martens, 2012). Recently, one of the most surprising results has been the exceptionally high cryptic (or hidden) ostracod diversity, which has been detected by sequencing the mitochondrial COI gene. In the morphospecies Eucypris virens (Jurine, 1820), for example, Bode et al. (2010) identified more than 40 cryptic species from populations in Europe and Northern Africa, and Schön et al. (2012) found up to eight cryptic species in the darwinulid ostracod Penthesilenula brasiliensis (Pinto and Kotzian, 1961) at a global scale. Such cryptic species can mostly not be identified by morphological characters, although close morphological

inspection has provided small but consistent differences, for example, in Australian Bennelongia species (Martens et al., 2012). In any case, the detection of more cryptic or near-cryptic ostracod species will significantly increase current estimates of ostracod diversity, possibly by a factor of 10 or more.

Fossil Record Ostracode valves are commonly fossilized in sedimentary records. The bivalved, calcite carapace of the ostracode is easily preserved in many aquatic sedimentary environments, occasionally forming shell beds (coquinas) of valves. Although soft tissue is rarely preserved, many characteristics of the calcite valves are diagnostic to the species level, although, with long stretches of time, homoplasy becomes an important factor (Horne et al., 2005). The recent analysis by Oakley et al. (2012), which indicates that ostracodes are likely monophyletic, also suggests that the base of the crown Ostracoda lies in the Cambrian Period, some 500 mya. As currently understood, the earliest unequivocal fossil record of ostracodes occurs in Ordovician-aged marine sedimentary rocks (Siveter et al., 2001). In light of these new phylogenetic results, it is probable that the Cambrian records may be reexamined for any possible evidence of fossil podocopids or myodocopids of that age. The fossil record of nonmarine ostracodes begins in the Carboniferous (Maddocks, 1982). An overview of the age and phylogeny of nonmarine ostracodes is presented in Martens (1998), further discussed in Horne (2003), and summarized here (Figure 30.2). Based on the fossil record, the Cytheroidea emerged about 450 mya, whereas the Cypridoidea originated in Devonian time (about 400 mya). The Darwinuloidea also first appeared in the Devonian, around 360 mya. All three lineages originated as marine taxa, with invasion of continental habitats not beginning until the late Paleozoic. The Cypridoidea expanded with an explosive radiation beginning in mid-Jurassic (∼150 mya).

Distribution and Diversity Nonmarine ostracode species can be found from equatorial to polar regions, and include cosmopolitan as well as highly endemic forms. Ostracodes occur in almost all aquatic habitats, including interstitial and surface water bodies of all kinds, (semi-)terrestrial habitats, bromeliad leaf cups, thermal springs, caves, and oxygenated aquifers. It is important to note that they are also found in anthropogenic structures in which rain water or irrigation water can collect, in anything from small buckets or birdbaths to rice paddies and waste-water treatment plants. This remarkable distribution makes them valuable choices for environmental monitoring and paleoenvironmental reconstruction, yet they are often overlooked in environmental assessments. In a recent report

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FIGURE 30.2  Stratigraphic record of key podocopid lineages. Figure modified and redrawn from Martens and Horne, 2009, Figure 4.

of the Freshwater Animal Diversity Assessment (FADA) project, Martens et al. (2008) noted that there are ∼2000 species and about 200 genera of extant, nonmarine ostracodes (Figure 30.3). This is likely to be a conservative count, because significant areas of the globe have yet to be sampled. Roughly 50% of this diversity is accounted for by the family Cyprididae and another 25% by the family Candonidae (­Martens et al., 2008). Endemism plays an important role in nonmarine ostracode diversity. “At a specific level, nearly all families, including the large Cyprididae and Candonidae, have endemicity rates (at the level of zoogeographical Realms) of around 90%, meaning that only about a tenth of all species have intercontinental distributions” Martens et al. (2008, p. 188).

Taxonomic and Biogeographic Databases Several databases exist that provide distribution and diversity data for nonmarine ostracodes (Figure 30.4). At present, these databases are composed primarily of information about northern hemisphere taxa; however, new databases are in development that include southern hemisphere representatives as well. Perhaps the most well-known taxonomic database is the Kempf database housed within the University of Cologne and available in CD and book form (see http://ostracoda-on.tripod.com). This extensive and detailed database is a compilation of taxonomic references, including bibliographies and genus/species indexes, and is an invaluable source of information that

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can be used as a tool in a range of diversity and biogeographical studies. Other types of ostracode databases contain environmental and climate data. (It should be noted here that “ostracode” and “ostracod” are acceptable spellings in use in different countries, and both spellings are apparent in the different database titles.) In the Northern Hemisphere, the North American Nonmarine Ostracode Database (NANODe) provides species distributions and associated hydrochemistry and climate data for about 600 sites in the United States (www.kent.edu/nanode). These data are also available through a larger relational multiproxy database called Neotoma (www.neotomadb.org), which also includes pollen, plant macrofossil, diatom, and fossil vertebrate data for the Pliocene through modern time. A database of Canadian species and US species called the North American Combined Ostracode Database

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(NACODe) is also under development (Curry et al., 2012). NACODe links the Delorme Ostracode Autecological Database (c. 30,000 records from c. 5,000 sites), housed at the Canadian Museum of Nature in Ottawa, Ontario, Canada, with the NANODe database, forming a North American database of nonmarine ostracode distributions with associated hydrochemistry and climate data. The biogeographical database called Nonmarine Ostracod Distribution in Europe (NODE) (Horne et al., 1998a) contains about 10,000 living and 2000 Quaternary fossil records. These and other regional databases are in the process of being linked via the Ostracod Metadatabase of Environmental and Geographical Attributes (OMEGA), which will be made available through the BioFresh global information platform for freshwater biodiversity (Horne et al., 2011). The FADA (see above) database provides the

FIGURE 30.3  Diversity and endemicity of extant, nonmarine ostracodes presented as species/genus numbers, with endemic species/endemic genus numbers in parentheses. PA = Palaearctic, NA = Nearctic, NT = Neotropical, AT = Afrotropical, OL = Oriental, AU = Australasian, PAC = Pacific Oceanic Islands, ANT = Antarctic. Redrawn and modified from Martens et al., 2008, Figure 3.

FIGURE 30.4  Ostracode sampling localities in three different databases: NODE (Europe), NANODe (USA), and the Delorme Ostracode Autecological Database (Canada).

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taxonomic list of living nonmarine Ostracoda of the world on which the assessment by Martens et al. (2008) is based. This list (http://fada.biodiversity.be/group/show/18) is regularly updated.

GENERAL BIOLOGY When first viewing an ostracode under a microscope (­Figure 30.5), the hinged calcite shell (or carapace, which is divided into two valves), may be observed to be as thin and transparent as a curving shard of glass, or opaque and textured with faint or distinct ornamentation. The two valves of the shell are hinged along the dorsal margin and often scattered with and also fringed with tiny sensilla that emerge from pores. The median eye, if present, may be visible as a dark spot that sparkles in reflected light, located on the top of the head of the animal near the front of the dorsal hinge line. One may also see the adductor muscle scars, located laterally on the inner surface of each valve (but often visible as “lucid spots” from the outside). The traces of the paired ovaries or testes may be visible through the wall of the shell. If the observer is examining swimmers, the long, plumose swimming antennules may be visible (the antennules are the first pair of eight pairs of appendages; see below). If the species is benthic, then the antennules may be shorter, thicker, and adapted for crawling and digging in the sediment. The shell may

FIGURE 30.5  Typical view of an ostracode under the microscope. Note the central adductor scars, which are diagnostic of the superfamily. Small white arrow points toward front of animal. Shown is a female Dolerocypris sinensis, a cypridoidean. Scale bar = 1 mm.

FIGURE 30.6  Morphological features of the ostracode shell.

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be smooth or ornamented, pigmented with stripes and splashes of green, brown, black, or occasionally blue, or often merely light beige in color. Here, we give a brief introduction to the shell and appendages, drawing primarily from the detailed review of ostracode biology in Smith and ­Delorme (2010), and we encourage the reader to explore the key monographs on this subject, such as in the work of Meisch (2000).

Shell Morphology Ostracodes are mostly free living except for the Entocytheridae (Hart and Hart, 1974), which are commensal upon other crustaceans. The ostracode body, or visceral mass, is completely encased within a bivalved carapace formed by two flaps (duplicature) of the epidermis within which the bioprecipitation of low-magnesium calcite forms the shell. The only exception among nonmarine ostracodes is the Entocytheridae, which are very weakly calcified. The shell, like the body and appendages, has an outer cover of pseudochitinous epicuticle. When the ostracode molts, the complete exoskeleton of carapace and appendages is shed. Each valve of the shell is divided into two parts, the calcified outer lamella and inner lamella (Figure 30.6). The outer lamella is the major part of the shell, whereas the calcified part of the inner lamella bordering the posterior, ventral, and anterior margin is a mineralized part of the inner layer of the duplicature, strengthening the free marginal region of the shell. It forms a part of the free margin and projects toward the center of the shell. The outer lamellae contain pores through which project sensilla, which are sensitive to touch. Where such pores pass through the fused zone, where the calcified inner and outer lamellae are in contact, they form radial or marginal pore canals. Inward from the free margin, the calcified inner and outer lamellae may be separated in some regions, forming anterior and posterior vestibula. The inner edge of the calcified inner lamella is referred to as the inner margin (Figure 30.6). Ridge structures (selvage and lists) on the surface of the calcified inner lamellae are important in taxonomy down to the species level. Pustules, teeth, or crenulations may appear on the

SECTION | VI  Phylum Arthropoda

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outer margin of the duplicature. The shell may contain additional structures on the outer lamella, such as lateral depressions called sulci. Raised areas variously termed alae, knobs, bosses, papillae, or pustules, depending on their shape, size, and orientation, may also be found on the shell surface. The surface of the shell may also be pitted, punctate, or wrinkled, or may have a reticulate surface. These ancillary structures of the shell are very often used for generic and specific identification. Structures on the internal surface of the shell may also be used in ostracode systematics (Horne et al., 2002). Many ostracode muscles are attached directly to the calcareous shells because the chitinous exoskeleton cannot provide a rigid surface to anchor many of the muscles. The points of attachment of these muscles leave scars as raised or depressed areas on the interior of the shell. For example, the closing or adductor muscles form a distinctive pattern of scars (Figure 30.7) often associated with a pair of mandibular scars, which are attachment points of sclerotized rods that partly support the pivoting dorsal end of the mandibular coxa where it butts against a fulcral point on the inside of the shell. In the Cypridoidea, traces of the ovaries and testes can also be seen laterally in the posterior of the shell (Figure 30.8). Collectively, these scars provide a good diagnostic tool with which the aquatic biologist can determine superfamily and

FIGURE 30.7  Central muscle scar (adductor muscle and mandibular scars) patterns of podocopid ostracodes. (Arrows point toward anterior of carapace): (a) Cytheroidea; note the vertical stack of four to five scars; (b) Darwinuloidea; note the circular rosette of approximately 10–12 segments; (c) Cypridoidea; note elongate scars, openly arranged; (d) Cypridoidea, family Candonidae; note the more tightly arranged “pawprint.” From Smith and Delorme, 2010, Figure 19.3.

gender. Juvenile valves may have a different morphology very different from adult valves, and have, in the past, even been described as different species, even in different genera. De Deckker and Martens (2013) provide extensive examples from the Australian ostracod genus Bennelongia.

Body and Appendage Morphology Appendages The ostracode body and appendages are suspended from the dorsal region in an elongate chitinous pouch. There is no clear distinction between the thorax and abdomen. Instead, the posterior of the body tapers off bluntly and ends in a pair of uropodal rami (commonly known as the furca, although not homologous with the furca of other crustaceans). In the adult form, the head region contains four pairs of appendages that are used for swimming, walking, and feeding. The thoracic region features three pairs of appendages that are used or adapted for feeding, creeping, and cleaning of the shells. Flexible branchial plates on some appendages (at least always the fourth appendage, the maxillula) are used to generate a flow of water through the domiciliar space inside the shell, for respiratory purposes. Ancillary cuticular structures such as setae, claws, and pseudochaetae, found on most limbs, are recognized as important in functional morphology and systematics. Appendages are shown in Figures 30.9–30.11 and are listed, with common terminology,

FIGURE 30.8  Interior views of right valves of two adult Candona elliptica, showing trace scar of (a) ovaries in female and (b) testes in male, as indicated by arrows inside valves. Also visible are adductor scars. Arrows in upper left of photographs indicate anterior direction.

Chapter | 30  Class Ostracoda

765

in Table 30.2. Historically, different terminologies were developed to describe these appendages. Here, we follow the terminology of Meisch (2000). As ostracodes develop and mature, they pass through molt stages, and at each stage they develop additional appendages until they reach the final adult stage and sexual maturity. The general order of appearance of the appendages is listed in Table 30.3, although it must be noted that there are exceptions in several groups (Smith and ­Martens, 2000; Smith and Kamiya, 2003, 2008).

Head Region The head region is composed of the forehead, the upper lip, and the hypostome. The four pairs of appendages in the head region are the antennulae (A1), antennae (A2),

(a)

0.1

0

0.1 0.2 mm. Eye

mandibulae (Md), and maxillulae (Mx1). The antennulae are curved up and backward. When the distal four podomeres of the antennulae have long, plumose setae, it means that the ostracode is a good swimmer (e.g., Bennelongia, Heterocypris, Cypridopsis); if these long setae are absent, as, for example, in the darwinulids (Figure 30.10) and the limnocytherids (­Figure 30.11), the ostracode cannot swim. The antennae extend down and curve backward. These appendages are robustly constructed and have strong claws for walking and climbing. Swimming setae may be present on the first podomere of the endopodite, to aid the swimming action of the antennulae. Ostracodes that swim maintain a sustained motion. The swimming setae of the first antennae used in propelling the animal forward must be in constant motion or the animal will sink to the substrate.

Adductor muscle scar Zenker’s organ Mandible Third thoracic leg Seminal vesicle L7 Hemipenis

antennule A1

Uropodal ramus

Antenna maxillula A2 Branchial plate Prehensile palp of first thoracic leg Second thoracic leg L5 L6

(b)

Third thoracic leg L7 Adductor muscle scar Mandible Ovaries 0.1 0 0.1 0.2 mm. Antennule Eye A1

Maxillula Antenna A2

Branchial plate First thoracic leg L5

Uropodal ramus Second thoracic leg L6

FIGURE 30.9  Sketch of the internal morphology of (a) male and (b) female Candona suburbana Hoff (Candonidae). From Kesling (1951 and modified from Smith and Delorme, 2010, Figure 19.4).

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Egg

Mandible Adductor muscle scars

TABLE 30.2  Appendage Terminology for Podocopida in Common Use Depending on Superfamily (Listed in Order From Front to Back of the Animal)

Eye Antennule A1

Maxillula First thoracic leg Third thoracic leg Branchial plate L7 Second thoracic leg L6 0.2 0.3 mm

Antenna A2 0

0.1

FIGURE 30.10  Sketch of the internal morphology of a female Darwinula stevensoni (Brady and Robertson) (Darwinulidae). From Kesling 1951 and modified from Smith and Delorme, 2010, Figure 19.5.

Mandible Antennule

A1

Eye

Adductor muscle scars Branchial plate

0

0.1

0.2

0.3

Most Common Terminology

Alternate Terminology in Use

1.

A1: antennula

Antennula, or first antenna

2.

A2: antenna

Second antenna

3.

Md: mandibula

Mandibula

4.

Mx1: maxillula

Maxillula, or first maxilla

5.

L5: fifth limb

First thoracic leg (T1), maxilla, second maxilla, or walking leg

6.

L6: sixth limb

Second thoracic leg (T2), or ­walking leg

7.

L7: seventh limb

Third thoracic leg (T3), walking leg, or cleaning leg

8.

uropod (uropodal rami)

Furca, furcal rami, or caudal rami

Hemipenis

Compiled from Meisch (2000) and Martens and Horne (2009).

Thoracic Region Maxillula Antenna

A2

First thoracic leg L5 Third thoracic leg L7 Second thoracic leg

L6

FIGURE 30.11  Sketch of the internal morphology of a male Limnocythere sanctipatricii (Brady and Robertson) (Limnocytheridae). From Kesling (1951 and modified from Smith and Delorme, 2010, Figure 19.6).

Because swimming is energetically quite expensive, adult ostracodes can swim only short distances (usually between adjacent plants). Juveniles in the early ontogenetic stages can travel farther, behaving almost like plankton. In most species, well-developed teeth on the distal ends of the mandible coxae meet in the mouth between the upper lip and the hypostome to grind the food before it is ingested. A small respiratory (branchial) plate extends from the mandibular palp, in some groups reduced to one or two setae. Located ventrally and behind the mouth is a keel-shaped structure called the hypostome. Rake-like organs are situated at the rear of the mouth. They consist of chitin shafts with terminal toothed structures and are used for straining and feeding. The fourth and final set of cephalic appendages is the maxillulae, located on either side of the hypostome posterior of the mandibles, which are made up of one two-segmented palp plus three endites anteriorly and, posteriorly, a large branchial (ventilatory) plate. The maxillulae pass food particles toward the mouth, and the branchial plate aids respiration.

There are three pairs of thoracic appendages. In the Cytheroidea, the first pair of thoracic legs, which constitute the fifth pair of limbs (L5) if one counts from the front of the animal, are robustly developed as legs, with long endopodites bearing terminal claws and used for walking, climbing, and clinging to surfaces. In Cypridoidea and Darwinuloidea, they are reduced, bearing small branchial plates and brush-like anterior endites; the endopodite is reduced to a palp in females but is developed as (mostly asymmetrical) claspers in males. In all three superfamilies, the second pair of thoracic legs (L6) is for locomotion, and each appendage usually terminates in a robust claw. The third pair of thoracic legs (L7) is used for walking in the Cytheroidea and Darwinuloidea, but in the Cypridoidea the L7 is a cleaning limb with terminal setae (with a pincer modification in Cyprididae) used to remove foreign objects from the posterior domiciliar space of the carapace. The commensal entocytherids have modified the L5–L7 appendages for grasping onto the gills of a crayfish host (Hart and Hart, 1974). Paired hemipenes are found in syngamic species behind the third thoracic legs (L7) on the ventral side of the thorax. Paired uterine openings lie behind and inside the vaginal openings of sexual and asexual females. The distal part of the thorax terminates in paired uropodal rami (the furca). In some groups, these are reduced to whip-shaped structures, whereas in others they are well developed and armed at the end with long serrated claws. Meisch (2007) argued that the ostracode caudal rami evolved from uropodal plates and are best termed uropodal rami in the Podocopida, and this terminology is accepted here.

Chapter | 30  Class Ostracoda

767

TABLE 30.3  General Development of Appendages With Each Instar of of a Typical Cypridoidean Ostracode Instar

1

2

3

4

5

6

7

8

9 (Adult)

Antennula A1

+

+

+

+

+

+

+

+

+

Antenna A2

+

+

+

+

+

+

+

+

+

Mandibula Md

−

+

+

+

+

+

+

+

+

Maxillula Mx1

−

+

+

+

+

+

+

+

Uropod

−

−

−

+

+

+

+

+

−

+

+

+

+

+

−

+

+

+

+

−

+

+

+

−

+

+

−

+

Leg L5 Leg L6 Leg L7 Ovaries Testes

Note that exceptions to the above pattern are known (see text). A minus sign (−) indicates the anlage of a structure. From Smith and Delorme, 2010, Table 1.

Gender Identification Sexual dimorphism is common in ostracodes, and can often be quite pronounced in external shell features, as in the limnocytherids and candonids. Gender, in most cases, can be determined visually with a microscope. For the Cypridoidea, this is done by observing the presence or absence of testes and traces of the testes on the inner surface of the shell. Such features can be observed through the closed translucent shell in transmitted light. Upon separating the valves, Zenker’s organs (see section below), hemipenes, and ovaries, and modification of the first thoracic leg into prehensile palps, can be observed. Generally, the male is larger than the female in limnocytherids and candonines but smaller in other groups such as cypridids. In some groups (e.g., the Timiriaseviinae), the female carapace is greatly expanded posteriorly to form a brood pouch for eggs and small juvenile instars. An interesting feature of Cyprididae with mixed reproduction is that there are three genders: male, sexual females, and asexual females. Sexual and asexual females cannot be distinguished morphologically.

Internal Anatomy and Physiology Several extensive descriptions of ostracode anatomy and physiology are available (e.g., Maddocks, 1992; Meisch, 2000), and this section summarizes the discussion of this topic found in Smith and Delorme (2010).

1995). The branchial plates of some appendages move and renew oxygenated water past the inner surfaces of the animal. Large respiratory cells occur in the inner lamella, forming a respiratory epithelium over the valve cavity, which creates a blood sinus (Vannier and Abe, 1995) and has an osmoregulatory role (Keyser, 1990; Yamada et al., 2004).

Digestive System The digestive system of ostracodes consists of the mouth, esophagus, stomach, intestine, rear gut, and anus. The mouth is a large opening with the large teeth of the mandibular coxae on either side. The esophagus is muscular and leads into the stomach. The hepatopancreas lies within the duplicature and empties into the anterior part of the stomach. Most digestion takes place in the stomach, where the food is formed into balls that pass from the stomach through the rear gut for nutrient absorption and then exit the anus as fecal pellets. Symonova (2007) demonstrated two variants in the role of the hepatopancreas in digestive processes and nutrient absorption among the podocopids. In candonids, digestion occurs primarily in the foregut with digestive enzymes provided by the hepatopancreas; in other podocopids studied, digestion occurs farther along in the gut and involves the large secretory cells of the hepatopancreas.

Circulatory and Respiratory Systems

Visual System

The circulatory system of freshwater ostracodes lacks both heart and gills, and thus gaseous exchange takes place across the entire body surface and particularly the uncalcified inner lamella of the duplicature (Keyser, 1990; Vannier and Abe,

The single median eye of the podocopid ostracode is made up of three optic cups (ocelli) and lies in the anterior part of the body just below the rim of the valves. In some groups the lateral ocelli are connected to cuticular

768

lenses, integral with the valves, with the lenses forming anew during ecdysis (Tanaka, 2006).In a recent study of podocopid vision, Tanaka (2006) demonstrated the variation present in the light-gathering ability of the podocopid lens, the thickness of the lens being determined by the structure and thickness of the shell. In a study of the eye in Notodromas monacha (O.F. Müller, 1776), Andersson and Nilsson (1981) showed how the presence of reflecting crystals in the tapetal cells lining the optic cups create a mirror, which can focus an image on the retina. Subterranean-interstitial (hypogean) candonid ostracodes have no eye pigments, and thus are blind; the loss of vision in the hypogean ostracodes is an interesting aspect of the debate regarding the evolutionary history of the dispersal of these taxa in aquifer systems (Danielopol et al., 1994).

Reproduction Broadly speaking, ostracodes develop from eggs and pass through a molting process (ecdysis) involving eight juvenile instars to reach the (ninth) mature adult instar. They reach sexual or asexual adulthood in the final instar within a few months to more than 1 year. McGregor (1969) (Lake Michigan) and Ranta (1979) (Finland) record life cycles of up to 4 years for D ­ arwinula ­stevensoni (Brady and ­Robertson, 1870), but Van Doninck et al. (2003) showed that the same species needs only 1 year to complete a life cycle in more temperate conditions (­Belgium). ­Ecdysis ends in the adult stage. However, there is considerable variability in the details of how reproduction, egg development, and life cycle processes proceed. N ­ onmarine ostracodes reproduce both sexually (­ amphimixis) and asexually (apomictic parthenogenesis). Three ­reproductive modes have been identified in n­ onmarine taxa: fully sexual reproduction, mixed reproduction, and ancient parthenogenesis (Meisch, 2000). Parthenogenesis is not cyclic in ostracods (as it is in Cladocera), and reproductive mode is fixed in sexual and asexual females, although some evidence is found for reversals (sexual females on occasion producing ­asexuals). Sexual reproduction is common. A tally of examined extant North American ostracode species, for example, indicated that 80 species are asexual and 111 are sexual (Bell, 1982; Havel et al., 1990a). On the other hand, in a recent study of nonmarine ostracode species on Pacific Islands, only parthenogens were found (Schabetsberger et al., 2009). Full discussions of ostracode reproductive modes can be found in Martens (1998), Meisch (2000) and Schön et al. (2009). Ostracodes have paired reproductive organs in both sexes, and in many taxa, especially among the Candonidae and the Limnocytheridae, there are gender-specific shell shape and size characteristics. Space accommodations

SECTION | VI  Phylum Arthropoda

within the shell occur in the arrangement of the reproductive organs. In particular, the reproductive organs in males and females take up a lot of space within the shell—up to one-third of the volume of the entire animal (Danielopol et al., 1990a). In females of the Cytheroidea and Darwinuloidea, the gonads are located inside the body; whereas in the Cypridoidean females, the gonads are located within the duplicature. Female Reproductive Organs Females have paired vaginal openings, with oviducts housed in the shell cavity. The ovaries are found within the duplicature and comprise a continuous sequence of gametes. In the adult form, traces of the ovaries may be visible as markings on the inner surface of the calcified outer lamella, as in the Candoninae (Figure 30.8). In cypridoideans at least, the female copulatory organs may be made up of modified appendages from three to five body segments, as was shown by Matzke-Karasz and Martens (2005, 2007), who described remnants of the paired appendages of these segments in two South African genera (five pairs in Liocypris and three in Afrocypris). For sexually reproducing females, the spermatozoa are received into the seminal receptacles next to the ovaries. Male Reproductive Structures The testes are similarly placed in males as the ovaries in females, and traces of them can be seen on the inner and posterior region of the shell, as in the Candoninae ­(Figure 30.8). In some groups, such as the Cypricercinae (e.g., Bradleystrandesia), the testes form a spiral around the inner edge of the shell rather than being confined to the posterior area. The testes change into the vasa deferentia as they leave the epidermis and enter the body cavity. The vas deferens joins up with the ejaculation ducts or, in the case of the Cypridoidea, Zenker’s Organs. (Figure 30.9). These large organs are made up of longitudinal as well as radial muscles in the shape of a tube, on which are found radiating bristles covered in a chitinous sheath. The alternate contraction and expansion of these paired organs pump the sperm into the hemipenes during copulation. Zenker’s organs lie in a nearly horizontal position in the posterior part of the body cavity and are visible through transparent shells in many live specimens. Zenker’s organs are absent in the ­Cytheroidea, where part of the hemipenis itself performs the function of a sperm pump, and in the Darwinulidae, which might indicate that the exceptionally rare males in this latter group are nonfunctional (Smith et al., 2006). The spermatozoa of ostracodes are among the largest produced in the animal kingdom (exceeded only by certain species of Drosophila (Pitnick and Marcow, 1994) and, in many cases, greatly exceed the length of the ostracode carapace (see Matzke-Karasz, 2005 and references therein).

Chapter | 30  Class Ostracoda

For example, Meisch (2000) reported that for Cyclocypris ovum (Jurine, 1820) (carapace length of 0.5–0.7 mm), the sperm length is 6 mm, approximately 10 times the length of the animal! For males, the production and presentation of such large sperm requires extensive coiling of the testes to fit them within the carapace. In transmitted light, the sperm are often visible within the female seminal receptacle and are identifiable by a spiraled coiled line (Matzke-Karasz, 2005). Both hemipenes function during copulation, and multiple sperm are delivered into the female receptacle (see below), as has been observed by Vandekerkhove et al. (2007) in dissections of Eucypris virens (Jurine, 1820). Egg and Brood Care The freshwater ostracode egg is a double-walled spheroid of chitin and calcium carbonate. The space between the two spheroids is occupied by a fluid. These two characteristics of the egg allow it to withstand desiccation and freezing, although not all ostracode species produce desiccationresistant eggs. Most ostracode eggs are white, but some are green (Cypridopsis vidua) or bright orange (Heterocypris incongruens (Ramdohr, 1808), Potamocypris smaragdina Vavra, 1891). Egg size and shape can be quite variable within the same clutch. Dumont et al. (2002) describe at least two types of eggs of H. incongruens in the same clutch. These two types might also have different functionality (e.g., diapausing and subitaneous eggs), suggesting a risk-reduction feature that may or may not be environmentally induced. Eggs are slightly flexible, with a thin (2-μm) chitinous coating, and, in the case of the common Pseudocandona marchica Hartwig, 1899, have an average diameter of about 90 μm (Matzke-Karasz, 2005). Some podocopid lineages exhibit brood care, with females having distinct brood chambers, as seen in Cytheridella ilosvayi Daday, 1905. Internal brooding is always present in the Darwinuloidea and in a few groups of Cytheroidea (among nonmarine groups the Timiriaseviinae and Cyprideis), but is absent in the Cypridoidea. In the cases of brood care, eggs are retained within the posterior portion of the female carapace until hatching and often through the first two or three instars. Far more typical, however, are those species that do not brood, and instead place and glue eggs onto a suitably firm surface, such as vegetation, shells, or sand grains, and then abandon them (Horne et al., 1998a). An intriguing situation occurs in some South African species of the genus Gomphocythere, in which brood care occurs in externally visible brood pouches in the female, but where also diapausing eggs must be formed, as some species occur in fully desiccating temporary rock pools. Brood selection has been identified in parthenogenetic freshwater invertebrates as one process that can reduce the accumulation of damaging mutations in subsequent generations (Lively and Johnson, 1994). What appears to be brood selection in ostracodes, the process of removing

769

defective eggs or embryos from the brood chamber, has been observed in Darwinula (Horne et al., 1998b) and Penthesilenula (Pinto et al., 2007). Horne et al. (1998b) noted that adult Darwinula stevensoni (Brady and Robertson 1890) carry a clutch of about 10 eggs (occasionally more) within the brood chamber, and that some of these eggs hatch earlier than others. The brood chamber is thus occupied by eggs and early instars, the latter of which actively move about. The adult can select, manipulate, and remove certain eggs and also molted valves of instars, while preventing the escape or loss of the active juveniles in the chamber. Pinto et al. (2007) found a similar behavior in Penthesilenula brasiliensis Rosetti and Martens, 1998, and, by culturing the discarded eggs, showed that some of them did hatch, although none reached maturity. Pinto et al. (2007) raised the question of whether this process of removing eggs in Penthesilenula is, in fact, brood selection or bet hedging, in which the adult maximizes the likelihood of producing a continuous supply of viable offspring but is not necessarily aborting defective embryos. Many questions remain to be answered in this interesting area of reproductive success. Is brood selection one way to reduce co-evolved endo-parasites in these parthenogenetic forms, as has been suggested by Horne et al. (1998b)? Is this process observable in syngamic nonmarine ostracode species? What processes govern the timing of egg hatching within the brood chamber? How can the adult recognize the presence of a defective embryo? Are the discarded eggs in fact defective? New approaches to these questions will likely involve detailed molecular studies and culturing experiments.

GENERAL ECOLOGY AND BEHAVIOR Habitat Selection Nonmarine ostracodes are benthic swimmers, or epibenthic or infaunal, or interstitial. They generally show a patchy distribution, and so assumptions cannot easily be made about their population density. Ostracodes can be collected in most aquatic habitats, including springs, streams, lakes, wetlands, rivers, oxygenated aquifers (especially those with fracture flow), hydrated soils, rainforest litter, bromeliad tanks, and human-constructed water impoundments of all sizes, from birdbaths to reservoirs. Different assemblages will be found in these habitats, even within a small spatial area. The mosaic of species assemblages encountered is a reflection of the hydrologic characteristics of the habitat. For example, the ostracode fauna recovered from the hyporheic zone of a stream will have only a small overlap with the fauna from the stream benthos, or with the wetlands and springs fauna associated with that stream. A detailed discussion of collection methods in various aquatic habitats is presented in the next section.

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770

At a greater spatial scale, the distribution of nonmarine ostracodes across continents and islands has produced a seemingly endless theorizing on dispersal and habitat selection. This rich area of research is fueled in part by the tightly interconnected factors of reproductive strategies (parthenogenesis, amphimixis, and characteristics of the ostracode eggs), opportunities for passive dispersal (birds, wind, water), and the range of geologic and anthropogenic landscape transformations. The result of these factors is that nonmarine ostracodes can be found in just about every kind of water body. What is perhaps startling is that, given all the possible ways in which ostracodes can disperse, there are comparatively few species that are cosmopolitan. Martens et al. (2008) reported that about 90% of nonmarine ostracode taxa are regionally or locally endemic. Furthermore, the diversity “hotspots” for nonmarine Ostracoda are rich in endemics and include oxygenated aquifers, ancient lakes (e.g., Baikal and Tanganyika), and shallow temporary pools in areas such southwestern Africa and western Australia (Martens et al., 2008). Such a high number of regionally endemic fauna relative to cosmopolitan fauna suggests that successful dispersal mechanisms are not enough to establish successful populations, and that physiological and environmental constraints account for a significant impact on population success. De Meester et al. (2002) have proposed that populations of aquatic organisms with high potential passive dispersal capabilities but low gene flow are strongly influenced by rapid adaptation in founder events (monopolization hypothesis). Nonmarine ostracode distributions present good test cases for this hypothesis.

Environmental Constraints on Distribution Setting aside biological constraints such as predation and competition, the habitats that prove to be successful for ostracode populations are governed by a few physical and chemical constraints, including pH, oxygen, temperature, salinity, solute composition, hydrology and sediment type (Mesquita-Joanes et al., 2012). In a regional view, these constraints are related to climate and landscape features, with the result that regional endemism, so predominant in ostracode distributions, is at least in part a function of regional climate and landscape. For example, ostracode fauna of the arid northern Great Plains of the United States has little overlap with fauna in the forested Great Lakes region (NANODe, www.kent.edu/NANODe; Curry et al., 2012), although the winds and bird migration pathways connect the two regions. Here, we summarize the physical and chemical constraints that govern ostracode distributions.

semi-terrestrial species can exist in hydrated soils and leaf litter, where no standing water is visible, or among the wet mosses and charophytes of fen wetlands (Smith and Delorme, 2010). However, most ostracodes live in fully aquatic environments. The composition and concentration of major ions in water are important factors determining species distribution (NANODe, www.kent.edu/NANODe; Smith and ­Delorme, 2010). In dilute water below approximately 300 mg/L, bicarbonate, calcium, and magnesium are the most common major ions. With increasing salinity (e.g., in evaporative conditions or solute input from ground water), these ionic concentrations rise until calcite saturation is reached, and calcite precipitates at a total ionic concentration of approximately 300 mg/L. This is the calcite branch point, the first mineral branch point occurring in natural waters (Eugster and Jones, 1979). Beyond that point, the water becomes depleted in calcium and enriched in bicarbonate, or vice versa, resulting in a solute path toward either bicarbonate-enriched, calciumdepleted saline water or bicarbonate-depleted, calciumenriched saline water (Figure 30.12). Ultimately, other mineral branch points (such as gypsum, for example) will be reached, and further changes in major ion composition will occur. Hydrochemically, the fate of a water body in an evaporative setting is determined at the calcite branch point (Eugster and Jones, 1979), although changes in the solute path can occur if hydrochemically different water is added. The major ion composition of water that has passed beyond the calcite branch point, rather than salinity, plays a critical role in determining which species will be present. For example, Carbonel and Peypouquet (1983) and Forester (1983, 1986) noted that different species of Limnocythere could be found in lakes with the same salinity but of different ionic composition. There are boundaries, which Forester identified as hydrochemical ecotones, which define the habitats for these species (Figure 30.12(b)). This observation was confirmed and further developed (Smith, 1993; Curry, 1999). Although Limnocythere remains the most clear-cut example of this hydrochemical partitioning, many other ostracode species respond to these solute differences (Radke et al., 2003). Thus, knowledge of which species respond to different solute paths is very useful in reconstructing hydrochemical changes within lakes. Nonmarine ostracodes can tolerate extremely low salinities, with total concentrations of a few milligrams per liter (www.kent.edu/NANO; Forester et al., 2005) to extremely high concentrations of 170 g/L, well above ocean salinity levels (De Deckker, 1981).

Major Ion Composition and Concentration

Dissolved Oxygen

Ostracodes are aquatic organisms, yet have a range of tolerances in the ionic composition and concentration, and dynamics, of the aqueous environment. Some

The extent, duration, and timing of oxygenated conditions in relation to an ostracode’s life cycle are critical survival requirements. Some lacustrine systems may undergo

Chapter | 30  Class Ostracoda

Alkalinity / Calcium (meq/L)

(a) 100000

771

All NANODe Sites

10000 1000 100 10 1 0.1 0.01

1

10

100

1000

10000 100000 1000000

Total Dissolved Solids (TDS) (mg/L)

Alkalinity/ Calcium (meq/L)

(b) 100000 10000 1000 100 10 1 0.1 0.01

1

10

100

1000

10000 100000 1000000

Total Dissolved Solids (TDS) (mg/L) All NANODe Sites Sites with C. ohioensis Sites with L. ceriotuberosa Sites with L. staplini

Sites with C. elliptica Sites with L. itasca Sites with L. sappaensis

FIGURE 30.12  Major ion hydrochemistry of approximately 600 surface water sites in NANODe (Forester et al., 2005), showing: (a) solute distribution of alkalinity/calcium in meq/L versus increasing concentration as total dissolved solids in mg/L and (b) distribution of selected ostracode species showing preferences for ranges of ionic compositon as well as concentration. From www.kent.edu/NANODe.

seasonal anoxia or oxygen depletion at depth, yet still maintain an ostracode population, because the animal’s life cycle may be short enough to accommodate such a seasonal change. For example, Delorme (1978) found that Candona caudata (Kauffmann, 1900) could survive at depth in Lake Erie, the shallowest of the North American Great Lakes, because its short life cycle allows it to produce eggs prior to seasonal anoxia. Species with long life cycles of 1 year or more, such as Cytherissa lacustris (Sars, 1863) which has a

minimum oxygen requirement of 3 mg/L (Danielopol et al., 1990b; Smith and Delorme, 2010), cannot live in lakes with seasonal anoxia or oxygen depletion. The mean requisite for dissolved oxygen by ostracodes falls within a very narrow margin of 7.3–9.5 mg/L (Smith and Delorme, 2010). Candona subtriangulata Benson and MacDonald, 1963 has the highest minimum oxygen requirement known for North American species of 5.6 mg/L. This is a species that lives today in the northern North American Great Lakes, although not in Lake Erie, where oxygen levels at depth rarely reach that minimum threshold. A high number of species (34 of 43) from Canadian habitats in the Delorme Ostracode Autecological Database in NACODe (Curry et al., 2012) can tolerate low dissolved oxygen concentrations. In aquifers, oxygen is the primary limiting factor in ostracode distributions. An interesting adaptation to occasional and erratic anoxia in South American floodplains occurs among ostracodes and other invertebrates living in the “pleuston” of floating plants such as Eichhornia crassipes (Mart.) Solms, 1883 (e.g., Higuti et al., 2007). Here, normally benthic (swimming as well as nonswimming) animals live in the root systems of such floating plants, which, in general, also capture considerable amounts of sediment. In the channels, rivers, and open and closed lakes of such floodplains, water levels can rise up to 6 m overnight, rendering the sediments anoxic. However, the floating plants just follow the rising water levels, so that the pleuston communities do not suffer from these anoxic events. Oxygen is also related to depth in several large and ancient lakes. For example, Lake Baikal is ca 1700 m deep and oxygenated to its deepest point, and benthic ostracods can occur there. The east African Lake Tanganyika is close to 1500 m at its deepest point, but below 150–200 m the lake is completely anoxic, so there are no truly abyssal ostracods in Lake Tanganyika (Martens, 1997).

Temperature The water temperature of the aquatic habitat plays an important role in the habitat, seasonal, and geographical distribution of ostracodes. The temperatures in shallow water habitats can range from 0 °C to greater than 30 °C, depending on latitude, altitude, and water depth. Species living in this niche must have the ability to tolerate this broad range in one or more life stages. In North America, species such as Candona acutula Delorme, 1967, C. ­candida (O.F. M ­ üller, 1776), C. ohioensis Furtos, 1933, Cyclocypris ampla Furtos, 1933, C. sharpei Furtos, 1933, Cypria ophtalmica (Jurine), 1820, and Limnocythere staplini Gutentag and Benson, 1962 exhibit this range for bottom water temperature. The exceptions to the above are shallow groundwater-fed streams, which may have a nearly constant temperature that approximates mean annual air

772

temperature. The concept of temperature as a limiting factor on ostracode distribution forms the basis of methodologies for using nonmarine ostracodes as paleotemperature proxies (e.g., MOTR in Horne, 2007). Ostracodes show a remarkable adaptation to extremes of the temperature range found in natural waters. Korschelt (1915) experimentally froze C. vidua and Cypria ophtalmica in ice for several hours, and reported that most specimens survived after thawing. At the other temperature extreme, Cypris balnearia (Moniez, 1893) has been collected from thermal springs at temperatures of 45–50.5 °C. Wickstrom and Castenholz (1973) recovered Potamocypris from an algal-bacterial substrate of a hot spring, in Oregon (USA), at temperatures ranging between 30 °C and 54 °C. Chlamydotheca arcuata (Sars, 1901) has been found in warm springs in arid regions of the USA and Mexico, where the water temperature varies between 24 °C and 39 °C (­Forester, 1991), and Thermopsis thermophila Külköylüoğlu, Meisch and Rust, 2003 has been recorded from a hotspring of 40–50 °C in Nevada, USA (Külköylüoǧlu et al., 2003).

Hydrology The hydrology of an aquatic habitat is of primary significance in governing the physical and chemical conditions of the ostracodes’ environment. Few natural aquatic settings are entirely dominated by one water source. Typically, groundwater and surface water combine in some way to create the habitat. This may involve subsurface as well as surface flow, numerous sources of water, and changes in the entire combination on a seasonal, decadal, or longer time scale. Aquatic habitats that are in close geographic proximity may have quite different ostracode fauna, because they are hydrologically different even though they are subject to the same climatic conditions. For example, on a single groundwater flow path, one is likely to encounter some combination of recharge dominated lakes, through-flow lakes, and discharge-dominated lakes (Figure 30.13). Different faunal assemblages will be found in them, because hydrologic characteristics govern the major ion composition and concentration, the flow, and the seasonal or long-term variability of the aquatic system. This relationship of hydrology to ostracode assemblages is also important in understanding the variability in isotope and trace element geochemistry of nonmarine ostracode shells (Ito and ­Forester, 2009; Smith and Palmer, 2012). A very important habitat in which ostracods abound are temporary pools. These can come in a great variety, from large but shallow and turbid pans, to small but deep (>1 m) rockpools on mountainous outcrops. Life in such temporary habitats requires certain biological trait-adaptations, such as the production of desiccation resistant eggs, parthenogenesis (leading to exponential population growth), short life cycles and high fecundity, and others. Most of these traits

SECTION | VI  Phylum Arthropoda

FIGURE 30.13  Groundwater flow as an important influence on a lake’s ostracod assemblage and shell geochemistry record. Lakes within the same climatic and geological regime and that are geographically in close proximity may have different palaeolimnological records due to groundwater flow paths. From Smith and Palmer, 2012, Figure 11.3.

occur in Cyprididae, which constitute half of all living nonmarine ostracods. Indeed, Cyprididae are most common in temporary habitats, and must have radiated in these habitats. Large ostracods, such as the African Megalocypridinae, which easily attain lengths of 3 mm (Megalocypris princeps Sars, 1899, even 7–8 mm, have life cycles of ca 3 months, even in warm climates, and thus require pools with longer hydroperiods (duration of inundation and standing water) to complete their life cycles and to lay eggs for the next aquatic phase of the pool. Also, most large ostracods occur in temporary pools or lakes (Megalocypridinae, Liocypridinae, etc.), including rice paddies, as such habitats tend to be free of predatory fish, a feature that they share with (hyper-) saline lakes in which the giant Australian Mytilocypridinae have radiated. Worldwide, ephemeral water bodies can be found that are high in ostracode species diversity and rich in numbers of endemic taxa (Table 30.4). Species associated with these habitats have been found to produce varying percentages of resting eggs within a single clutch (Rossi et al., 2012). Resting eggs may undergo multiple desiccation events before eventually hatching during a subsequent inundation. Recent controlled experiments indicate that, for a species commonly found in ephemeral water bodies, H. incongruens, the age of the resting eggs is not a significant factor in hatching success (Rossi et al., 2012). Many questions remain to be answered in this area of ostracode research. What conditions or processes are involved in triggering the hatching of resting eggs? Why is ostracode diversity so high in these systems? What kinds of selection pressure govern the production of resting eggs? Because of the limited spatial and temporal nature of ephemeral water bodies, they provide excellent natural laboratories for testing hypotheses concerning ostracode autecology, phylogenetics, and diversity. In all ephemeral systems, hydroperiod, photoperiod, and water depth play important roles. The hydroperiod is strongly correlated with water depth, in that shallow pans or depressions tend to evaporate quickly (Brooks and Hayashi, 2002). Reproductive strategies and population dynamics are

Chapter | 30  Class Ostracoda

773

TABLE 30.4  Examples of Studies of Ostracoda in Temporary Water Bodies Examples of Temporary Water Body Habitat Types and Hydroperiods

Ostracode Genera

Reference

Desert playas: seasonal, weeks

Candona, Cypriconcha, Cypridopsis, Heterocypris, Limnocythere, Physocypria, Potamocypris

Horne, 1996

Vernal pools: seasonal, weeks to months

Bradleystrandesia, Candona, Cypris, Eucypris, Heterocypris, Limnocythere

King et al., 1996

Arctic polygon ponds: seasonal, weeks

Candona, Fabaeformiscandona, Cyclocypris, Cypria

Schneider et al., 2012

Prairie potholes: seasonal, weeks to months

Candona, Cypriconcha, Cypridopsis, Heterocypris

Smith, 1993

Bromeliad tanks: seasonal, months

Elpidium, Candonopsis s.l.

Jocque et al., 2013

Rice paddies: human-induced, months

Bradleystrandesia, Candona, Chlamydotheca,Chrissia, Cypretta, Cypridopsis, Cypris,Dolerocypris, Isocypris, Fabaeformiscandona, Herpetocypris, Heterocypris, Ilyocypris, Limnocythere, Stenocypris, Tanycypris, Trajancypris

Rossi et al., 2003

High-elevation temporary water bodies: seasonal, weeks

Eucypris, Ilyocypris, Heterocypris, Limnocythere

Cuzminsky et al., 2005

Semi-desert, temporary pools: weeks to months

Heterocypris, Ilyocypris, Tonnacypris, Eucyprinotus, Potamocypris

Eitam et al., 2004

strongly influenced by hydroperiod. For example, Martins et al. (2008) examined the effect of hydroperiod variability on the reproductive strategies and population success of Eucypris virens, a geographic parthenogen. They showed that, in conditions of variable hydroperiod, eggs produced by syngamic Eucypris virens were more likely to hatch than those produced by the parthenogenetic form. However, in conditions of stable hydroperiod, the parthenogenetic forms tended to hatch earlier and simultaneously, a successful strategy that outperformed the syngamic population. The photoperiod (light phase of a light/dark cycle) can change substantially with season, and is also very important, particularly with latitude. Other factors, including features of the water chemistry such as specific conductance, temperature, pH, and alkalinity are also important, as is the substrate (King et al., 1996). Also, as ostracode populations rise in a small temporary pool, metabolites of the adults accumulate and may trigger chemical cues to the resting eggs (Rossi et al., 2012). Nonmarine ostracodes have evolved adaptations that allow them to inhabit ephemeral water bodies, including drought-resistant eggs and torpidity, which also facilitate passive dispersal via winds, water and animals, especially birds (Brochet et al., 2010). In tropical ephemeral systems such as in bromeliad tanks, phoresy is observed to be a significant method of dispersal. Ostracodes of the genus Elpidium have been observed clinging to the skin of tree frogs as these animals make their way from one bromeliad plant to another: the ostracode pinches a bit of skin between the

tightly closed valves, and is transported by phoretic dispersal (Sabagh et al., 2011). Some widely distributed ostracode species have been shown to pass through the gut of waterfowl alive. Eggs have also been shown to be viable in the same circumstances. Cosmopolitan species that accomplish this feat include C. vidua (probably one of the most widespread species of nonmarine ostracode), H. incongruens, and species of Physocypria (Proctor, 1964; Proctor et al., 1967). A discussion of these kinds of dispersal mechanisms can be found in Mesquita-Joanes et al. (2012).

Feeding Behavior The diet of most ostracodes is restricted to algae (phytophilic) and organic detritus. Strayer (1985) observed that diatoms are a significant part of the diet for Cypria turneri Hoff, 1942. Grant et al. (1983) reported that Heterocypris carolinensis (Ferguson, 1958) feeds on the blue-green algae Nostoc. Campbell (1995) calculated that tree pollen made up 8% and organic material 42% of the diet for Australocypris insularis (Chapman, 1966). Roca et al. (1993) used Chara with alterable amounts of periphyton covering the stems as a food source for ostracodes in culture. When feeding, ostracodes sweep organic particles into the mouth using the maxillulae. The mandibular teeth grind large organic particles into a smaller size before they enter the mouth. Although ostracodes are predominantly herbivores and detritivores, a few have shown carnivorous characteristics (for a detailed summary, see Wilkinson et al., 2007)).

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Ostracodes have been observed attacking and eating the soft tissue of certain snails (Sohn and Kornicker, 1975). Some species (Cypridopsis hartwigi G.W. Müller, 1900; C. vidua, Cypretta kawatai Sohn and Kornicker, 1972, and H. incongruens) are known from experiments, by the above authors, to eat the soft tissue of snails, such as the mantle and antennae, and to crawl into the respiratory organs. Yousif et al. (2013) recently demonstrated the controlling effect on the transmission of the blood fluke Schistosoma mansoni Sambon, 1907 by the predatory carnivory of Candonocypris novaezelandiae (Baird, 1843). Candonocypris novaezelandiae consumed the eggs and juvenile forms of the gastropod Biomphalaria alexandrina (Ehrenberg, 1831), the intermediate host for the parasite S. mansoni. In addition, C. novaezelandiae also consumed the free-living larval stages (miracidia and cercariae) of S. mansoni. Heterocypris incongruens, Cypretta kawatai, and to a lesser extent, C. vidua, have also been shown to prey upon the gastropod Biomphalaria glabrata Say, 1818, another intermediate host for S. mansoni (Sohn and Kornicker, 1975). Havel (1993) used Paramecium sp. as a source of food for Eucypris sp. in experiments on effects of area and patchiness on species richness. Campbell (1995) provided detailed information on the preying habits of Australocypris insularis. These large ostracodes feed on smaller ostracodes such as Diacypris compacta (Herbst, 1958), as well as on copepods and other microinvertebrates. This large species was also found grouped around drowned bee carcasses, presumably feeding on them, with terrestrial invertebrate detritus making up 8% and insects 3% of the diet. Rossi et al. (2011) recorded predation and cannibalism in populations of H. incongruens. In that study, a small population of parthenogenetic H. incongruens was released into a pond and monitored for more than 1 year. H. incongruens attacked chironomid and mosquito larvae, as well as each other. Several ostracod groups are known to be active filter feeders, and the long setae of the mandibular palps in Darwinulidae seem to form a functional “basket” to capture food particles during filter feeding. Other ostracods, such as species of Bennelongia, have been observed appearing to filter particle rich water, but do not show such morphological adaptations in the soft parts.

Predators and Parasites Ostracodes and their eggs are preyed upon in several ways: they are consumed rather indiscriminately along with aquatic plants and sediment by waterfowl; they are sought after and consumed by fish; and they fall prey to the attacks of other crustaceans, especially amphipods, and the aquatic larvae of insects such as dytiscid beetles. Some species have been observed to avoid predation attempts by retreating into subaquatic vegetation. In a controlled study by Roca et al. (1993), C. vidua was cultured in tanks containing

the macrophytic alga Chara. When water retrieved from a fish tank containing juvenile cyprinid fish was added to the culture, C. vidua was observed to respond to the chemical cues of the presence of fish by retreating into the clumps of Chara. Although numerous studies exist documenting predation by fish, waterfowl, amphibians, dipterans, coleopterans, and odonotans, few quantitative data are available (for a summary of these studies, see Mesquita-Joanes et al., 2012). Ostracodes can be hosts for commensal, in some cases parasitic organisms, including bacteria, ciliates, fungi, nematodes, copepods, and isopods. Data available on these relationships are typically linked to the studies of pathogens, although very little is known about the nature of the ostracode–pathogen interactions (Mesquita-Joanes et al., ­ 2012). Ostracodes are observed to be the intermediate hosts for a variety of parasites. For example, Mytilocypris ­henricae Chapman, 1966 is thought to be the intermediate host for cercaria of trematodes, which have waterfowl as an end-host (Martens et al., 1985), and Potamocypris unicaudata Schafer, 1943 is the obligatory crustacean intermediate host for a fungal parasite that is pathogenic for several species of mosquito larvae (Whisler et al., 2009). Nonmarine ostracodes, with the exception of the Entocytheridae family, which are commensal on crayfish, are not parasites or commensals of other taxa (Aguilar-Alberola et al., 2012). Peritrich ciliates are occasionally observed covering the carapaces of nonmarine ostracodes, although these are ­epibionts rather than parasites (Griffiths and Evans, 1994). One important aspect of parasitism in crustaceans is the effect on reproduction. Some internal parasites, such as the bacteria Wolbachia, can induce parthenogenesis and selectively kill male crustacean hosts through intracellular infection. Such internal parasitism has long been hypothesized to contribute to geographic parthenogenesis. A recent study by Bruvo et al. (2011) on the effects of Wolbachia examined natural asexual and sexual populations of the geographic parthenogen Eucypris virens in western and central Europe. They found no evidence of Wolbachia infection among E. virens, suggesting that other mechanisms (or possibly, other intracellular parasites) may be involved in the evolutionary dynamics of geographic parthenogenesis for this species.

COLLECTING, CULTURING, AND SPECIMEN PREPARATION Field Collection Ostracodes are not typically caught in a plankton tow or trap, and over the years this has led to the incorrect notion that ostracodes are a rare component of the aquatic habitat. They do not move vertically near the surface of open water, nor do they typically swim or float in open water in the

Chapter | 30  Class Ostracoda

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middle of a lake, and thus most plankton traps will not collect them. The exception to this is in the occurrence of early juvenile stages of Cyprididae, which may appear in plankton traps. They generally show a patchy distribution, and so assumptions cannot easily be made about their population density. Depending on the habitat (lentic, lotic, or interstitial), different sampling strategies are required. Here, we summarize collection methods for different habitats, drawing on the discussion in Smith and Delorme (2010).

Wetlands Ostracodes can be collected from hydrated soils, on mosses or charophytes, and among the roots of cattails and bullrushes in fen (alkaline dominant) wetlands (Havel et al., 2000), even if there is no standing water present. These can be collected easily by collecting sediment and plant samples, or, if the sediment is slightly submerged, by using a D-frame net or hand-net with an attached collecting bottle. The mesh openings of the net should be between 100 and 150 μm (Figure 30.14).

Springs Surber samplers and D-frame nets with attached collecting bottles are most useful in collecting ostracodes from rheocrene (flowing) springs. It should be noted that springs

(a)

often occur within a lacustrine setting, and these sites can frequently be identified along the lake shore by examining the vegetation. Stands of bullrushes or charophyte thickets are indicators of groundwater discharge, and a temperature probe is a simple method of confirming groundwater seepage into the lake. Often, after heavy rains, springs linked to shallow unconfined groundwater may show higher rates of flow, and these are opportunities to collect individuals that have been flushed out of the sediment matrix by higher flow velocities.

Lakes Within lakes, ostracodes are most commonly found in the littoral zone, in the sediment, and in periphytic (around and amidst submerged aquatic vegetation). However, in large, deep, oxygenated lakes, populations of benthic species can be found at considerable depths. The Ekman and Ponar grabs or Hongve sampler are most useful for deep lake benthos sampling (Figure 30.14), whereas the D-frame hand-net is most useful in the littoral zone. For otherwise unaccessible sites, the Bola pipe, fitted with a screen or a net, is useful (Figure 30.14), because it can be swung out a considerable distance and drawn back in to shore, collecting a sampling of the benthic taxa. However, going physically into the lake with waders and a D-frame hand net still gives higher numbers of specimens in the sample. For small, shallow ponds, and temporary pools or puddles, a sampling device based on a common entomologist exhauster can be used that will not disturb the sediments (Viehberg, 2002). For very large and deep lakes, oceanographic equipment such as a Reineck boxcorer is advisable.

Aquifers

(g) (c)

(f)

(d) (e)

(b) FIGURE 30.14  Field equipment for collecting nonmarine ostracodes: (a) D-frame net with collecting jar; (b) Bola pipe with net (can also be made with a screen); (c) Hongve sediment sampler; (d) ordinary turkey baster for siphoning water; (e) Ekman grab for sediment sampling; (f) minipiezometer for shallow groundwater prospecting; (g) Bou-Rouch pump for sampling the hyporheos of streams. Image from Smith and Delorme, 2010, Figure 19.11.

An aquifer can be sampled by pumping water into a sieve directly from wells. Remember, however, that aquifer ostracodes are small (5.5 and calcium concentrations >2.5 mg/l (Olden et al., 2006). Life cycles are similar, and all three achieve their greatest abundance in lakes with abundant cobble substrate in the littoral zone (Lodge and Hill, 1994). All three species prefer cobble substrate over unvegetated sand/mud or vegetated sand, but both virile and northern clearwater crayfish are excluded from their preferred habitats by rusty crayfish (Hill and Lodge, 1994). In the absence of predatory fish, competition for food between rusty crayfish and the other congeners results in virile crayfish suffering decreased growth and northern clearwater crayfish experiencing increased mortality compared to allopatric situations (Hill and Lodge, 1999). Both competition for shelter and competition for food interact strongly with predation. In the presence of predatory fish, crayfish of any species that are either without shelter or of a small body size are vulnerable to attack (DiDonato and Lodge, 1993). Because the rusty crayfish displaces both O. propinquus and O. virilis from shelter and has a larger maximum size than both congeners (Hill et al., 1993), it has increased advantages in the presence of predatory fish (Hill and Lodge, 1999). Rusty crayfish also suffer lower sublethal effects from the presence of predators than do the two congeners (Hill and Lodge, 1995). Thus, increasing the abundance of predatory fish is likely to decrease the absolute

D

E

F

FIGURE 32.35 (a) Orconectes virilis; (b) O. propinquus; and (c) O. rusticus, three crayfish species for which ecological interactions and hybridization (between O. propinquus and O. rusticus) have been well studied in the Upper Midwest (Wisconsin and Michigan). Photographs courtesy of Chris Lukhaup.

abundance of all three species, but is likely to increase the relative abundance of rusty crayfish (Hill and Lodge, 1999; Garvey et al., 2003). In addition to the ecological advantages enjoyed by rusty crayfish over O. propinquus and O. virilis, the rusty crayfish has another advantage with respect to northern clearwater crayfish. Rusty crayfish hybridize with O. propinquus but not with O. virilis (Perry et al.,

Chapter | 32  Class Malacostraca, Order Decapoda

2001a,b). Hybrids result primarily from mating between rusty crayfish females and O. propinquus males (Perry et al., 2001a,b). Hybrids are competitively superior to both parental species in competition for food (Perry et al., 2001a,b). Hybridization increases the rate of displacement of O. propinquus by rusty crayfish by 5–36%, although the introgressed genes of O. propinquus are likely to persist in the resulting crayfish populations that consist of individuals that are morphologically and ecologically more like rusty crayfish (Perry et al., 2001a,b). When rusty crayfish have been introduced into lakes where O. virilis, O. propinquus, or both are established, the rusty crayfish displace both species. This is consistent with the ecological superiority of rusty crayfish with respect to both O. virilis and O. propinquus, and with the genetic superiority of rusty crayfish over O. propinquus (Olsen et al., 1991). In most lakes, both O. virilis and O. propinquus become functionally extirpated within a few years following the establishment of rusty crayfish (Wilson et al., 2004; Olden et al., 2006). The results have also been dramatic for the rest of the food webs in these lakes. As the rusty crayfish becomes abundant in lakes, the slow-moving, large macroinvertebrates, especially snails, are either reduced or eliminated, and macrophytes (especially the submersed species) are also either reduced or eliminated by both herbivory and by nonconsumptive destruction (Lodge et al., 1994, 1998; Rosenthal et al., 2005; Peters et al., 2008). In addition, reductions in the population size of snails and other grazers often produces increases in algae growing on cobbles, other abiotic substrates, and macrophytes (Lodge et al., 1994). Populations of some bottom-nesting fish, such as bluegill and pumpkinseed, are reduced by crayfish that eat fish eggs (Wilson et al., 2004). However, the complex, potentially reciprocal relationships between rusty crayfish and sunfish, and between rusty crayfish and other fish, require more research attention (Roth et al., 2007; Bampfylde et al., 2009). The changes that rusty crayfish cause in lakes (especially reductions in sunfish populations), have economic consequences that make it beneficial to invest in the prevention of the spread of rusty crayfish to other lakes, and it would be cost-effective to develop methods to reduce the abundance of nuisance crayfish populations in general (Keller et al., 2008).

Other Case Studies of Crayfish Impacts The impact of invasive North American crayfish on aquatic communities and ecosystems has been quantified by studies of rusty crayfish in streams in the United States (Kuhlmann et al., 2008); of red swamp crayfish (P. clarkii) in invaded parts of North America (Lodge et al., 2000; Rudnick and Resh, 2005), Europe (Lodge et al., 2000; Gherardi, 2006; Gherardi and Acquistapace, 2007), and Kenya (Rosenthal

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et al., 2005; Lodge et al., 2005; Foster and Harper, 2006b); and of spinycheek crayfish (Orconectes limosus) in Europe (Kozák et al., 2007), and signal crayfish (P. leniusculus) in Scandinavia (Lodge et al., 2000; Nyström, 2002). These case studies illustrate the central role that crayfish play in aquatic communities, and the importance of the threat to other crayfish posed by invasions by some crayfish species.

COLLECTING, CULTURING, AND SPECIMEN PREPARATION Collecting and Culturing Shrimp Freshwater shrimp occupy both lotic and lentic environments and are most readily collected with the aid of a sturdy, long-handled, fine-meshed dip net or a small-meshed seine. When a particular locality is choked with vegetation or if the water is greater than 1.5 m deep, shrimp can be caught in funnel minnow traps (fine-meshed cylindrical wire traps with an inverted wire mesh cone on each end) baited with meat (fish or mammal). In clear to low-turbidity waters, shrimp may be collected at night with the aid of a headlight and dip net (the ruby-colored reflection of their eyes makes for easy location of individuals). An entertaining account of several methods of collecting shrimp and crab in freshwater is presented in Chace and Hobbs (1969, pp. 45–47). Shrimp can be raised in much the same manner as described below for crayfish. However, shrimp require more vegetation, deeper water, and well-aerated tanks, and females with eggs should be isolated. When the young shrimp hatch, they remain attached to the pleopods of the female for up to several weeks, after which they leave the mother. The female should then be separated from her offspring in order to prevent parental cannibalism. As the young undergo molting and growth, some sibling cannibalism will probably occur. This can be reduced significantly by providing cover for the molting young or by dividing them among several containers.

Crayfish Methods for collecting crayfish are similar, wherever they are found in the world. For small habitats like ditches, mudbottomed pools, and in thick littoral vegetation, a sturdy dip net is useful. This type of net (delta net preferred) can also be used to collect individuals from rocky substrata in lakes and streams. Hand collecting by turning over shelter rocks is very effective in shallow streams that have little vegetation. Snorkeling can improve collection rates dramatically where the waters are sufficiently deep, and a small-meshed seine net (2–5 m long) is also very useful. Deep sections or pooled areas of streams, as well as shallow ponds and the littoral zone of lakes, may be sampled by pulling a seine

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net (5–8 m length) through the water, taking care to ensure that the weighted margin of the seine remains in contact with the substratum, and that crayfish are disturbed from their refuges. Electroshocking can facilitate the collection of crayfish by using seines. Baited wire traps are more effective than seines in vegetation-choked habitats and when sampling deep waters of lakes. Funnel minnow traps (as described above for shrimps) with enlarged openings (4–5 cm in diameter) are very effective. Catches are much higher at night. A residence time of one to two nights, using meat as bait (e.g., chicken and fish scraps, beef liver, canned cat food, and so on), is a productive approach to this sampling technique. Traps are much less effective in flowing waters. Collecting semiterrestrial burrowing crayfish requires strategies different from those described above. The diameter of the burrow tube and size of the chimney pellets provide good indicators of the size of the crayfish inhabiting the burrow (Hobbs and Jass, 1988), with the smaller openings containing juveniles and the wider ones adults. Note, however, that the adults of some burrowing crayfish and their burrow diameters are quite small (e.g., Cambarellus puer (swamp dwarf crayfish), Faxonella clypeata, and P. pygmaeus (Christmas tree crayfish)). Capturing burrowing crayfish can be fruitful on warm, humid, especially rainy, nights when they often leave their burrows to forage (Norrocky, 1984). A small aquarium net is generally adequate to capture stygobiotic shrimps and crayfish in subterranean streams. These decapods are generally white or translucent and very visible. Hand-grabbing is also a fairly productive means of collecting them once they have been disturbed and are swimming in open water. Funnel minnow traps can also be used, but must be checked often to avoid overexploiting a population. Cave populations are usually small, maximum population growth rates are very low, and therefore care must be taken not to collect too many individuals from any cave. When conducting genetic studies of stygobiotic (or any other species), removal of a pereiopod (walking leg) is usually sufficient to provide an adequate tissue sample. The crayfish can then be released immediately and will regenerate the appendage. All the methods described above can provide qualitative data, for example, to document the presence of a species. If employed appropriately, some of the same methods, along with additional methods, can provide quantitative data on relative, if not absolute, abundance. Relative abundance measures that can be compared between habitats can be derived from the methods described above if the applications are standardized for sampling effort (e.g., by time or area covered). Trapping can be easily standardized, and its biases are well quantified in some circumstances (Steucheli, 1991). Crayfish traps may tend to oversample large individuals and males, but habitats with water clear enough for good visibility, then snorkeling or scuba diving to hand collect all crayfish in a quadrat or other defined area, provides

the least-biased population estimates (Charlebois and Lamberti, 1996). For mark–recapture population estimates or studies of individual movements, various marking and tracking techniques have been used. Crayfish are aggressive, antisocial, and cannibalistic, so rearing and maintaining more than a small number can be problematic. On the other hand, crayfish are scavengers and detritus feeders and will feed on almost anything organic. One system is to maintain each crayfish in a single container (e.g., a 20 × 8-cm stackable glass bowl) kept at room temperature. No aeration is required, provided that the water does not become fouled with food or wastes. The bowls should have sand or small-grain gravel substrata and/or a shelter, and the water should be sufficiently deep to cover the crayfish, but does not need to be deeper. If multiple crayfish are kept in a single (larger) container, it is essential to provide a shelter per crayfish not only because they are territorial and will attack other crayfish in their space, but also because they like to hide during the day and emerge at nightfall. Shelters can consist of rocks, flowerpots, bricks with holes, or sections of PVC pipe. Individuals should be fed two to three times per week, and the water should be changed approximately every other week. Food can range from aquatic plants (Potamogeton or Elodea), dry fish food, scraps of meat, and earthworms to dried cat food. Aquaria should be completely covered with no holes in the tank lid because crayfish will try to escape by climbing up the airline tubing, the filter, or even the plants. If they do escape, most species of crayfish can survive for several hours out of the water. In some parts of Louisiana and other southern states in the United States, crayfish production is economically important. Large, drainable ponds with water depths of 1–1.5 m are often used. Considerable research has been directed toward increasing the yields of farmed decapods such as the red swamp crayfish (P. clarkii) (Avault, 1993; Daniels et al., 1994). In various countries in Europe, both native European crayfish and introduced North American crayfish are valued as food. In China, P. clarkii is cultured extensively and exported to the United States and elsewhere. North American crayfish species with known or potential economic importance in North America include: P. clarkii and P. zonangulus, the burrowers Cambarus diogenes (devil crawfish) and F. devastator, and the stream- and lakedwelling O. rusticus. For additional information on crayfish aquaculture, see Avault (1993) and Huner (1995).

Crabs The collection of freshwater crabs can be problematic because these animals are nocturnal and secretive, but they can usually be captured during the day from rivers and streams by turning over flat rocks and vegetation. Ng (1988) recommends positioning a net from bank-to-bank downstream of the collection site to catch any crabs that may escape in the turbid water

Chapter | 32  Class Malacostraca, Order Decapoda

created when the rocks are turned. Semiterrestrial species living in burrows can be dug from their holes that can be up to a meter deep. Pitfall traps (consisting of a wide-mouthed, deep container placed in a hole so that the rim is level with the ground) can be used to catch semiterrestrial burrowing species that only forage at night. A small amount of glycerol (rather than the preservative ethanol) placed in the bottom of the container of the pitfall trap is recommended because glycerol does not cause captive crabs to lose their limbs when struggling to escape. Crabs living in large, fast-flowing rivers and in large lakes are best caught using baited traps like those described above for crayfish, or by using larger traps with larger funnel openings, depending on the size of crabs to be collected. Dip nets and seines may also be useful in some situations. In the more arid parts of the tropics and subtropics where freshwater crabs are found, the seasonal rainfall patterns bring about seasonal changes in surface waters; in the wet season, crabs can be collected by hand from wetlands, streams, and rivers, whereas during the long dry season, crabs can be dug from their burrows. In the rain forests, crabs can be caught by hand in streams or in baited basket traps set overnight in rivers and larger water bodies. Other tree-living semiterrestrial species of freshwater crabs that live in primary in rain forests in Africa (e.g., G. macropus, Potamonautes raybouldi) can be collected from holes in trees where they live when they emerge to feed (Cumberlidge, 1991, 1999; Bayliss, 2002).

Preservation of Shrimps, Crayfish, and Crabs After collection, shrimps, crayfish, and crabs should be preserved in 70–95% ethanol. Unlike formalin, ethanol keeps decapod joints flexible and (importantly) preserves DNA. If epizooites (e.g., protozoans, copepods, entocytherids, branchiobdellids) are to be saved, the solution in which the decapods were killed should be filtered, and crayfish exoskeletons should be rinsed off and the rinse water poured through a fine seive. Any symbionts should be preserved in 70–75% ethanol and carefully labeled. Shrimps, crayfish, and crabs should be stored in clamp-top glass jars with gaskets. The organisms should be placed in the jar head-down with adequate ethanol preservative. The specimens should be accompanied by a 100% rag paper label on which collection data are clearly printed using insoluble black ink.

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SECTION | VI  Phylum Arthropoda

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Chapter | 32  Class Malacostraca, Order Decapoda

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SECTION | VI  Phylum Arthropoda

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Chapter | 32  Class Malacostraca, Order Decapoda

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Wheatly, M.G., de Souza, S.C.R., Hart, M.K., 1996. Related changes in hemolymph acid-base status, electrolytes, and ecdysone in intermolt crayfish (Procambarus clarkii) at 23°C during extracellular acidosis induced by exposure to air, hyperoxia, or acid. J. Crust. Biol. 16, 267–277. Wiens, A.W., Armitage, K.B., 1961. The oxygen consumption of the crayfish Orconectes immunis and Orconectes nais in response to temperature and to oxygen saturation. Physiol. Zool. 34, 39–54. Willman, E.J., Hill, A.M., Lodge, D.M., 1994. Response of three crayfish congeners (Orconectes spp.) to odors of fish carrion and live predatory fish. Am. Midl. Nat. 132, 44–51. Wilson, K.A., Magnuson, J.J., Lodge, D.M., Hill, A.M., Kratz, T.K., Perry, W.L., Willis, T.V., 2004. A long-term rusty crayfish (Orconectes rusticus) invasion: dispersal patterns and community change in a north temperate lake. Can. J. Fish. Aquatic Sci. 61, 2255–2266. Wowor, D., Muthu, V., Meier, R., Balke, M., Cai, Y., Ng, P.K.L., 2009. Evolution of life history traits in Asian freshwater prawns of the genus Macrobrachium (Crustacea: Decapoda: Palaemonidae) based on multilocus molecular phylogenetic analysis. Mol. Phylogenet. Evol. 52, 340–350. Wu, H.X., Xue, J.Z., Cumberlidge, N., 2010. An extra embryonic phase in the true freshwater crab Sinopotamon yangtsekiense Bott, 1967 (Decapoda, Potamidae). Chin. J. Oceanol. Limol. 28, 725–730. Xue, J., Liu, Y., Cumberlidge, N., Wu, H.X., 2010. First report of developmental changes inside the eggs of the Chinese freshwater crab, Sinopotamon yangtsekiense Bott, 1967 (Potamoidea, Potamidae), with comments on its evolutionary significance. Contri. Zool. 79, 79–84. Yeo, D.C.J., Ng, P.K.L., 1996. A new species of freshwater snapping shrimp, Alpheus cyanoteles (Decapoda: Caridea: Alpheidae) from Peninsular Malaysia and a redescription of Alpheus paludicola Kemp, 1915. Raffles Bull. Zool. 44, 37–63. Yeo, D.C.J., Shih, H.-T., Meier, R., Ng, P.K.L., 2007. Phylogeny and biogeography of the freshwater crab genus Johora (Crustacea: Brachyura: Potamidae) from the Malay Peninsula, and the origins of its insular fauna. Zool. Scr. 36, 255–269. Yeo, D.C.J., Ng, P.K.L., Cumberlidge, N., Magalhães, C., Daniels, S.R., Campos, M.R., 2008. Global diversity of crabs (Crustacea: Decapoda: Brachyura) in freshwater. Hydrobiologia 575, 275–286. Zakšek, V., Sket, B., Trontelj, P., 2007. Phylogeny of the cave shrimp Troglocaris: evidence of a young connection between Balkans and Caucasus. Mol. Phylogenet. Evol. 42, 223–235. 435. Zulandt, T., Zulandt-Schneider, R.A., Moore, P.A., 2008. Observing agonistic interactions alters subsequent fighting dynamics in the crayfish, Orconectes rusticus. Anim. Behav. 75, 13–20.

Chapter 33

Hexapoda—Introduction to Insects and Collembola James H. Thorp Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, Lawrence, KS, USA

Brian J. O’Neill Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, Lawrence, KS, USA

Chapter Outline Introduction to Hexapoda 849 General Introduction to Hexapoda 849 Phylogenetic Relationships 850 Introduction to the Class Insecta 851 Evolutionary Diversification of Aquatic Insects 851 Additional Literature Sources 851 General Biology of Aquatic Insects 851 Tagmatization852 Head852 Thorax852 Abdomen852 Respiration and Circulation 852 Respiratory System 852 Circulatory System 853 Excretion and Osmotic Balance 854 Reproduction and Development 855 Generation Time 855 Metamorphosis855 General Ecology and Behavior of Aquatic Insects 855 Habitat Selection 856 Ephemeral versus Permanent 856 Oceans and Saline Lakes versus Freshwater Habitats 857 Lotic versus Lentic 857 Planktonic versus Benthic 859 Surface, Littoral Zone, and Deep-Water Habitats 860 Infauna versus Epifauna in Littoral Areas 861 Epigean versus Hypogean 861

INTRODUCTION TO HEXAPODA General Introduction to Hexapoda Over 1 million species of hexapods have been described— or approximately 70% of all described species—and this may represent only 10% at best of the actual biodiversity (Berenbaum, 2009). Fewer than 10,000 of these hexapod

Physiological Constraints 861 Osmotic and Ionic Constraints 861 Thermal Maxima and Minima 862 Acidity/Alkalinity862 Oxygen Tension 863 Feeding Behavior 863 Predators and Parasites 864 Introduction to Collembola 864 Introduction864 Phylogeny and Species Diversity 865 General Biology of Collembola 865 External Anatomy 865 Reproduction and Development 866 General Ecology of Collembola 867 Habitat867 Physiological Adaptations and Constraints 867 Feeding Behavior 868 Predators and Parasites 868 Collecting, Culturing, and Specimen Preparation of Insects and Springtails 868 Collecting and Culturing 868 Collecting868 Culturing868 Specimen Preservation and Preparation 869 Acknowledgments869 References869

species are not insects. The group dates back at least to the protoinsects (e.g., Rhyniognatha hirsti Tillyard 1928) of 400 mya in the early Devonian period. Since that time, they have come to dominate the terrestrial environment and be important ecological players in inland water ecosystems. However, only a few groups have managed to invade marine environments, and those are primarily limited to estuaries and ocean margins.

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00033-4 Copyright © 2015 Elsevier Inc. All rights reserved.

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Hexapods have exploited all inland water ecosystems including ephemeral wetlands to the deepest lakes, intermittent streams to great rivers, subterranean streams to interstitial, hyporheic groundwaters, saline lakes to thermal pools, and phytotelmata from wetlands (e.g., pools within pitcher plants) to high in-forest canopies (e.g., in bromeliads). Some species are only marginally members of aquatic systems (the surface-dwelling springtails), whereas others are wholly aquatic as larvae or in all life stages. They play valuable roles in these ecosystems as detritivores, herbivores, and predators, as well as food web prey items for other invertebrates and aquatic and terrestrial vertebrates. The diagnostic characteristics of this major taxon within the phylum Arthropoda include: (1) three pairs of jointed legs in at least one life stage; and (2) fusion of adjacent segments into more-or-less three body tagmata (head, thorax, abdomen). The head features a gnathal region with mandibles, maxillae, and labium; eyes are usually present. The thorax contains three segments, each bearing a pair of jointed legs; the thorax of extant species may contain up to eight podomeres. The abdomen in a primitive state contains 11 segments, a telson or homologous structure, and in some cases, weak legs. All winged arthropods are hexapods (most living species of insects in some life stage), but some groups of hexapods lack wings in all life stages (e.g., Collembola). This chapter has been designed with three components: (1) broad-based coverage of Hexapoda as a whole; (2) introduction to the class Insecta, as a prelude to chapters on individual orders in Volume I; and (3) description of characteristics of the group known as springtails (Collembola), with a focus on aquatic taxa. Keys for identifying these arthropods are given in subsequent volumes devoted to individual zoogeographic regions.

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related to the other major subphylum Crustacea (see Chapter 27). Some authorities consider Hexapoda to be a sister group with all or a subset (either Malcostraca or Branchiopoda) of Crustacea, depending on whether the latter subphylum is considered mono- or polyphyletic (e.g., Cranston and Gullan, 2009). Within this subphylum are the class Insecta (insects) and two other groups: (1) Collembola (springtails) and Protura (known either as proturans or more rarely “coneheads”; Figure 33.1), and (2) Diplura (two-pronged bristletails; Figure 33.2). Springtails and coneheads are usually considered as either separate but closely related classes in the subphylum Hexapoda or as orders within the class Parainsecta, depending on whether Parainsecta is viewed as being composed of paraphyletic or monophyletic taxa. The evolutionary position of Diplura is uncertain. It was traditionally placed with Collembola and Protura in the class Entognatha, because the mandible and maxilla of all three groups are partly contained within the head capsule (as opposed to the external position in insects), but it is more commonly now treated as the sole order within the class Entognatha. However, some authors still consider Collembola to be a member of the class Entognatha. In the remainder of this chapter, we will focus on the general biology and ecology of both springtails (with an emphasis on aquatic species) and the class Insecta as a whole. Please consult Chapters 34–41 for more detailed

Phylogenetic Relationships As discussed in Chapter 24, the higher taxonomy of Arthropoda is somewhat controversial at present (e.g., Akam, 2000; Giribet and Edgecombe, 2013). More recent systematics (e.g., Dohle, 2001; Dunn et al., 2008; Regier et al., 2010; Giribet and Edgecombe, 2013) divide the phylum Arthropoda into the subphyla Trilobita, Chelicerata, Myriapoda, and Tetraconata (or Pancrustacea), although the last clade may be elevated to include multiple subphyla of its own. The presence of the hexapods within Tetraconata indicates the evolutionary origin of insects from crustaceans. Because the taxonomic level of Hexapoda—and all levels above family—have not been resolved yet from this potential change in the status of the former subphylum, we will continue to refer to the group as a subphylum in this chapter. Hexapoda is the largest subphylum (or superclass in some taxonomic schemes) within the huge phylum Arthropoda (see Chapter 24), where it is most closely

FIGURE 33.1  Springtails courting. Collembola: Sminthurides lepus. Photo courtesy of Dr F. Soto.

FIGURE 33.2  Drawing of the dipluran Parajapyx sp. K.A. Justus in Markus Koch’s section on Diplura, from Resh and Cardé (2009a).

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information on the general biology and ecology of insects in orders with aquatic species. Taxonomic information at the family level is available for insects and springtails in the taxonomic volumes of this series. Protura and Diplura will not be discussed further in this chapter because these groups lack aquatic members.

INTRODUCTION TO THE CLASS INSECTA Evolutionary Diversification of Aquatic Insects If recent theories have correctly concluded that insects evolved from a marine or perhaps freshwater crustacean ancestor, then a natural assumption might be that the earliest insects lived in freshwater, estuarine, or marine habitats and only later colonized and diversified on land. However, no firm evidence supports this conclusion, and, in fact, the earliest, definitive evidence of aquatic hexapods is from the late Triassic (c. 230 mya), which is approximately 200 mya younger than the earliest known terrestrial insects linked to the Devonian and early Carboniferous periods (e.g., Bradley et al., 2009; Cranston and Gullan, 2009). The most primitive aquatic insects identified from fossil remains—aquatic heteropteran bugs—show evolutionary adaptations from a terrestrial existence, such as evaginated trachea in the form of external aquatic gills. The oldest aquatic insects are in the paleopteran group, which comprises extant orders (Ephemeroptera and Odonata) and one the extinct, nonaquatic order (Paleodictyopterida). More controversy surrounds the evolutionary appearances and relationships of the remaining orders with aquatic species. Aquatic insects occur within 14 orders, but only 5 of these are wholly or mostly aquatic: Ephemeroptera (mayflies), Odonata (dragonflies and damselflies), Plecoptera (stoneflies), Trichoptera (caddisflies), and Megaloptera (e.g., hellgrammites and dobsonflies). Of the other nine orders with aquatic species, three are extremely abundant on an absolute scale in inland water systems but are vastly more speciose in terrestrial habitats: Hemiptera (true bugs), Coleoptera (beetles), and Diptera (aquatic flies and midges). Another six orders have relatively and absolutely low numbers of aquatic species: Orthoptera (aquatic grasshoppers), Neuroptera (spongillaflies), Hymenoptera (parasitoid wasps attacking aquatic larvae of other orders), Lepidoptera (aquatic moths), Mecoptera (scorpionflies), and Blattodea (aquatic cockroaches). Within this last group of six, only Neuroptera and Lepidoptera contain species for which the larvae are free-living residents within water. Among the lifestyle groups that apparently never evolved or adapted to an aquatic existence are the eusocial insects. This is reflected in the total absence of aquatic ants, wasps, bees, and termites, with the exception of parasitoid

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wasps that lay their eggs in aquatic larvae and then leave the water or drown.

Additional Literature Sources The popularity and importance of studying insects are responsible for the huge literature on this subject. Chapters 34–41 in the present volume describe aspects of the ecology and general biology of the various orders of aquatic insects, and they provide detailed literature backgrounds of each order. For a broader perspective on insects, Resh and Cardé (2009a) is an excellent source for most aspects of insect biology but includes only limited aquatic coverage. This subject is covered in many scientific journals and several scientific organizations, such as the Society for Freshwater Science (formerly the North American Benthological Society), the European Federation for Freshwater Sciences, and various freshwater groups in other regions of the world. Aquatic insect ecology has also been the subject of numerous books (e.g., Resh and Rosenberg, 1984; Ward, 1992), but many of them are somewhat dated. The taxonomic literature on aquatic insects is broad. Chapter 16 in Volumes II and III of Thorp and Covich’s Freshwater Invertebrates includes keys to families of aquatic insects in the Nearctic and Palaearctic regions, respectively, and future volumes are expected to include keys at the family level or lower for insects in different zoogeographic zones. Keys to aquatic insects and collembolans have been published for various zoogeographic regions and vary widely in quality, accessibility, and language. Some examples are the following. The best for North America is edited by Merritt et al. (2008), which at this writing is in its fourth edition. The multivolume series Aquatic Biodiversity in Latin America contains excellent coverage of various groups; it is published in English, Spanish, and Portuguese by a Russian publishing group. The Australasian and Oceana fauna are covered in the Cooperative Research Centre for Freshwater Ecology: Identification and Ecology Guides, but many of these are out of date or limited to certain parts of Australia. Many volumes have been published on Asian fauna, but these are primarily regional keys and vary from very high to very low quality. Several volumes (but less than the original 20 planned) of the Guides to the Freshwater Invertebrates of Southern Africa were published by the Water Research Commission. A few field guides are available for students and amateur naturalists, most notably two by Voshell (2002) and Thorp and Rogers (2011).

GENERAL BIOLOGY OF AQUATIC INSECTS The following sections summarize first the general biology of insects with an emphasis on larvae. For more specific information, consult Chapters 34–41 or other reliable sources, such as the Encyclopedia of Insects (Resh and Cardé, 2009).

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Tagmatization The basic insect body contains three regions or tagma— head, thorax, and abdomen—each composed more or less of fused segments with disparate functions (Figure 33.3). The distinctiveness of these tagma and elaboration of ­different structures vary widely among the large morphological and functional diversity of larva insects.

Head The anterior tagma forms a mostly upwardly oriented head. It contains dorsally one pair of antennae (vs. two in the related subphylum Crustacea), two compound eyes, and three or fewer simple eyes (ocelli). The ventral side features feeding structures, some of which have a defensive function in a few insects. These mouthparts include a labrum (upper lip), one pair each of mandibles and maxillae, and a labium (lower lip). Mandibles often are heavily sclerotized, which enables them to function as a toothlike structure. Maxillae and the labium possess palps, which are divided into two or more palpomeres whose function is sensory. The mouth structure reflects whether food is obtained by biting animal prey, scraping algae, manipulating dead organic matter, or sucking liquids from plants or animals. Aside from the head’s role in obtaining and initially processing food, it is the primary site of neural activity. External sensory data (e.g., from the eyes and tactile structures) and internal functional information are processed by a socalled “brain” (Strausfeld, 2009). This structure consists of: (1) a fused, tripartite supraesophageal ganglion (proto-, deuto-, and tritocerebrum); and (2) a postoral, subesophageal ganglion. The former processes information received from throughout the body, including from paired segmental

ganglia and a ventral nerve cord passing through all tagma. The subesophageal ganglion primarily processes information from the mouthparts and associated structures and glands.

Thorax The middle tagma, or thorax, comprises three large segments: prothorax (anterior most segment), mesothorax (middle), and metathorax (posterior) (Figure 33.3). Each segment contains chitinous plates, or sclerites, which provide support and some protection. These sclerites are called the notum, sternum, and pleura (singular = pleuron) for the dorsal, ventral, and lateral sclerites, respectively. The meso- and metathorax of hemimetabolous larvae and paurometabolus nymphs may contain dorsal wingpads (precursor of adult wings). The most prominent thoracic structures in most insect larvae are a pair of segmented legs on each thoracic segment. Each leg is articulated into five divisions: coxa (most proximal leg segment), trochanter, femur and tibia (both often elongated), and tarsus (most distal segment). The tarsus, which is often subdivided, terminates in zero to two claws depending on which leg is examined and the taxonomic group. For insect larvae with legs (the vast majority), these appendages primarily function in locomotion but are used in digging in some taxa. Adults of most insects also have functional wings on the thorax; these develop in many cases from larval wingpads. The thorax as a whole has many functions other than locomotion, including digestion and in some cases respiration. The digestive system consists of a fore-, mid-, and hindgut. However, these do not correspond exactly in position with the three tagma, although the foregut starts in the head and the hindgut terminates in the abdomen.

Abdomen The last tagma is the abdomen (Figure 33.3). It is primitively subdivided into 11 segments, although the last 2 may be fused and thus indistinguishable. Sclerites on these segments are arranged as in the thorax. One to three caudal appendages occur in some taxa. Simple or branched, tubular gills may be present on the first two anterior segments (as in stoneflies and mayflies) or occur on the terminal segment (as in damselfly nymphs). Functions of the abdomen include respiration, absorption of nutrients, water and salt balance (via Malpighian tubules and rectal glands), and reproduction in adults.

Respiration and Circulation Respiratory System FIGURE 33.3  Major regions of insect anatomy. Drawn by Brian O’Neill.

Obtaining oxygen is relatively easy from air but challenging from water because air holds approximately

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200,000 ppm of O2, whereas saturated cold water retains only 12–15 ppm; moreover, oxygen molecules diffuse more than 300,000 times faster in air compared with water (Resh et al., 2008). Under nonpolluted conditions, oxygen levels are higher in cold versus warm water, lotic versus lentic systems, turbulent versus laminar flow conditions, oligotrophic versus eutrophic systems, low versus high detrital abundance, and surface versus groundwaters. Low-oxygen conditions can affect the distribution of species, especially in lentic habitats. For these reasons, aquatic adult and larval insects have commonly abandoned primary dependence on the internal tracheal system found in most terrestrial insects and instead depend on external gills, absorption across the external integument or through the anus, transported air bubbles, or oxygen sucked from underwater plants. Internal tracheal systems (Figure 33.4) consist of a network of air-filled tubes that transport gases by diffusion between outside air (free or bound within an air bubble held by the insect under water) and internal cells. The latter range from large tracheae to minute (2–5 μm) tracheoles, which contact individual cells. Air enters the tracheal system through many (polyneustic systems) or few (oligopneustic) pairs of contractile spiracles or across the outer cuticle in closed (apneustic) systems. Because of this ramifying network of tubes, transport of oxygen within the circulatory system is relatively unimportant in most species. Oxygen for the tracheal system can be obtained by aquatic insects from: (1) the air by flying adults and surface-dwelling adults and larvae; (2) air tubes of underwater plants by “plant breathers”; or (3) an underwater air bubble. Some plant breathers in the orders Coleoptera and Diptera have highly modified spiracles capable of piercing roots and stems of aquatic vascular plants to obtain oxygen from the plant’s air-filled tubules (aerenchyma). Underwater air bubbles are carried by many aquatic beetles and bugs. These transportable air supplies occur in temporary and permanent (plastron) air storage sites carried under the insect’s

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body. Several families of beetles capture an air bubble from the surface and then carry it underwater as a temporary storage or as a physical gill (Figure 33.5) before replenishing the bubble periodically at the surface. In both cases, oxygen enters the bubble from the surrounding water (especially from colder, more oxygenated water) by diffusion, whereas waste gases diffuse outward from the bubble. Some beetles (e.g., riffle beetles), naucorid bugs, and aquatic moths living in more oxygenated habitats use a permanent or at least longer lasting storage device called a plastron. Hydrofuge hairs of the plastron hold a nitrogen-rich layer of air between the insect body and an outer air bubble. Oxygen diffuses continuously into and through the outer bubble into the plastron and then into the tracheal system. Waste gases can pass in the opposite direction. Filamentous and platelike tracheal gills (Figure 33.6) are used on the thorax or abdomen by members of many insect orders, such as Ephemeroptera, Odonata, and Plecoptera to name a few. Damselfly nymphs (Odonata, Zygoptera) have prominent caudal gills, whereas dragonfly nymphs (Odonata, Zygoptera) have internal gills in the caudal area of the alimentary tract. In the latter case, water is pumped into and out of the anus and around the gills. Despite their name, tracheal gills may have an equal or even dominant role in osmoregulation and may function as gills only under low-oxygen conditions; their primary role remains controversial. Insects with high surface-to-volume ratios (e.g., most dipteran and aquatic moth larvae) and little external, relatively impermeable chitin commonly respire entirely across the body surface by cutaneous respiration. Even some insects with large tracheal gills may obtain most of their oxygen by passage of gases across the outer integument. Oxygen-binding respiratory pigments are less important in most Hexapoda because of the ramifying nature of the tracheal system, but two pigments occur in the hemolymph of some groups. Hemocyanin, which is common in crustaceans, is present in more primitive hexapod groups, including Collembola and the insect order Plecoptera. Hemoglobin occurs in the hemolymph of some larval and pupal midges (Diptera, Chironomidae), particularly in the bright red “bloodworms” (Chironomus), and in two genera of backswimmers (Hemiptera, Notonectidae, Anisops and Buenos). Hemoglobin enables bloodworms to occupy lowoxygen environments (e.g., detrital sediments of lakes) and backswimmers to transfer oxygen from an attached air bubble into the bug’s body, thereby stabilizing buoyancy and extending dive time.

Circulatory System FIGURE 33.4  Internal tracheal system of an insect. Drawn by Brian O’Neill.

Characteristics of the open circulatory system vary with the insect’s size and developmental stage. In more complex forms, it consists of: (1) a dorsal vessel running most of the body

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FIGURE 33.5  Ventral air bubble of the backswimmer Neoplea (Heteroptera: Pleidae). Resh et al. (2008).

FIGURE 33.7  Open circulatory system of generalized insect. Open arrows show flow direction of hemolymph. Drawn by Brian O’Neill.

Excretion and Osmotic Balance

FIGURE 33.6  Plate-like tracheal gills (Ephemeroptera). Photo courtesy of Matthew A. Hill.

length, with the anterior section called the aorta; (2) a dorsal tubular heart; (3) an open hemocoel; and (4) paired ostia and contractile vesicles in the abdominal segments (Figure 33.7). Although the heart and some accessory structures have some contractile abilities, general body movements are responsible for most circulation, especially in smaller individuals.

Although terrestrial insects have adapted to reduce water loss by excreting nitrogen wastes as uric acid, aquatic forms primarily release waste nitrogen as ammonia, which is a highly toxic product that requires more water for dilution but is energetically less expensive to produce than uric acid. Wastes products are released from the hemolymph into the gut for expulsion through the Malpighian tubules. These are slender branching tubes connected to the alimentary tract between the midgut and hindgut that also absorb solutes and water as needed. The product they produce is generally isosmotic to external water; thus, these do not always play a role in maintaining internal osmotic conditions. The isomotic and osmotic state of aquatic insects depends on the external habitat. Those living in soft or hard “fresh” water habitats are hyperosmotic to the external water, meaning that they tend to lose ions and gain water. They replace these ions by absorption through the gut and

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release excess water by excretion. In contrast, those living in hypersaline environments, such as Mono Lake (­California, United States), are constantly losing water and gaining salts. Among the most efficient ionic and osmotic regulators of insects are the brine and shore flies (Diptera, Ephyridae), such as Ephydra hians, which flourishes in waters at least 3 times as concentrated as the ocean as well as being extremely basic (pH ~10) and having high concentrations of sulfate and bicarbonate anions. This brine fly maintains a normal osmotic concentration (~300 mOsm, similar to other aquatic species) even when external concentrations reach 2000 mOsm. To survive in this extreme environment, the fly must drink large amounts of liquids, save the water, and excrete the salts. Its hyperosmotic urine is produced in the hindgut anterior to the rectum, whereas mosquitoes in saline environments produce hyperosmotic urine in their extensive rectum.

Reproduction and Development Generation Time Aquatic insects generally have one (univoltine), two (bivoltine), or multiple (multivoltine) generations per year, depending on the order, species, and climatic conditions. However, in environments with very low annual temperatures (e.g., Arctic) or low food availability, an individual generation may require 2 (semivoltine) or more years. Indeed, two species of true midges are reported to have 7-year life cycles in Alaska. Huryn et al. (2008) included an extensive table showing the generation patterns for all orders and many species of aquatic insects. Given that reproduction primarily occurs in the terrestrial phase of insects, this process will not be considered further in this chapter. However, insect reproduction is discussed for each order in Chapters 34–41.

Metamorphosis Insects undergo gradual to radical changes from egg to adult as an animal grows and develops. For these processes of metamorphosis to proceed, the old cuticle must be shed and replaced with a larger exoskeleton in a process termed a molt or ecdysis. The older skin, or exuviae, is discarded or sometimes consumed by the developing arthropod as a source of calcium. Ecdysis can be physically challenging and dangerous for the insect, especially if a predator locates the temporarily soft-bodied and mobility-challenged prey. Aquatic hexapods undergo one of three basic patterns of metamorphosis: ametabolous, hemimetabolous, and holometabolous development. Ametabolous metamorphosis involves gradual changes between instars with increasing growth and eventual reproductive capability; no dramatic changes in body form occur between immature larvae and mature adults. Collembola are the only aquatic hexapods

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exhibiting this form of development. Hemimetabolous development is also termed incomplete or gradual metamorphosis because only three basic stages are involved: egg, nymph, and adult (with a preadult subimago in mayflies). That is, pupation does not occur. The nymph may resemble the adult somewhat and change only marginally between molts (­paurometabolous development), as in aquatic bugs, or it may be very distinct (e.g., in ­Odonata). Well-developed antennae, eyes, and legs are evident early in the nymphal stage, as are rudimentary wings. This pattern of h­emimetabolous development is shown among the following orders with aquatic species: Ephemeroptera, ­Hemiptera, Odonata, and Plecoptera, as well as in ­Orthoptera with its very few and only marginally aquatic species. ­Holometabolous (complete) development includes egg, larva, pupa, and winged adult. This metamorphic pattern is typical of the orders ­Coleoptera, Diptera, ­Lepidoptera, Megaloptera, Neuroptera, and Trichoptera as well as the parasitoid Hymenoptera. Stages of metamorphosis differ among insect orders, and the names describing them vary somewhat among countries and between insects and other arthropods (Stehr, 2009). For purposes of this book, we generally refer to the stages of insect development as egg, larva, pupa, and adult (imago), with each separated by one or more (in larvae) molts. However, not all orders go through all of these stages, as explained below (Figure 33.8). The separate stages are also called instars. Some controversy exists about the name for the stages between egg and either pupa, subimago, or adult. The general term “larva” is used widely for all taxa at this point in development, but the terms nymphs or naiads have been used for juvenile instars of insects with direct development, as in Ephemeroptera, Hemiptera, Plecoptera, and Odonata. The most dramatic development occurs during pupation for species exhibiting this metamorphic stage. The short subimago stage, which is present only in Ephemeroptera, resembles an adult but is sexually immature. Aquatic insects typically pass through 4–6 instars, but up to 30 have been reported for some Ephemeroptera, Odonata, and Plecoptera. During adverse environmental conditions, the insect may exist as a resistant egg or pupa before eclosion produces a larva at hatching or an adult from the protective pupal case.

GENERAL ECOLOGY AND BEHAVIOR OF AQUATIC INSECTS The literature of the ecology of aquatic insects is vast; thus, no one book—much less one chapter—can hope to even touch on all of the major issues. Indeed, there are scientific organizations (e.g., Society for Freshwater Science) that have aquatic insects as a de facto scientific emphasis and probably thousands of papers are published annually in various journals with some relevance to these organisms. The second and third editions of the Ecology and Classification

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FIGURE 33.8  Hemimetabolous and holometabolous development. Drawn by Brian O’Neill.

of North American Freshwater Invertebrates briefly covered aspects of aquatic insect ecology in a single, excellent chapter (e.g., Hershey et al., 2010). This has been replaced in the fourth edition by expanded coverage on the ecology of mostly individual insect orders in Chapters 34–41. Consequently, the current chapter merely summarizes some important aspects and focuses more on general patterns.

Habitat Selection Some general patterns of insect diversity within pristine habitats include a direct relationship with resource abundance, habitat permanency (seasonal to evolutionary time periods), and habitat complexity and an indirect association with latitude, altitude, salinity, and distance (e.g., islands) to large land masses (continents). Although diversity tends to be reduced by fish predation, other characteristics in those habitats containing fish may reverse the overall relationship. However, it is important to note that not all taxonomic groups follow these general rules. Moreover, some positive relationships (e.g., favored by higher oxygen content) may be hidden by effects of other factors associated with opposite trends (e.g., temperatures at high altitudes and latitudes). More information about different aquatic habitats can be found in Chapter 2 of this volume.

Ephemeral versus Permanent The abundance and diversity of aquatic insects is almost always greater in relatively permanent aquatic systems than in ephemeral streams and wetlands. The magnitude of the difference is related greatly to the predictability, periodicity

FIGURE 33.9  Ephemeral wetland commonly called Playa Lake, Pawnee National Grassland, Colorado, United States. Photograph by Brian O’Neill.

(seasonal timing), and length of the wetted stage (weeks to months or years). Insect diversity tends to increase with longer wetted periods up until the point that the wetland can support insectivorous fish. Ephemeral wetlands, such as playa lakes (Figure 33.9), are dominated by branchiopod crustaceans, such as fairy shrimp (Figure 33.10) and cladocera, but they also contain diverse insect populations that can reproduce rapidly or fly to other wetted habitat when the initial pool dries. Species with multiyear life cycles are essentially excluded from ephemeral lentic systems unless they use a dormancy stage. The relative abundance of insects and crustaceans varies with time of the year, geographic location, and characteristics of individual wetlands.

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FIGURE 33.10  Representative fairy shrimp. Crustacea: Anostracta: Chirocephalidae. Photograph courtesy of Denton Belk.

FIGURE 33.11  Brine fly from the Great Salt Lake, Utah, United States (Diptera: Ephydra cinerea). Photo courtesy of Wayne Wurtsbaugh.

The ecological effects of drying in ephemeral headwater streams and even large rivers in arid biomes are typically different from environmental constraints in wetlands. When a stream dries, aquatic stages of insects can travel downstream, survive within deeper pools, move upstream into more permanent springs where present, or dig vertically or laterally into hyporheic waters. Adult insects can emerge as adults and then either travel to a more suitable habitat or leave behind resistant eggs. Nonetheless, insect diversity is still lower in ephemeral than permanent lotic ecosystems.

include challenges of dispersing by winged adults in windy habitats and difficulties for late instars emerging on either wave-swept shores or rough ocean surfaces from depths much greater than present in rivers and almost all lakes (passing through a gauntlet of predators in the process). Given the paucity of aquatic insects in estuaries and ocean environments, it should not be surprising that insects are vastly more abundant in freshwater than inland saline environments. Dipterans are the most commonly occurring insects in saline lakes. For example, the insects found in and along the shoreline of the Great Salt Lake of the western United States consist of two species of brine flies (Ephydra cinerea and Ephydra hians; Figure 33.11), and another possible permanent resident is the water boatman Tricorixa verticalis (Figure 33.12). This hemipteran apparently feeds on Artemia brine shrimp (Crustacea, Branchiopoda) and larvae of the dipteran flies. See a subsequent section on physiological constraints on life in saline waters.

Oceans and Saline Lakes versus Freshwater Habitats Although dipterans (especially mosquitoes, true midges, and biting midges) and members of a few other insect orders colonize estuarine environments and some intertidal areas, they are entirely absent from the open ocean except for the surface-dwelling water striders in the genus Halobates (Hemiptera). Although osmoregulation was previously the most common explanation for this depauperate marine fauna, it has lost popularity because of the presence of estuarine species and a few insects in saline lakes. The currently most popular explanation is the presence of more ancient, diverse, and superior competitors in the ocean, especially the very diverse assemblage of benthic polychaete worms (Gullan and Cranston, 2005; Huryn et al., 2008). However, this and other explanations are merely conjecture at this point. We suspect that no single cause for the near absence of marine insects exists. Rather it could be some combination of: (1) osmoregulatory limits for most extant insect taxa; (2) interspecific resource competition (especially with polychaetes and the highly diverse crustacean assemblage) and occasional interference competition (e.g., with snails and coral); (3) intense and diverse benthic through nektonic predation by fish and invertebrates (e.g., cnidarians); (4) the general absence of insects from planktonic habitats (see later section); and (5) other life-history constraints. The last could

Lotic versus Lentic Faunal Differences The invertebrate fauna of lakes and wetlands differs considerably from that in headwater streams and the main channels of rivers but less so from the lateral slackwaters and true backwaters of multichanneled rivers. The differences are likely primarily due to the combined effects of: (1) hydraulic forces from flowing water (occasionally stressful in lotic systems but mostly benign in lentic habitats except for wave action near shore); (2) thermal/dissolved oxygen characteristics (typically less temperature fluctuations and higher oxygen in flowing water in general); (3) substrate characteristics (often larger particle size and higher oxygen levels in lotic systems); and (4) food provisioning (filter feeding easier for insects in lotic systems) and type (autotrophic production commonly greater in lentic systems, at least in the epilimnion). In addition, flowing water accumulates less ice than lentic systems in the

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FIGURE 33.13  Epigean sprawler-dragonfly nymph in the Libellulidae family. Photo courtesy of Matthew A. Hill.

FIGURE 33.12 Waterboatmen, Tricorixa verticalis, that frequents the Great Salt Lake, Utah, United States. Photo courtesy of Katherine Sirianni.

same region. When this lake ice is near shore, it scours the bottom and any resident invertebrates in wetlands, ponds, and lakes as it melts on windy days. However, stream ice, when present, can be potentially more dangerous because of the scour from water currents moving ice downstream. Finally, the catastrophic and behavioral drift common in many stream insects transports them downstream, requiring mechanisms to repopulate upstream areas—a problem not encountered in lakes. The results of these physicochemical differences are that the diversity and type of stream insects vary between lotic and lentic systems. Pleustonic and other water-surface-dwelling insects (e.g., gyrinid beetles) and springtails are more common in numbers and diversity in lentic environments, although they inhabit the true backwaters of rivers and other sheltered environments of all lotic systems. Epigean sprawlers (e.g., many dragonfly nymphs; Figure 33.13) are typically most abundant in lentic systems. Species living within organic sediments (e.g., chironomid midges) are more common in lakes whereas hyporheic insects requiring coarser, more oxygenated sediments (e.g., specialized stoneflies) are more abundant along rivers. Filter-feeding insects (e.g., net-building caddisflies; Figure 33.14) are more diverse in lotic systems because food is constantly arriving on the currents. Finally, species

FIGURE 33.14  Caddisfly net used for filter feeding (Trichoptera). Photo courtesy of Garold Sneegas.

requiring more oxygen (e.g., many stoneflies) occur predominately in lotic systems, especially at higher elevations where temperatures are lower and oxygen tensions are greater. Behavioral and Catastrophic Drift One common characteristic of lotic systems not found to any extent in lakes is insect drift. Organisms depart the bottom substrate and enter the water current for transport downstream as a result of behavioral or catastrophic drift. In the former case, an insect will release its hold on a rock, log, or other substrate and be propelled downstream. However, in catastrophic drift, the insect has not chosen to enter the water current and instead is impelled downstream by

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FIGURE 33.15  Phantom midge (Insecta: Diptera: Chaoborus). Photo courtesy of Matthew A. Hill.

strong currents ripping it off of the substrate or as a result of another organism bumping it off a rock or log. Drifting occurs most frequently during the night. This was formerly used as evidence of the importance of behavioral over catastrophic drift, but another explanation is that aquatic insects tend to be more active in general at night when fish are visually constrained, and this activity may place them on exposed surfaces where currents may tug them off. However, it seems clear that some behavioral drift occurs when the insect is seeking more food or escape from a benthic competitor or predator. Not all insect orders or families tend to drift, and larger insects are less likely to drift than smaller ones, probably because of vulnerability to fish predators. For example, mayfly nymphs (Ephemeroptera; Figure 33.6) are frequent drifters, whereas dragonflies (Odonata) rarely drift. One potentially puzzling question might be why is the upstream community not depopulated by downstream drifting of resident insects? The answer seems to be that adults tend to fly upstream to lay their eggs after emerging and mating, and their progeny benefit in lower population densities compared with downstream assemblages. Moreover, some taxa in their aquatic stages tend to move progressively upstream on an individual basis over time, and a few invertebrate taxa undertake mass migrations (e.g., some crustacean amphipods).

Planktonic versus Benthic Most aquatic insects have a benthic larval stage and a winged adult stage capable of flying for a short (e.g., mayflies) or long period (e.g., dragonflies). The very rare exceptions include stoneflies from ancient, deep-water lakes that lack a winged adult stage: Capnia lacustra in Lake Tahoe and Baikaloperla elongata in Lake Baikal (both in Plecoptera, Capniidae). Although both reside in deep-water habitats, they are primarily benthic shredders feeding on aquatic vegetation. In a very different environment, the neotenous stonefly Isocapnia sp. lives in the hyporheic zone adjacent to gravel bed rivers in Montana and never develops a winged stage (Geoffrey Poole, personal communication).

The overwhelming focus on a zoobenthic existence by aquatic insects means that none are true members of the pelagic “holoplankton” (i.e., organisms that spend their entire life cycle in the water column of open-water areas). True holoplanktonic invertebrates in freshwaters are, instead, microcrustaceans (copepods and cladocera) and rotifers, all of which are small on an absolute scale and relative to almost all insects. Pelagic, meroplanktonic invertebrates—organisms that spend only a portion of their lives in the plankton of open waters—occur, but they are still rare in freshwaters. Aside from short-lived, early instar stages of meroplanktonic insects (mostly chironomid midges), which spend most of their larval existence in the benthos, the only true meroplanktonic insects are larval dipterans, such as Chaoborus (phantom midges; Figure 33.15). These migrate vertically on a daily basis from deep, dark waters during the day to shallow, dark waters at night. Although very few insects are planktonic and pelagic, many insects live above the bottom in areas where fish predators are absent or are less-effective foragers, with the latter primarily being areas with rooted vegetation in the littoral zone. Some other insects inhabit the near-surface area inside and outside of the protective littoral zone when fish predators are temporarily or permanently absent. These taxa may dive toward the bottom in search of food or live just below or on the water surface. Some examples are mosquito larvae (Diptera, Culicidae), backswimmers (Hemiptera, Notonectidae), and water striders (Hemiptera, Gerridae). Ephemeral pools and salt lakes lack fish predators; thus, they can support insects outside of the littoral zone. Several possible abiotic and biotic characteristics may explain the paucity of planktonic insects. From a physical perspective, being very small (20 m usually) and open ocean (Hopkin, 1997). “Aquatic” Collembola are actually only semiaquatic, with most occupying moist terrestrial habitats, including most of the nearly 700 species reported from North America alone.

Phylogeny and Species Diversity Collembola are among the most ancient hexapods, with fossil forms stretching at least back to the early Devonian 400 mya. Truly aquatic species are rare, with most aquatic species reported from the water surface or on floating aquatic plants. For example, Waltz and McAfferty (1979) considered that only 10 North American species were semiaquatic and 5 more were riparian with a propensity to venture onto the water surface. Even within these 15 species, the aquatic adaptations varied widely. Most species in the Nearctic are either Podura aquatica or members of the genus Sminthurides (Christiansen and Snider, 2008). Taxonomic keys to families of semiaquatic Collembola in various zoogeographic regions appear in subsequent volumes of the fourth edition of Thorp and Covich’s Freshwater Invertebrates In addition, Christiansen and Snider (2008) briefly discussed the biology of Collembola and provided a key to families and genera for North America.

General Biology of Collembola External Anatomy Collembola are small, wingless hexapods, usually less than 6 mm long, with aquatic species usually less than 3 mm

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(Figure 33.1). They differ from insects in having only 6 abdominal segments instead of as many as 11 in primitive insects (fewer in more modern species of insects and typically combined into abdominal tagma). In addition, their maxillae and mandibles of the head are concealed under the head capsule into which their labium is fused. The tibia and tarsus on each leg are fused into a single segment in springtails. Their most distinguishing features are the collophore (described previously) and the abdominal furcula (= furca) on the fourth abdominal segment. The furcula is present in most species and all taxa considered at least semiaquatic. The furcula consists of a basal piece (manubrium) and two arms (Figures 33.22–33.25). Each arm consists of a basal (dens) and an apical (mucro) segment. The furcula is typically folded under the abdomen by a midventral, clasplike structure (tenaculum) on the third abdominal segment. When the tenaculum releases the furcula, the latter springs backward in as little as 18 ms and propels the animal several centimeters through the air—hence, the common name “springtail.” Most truly aquatic Collembola, such as Podura aquatica, share some defining characteristics, the most obvious being the shape of the furcula. Unlike the laterally flattened furculae of terrestrial species, the furculae of aquatic species are dorsoventrally flattened. The mucro is often enlarged (Deharveng et al., 2008), creating a paddle-shaped structure (Figure 33.26), and some species have setae along the “handle” of the paddle. The latter presents more surface area on the water, thereby allowing the springtail to make a more efficient strike to the water’s surface (Guthrie, 1903) that is less likely to break through the surface tension. These remarkable adaptations to the aquatic habitat (Maynard, 1951) are even inspiring engineers to create extremely small machines that walk and jump on the surface of the water (Hu et al., 2007). The spring mechanism in Collembola is not designed for normal locomotion per se, but it is instead a device for jumping away from danger. The resulting direction is somewhat unpredictable for the collembolan and its intended

FIGURE 33.22  Sminthuridae (lateral view) showing furcula (F) and collophore (C). Peckarsky et al. (1990). FIGURE 33.24  Poduridae; furcula of Podura (dorsal view) showing manubrium (MA), dens (D), and mucro (MU). Peckarsky et al. (1990).

FIGURE 33.23 Poduridae; Podura (lateral view) showing furcula (F) and collophore (C). Peckarsky et al. (1990).

FIGURE 33.25  Isotomidae anterior portion (lateral view). Redrawn from Peckarsky et al. (1990).

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FIGURE 33.26  Furcula of aquatic collembolan showing enlarged mucro and setae that increase surface area. Drawn by Brian O’Neill.

predator, but the relative great distance traveled in comparison to the collembolans size often provides a suitable margin of safety for the springtail. Aquatic species also use their ungues (claws) and ventral tube (the only wettable areas of the cuticle) to anchor themselves to the water’s surface (Baudoin, 1955; Paclt, 1956). The claws break through the surface film of the water, giving lateral purchase and traction, whereas the unwettable nature of the tibiotarsus provides the buoyancy to keep the springtail above the water (Noble-Nesbitt, 1963). Iostoma riparia has broadened claws that facilitate movement on the water surface (Simonsen et al., 1999).

Reproduction and Development Springtails do not copulate in the manner of insects. After courtship, the male in most species deposits stalked or sessile spermatophores that are then retrieved by a female with no further contact with her partner. However, given an opportunity, the males of all species of Collembola will eat the spermatophores of other males, regardless of species, and replace them with their own, thereby reducing competition (Waltz, 1979; Hedlund et al., 1990). Even more vulnerable are the spermatophores of species that deposit them directly on the water surface. Consequently, direct sperm transfer (Hopkin, 1997) and morphological modifications to facilitate sperm transfer are more common in aquatic species (Guadalupe Palacios Vargas and Castaño-Meneses, 2009). Strebel (1932) described indirect and direct sperm transfer in Podura aquatica. Direct mating consisted of both sexes touching abdomens and transferring sperm. Similar behavior has been observed in Sminthurides aquaticus (Hopkin, 1997). Indirect mating in P. aquatica begins with males performing a courtship

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FIGURE 33.27  Floating spermatophore of Podura aquatica. Drawn by Brian O’Neill.

dance and depositing a stalked spermatophore with a floating plate on the water surface (Figure 33.27). They then push and bump the female to the spermatophores (Schliwa and Schaller, 1963). The female then takes up the spermatophore through a set of vulvae at the genital pore (Waltz, 1979). Eggs of aquatic species are generally hatched in the genital opening of the female within a few weeks, and the young are then carefully laid onto a moist surface (Chang, 1966). Egg-laying can be delayed by dry external conditions or the female’s diapause. For eggs that were expelled or oviposited, as in other species, the eggs sink to the bottom and hatch normally. Suitable habitats for egg-laying may include habitats below the water surface in at least one species—making the newly hatched young truly aquatic until they pass through the water surface film and become semiaquatic. Young collembolans reach sexual maturity within a few weeks and molt from 3 to 7 times before reaching maturity. The young of some species may enter diapause. If they are hatched underwater, immature springtails may briefly feed on submerged matter. Once they are exposed to the air-water surface, they develop an unwettable cuticle and never again become submerged (Chang, 1966; McCafferty, 1981). Collembola undergo a gradual metamorphosis. As a result, young springtails closely resemble adults except for the former’s smaller size and lack of genitalia. Molting will continue throughout the adult’s life of a few weeks to a few months, with at least one species reported to molt 50 times whereas other species may molt as little as only twice during the adult stage. Most species overwinter in a diapausing stage, but some are cold hardy and active at temperatures approaching freezing.

Chapter | 33  Hexapoda—Introduction to Insects and Collembola

General Ecology of Collembola The ecology of aquatic (semiaquatic) Collembola is poorly known in general (Deharveng and Lek, 1995). Thus, the following discussion is relatively sparse and subject to revision as more data slowly come in from investigators around the world.

Habitat Although approximately 45% of Collembola are either hydrophilous (Deharveng and Lek, 1995), riparian species, or associated with wet habitats, the truly aquatic Collembola constitute a relatively small proportion of the entire order. However, Collembola often constitute the most abundant and diversified arthropods in a large range of wet habitats (Deharveng and Lek, 1995) to the point that they can cover the water and completely obscure the pool below (Maynard, 1951). “Almost any quiet body of water has its quota skipping about on the surface, while the vegetation along the shore will hide others. Even the surfaces of wells and cisterns may be populated by several species” (­Maynard, 1951). However, keep in mind that many species are found on water accidentally (McCafferty, 1981), and very few lineages have adapted to this environment (­Deharveng and Bedos, 2004) from their terrestrial edaphic origin (D’Haese, 2002, 2003). Springtails are most often associated with calmer lentic freshwater and more rarely with lotic habitats, where they are more likely found in slackwater habitats including among floating vegetation. Nonetheless, DeWalt (unpublished) reported large numbers in drift samples from the Brazos River in Texas. Three common species living in open water are P. aquatica, Isotomurus palustris, and Sminthurides aquaticus. Among these species, P. aquatica is associated with ponds, bogs, drains, and is “invariably present on the surfaces of puddles” in England (Hopkin, 1997). Springtails are also found in snowmelt puddles (Deharveng et al., 2008) and springs (Maynard, 1951). Although a surface habitat is more common, some collembolans occur in lake waters as deep as 20 m (Christiansen and Snider, 2008) whereas others are found living interstitially in various habitats or in the surface film of cave habitats (Vandel, 1965), especially Arrhopalites (Christiansen and Snider, 2008). No species occupy the open ocean waters, but some occur along the ocean terrestrial margins. They are well-suited to life on open water surfaces, but some species live instead on the surface of aquatic plants, such as duckweed (e.g., Lemna and Wolffia; Maynard, 1951) and watercress (Nasturtium officinale), or on leaves and other detritus along the water’s edge (Guthrie, 1903). Furthermore, at the extremely small scale at which soildwelling Collembola live, many traditionally “terrestrial” species experience aquatic environments when interstitial spaces are filled with water, and some Collembola are

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found in humid sandy sediments at the level of the water table (Deharveng et al., 2008). When surrounded by water, these species often use hydrophobic hairs, a type of wax, and a specific surface geometry (Noble-Nesbitt, 1963) to create an attached air bubble. Oxygen can diffuse into the air bubble and the springtail can respire across the entire body cuticle (Marx and Messner, 2012). This also increases buoyancy and allows a rapid return to the water surface in a bubble-like fashion (Rapoport and Sánchez, 1963). One marine species, Anurida maritime, uses this property during high tide when it crawls into crevices and retains a bubble of air sufficient for survival underwater for several days if necessary (Maynard, 1951). However, some Collembola may drown when their habitat floods (Mertens et al., 1983). Collembola are constrained in their ability to actively cross water bodies because they lack wings, yet they can float passively downstream (Mouloud et al., 2007) or be blown long distances (Rapoport and Sánchez, 1963).

Physiological Adaptations and Constraints Some species occur at high elevations and in northern latitudes, and they may be active during winter (Guthrie, 1903). Indeed, Cryptopygus antarcticus and Crytopygus sverdrupi, two Antarctic species, can survive temperatures as low as −30 °C (Block, 1981; Somme, 1986). Collembola sometimes face problems from exposure to a wide range of salinities, despite their waterproof integument, which is relatively impermeable to salts (Witteveen, 1988). These problems result because they absorb water and ions via their mouths and abdominal vesicles (Verhoef and ­Witteveen, 1980; Eisenbeis, 1982). However, at least one aquatic species from the genus Anurida can thrive in habitats 2.2 times saltier than seawater (Ring, 1991). Although some Collembola can be highly restricted to the plant or soil environment in which they typically live (Blackith, 1974), P. aquatica has an extremely wide distribution, which may be due to its large tolerance to water temperature and pH (von Heyner, 1972). Wherever they live, springtails are typically very sensitive to desiccation. Indeed, some troglobitic species in the ­ family Hypogastruridae cannot survive more than a few hours below 96% humidity (Thibaud and Christian, 1997). Many terrestrial species have adaptations, behaviors, and strategies that allow for survival in arid climates. For example, some species have very short life histories to avoid arid periods (Greenslade, 1981), desiccation-resistant eggs ­(Poinsot, 1971), anhydrobiosis (Testerink, 1983; ­Verhoef and Li, 1983), and ornamented cuticles that trap ­water-saturated air, resulting in decreasing water loss (King et al., 1990). In addition, aquatic Collembola are not always tied to aquatic environments, and many are found on the terrestrial side of the soil-water ecotone, and vice versa, showing that the ecotone can be “permeable” (Mouloud et al., 2007).

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Most Collembola float on the surface of a liquid (­Hopkin, 1997), even under high pressure and with the addition of surfactants (King et al., 1990) or ethyl alcohol (Rapoport and Sánchez, 1963), because of the properties of their hydrofuge (unwettable) cuticle (Noble-Nesbitt, 1963).

Feeding Behavior Collembolans feed primarily on algae (Moen and Ellis, 1984), detritus (Rapoport and Sánchez, 1963), and other organic material caught on the water surface film or on floating or emergent aquatic vegetation. Consumption of bacteria is also likely, especially given their detrital feeding. One study of a hydrophilous, but nonaquatic springtail used stable isotopes to trace nutrients from added midge carcasses through the springtail detritivores into the remainder of the food web. No other detritivore was found to use these midge carcasses as food, suggesting that aquatic Collembola may occupy an important link in the greater aquatic food web (Hoekman et al., 2012), much like their terrestrial relatives, which are very important detritivores.

Predators and Parasites Collembola are attacked by various invertebrate predators and mostly depend on their springing ability to escape. However, P. aquatica, among many other springtails (see review in Messer et al., 2000), has defensive chemicals (Larsen, 1959; Weinreich, 1968) that can deter some predators, such as the surface-feeding mosquito fish Gambusia affinis, the water strider Microvelia reticulata (Messer et al., 2000), and the staphilinid beetle Stenus comma (Bauer and Pfeiffer, 1991). However, these chemicals have no apparent effect on the European Fire-Bellied Toad, Bombina bombina (Lác, 1958), the European common frog Rana temporaria (Messer et al., 2000), and the fishing spider Dolomedes triton (Zimmermann and Spence, 1989). When young springtails in aquatic habitats subsequently molt, they lose their nonwetting properties for a short time and can be invaded by microorganisms from the surface film (Rapoport and Sánchez, 1963), including a fungal parasite that infects the rectum of P. aquatica (Manier, 1979).

COLLECTING, CULTURING, AND SPECIMEN PREPARATION OF INSECTS AND SPRINGTAILS Collecting and Culturing Collecting Techniques for collecting and preserving springtails and the great diversity of aquatic insects are surprisingly similar, and these are discussed in more detail in Chapters 34–41 of the current volume. Most of the basic information on collecting

any aquatic organism can be found in Chapter 3 of the current volume and in “Chapter 2: General Techniques for Collecting and Identification” in Thorp and Rogers (2011). The first step in any collecting program is to make certain you are collecting legally. Know the laws for collecting on public lands (especially national parks) and get permission before sampling aquatic systems on private lands. The second step is to make certain you are collecting safely. This is especially critical if you are collecting by yourself. Be prepared for accidents and getting wet, particularly if you sample in cold environments. Make sure someone knows where you are going and when you plan to return. Read in advance about the habits of the organisms you seek so that you can efficiently seek the microhabitats where they are likely to occur. For example, this might include seasonality, nocturnal versus diurnal location, current velocity and substrate preferences (on, in, or under rocks, wood, sediment, etc.), depth and distance from shore for aquatic stages, or substrate preferences and distance from water for some adult stages. Choose the right tool for collecting if you are using more than your bare hands. The choice of samplers will depend on the target organism, specific locality, and the need for quantitative or qualitative information. The most common tools you will use are: (1) sweep nets (of various mesh sizes) you purchase from a wildlife supply company or assemble on your own for aquatic and terrestrial insects; (2) mechanical dredges or grabs (e.g., Ekman or Ponar) for shallow or deep water; (3) drift nets to collect insects caught in stream currents; (4) commercial bottom samplers for flowing water (e.g., Surber sampler) or littoral areas (various types of four-sided boxes); and (5) traps for emerging or flying insects. Collected samples may be sorted in the field or later in the laboratory on a white tray. Sometimes it is useful to include a dye (e.g., Phloxine B or Rose Bengal), which will be absorbed by invertebrates, thereby making them easier to see and extract from a background of plant matter or detritus. Collembola can be a challenge to collect because of their springing ability. One way DeWalt et al. (2010) recommended collecting aquatic species was to force them to jump into a white pan containing either 95% ethanol with 3% glacial acetic acid or water with detergent to prevent them from jumping from the pan. Floating sticky traps can also be used to capture them. Preserve springtails in 80–95% ethanol, with 1–2% glycerin added to prevent accidental desiccation (Christiansen and Bellinger, 1980).

Culturing Culturing techniques vary somewhat with insect order but more so with organismal life history, especially developmental patterns and feeding behavior. Chapter 3 discusses

Chapter | 33  Hexapoda—Introduction to Insects and Collembola

some general rules that are applicable to insects and springtails, but consult Chapters 34–41 for more detailed advice.

Specimen Preservation and Preparation In most cases you should preserve aquatic stages in 70–75% ethyl alcohol—commonly called ethanol and abbreviated as ETOH or EtOH. The amount used and whether it is replaced a day later will depend in part on the presence of easily decomposable detritus in the sample. Isopropyl alcohol can also be used if ethanol is not available. It is somewhat less desirable than ethanol because extracting useful DNA from isopropyl-preserved specimens is more difficult; some also feel that ethanol has a less disagreeable odor. However, isopropyl alcohol can typically be obtained more easily at the last minute from local pharmacies or other public stores, at least in the United States. See Chapter 3 for instructions on preparing labels for specimens. Make sure that your specimens are properly labeled for future consultation, especially if you plan to place the specimen in a museum. Consult Chapter 3 of this volume for details. Although you will preserve most aquatic insects in ETOH, adult insects in many orders (e.g., Odonata (dragonflies and damselflies; Chapter 35) and some Coleoptera (beetles; Chapter 39)) are more appropriately pinned as dry mounts, as described in the relevant chapters. Very small insects and springtails may be preserved on glass slides for better viewing and identification. For example, to identify the genus of a larval midge (Diptera, Chironomidae; Chapter 40), the head capsule needs to be mounted on a slide and the mouthparts spread. Some insects can be identified to species at the larval level (e.g., many caddisflies) whereas others require adult stages for species identification (e.g., most Diptera). Identification of species from larval specimens requires the presence of distinguishing characteristics in the larvae and a published link between larval and aquatic stages. This latter link is missing in many species, especially dipterans.

ACKNOWLEDGMENTS Some of the information on springtails was derived from Chapter 16 “Diversity and Classification of Insects and Collembola” in the preceding (third) edition of the invertebrate volume edited by Thorp and Covich (2010). We appreciate access to that fine chapter that was written by R. Edward DeWalt, Vincent H. Resh, and the late William L. Hilsenhoff.

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Manier, J.F., 1979. Orchesellaria podurae n. sp. (Trichomycete, Asellariale) parasite de Podura aquatica L. (Insecte, Apterygote, Collembole). Rev. Mycol. 43, 341–350. Marx, T.M., Messner, B., 2012. A general definition of the term “plastron” in terrestrial and aquatic arthropods. Org. Divers. Evol. 12, 403–408. Maynard, E., 1951. The Collembola of New York State. Comstock Publishing Company, Inc, Ithaca, NY. McCafferty, W.P., 1981. Aquatic Entomology: the Fishermen’s and Ecologists’ Illustrated Guide to Insects and Their Relatives. Jones and Bartlett Publishers, Sudbury, MA. 448 pp. Merritt, R.W., Cummins, K.W., Berg, M.B. (Eds.), 2008. An Introduction to the Aquatic Insects of North America, fourth ed. Kendall/Hunt, Dubuque, IA, p. 1158. Mertens, J., Coessens, R., Blancquaert, J.P., 1983. Reproduction and development of Hypogastrura viatica in relation to temperature and submerged condition. Rev. d’Écologie Biol. sol. 20, 567–577. Messer, C., Walther, J., Dettner, K., Schulz, S., 2000. Chemical deterrents in podurid Collembola. Pedobiologia 44, 210–220. Moen, P., Ellis, W.N., 1984. Morphology and taxonomic position of Podura aquatica (Collembola). Entomol. Generalis 9, 193–204. Mouloud, S.A., Lek-Ang, S., Deharveng, L., 2007. Fine scale changes in biodiversity in a soil-water ecotone: Collembola in two peat-bogs of Kabylia (Algeria). Vie Et. Milieu – Life Environ. 57, 149–157. Noble-Nesbitt, J., 1963. Transpiration in Podura aquatica L. (Collembola, Isotomidae) and the wetting properties of its cuticle. J. Exp. Biol. 40, 681–700. Peckarsky, B.L., 1982. Aquatic insect predator-prey relations. BioScience 32, 261–266. Peckarsky, B.L., Fraissinet, P.R., Penton, M.A., Conklin, Jr., D.J., 1990. Freshwater Macroinvertebrates of Northeastern North America. Cornell University Press, Ithaca, NY. Paclt, J., 1956. Biologie der flügellosen insekten. Gustav Fischer Verlag, Jena. Poinsot, N., 1971. Ehtologie de quelques espéces de Collemboles Isotomides De Prevence. Ann. l’Université Provence, Sci. 45, 33–53. Rapoport, E.H., Sánchez, L., 1963. On the epineuston or the superaquatic fauna. Oikos 14, 96–109. Regier, J.C., Shultz, J.W., Zwick, A., Hussey, A., Ball, B., Wetzer, R., Martin, J.W., Cunningham, C.W., 2010. Arthropod relationships revealed by phylogenomic analysis of nuclear protein-coding sequences. Nature 463, 1070–1084. Resh, V.H., Cardé, R.T. (Eds.), 2009a. Encyclopedia of Insects. second ed. Academic Press, San Diego, CA. Resh, V.H., Cardé, R.T., 2009b. Insecta, overview. In: Resh, V.H., Cardé, R.T. (Eds.), Encyclopedia of Insects. Academic Press, San Diego, CA, pp. 501–502. Resh, V.H., Rosenberg, D.M., 1984. The Ecology of Aquatic Insects. Praeger, NY. 626 pp. Resh, V.H., Buchwalter, D.B., Lamberti, G.A., Eriksen, C.H., 2008. Aquatic insect respiration. In: Merritt, R.W., Cummins, K.W., Berg, M.B. (Eds.), An Introduction to the Aquatic Insects of North America, fourth ed. Kendall Hunt, Dubuque, IA, pp. 39–54. Ring, R.A., 1991. The insect fauna and some other characteristics of natural salt springs on Saltspring Island, British Columbia. Memoirs Entomol. Soc. Can. 155, 51–61. Ross, H.H., 1967. The evolution and past dispersal of the Trichoptera. Annu. Rev. Entomol. 12, 169–206. Schliwa, W., Schaller, F., 1963. Die paarbildung des springschwanzes Podura aquatica (Apterygota [Urinsekten], Collembola). Naturwissenschaften 50, 698.

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Chapter 34

Order Ephemeroptera Michel Sartori Museum of Zoology, Palais de Rumine, Lausanne, Switzerland

John E. Brittain Natural History Museum, University of Oslo, Oslo, Norway

Chapter Outline Introduction To Mayflies (Ephemeroptera) 873 Brief History and Paleontology 873 General Systematics 874 Phylogenetic Relationships 874 Distribution, Diversity, and Endemism 876 General Biology 877 External Anatomy of Imagos and Nymphs 877 Winged Stages 877 Nymphs878 Internal Anatomy of Nymphs 880 Life Cycle 880 Behavior of the Winged Stages 881 Emergence881 Subimaginal Stage 883

INTRODUCTION TO MAYFLIES (EPHEMEROPTERA) Brief History and Paleontology Extant Ephemeroptera represent what is left of a much diversified group of primitive flying insects (Ephemerida), the origin of which goes back to the Carboniferous. Permian data confirm that the group was already present at the end of the Paleozoic. Ephemerida reached their greatest diversity during the Mesozoic, mainly in the Jurassic and Cretaceous. All of these species share the presence of a costal brace at the base of the forewing and a reduction in the anal region of the hindwing with modern mayflies. However, contrary to them, they had homonomous wings (i.e., fore- and hindwing of the same size), and their aquatic stages could possess up to nine pairs of abdominal gills (compared with a

Flight Activity 883 Reproduction884 General Ecology and Behavior 884 Habitat Selection 884 Physiological Constraints 885 Feeding Behavior 885 Other Relevant Behavior 885 Predators886 Parasitic and Commensal Relationships 886 Environmental Changes and Human Effects 886 Mayfly Interactions with Humans 887 Collecting, Rearing, and Specimen Preparation 888 References888

maximum of seven in extant species). Some species also had a wing span over 90 mm. All of these lineages, including Permoplectoptera (e.g., Protereismatidae or Misthodotidae), went extinct by the end of the late Jurassic. A recent study described adults and nymphs of a peculiar fossil insect order, the Coxoplectoptera, which could be the true sister group of modern Ephemeroptera (Staniczek et al., 2011). Although the adults have homonomous wings, the nymphs possess seven pair of gills as in the modern mayflies, a single tarsal segment (compared with five tarsal segments in the nymphs of Protereismatidae), and a single pretarsal claw (compared with paired claws in Proteiresmatidae). Heteronomous mayflies with reduced hindwings had appeared by the end of the Jurassic. The Tertiary fauna, as documented by fossils in Baltic or Dominican amber, is definitely contemporary with the presence of extinct and living genera of modern families.

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385026-3.00034-6 Copyright © 2015 Elsevier Inc. All rights reserved.

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General Systematics Ephemeroptera constitutes a small order of extant insects, with approximately 40 families, 440 genera, and 3330 species (Table 34.1). The state of our knowledge varies considerably depending on the geographic region. Some areas of North America and Europe are well known, whereas others, such as Southeast Asia or the Neotropics, still harbor numerous undescribed species. Within the period 2009–2011, almost 150 new species were described: more than 70 from South America and only 8 from North America. This can also be seen by comparing Table 34.1 with those published in Barber-James et al. (2008) and Brittain and Sartori (2003, 2009). We estimate that around 1000 species are still waiting to be described worldwide. The increasing use of genetic tools is also bringing new insight into mayfly systematics, which may potentially boost the number of taxa (Monaghan and Sartori, 2009). The supraspecific taxonomy has been the subject of major changes during the last 20 years, mainly because of the gathering of species into a more phylogenetic system, leading to a narrower concept of genus and family. As a result, the number of genera increased by 30% during this period because of the phylogenetic rearrangement of species groups, together with the discovery of new taxa, especially in the tropics. As shown in Table 34.1, the distribution of species among families is rather unequal. Because of their ancient origin (see previous section), we can represent ephemeropteran evolution as a baobab tree, with a large trunk, numerous broken ramifications, but few actual branches, some with very few leaves and others with bunches of boughs and dense foliage. Hence, fewer than 50% of the known species and genera belong to the families Baetidae and Leptophlebiidae, whereas 17 families are monogeneric, 8 of which are also monospecific. Several genera are particularly species rich, and the 12 richest encompass approximately one third of all known species (Table 34.2).

Phylogenetic Relationships The relationships of Ephemeroptera, Odonata, and Neoptera are one of the major unsolved problems in entomology (Blanke et al., 2012). Traditionally, Ephemeroptera and Odonata were clustered in the clade Paleoptera, characterized by wings unable to be folded against the body at rest. This clade was considered as the sister group of Neoptera. However, alternative theories have also been proposed, including the Metapterygota and Chiastomyaria hypotheses. The former suggests a basal position of Ephemeroptera compared with Odonata + Neoptera, whereas the latter hypothesizes that Odonata occupies a basal position compared with Ephemeroptera + Neoptera.

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All of these hypotheses were proposed on morphological and/or molecular data, but no consensus exists at the moment, although recent studies bring new evidences for the Paleoptera hypothesis (Blanke et al., 2013; Thomas et al., 2013). Deciphering the relationships among extant Ephemeroptera still attracts attention, and the higher classification is also now a matter of debate. McCafferty and Edmunds (1979) proposed two suborders: Schistonota (nymphs with free wing pads) and Pannota (nymphs with basally fused wing pads). It soon transpired that the Schistonota was paraphyletic, and McCafferty (1991) proposed a new classification, including three suborders (Setisura, Pisciforma and Rechtracheata, with the infraorder Pannota). Several modifications were put forward by McCafferty in subsequent years, in addition to the work of Kluge (e.g., 2004). All of these studies were based on morphology. The results were quite congruent and are summarized in Ogden and Whiting (2005) and Ogden et al. (2009). McCafferty and Kluge recognize a basal suborder, including the families Baetiscidae and Prosopistomatidae, characterized by the development of a mesonotal shield in the nymphs (hence the name Carapacea given by McCafferty) and the peculiar position of the anal veins in the forewings (hence the name Posteritorna given by Kluge). Three other suborders were then redefined as Furcatergalia (including Pannota, Ephemeroidea and Leptophlebiidae), Setisura (Heptagenioidea) and Pisciforma (including Baetoidea and Siphlonuroidea). Ogden and Whiting (2005) proposed the first molecular phylogeny, followed by Ogden et al. (2009), who performed a combined analysis of five genes (5880 bp) and 101 morphological characters on 112 species in 107 genera and 42 families. The results are quite different from those that are based solely on morphology, although taxa such as Furcatergalia or Pannota were found to be monophyletic (Figure 34.1). The suborder Carapacea is not basal, but nested with the family Oligoneuriidae at the base of Furcatergalia, whereas the suborders Setisura and Pisciforma are highly paraphyletic, as is the wellestablished superfamily Baetoidea (Baetidae + Siphlaenigmatidae). Three families are basal to the leftover taxa: Siphluriscidae, Baetidae (Pisciforma), and Isonychiidae (Setisura). The nymph of Siphluriscus chinensis Ulmer, 1920 (Siphluriscidae) has been described recently and exhibits archaic morphological conditions, making it a good candidate to represent the oldest extant lineage (Zhou and Peters, 2003), confirmed by its position as the sister taxon to all other Ephemeroptera. The position of Baetidae and Isonychiidae is contradictory to previous hypotheses; however, as demonstrated by Ogden et al. (2009), the inclusion of these families in Pisciforma and Setisura, respectively, is based on homoplasies or plesiomorphic conditions. One of the main conclusions is that

Chapter | 34  Order Ephemeroptera

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TABLE 34.1  List of the Extant Families, with Number of Genera and Species in the Different Biogeographic Realms, According to Barber-James et al. (2008), as of July 2012 Family

# Genera

PAL

NEA

NEO

ORI

AFR

AUS

PAC

# Species

Distribution Complement

Acanthametropodidae

2

1

2

0

0

0

0

0

3

Ameletidae

2

17

35

0

5

0

0

0

56

Ameletopsidae

4

0

0

2

0

0

4

0

6

Ametropodidae

1

1

3

0

0

0

0

0

3

Austremerellidae

1

0

0

0

0

0

1

0

1

104

212

137

239

150

194

43

3

956

Baetiscidae

1

0

12

0

0

0

0

0

12

Behningiidae

3

3

1

0

2

0

0

0

6

26

43

36

37

33

61

13

1

221

Coloburiscidae

3

0

0

1

0

0

6

0

7

Amphinotic

Coryphoridae

1

0

0

1

0

0

0

0

1

Brazil

Dicercomyzidae

1

0

0

0

0

4

0

0

4

Continental Africa

Dipteromimidae

1

2

0

0

0

0

0

0

2

Japan

Ephemerellidae

22

53

65

0

32

0

0

0

148

Ephemeridae

7

11

16

3

39

13

0

0

80

Ephemerythidae

1

0

0

0

0

3

0

0

3

Euthyplociidae

7

0

0

7

3

10

0

0

20

Heptageniidae

33

306

128

5

140

21

1

0

598

Ichthybotidae

1

0

0

0

0

0

2

0

2

Isonychiidae

1

14

16

1

4

0

0

0

34

Leptohyphidae

14

0

29

124

0

0

0

0

145

Leptophlebiidae

141

58

69

247

68

52

120

38

643

Machadorythidae

1

0

0

0

0

1

0

0

1

Continental Africa

Melanemerellidae

1

0

0

1

0

0

0

0

1

Brazil

Metretopodidae

3

4

9

0

0

0

0

0

11

Neoephemeridae

3

4

4

0

6

0

0

0

14

Nesameletidae

3

0

0

1

0

0

6

0

7

Oligoneuriidae

12

12

8

23

2

11

0

0

55

Oniscigastridae

3

0

0

2

0

0

6

0

8

Palingeniidae

6

10

0

0

14

4

4

0

32

Polymitarcyidae

6

6

7

60

12

2

0

0

85

Potamanthidae

3

7

4

0

13

0

0

0

24

Prosopistomatidae

1

3

0

0

11

6

2

0

22

Rallidentidae

1

0

0

0

0

0

1

0

1

New Zealand

Siphlaenigmatidae

1

0

0

0

0

0

1

0

1

New Zealand

Siphlonuridae

4

25

25

0

0

0

0

0

48

Baetidae

Caenidae

Amphinotic

Australia

Continental Africa

New Zealand

Amphinotic

Amphinotic

Continued

SECTION | VI  Phylum Arthropoda

876

TABLE 34.1  List of the Extant Families, with Number of Genera and Species in the Different Biogeographic Realms, According to Barber-James et al. (2008), as of July 2012—cont’d Family

# Genera

PAL

NEA

NEO

ORI

AFR

AUS

PAC

# Species

Siphluriscidae

1

1

0

0

0

0

0

0

1

Teloganellidae

1

0

0

0

1

0

0

0

1

Teloganodidae

8

0

0

0

13

8

1

0

22

Tricorythidae

5

0

0

0

7

29

1

0

37

Vietnamellidae

1

0

0

0

6

0

0

0

6

441

793

606

754

561

419

212

42

3328

Total

Distribution Complement China

Southeast Asia

PAL, Palaearctic; NEA, Nearctic; NEO, Neotropical; ORI, Oriental; AFR, Afrotropical; AUS, Australasian; PAC, Pacific Islands. Note that the total numbers of species in rows and columns are not necessarily equal because of the occurrence of some species in several realms.

TABLE 34.2  The Twelve Most Speciose Mayfly Genera as of July 2012 Genus

Family

Baetis Leach, 1815

Baetidae

152

Worldwide

Rhithrogena Eaton, 1881

Heptageniidae

152

Holarctic and Oriental

Caenis Stephens, 1835

Caenidae

141

Worldwide

Epeorus Eaton, 1881

Heptageniidae

93

Holarctic and Oriental

Cloeon Leach, 1815

Baetidae

74

Worldwide

Pseudocloeon Klapalek, 1905/Labiobaetis Kluge and Novikova, 1987

Baetidae

73

Worldwide except Neotropical

Afronurus Lestage, 1924

Heptageniidae

64

Oriental; Afrotropical and Palaearctic

Ecdyonurus Eaton, 1868

Heptageniidae

61

Holarctic and Oriental

Tricorythodes Ulmer, 1920

Leptohyphidae

59

Panamerica

Thraulodes Ulmer, 1920

Leptophlebiidae

55

Panamerica

Paraleptophlebia Lestage, 1917

Leptophlebiidae

54

Holarctic and Oriental

Ameletus Eaton, 1885

Ameletidae

53

Holarctic and Oriental

Total

our understanding of mayfly phylogeny is hampered by morphological convergences in many features. The exact status of “­Siphlonuroidea” and “Heptagenioidea” still needs to be resolved, and further studies including more genes are necessary.

Distribution, Diversity, and Endemism Mayflies are distributed throughout the world, colonizing freshwater and sometimes brackish waters on all continents except Antarctica. Their presence on islands is explained on one hand by vicariance processes, induced, for instance, by the

Species #

Distribution

1031

Gondwana break up (New Caledonia, Seychelles, Sri Lanka), but also by dispersal events on continental islands (i.e., Madagascar) or oceanic islands (e.g., Macaronesia, la Reunion). Dispersal by mayflies and colonization of new habitats has long been considered a rare phenomenon, but there is a growing record of data that prove some species can disperse over at least 700 km. Thus, mayflies are only absent from remote islands, such as the Tristan da Cunha Archipelago and Gough Island (Barber-James, 2007) in the southern Atlantic Ocean or the Galapagos Islands and Polynesia in the Pacific Ocean, where distance and the lack of suitable habitats explain their probable absence. Accidental introduction by human activities

Chapter | 34  Order Ephemeroptera

FIGURE 34.1  Phylogenetic relationships among extant mayfly families; nodes with former appropriate names are mentioned. From Ogden et al., 2009.

877

Endemism in mayflies is a function of the history of the lineage under study and the ecological requirement of their nymphs. A good example is provided by Malagasy fauna, in which the Baetidae have been genetically studied (Monaghan et al., 2005a). If all but one species are endemic to the island, the lineages that are issued from vicariant processes are composed of endemic genera (paleoendemism), whereas those issued from dispersal processes included cosmopolitan or tropical genera (neoendemism). The nonendemic species is member of the genus Cloeon, the nymphs of which inhabit pools, swamps, or even water tanks. The females are ovoviviparous, living unusually longer than other species and thus are more amenable to disperse actively or passively. Species endemism is high in mayflies, reaching almost 100% in Australasia and Africa. Fewer than 60 species (180 spp. (3/43 in Nearctic); Dixa, Dixella

3 genera, ≈80 spp. (3/16 in Nearctic); Dilated tarsomere aids in aerial drift dispersal (bittacomorBittacomorpha, Ptychoptera phines)

12 genera, ≈50 spp. (2/4 in Nearctic); Presumed rare, but may be locally abundant Mischoderus, Protanyderus, Protoplasa

Systematics and Exemplar Genera

Cosmopolitan, ≈35 genera, ≈330 spp. (4/40 in mountainous regions Nearctic); Agathon, Blepharicera

Cosmopolitan

Cosmopolitan

Cosmopolitan

Holarctic, Australasian, Neotropical

Cosmopolitan

Mostly tropical and subtropical

Cosmopolitan

Holarctic, Australasian, Neotropical, Oriental

Holarctic, Oriental, Afrotropical

Nearctic, Australasian Neotropical, Afrotropical

Distribution

Chapter | 40  Order Diptera 1045

Common Name

Larval Habitat

Moth flies Sand flies

Crane flies

Psychodidae

Tipuloidea

Pelecorhynchid flies

Horse flies Deer flies

Oreoleptid flies

Water snipe flies

Soldier flies

Pelecorhynchidae

Tabanidae

Oreoleptidae

Athericidae

Stratiomyiidae

Brachycera

Axymyiid flies

Axymyiidae

Distribution

Predator of other invertebrates

Predator of other invertebrates

Diverse: Predators, shredders, collector– gathers

Scrapers, collector– gatherers

Collector–gatherer

Known only from W. Nearctic

Cosmopolitan

Holarctic, Australasian

Cosmopolitan

Cosmopolitan

Holarctic, rarely collected

Collector–gatherers

Cosmopolitan

Cosmopolitan? Lotic, often assoc. With Predator of other insects (e.g., chironofilamentous algae mids) Hot springs, madicolous, terrestrial

Systematics and Exemplar Genera

Lays eggs above H2O, dies— others repeat, get large ball Adults capable of hovering in one spot for extended periods

>380 genera, ≈2700 spp. (13/≈190 aquatic spp. in N America); Euparyphus, Odontomyia, Oxycera

Larvae pupate in areas above water-line at emergence 12 genera, >130 spp. (2/4 in Nearctic); Atherix, Suragina

1 species, Oreoleptis torrenticola

Female hematophagous on warm and cold-blooded vertebrates

>150 genera, ≈4400 spp. (14/>330 aquatic spp. in Nearctic); Chrysops, Hybomitra, Silvius, Tabanus

Largest Diptera “family”

≈250 genera, >16,000 spp. (≈30/≈300 aquatic spp. in Nearctic); Antocha, Hexatoma, Limonia, Lipsothrix, Ula

Some adults feed on flowers

Important in trickling filters

≈140 genera, >3000 spp. (6/67 aquatic spp. in Nearctic); Clogmia, Maruina, Neotelmatoscopus, Pericoma

2 genera, ≈50 spp. (1/7 in Nearctic); Glutops

Locally abundant in small streams and seepages

Shed wings to oviposit underwater; mating pairs couple for life

Miscellaneous Information

3 genera, 6 spp. (2 in Nearctic); Axymyia

Grazers (?), Collector– Oriental, E. Nearctic, 1 genus, 8 spp. (2 in Nearctic); W. Palaearctic, gatherers Nymphomyia “uncommon”

Ecological Role of Larvae

Lotic, coarse-bottomed Predator of other invertebrates riffles; some from groundwater

Wetland soils, stream margins

Gravelly streambeds, saturated soil of swamps

Diverse: Lentic, lotic, marine intertidal, wood, terrestrial

Marshes, tree holes, madicolous

Saturated wood that is partly submerged

Nymphomyiidae Nymphomyiid flies Small stenothermal streams, mosscovered rocks

Taxon

TABLE 40.1  Synopsis of Aquatic/Semiaquatic Diptera—cont’d

1046 SECTION | VI  Phylum Arthropoda

Dance flies

Long-legged flies

Flower flies Hover flies

Seaweed flies

Dryomyzid flies

Snail-killing flies

Shore flies

Beach flies

Flesh flies

House flies

Empidoidea

Dolichopodidae

Syrphidae

Coelopidae

Dryomyzidae

Sciomyzidae

Ephydridae

Canacidae

Sarcophagidae

Muscidae

Widespread, esp. Around Pacific

Cosmopolitan

Cosmopolitan

Holarctic

Holarctic, Australasian

Cosmopolitan

Cosmopolitan

Cosmopolitan, Fletcherimyia Fletcherimyia widecollector–gatherers spread but disjunct or scavenger of trapped invertebrates

Marine species detritivores, FW spp. grazers (scrapers)

Diverse, but usually collector–gatherers

Predator of snails, slugs, snail eggs

Feed on seaweed or predaceous on barnacles

Assumed to be collector–gatherers of rotting seaweed

Aquatic spp. lentic and Collector–gatherers lotic–depositional, moss or predators (latter = Limnophora)

Mostly terrestrial; Fletcherimyia a burrower/miner in pitcher plants

Marine intertidal rotting seaweed

Diverse lentic, brackish, alkaline

Marshes, banks of ponds, streams

Marine intertidal rotting seaweed inside barnacles

Marine intertidal rotting seaweed

Cosmopolitan

Predator of other Cosmopolitan insects (e.g., culicids)

Predator of other insects (e.g., simuliids)

Usually collector– Lentic, tree holes, detrital accumulations, gatherers saturated wood, many terrestrial

Lake and stream margins, depositional zones

Lentic, lotic– depositional, moss, terrestrial

Helaeomyia petrolei breeds in pools of crude petroleum Few FW species known (all from torrential streams) Viviparous or ovoviviparous

≈130 genera, ≈2000 spp. (72/≈470 spp. in Nearctic); Hydrellia, Parydra 28 genera, ≈320 spp. (5/14 in Nearctic); Canace, Nocticanace >170 genera, ≈3100 spp. (2/7 aquatic spp. in Nearctic); Fletcherimyia, Sarcophaga ≈190 genera, >5200 spp. (7/>270 aquatic spp. in Nearctic); Limnophora

Puparium often adapted to fit in snail

Rat-tailed maggots in liquid manure

66 genera, ≈620 spp. (20/175 in Nearctic); Sepedon, Tetanocera

6 genera, 30 spp. (2 aquatic spp. in Nearctic); Oedoparena

14 genera, 35 spp. (2/5 in Neartic); Coelopa, Coelopina

≈210 genera, ≈6100 spp. (20/>130 aquatic spp. in Nearctic); Eristalis, Chrysogaster, Copestylum, Temnostoma

≈270 genera, ≈7400 spp. (44/1130 in Mating behavior of some Nearctic); Hydropharus, Tachytrechus species similar to empidoids

7 subgroups (Families), >210 genera ≈5400 spp. (15/≈300 aquatic spp. in Nearctic); Chelifera, Oreogeton

Chapter | 40  Order Diptera 1047

SECTION | VI  Phylum Arthropoda

1048

E

D

F

G

H

I

J

K

L

FIGURE 40.2  Exemplar larvae of aquatic nematocerans: (a) Deuterophlebia (Deuterophlebiidae) dorsal view; (b) Blepharicera (Blephariceridae) dorsal view; (c) Cnesia (Simuliidae); (d) Bezzia/Palpomyia (Ceratopogonidae) head, thorax, and abdominal segment 1, ventral view; (e) Bittacomorpha (Ptychopteridae) head, thorax, and abdominal segments 1–3, lateral view; (f) Eucorethra (Chaoboridae) head, thorax, and abdominal segment 1, ventral view; (g) Maruina (Psychodidae) dorsal view; and (h) Toxorhynchites (Culicidae), lateral view; (i) Axymyia (Axymyiidae), dorsal view. Images by S.A. Marshall (a–c, g) and G.W. Courtney (d–f, h, i)

Chapter | 40  Order Diptera

1049

(b)

(a)

(c)

(d)

(f)

(e)

(g)

FIGURE 40.3  Exemplar larvae of aquatic Brachycera: (a) Tabanus (Tabanidae), lateral view; (b) Odontomyia (Stratiomyiidae), oblique dorsal view; (c) Atherix (Athericidae), dorsal view; (d) Eristalis (Syrphidae), lateral view); (e) Fletcherimyia (Sarcophagidae), lateral view; (f) Ephydra (Ephydridae), lateral view; and (g) Limnophora (Muscidae), dorsal view. Images by G.W. Courtney

The body of most Diptera larvae is soft and flexible, comprising three variably distinct thoracic segments, with the abdomen having eight to nine segments (Figures 40.2 and 40.3). In most Chironomidae, Tipuloidea, and Simuliidae, the

body is subcylindrical (Figure 40.2(c, e, and h)). The body of other dipterans, especially those from interstitial (hyporheic) habitats, is elongated/serpentine (Figure 40.2(d)). Cyclorrhaphan larvae are maggot-shaped (Figure 40.3(e and g)).

SECTION | VI  Phylum Arthropoda

1050

Because larvae lack jointed thoracic legs, their locomotion requires controlled turgor pressure, including via creeping welts, parapods (prolegs), friction pads, and suctorial discs. Creeping welts, characteristic of certain Tipulidae, Tabanidae, Empididae, and Ephydridae (Figure 40.3(a, e, and f)), are transverse, swollen areas (ridges) that bear modified setae or spines. Parapods usually are paired, round, elongate, fleshy, retractable, and bear apical spines or crochets. These are typical of larvae in the families Chironomidae, Deuterophlebiidae, Nymphomyiidae, Simuliidae, Thaumaleidae, Athericidae, Oreoleptidae, and some others (e.g., Figure 40.2(a, b, and e)). Several larval Psychodidae (e.g., Figure 40.2(g)) possess friction pads (modified ventral body cuticle), as do certain hygropetric Ephydridae. Larval Blephariceridae resist torrential water flow with true suction devices (ventral discs). The variety of mechanisms for gas exchange in larval aquatic Diptera reflects the physics of life in fluid or semifluids. The basic system comprises external spiracles (openings) connected to internal tracheae (tubes) and finer tracheoles. Gaseous exchange may be direct with the atmosphere, via oxygenated fluids, or even derived from plant tissues. Hemoglobin in the hemolymph of some larval Chironomidae extracts and stores oxygen from oxygendepleted habitats. Many aquatic larvae, particularly those from well-oxygenated streams, lack spiracles and all gas exchange is transcuticular. Larvae of some families (e.g., Psychodidae) possess spiracles on the prothorax and last abdominal segment, whereas others (e.g., most cyclorrhaphans) have spiracles only terminally. In larval Culicidae, Ptychopteridae, many Ephydridae, and some Syrphidae (Figures 40.2(h and i) and 40.3(d and f)), posterior spiracles are located apically on a siphon (which may be retractile), allowing independence from dissolved oxygen. Unusually in larval Coquillettidia (Culicidae), a specialized siphon pierces the vascular system of submerged aquatic plants to obtain oxygen.

Pupal Morphology A pupa forms following completion of larval development and contains the metamorphosing adult (in pharate “cloaked” condition). The pupa shows features of both larva and adult, or appears relatively formless. Pupal nematocerans have an identifiable head, thoracic, and abdominal segments. The head and thoracic appendages (sheaths of pharate adult antenna, legs, and wings) are often fused to the body (Figure 40.4(d and f)). Externally nematoceran pupa may be adorned with spines, often transversely arranged on ridges, gill-like gas-exchange devices, and locomotory paddles (Figure 40 4(a–d)). Gas exchange organs vary and may include elongate lobes (e.g., in most Tipuloidea, Culicomorpha, Psychodidae), branched appendages (e.g., Deuterophebiidae, Simuliidae, many Chironomidae and some

Tipuloidea), or plate-like lamellae (e.g., most Blephariceridae). The thoracic organs of pupal Ptychopteridae are asymmetric, with one greatly elongated, allowing pupal submergence in anoxic sediments. Pupal gas exchange organs may contain specialized struts to maintain a layer of gases (i.e., an incompressible gill or plastron). Pupae of many aquatic Diptera (e.g., Culicidae) are mobile, using locomotory structures (e.g., paddles on the posterior abdomen) to swim (e.g., Figure 40.4(b)). Other pupae remain attached to the substrate (e.g., Figure 40.4(a)) via either a cocoon or tube spun with salivary gland silk (e.g., Simuliidae, many Chironomidae, and Tipuloidea) or by specialized adhesive discs (e.g., Blephariceridae, Deuterophlebiidae, and certain Psychodidae). Brachyceran pupae are more concealed and are compact or contracted (Figure 40.4(e–h)). In Cyclorrhapha (e.g., Syrphidae, Ephydridae), the pupa is a “puparium” comprising hardened, tough, desiccation-resistant cuticle of the final larval instar (Figure 40.4(g and h)). The enclosed (pharate) adult eventually breaks out by inflating a balloon-like structure (the ptilinum) on the anterior head. Few external features are evident on the puparium, although spiracles and surface spines are present.

Life Cycles Development is holometabolous with complete metamorphosis—the dipteran life-cycle comprises distinct ­ stages or instars separated by molts. A typical life-cycle consists of an egg, at least three larval instars (typically three in Brachycera and four in nematocerous Diptera, but more in Simuliidae, Thaumaleidae, Tabanidae and a few others), a pupa, and then an adult (imaginal) stage (Figure 40.5). The aquatic dipteran egg has microstructure typical for an insect, comprising an external shell of vitelline membrane and chorion, whose structure varies with the environment (e.g., whether laid terrestrially or within the water). Larval development within the egg varies according to temperature and physiological constraints, including the existence of a delay (diapause) in some taxa. The first larval stage—the larvula—develops within the egg capsule and often has doubled back on itself by the time it hatches. An “egg-burster” or “egg tooth” may aid the mandibles in breaking the chorion. For flies that lay massed eggs (a raft), the surrounding gelatinous matrix feeds the emerged larvula, which is a short-lived, dispersive stage. Larval development depends on environmental conditions, lasting as briefly as a few days to as long as several years; it can be interrupted by diapause in adverse conditions. At consecutive molts, larval size increases predictably (following Dyar’s Law). In the final larval instar, the developing pupa includes adult primordia (buds) of wings and legs, cephalic features, and prominent gas exchange organs such as the pupal thoracic “gills.” The contents of

Chapter | 40  Order Diptera

1051

(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

FIGURE 40.4  Exemplar pupae of Diptera: (a) Blepharicera (Blephariceridae), dorsal view; (b) Toxorhynchites (Culicidae), lateral view; (c) Tipula (Tipuloidea) head, thorax, and abdominal segments 1–3, lateral view; (d) Tipula (Tipuloidea) head, thorax, and abdominal segments 1–3, ventral view; (e) Tabanus (Tabanidae) lateral view; (f) Tabanus (Tabanidae) head, thorax, and abdominal segments 1–2, ventral view; (g) Ephydridae puparia, lateral view; and (h) Acalyptrate (Sphaeroceridae?) puparia, dorsal (above), and ventral (below) views. Images by G.W. Courtney

SECTION | VI  Phylum Arthropoda

1052

FIGURE 40.5  Schematic drawing of the life cycle of a common midge (Chironomidae) showing the various events and stages of development. From Gullan and Cranston, 2010

the late, final larval instar is a pharate (cloaked) pupa, which is revealed fully only after the pupation molt. The final molt may involve no relocation by the final larval instar, but often a specific and different microhabitat is sought. In lotic flies, this may include depressions on the upper, currentexposed faces of rocks or the splash zones on emergent rocks. In some rheophiles (e.g., Blephariceridae, Simuliidae), streamlined pupae orient with the thick, anterior end directed downstream, generating a downstream, oxygenated water vortex across gas exchange organs. The duration of the pupal stage varies with taxon and temperature, but this can be a day in very small Chironomidae to several weeks in larger and cryptic taxa such as Tabanidae. Emergence of the adult (imago) in some groups (e.g., Blephariceridae, Simuliidae) requires the pupa to be attached firmly to the substrate. The adult emerges by splitting the pupal cuticle along lines of weakness (the ecdysial lines) by applying downward pressure against the substrate. Observations of laboratory-reared adults indicate that some adults (e.g., some Blephariceridae, males of all Deuterophlebiidae) must rely on flow and/or an air bubble. Emergence in running water is rapid; most fly immediately with wings fully inflating within the pupa, whereas the creased wings of others unfold during emergence. In some taxa, balloon-like teneral wings containing hemolymph can carry the emerged adult to a resting site.

Reproduction Swarming and Mating Diptera swarms usually consist of male imagoes. These commonly seen aggregations lure prospective female mates, and are located with respect to markers. These markers may be along roadsides, above sunlit pools or riffles along streams, over selected trees or bushes, at hilltops, in sunny

gaps of forest canopies, or at any number of other visible features. The contrast between the swarm marker and background seems to involve polarized light rather than specific levels of illumination; this may be critical in determining the species-specific swarm location. Swarming and aerial mating are probably fundamental features of the Diptera, and their two-winged condition may relate to swarming and aerial mating. Male flies share features adaptive for swarming, including plumose antennae and Johnston’s organ for sound reception (e.g., many Culicomorpha) and divided, holoptic eyes (e.g., most Simuliidae, Blephariceridae, and Bibionidae). Females perceive the male-dominated swarms and mate association takes place, with aerial pairing before exiting the swarm. Swarming is well developed in certain brachycerans, notably in dance flies (Empididae) known for their predaceous habits, elaborate behaviors and “nuptial gifts.” Agile Syrphidae and Stratiomyiidae are adept at hovering. In extreme environments, aerial coupling may be replaced by surface mating; this is accompanied by wing reduction and genitalic rotation.

Oviposition The distribution of immature Diptera depends on site selection by ovipositing females. For many aquatic flies, eggs are laid in masses, located nonrandomly, close to the water surface, on emergent rocks or trailing vegetation. Many insects, including tabanids and chironomids, locate sites by horizontally polarized light. Unfortunately, this can cause them to be misled, for example, by wet roads, shiny vehicles, and newly-laid road tar. Unusual oviposition strategies include Athericidae laying enormous masses on overhanging terrestrial vegetation, or on man-made structures above running waters into which hatchlings drop. Similarly, some (perhaps many) Prosimulium (Simuliidae) lay egg masses (sometimes from several individuals and comprising up to

Chapter | 40  Order Diptera

three species) in terrestrial moss or bryozoans, with larval development following autumnal rains. In some lotic groups (e.g., Deuterophlebiidae and Nymphomyiidae, some Blephariceridae), females crawl underwater to oviposit, ensuring a suitable location for larval development; however, underwater oviposition may correlate with low fecundity. Other flies broadcast single eggs in flight or with the abdomen dipped into water from emergent structures. The behavior of Nearctic Nymphomyiidae is unusual, with adults coupling soon after emergence, shedding their wings during descent underwater in copula, and selecting a suitable moss-covered rock to lay an egg rosette around the couple, which then die. Hatching delay (egg diapause) is familiar in Aedes mosquitoes (Culicidae), whose eggs are laid in dry habitats that fill seasonally with rain water. Diapause is broken by lowered oxygen postinundation, but may not occur until several successive wettings.

Phenology A typical dipteran life-cycle follows a brief egg stage (usually days, but sometimes much longer), larval and pupal stages of varying length, and an adult stage lasting a few to many hours (Figure 40.5). The duration of the larvula is shortest, whereas the last larval stage—the major feeding stage—is much longer. All larval instars share the same habitat, but many Chironomidae have planktonic larvulae and benthic later-instars. Many aquatic Diptera are univoltine characterized by rapid growth. In seasonal systems with cold winters, immature insects, usually in an early instar, diapause until conditions are favorable. Postdiapause growth often begins with rising spring temperatures, although photoperiod and algal availability may be implicated. Time from egg-hatch to adult emergence varies between and sometimes within species, as does the presence of additional (bivoltine to multivoltine) generations. Tropical aquatic insects often continuously recruit and lack synchronized cohorts. In ephemeral, summer-dry systems, some fly larvae diapause in hyporheic sediments until surface flow returns. Nonfeeding, abbreviated adult life typify Deuterophlebiidae, Nymphomyiidae, and many Chironomidae. Deuterophlebiids have the shortest adult life of any Diptera, with females surviving a few hours and males perhaps 30–45 min. A brief, nonfeeding imaginal stage would be adaptive where larvae obtain resources for gamete production, or where environmental conditions adversely affect adult survival.

GENERAL ECOLOGY AND BEHAVIOR Dipteran ecological diversity reflects larval and adult differences, with larvae feeding and growing and adults reproducing and dispersing. Larval Diptera occur in all

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aquatic and semiaquatic habitats. The latter include damp sediments, saturated wood, decaying organic material, and the tissues of living organisms. True aquatic habitats range from clear, torrential streams to slow, silty rivers; lakes to ponds; cold and hot springs; seepages and groundwater; phytotelmata; and marine, inland saline and estuarine waters.

Habitat Selection Flowing Waters The Hyporheic, Depositional Zones, Springs, Seeps, and the Madicolous (Hygropetric) Zone Vast, underground water storages (the groundwater) connect with the surface hydrological system via a diversity of aquatic habitats including springs, seeps, madicolous (hygropetric) zones, and through depositional areas in flowing waters. A hyporheic zone of interstitial flowingwater broadly connects the groundwater to the surface waters. The hyporheic is frequented by many dipterans, especially larvae of the Ceratopogonidae and Chironomidae, and early instars of many other groups. The hyporheic provides an important reservoir against flooding and high mortality of the benthic community. An unusual member is the black fly Parasimulium, known only from subterranean stream channels of the Pacific Northwest of the United States. The immature stages share many similarities with typical cavernicolous organisms, including apparent blindness of larvae and unpigmented cuticle of larvae and pupae; however, Parasimulium is a typical simuliid in that larvae are filter-feeders with welldeveloped labral fans. Grading into the hyporheic zone are river sediments in the form of sand/gravel bars and deposits around obstructions (e.g., logs, large rocks). Where gravel and sand predominate (the psammon), interstitial oxygenated water is present, but in silty areas it is much more limited. Such habitats grade with the upper hyporheic, and are frequented by many larval dipterans with a narrow vermiform shape and snake-like “swimming” motion, such as many Ceratopogonidae and several Chironomidae (e.g., Stictocladius, Harnischia-complex members). Seasonally, early instars of others occur, presumably evading predation and providing an important refuge against scouring or desiccation. Where the water table is close to the surface, springs, seeps, and mires are found containing quite characteristic dipteran biotas, including many Tipuloidea and Ptychopteridae, notably in shaded seeps. Where thin water films (the madicolous/hygropetric zone) are formed, such as on cliff faces, at stream margins, and in splash-zones, larvae requiring atmospheric oxygen occur, such as those of Psychodidae (Figure 40.2(g)), Stratiomyiidae, and Thaumaleidae.

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Waterfalls, Riffles, and Rapids Turbulent running waters contain maximal dissolved oxygen; however, to take advantage of these conditions, insects must survive in high flow. Small larvae can use the boundary layer or microrefuges on immersed rock surfaces, but others (nematocerans such as Blephariceridae, Deuterophlebiidae, and Simuliidae, and brachycerans like Athericidae and Oreoleptidae) are specialized inhabitants (rheophiles) of current-exposed substrates. All lack spiracles and exchange gases directly through their cuticle. Highly specialized larval Blephariceridae frequent substrates where current velocities may exceed 2 m/sec. Their compact body has six primary divisions, the first comprising the fused head, thorax, and first abdominal segment (i.e., “cephalic” division, Figure 40.2(b)). Each primary division ventrally has a suctorial disc, allowing adherence to smooth, wet rocks. Larval Deuterophlebiidae share similar rocks and food resources (i.e., periphyton). Deuterophlebiid larvae are dorsoventrally flattened with seven pairs of elongate lateral prolegs that bear apical crochets (small, curved hooks; Figure 40.2(a)). Similar prolegs occur on the predaceous larvae of Athericidae and Oreleptidae. Larval Simuliidae also frequent current-exposed habitats, using a crochet-tipped posterior proleg and a silk pad to adhere to submerged rocks or vegetation, where their head extends into the current to strain microscopic food particles by using highly specialized labral fans (Figure 40.2(c)). The larvae of many lotic Diptera occur in more sheltered habitats beneath rocks (e.g., many Tipulidae and Chironomidae), inside moss (Nymphomyiidae and the muscid Limnophora), and among coarse benthic gravel (e.g., Tanyderidae, Tabanidae). Large Rivers Larger lowland rivers tend to have sediments of sand and gravel, and side channels often with slow current velocities and fine sediments where oxygen may be limited. These factors cause a natural shift in the fauna from headwaters to large rivers. In addition, many large rivers around the world have been modified by humans for navigation, power production, and water storage, with concomitant effects of dipteran diversity. Unfortunately, when rehabilitation projects are undertaken, the result can be a poorly diverse channel fauna dominated by larval Chironomoidae, with diversity highest in depositional areas. Rocky regions in levees and wing dams, aerated by riffles, can generate enormous numbers of pestiferous blackflies (Simuliidae).

Standing Waters Neuston (Surface Film) and Planktonic Larvae of Culicidae, Chaoboridae, and Dixidae lead active, exposed lives at or near the surface film, with larvae depending on atmospheric gas exchange via siphons. The larval abdomen usually bears specialized hairs, sclerites and/or lobes that enhance the larva’s ability to utilize the surface film for support. Many predatory adult flies feed at or close

to the surface film, seeking insects emerging or trapped on the film. Among the most common predatory flies are several Empidoidea and Ephydridae. Neustic insects may descend to the midwater, or even to the benthic level, but dominant amongst the midwater, vertical, migratory insect larvae are the Chaoboridae, which rise and fall on a diurnal rhythm. Benthic Most lentic Diptera are associated with benthic substrates. Larvae of many families that respire through spiracles and inhabit damp-to-saturated benthic sediments (e.g., ­Ptychopteridae, Tipulidae, Tabanidae, Stratiomyidae, and Dolichopodidae) must stay within reach of the air–water interface. Alternative strategies allowing independence of aerial oxygen by the use of cuticular gas exchange are small size (increasing surface-to-volume ratio), relative inaction (reducing oxygen demand) or, as in certain larval Chironomidae (“bloodworms”), possession of hemoglobin to absorb oxygen from near anoxic waters. The benthos of lakes, almost exclusively comprising Chironomidae among the insects, varies significantly in relation to nutrient status (chlorophyll levels) and average temperatures, providing data to allow past climate construction from communities sampled at intervals from the sediments (Dimitriadis and Cranston, 2001; Lotter et al., 2012).

Special Habitats Phytotelmata Plant-held aquatic habitats are microcosms with infauna, consisting in part of very specialized flies. In North America, these include “rot holes” in many trees and insectivorous pitcher plants, namely Darlingtonia and Sarracenia (Sarraceniaceae) (Figure 40.6(b and c)). Pitcher plants in North America almost predictably contain the chironomid Metriocnemus, the culicid Wyeomyia, and the sarcophagid Fletcherimyia. Rot holes may contain a high diversity at the family level, with not only Chironomidae but also some Ceratopogonidae, Psychodidae, Tipuloidea, and Syrphidae. Old World tropical pitcher plants (Nepenthes, Nepenthaceae) support larvae of many flies, as do bromeliads and tank plants (Bromeliaceae) in central and tropical southern America (Figure 40.6(a)). Notably, many tropical rainforest mosquitoes are restricted to such plant-container habitats. High Latitudes and Elevations: Cold and Glaciers Permanently frozen water seems unlikely to support aquatic Diptera, but diverse larval Chironomidae, notably subfamily Diamesinae and some Podonominae, thrive worldwide at the ice–water interface on glaciers. The elevational and low temperature record (active at −16 °C) is a flightless Diamesa studied at Yala Glacier in Nepal at a >5000-m elevation, where larvae graze on algae and bacteria growing on glacial ice. Elsewhere, chironomids develop in cold waters associated with snow-melt or glaciers in New Zealand,

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FIGURE 40.6  Special habitats for Diptera: (a) tank bromeliad (Vriesia); (b) pitcher plant (Sarracenia); (c) pitcher plant (Darlingtonia); (d) thermal spring (inset = stratiomyiid larva); (e) saturated wood showing axymyiid galleries; (f) saturated wood showing axymyiid tunnels; (g) seaweed rack (with pan traps); (h) margin of Great Salt Lake, with masses of ephydrid adults; and (i) Neocuripira (Blephariceridae) with phoretic Tonnoirocladius (Chironomidae). Images by G.W. Courtney

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North America, Eurasia, and Africa. The podonomines from melt-pools on the Antarctic Peninsula are the southernmost, free-living, holometabolous insects. At the other extreme, many flies are recorded from Lake Hazen, Ellesmere Island (81°N). A 7-year life cycle, the longest reported for an arctic insect, is proposed for two Alaskan Chironomus (Chironomidae) species. Thermal Pools At high temperatures, certain Diptera larvae tolerate springs and thermal pools (and associated unusual water chemistries) (Figure 40.6(d)). Species of larval Ephydridae (Ephydra spp.) and Stratiomyidae [Stratiomys, Hedriodiscus; Figure 40.6(d) (inset)] tolerate temperatures slightly above 50 °C in hot springs across North America. In warm waters of unusual acidity, certain Chironomidae and Ceratopogonidae may also thrive, and in warm alkaline waters, Stratiomyiidae may flourish. Saturated Wood Saturated wood (Figure 40.6(e and f)) may support the immature stages of several Diptera, including members of the Axymyiidae (Figure 40.2(i)) and various Tipuloidea (e.g., Epiphragma, Lipsothrix) and Syrphidae (e.g., Temnostoma), and in some regions, Tanyderidae. In North America (e.g., the Appalachian Mountain region), many taxa can occur in the same log. ­Other aquatic-to-semiaquatic groups with some xylophilic species are the Chironomidae (notably Stenochironomus and Xylotopus) and various mycetophiloids. Marine, Saline, and Intertidal Insects are minor components of marine and brackish-water diversity, yet some fly larvae prosper. Among those in estuarine marshes and intertidal pools are Ceratopogonidae, Chironomidae, Tipulidae, Tabanidae, Canacidae, and Ephydridae. The latter are particularly abundant in brackish-waters, including maritime marshes, tidal salt pools, and alkaline lakes of arid regions (Figure 40.6(h)). Others (e.g., Coelopidae, Helcomyzidae, and Heterocheilidae) frequent decaying seaweed on the world’s beaches (Figure 40.6(g)). Pontomyia (Chironomidae) are oceanic, and several orthocladiines such as Clunio and immature telmatogetonines (Chironomidae) inhabit the intertidal (and including the structures of oceanic vessels and oil platforms). Globally, inland saline habitats, including hypersaline and marine transitioning to fresh habitats, are prone to the development of nuisance populations of certain midges (Chironomidae) and brine flies (Ephydridae). Salt marshes and mangrove swamps can be rendered nearly uninhabitable by specialist blood-feeding Ceratopogonidae and Culicidae.

Feeding Behavior Diptera are important consumers and provide food for many other organisms in most ecosystems. Trophic diversity is reflected in larval feeding habits that include all functional

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feeding groups. In most Empididae, larvae and adults belong to the same trophic category (predators) but most adopt divergent feeding strategies. In a few of these dance (or dagger) flies, feeding can be restricted largely to the larvae. Some may feed on multiple food resources during the same life stage (e.g., larvae that can be both saprophagous and predaceous, and adults that are both nectarivorous and haematophagous). For example, larval Sciomyzidae may feed on dead or living mollusks, the same instar of Ephydridae larvae may consume algal, bacterial, or detrital resources, and Tanypodinae (Chironomidae) larvae tend to algivory early and predation later. Saprophagy is prevalent, especially in larval brachycerans feeding on decaying organic material with microorganisms providing nutrition. Decomposing plant fragments are consumed by the aquatic larvae of many Tipulidae, Ephydridae, and Chironomidae. These and others (e.g., Psychodidae, Syrphidae, Stratiomyiidae) also feed on fine organic matter. Culicidae and Simuliidae are mostly collector–filterers that consume fine organic matter and bacteria trapped by modified mouth-brushes or labral fans that direct water across the mouthparts (Figure 40.2(c)). In many saprophagous groups, (e.g., Stratiomyidae, Syrphidae) an internal, sieve-like pharyngeal filter concentrates food particles. Phytophagous groups, which consume (shred) live plants (including algae and fungi), are represented by larvae of Tipulidae and certain Chironomidae. Many fly larvae consume the thin biofilms of algae and organic matter. Notable biofilm grazers are larval Blephariceridae, Deuterophlebiidae, and certain Psychodidae, Ephydridae, and Chironomidae. Most predaceous Diptera attack other invertebrates as their primary food. Many families (e.g., Chironomidae, Tipulidae, and Ephydridae) contain a few predaceous species, whereas other groups (e.g., nearly all lower Brachycera) feed primarily or exclusively on invertebrates. In some, vertebrate prey (frogs and salamanders) can be part of the diet. Whereas predaceous larvae typically kill multiple hosts, parasitic and parasitoid larvae generally attack only one host. Of these, the Sciomyzidae are among the best known aquatic group. Parasitoids are rare among aquatic dipteran larvae, but examples include the Sciomyzidae, whose hosts are freshwater snails and fingernail clams, and Oedoparena sp. (Dryomyzidae) feeding on barnacles. Other relationships are uncertain; for example, diverse larval Chironomidae use other macroinvertebrates as substrate; and although this is often interpreted as phoresy (Figure 40.6(i)), some “phoretic” Symbiocladius and Nanocladius pierce their host cuticle to obtain hemolymph.

Predators, Parasites, and Parasitoids of Diptera The immature stages of aquatic Diptera can be summarized succinctly as food for predators. The diets of many insectivorous freshwater fish such as salmonids, at all ages and on all continents, are comprised in part of midge and blackfly larvae,

Chapter | 40  Order Diptera

pupae (especially if ascending through the water column), and spent adults from the surface. Mosquito fishes (Gambusia spp.), so called for their ability to eat numerous mosquito larvae, actually often eat other Diptera, especially adults fallen to the water surface. Other aquatic insects such as mayflies and stoneflies are important prey at certain times for carnivorous fish. Major invertebrate predators on immature aquatic dipterans include voracious odonate nymphs and megaloptera larvae. Swarming adult dipterans, such as midges and mosquitoes, are preyed upon by aerial specialists such as odonates, birds such as swifts, swallows, and martins by day, and bats from dusk onwards. These predators judge the altitude for their feeding according to the optimal heights at which their prey is swarming. The purple martin (Progne subis), is reputed to be a major predator of mosquitoes and for which “houses” are constructed, actually appears to prefer insects larger than mosquitoes and midges. The aquatic stages of Diptera can host parasites, notably water mites and mermithid nematodes. In developing larval chironomids and blackflies, Mermithidae can induce larvae to develop into intersex adults. Mymarid wasps (Hymenoptera: Mymaridae) may be egg parasitoids of certain aquatic Diptera, but this needs verification.

COLLECTING, CULTURING, AND SPECIMEN PREPARATION Collecting Collecting Adults Many adult Diptera are weak fliers: sweeping riparian vegetation or examining the underside of bridges, marginal leaves, or overhanging banks are usually productive activities. Adults can be sampled effectively by sweeping a net through mating swarms, collecting at light traps (especially for Chironomidae, Ceratopogonidae, and Culicidae), or using a Malaise (intercept) trap across a stream. Emergence traps placed over specific habitats can also be useful for collecting adults.

Collecting Immature Stages The phenomenon of drift in running waters can be used to collect many aquatic larvae and pupae. Deflection and interception of the water column into a fine-mesh net will trap (and concentrate) immature stages and also floating adults that have failed to emerge. A few may be adults with pupal skin attached, and pupae with the larval skin and head capsule attached, and thus all stages of a species can be associated without rearing. Collection of larvae and pupae for rearing to the morelikely named adult is more difficult for aquatic groups than for terrestrial Diptera. Only in the Simuliidae, Dixidae, Thaumaleidae, Chaoboridae, and Culicidae are the immature stages well known, but even there gaps remain. Some collections can be made by visual examination and with unspecialized equipment. In standing waters up to 50 cm deep, a fine-mesh pond

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net is effective for larvae living on or just above the benthos, such as Culicidae and Chaoboridae, and also for surfacedwelling larvae, including Dixidae. A sturdy pond net can be used for taking samples of a shallow bottom (benthos), but a grab sampler lowered from boat or jetty is essential for sampling deeper lakes. A pond net can be used in slower streams and ditches. A Surber samples quantitatively since the frame enclosing the substrate is of known dimensions, but also functions qualitatively for collecting larvae for rearing. Place the sampler in the stream with the frame on the bed and the net opening facing upstream. Both hands can then be used to clean upstream stones, or the bed can be kicked in front of the net and organisms can be concentrated in the net.

Rearing Diptera Larvae If the association of larvae and pupae with adults is intended, each larva must be separated, one in each container, to prevent the mistaken associations that can arise from rearing multiple, similar-looking larvae. Many fly larvae are killed by low oxygenation of the incubating medium, caused by oxygen demand from microorganisms, fungi, and algae. Provide a relatively large surface area of water for gaseous exchange, and many larvae will develop in a 50- by 12-mm-diameter tube. Simuliid larvae are filter feeders and require continuous current to bring particles to their mouthparts. This can be attained by a pump-driven air current directed through a nozzle placed just below the water surface close to the edge of the dish, causing a circular current of water in the dish. With smaller larvae, including many Chironomidae, Ceratopogonidae, and Psychodidae, the collection site water is adequate for rearing since it contains enough (but not too much) algae and detritus for the completion of larval development. With larger Chironomidae and Culicidae, the addition of some substrate and fish food may help, but in excess this encourages fungal development. Fungicides are not recommended, although occasional chlorinated tap water reduces fungal development without harming larvae. Many predaceous Diptera eat chironomid larvae, and these can be used as food, as can copepods and oligochaetes. Outdoor domestic water bodies, such as fish ponds and bird baths, contain potential food for predaceous fly larvae.

Pupae Most nematocerans with aquatic larvae also have aquatic pupae, and these should be left in the rearing vessel. If there is only one larva, the skin (exuviae) may be retained until all three stages are removed, but exuviae are devoured by water mites and other organisms. The same applies to the pupal skin—an unambiguous association must be maintained. Simuliid pupae, which darken with age, should be removed when tanned and placed on a piece of damp blotting paper in a tube to await the emergence of the adult.

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Among Tipuloidea and Tabanidae, the final-instar larvae leave the aquatic habitat to pupate in drier conditions. Pupal development often is faster than larval, and success from nonfeeding pharate larvae through pupation to adult can be high. Although usually requiring current to develop fully, pupae of Blephariceridae and Deuterophlebiidae can be reared in the laboratory. An effective method is to remove rocks with attached pupae from the stream and place them in a container maintained at high humidity but not inundated (Courtney, 1998). Frequent spraying of the rock with water and checking the container for pupal exuviae and emerged adults is critical to making associations.

Other Means for Associating Life Stages Molecular techniques now make the sequencing of DNA (mitochondrial, ribosomal, or nuclear) feasible and practical to link immature stages to unassociated (but named) adults (Krosch and Cranston, 2012). Procedures depend on existing databanks for validated adult sequences (the specimen’s DNA library) for the selected gene (predominantly a section of mitochondrial CO1 (cytochrome c oxidase subunit I), which is the so-called “barcode” region) (Ekrem et al., 2007). Despite being far-from-complete DNA libraries, inadequate vouchering, endosymbiont contamination, and naive population genetics underpinning analyses (including inadequate sampling of taxa and across species ranges) (Virgilio et al., 2010), this technique may gain utility in many areas of freshwater research including, potentially, environmental biomonitoring, as long as robust protocols and continued substantial funding are present.

Preserving Specimens for Later Identification It is rare that aquatic Diptera can be identified reliably in the field; and, even when possible, specimens should be returned to the laboratory for verification and vouchering. If material is reared, then the newly-emerged adult, which is teneral with a soft cuticle, must harden (to complete sclerotization) and only then should it be killed and preserved according to future use. Adult Chironomidae, Chaoboridae, Dixidae, Ceratopogonidae, Psychodidae, and all immature stages are collected best into 70–80% ethanol or propanol (higher strength is better if subsequent molecular study is envisaged, but causes brittleness of specimens). Other adult flies may be pinned or pointed dry. If reared specimens are preserved in fluid, all life-stages should be kept together, and any subsequent adult preparation should be referable unambiguously to its immature skins. For specimens needing subsequent microscope slide preparation, preservation in liquid is best, but dried specimens can be prepared as slide mounts. Storage of dried and liquid-preserved material should be in the cold and dark, with detailed location data labels as soon as it is practicable.

REFERENCES Colless, D.H., McAlpine, D.K., 1991. Diptera. In: Naumann, I.D. (Ed.), The Insects of Australia, vol. 2, second ed. Melbourne University Press, Melbourne, pp. 717–786. Courtney, G.W., 1998. A method for rearing pupae of net-winged midges (Diptera: Blephariceridae) and other torrenticolous flies. Proc. Entomol. Soc. Wash. 100, 742–745. Courtney, G.W., Merritt, R.W., 2008. Chapter 22: aquatic Diptera. Part one. Larvae of aquatic Diptera. In: Merritt, R.W., Cummins, K.W., Berg, M.B. (Eds.), An Introduction to the Aquatic Insects of North America, fourth ed. Kendall Hunt Publishing Company, Dubuque, Iowa, pp. 687–722. Courtney, G.W., Pape, T., Skevington, J.H., Sinclair, B.J., 2009. Chapter 9: Biodiversity of Diptera. In: Foottit, R.G., Adler, P.H. (Eds.), Insect Biodiversity: Science and Society. Blackwell Publishing, Oxford, pp. 185–222. Danks, H.V., Smith, A.B.T., 2009. Chapter 3: Insect diversity in the nearctic region. In: Foottit, R.G., Adler, P.H. (Eds.), Insect Biodiversity, Science and Society. Blackwell Publishing, Oxford, pp. 35–48. Dimitriadis, S., Cranston, P.S., 2001. An Australian Holocene climate reconstruction using Chironomidae from a tropical volcanic maar lake. Palaeogeogr. Palaeoclimatol. Palaeoecol. 176, 109–131. Ekrem, T., Willassen, E., Stur, E., 2007. A comprehensive DNA sequence library is essential for identification with DNA barcodes. Mol. Phylogenet. Evol. 43, 530–542. Gullan, P.J., Cranston, P.S., 2010. The Insects: An Outline of Entomology, fourth ed. Blackwell Publishing, Oxford. 505 p. Krosch, M., Cranston, P.S., 2012. Non-destructive DNA extraction from Chironomidae, including of fragile pupal exuviae, extends analysable collections and enhances vouchering. Chironomus 25, 22–27. Lotter, A.F., Heiri, O., Brooks, S., van Leeuwen, J.F.N., Eicher, U., Amman, B., 2012. Rapid summer temperature changes during termination 1a: high-resolution multi-proxy climate reconstructions from Gerzensee (Switzerland). Quat. Sci. Rev. 36, 103–113. Merritt, R.W., Webb, D., 2008. Chapter 22: aquatic Diptera. Part two. Pupae and adults of aquatic Diptera. In: Merritt, R.W., Cummins, K.W., Berg, M.B. (Eds.), An Introduction to the Aquatic Insects of North America, fourth ed. Kendall Hunt Publishing Company, Dubuque, Iowa, pp. 723–771. Pape, T., Blagoderov, V., Mostovski, M.B., 2011. Order Diptera Linnaeus, 1758. In: Zhang, Z.-Q. (Ed.), Animal Biodiversity: An Outline of Higher-Level Classification and Survey of Taxonomic Richness Zootaxa, vol. 3148. pp. 222–229. Virgilio, M., Backeljau, T., Nevado, B., De Meyer, M., 2010. Comparative performances of DNA barcoding across insect orders. BMC Bioinforma. 11, 206. Wiegmann, B.M., Trautwein, M.D., Winkler, I.S., Barr, N.W., Kim, J., Lambkin, C.L., Bertone, M.A., Cassel, B.K., Bayless, K.M., Heimberg, A.M., Wheeler, B.M., Peterson, K.J., Pape, T., Sinclair, B.J., Skevington, J.H., Blagoderov, V., Caravas, J., Kutty, S.N., Schmidt-Ott, U., Kampmeier, G.E., Thompson, F.C., Grimaldi, D.A., Beckenbach, A.T., Courtney, G.W., Friedrich, M., Meier, R., Yeates, D.K., 2011. Episodic radiations in the fly tree of life. Proc. Nat. Acad. Sci. 108. 5690–5695.

Chapter 41

Minor Insect Orders Matthew R. Cover Department of Biological Sciences, California State University Stanislaus, Turlock, CA, USA

Michael T. Bogan Department of Environmental Science, Policy, and Management, University of California Berkeley, Berkeley, CA, USA

Chapter Outline Introduction1059 Megaloptera1060 Introduction1060 Phylogenetic Relationships 1060 Distribution and Diversity 1061 Life History, Ecology, and Behavior 1062 Larval Morphology and Physiology 1063 Collecting, Rearing, and Specimen Preparation 1063 Neuroptera1064 Introduction1064 Phylogenetic Relationships 1064

INTRODUCTION This chapter provides an overview of the aquatic members of seven orders of insects: Megaloptera, Neuroptera, Blattodea, Hymenoptera, Lepidoptera, Mecoptera, and Orthoptera. The collection of these groups into one chapter is artificial and does not necessarily reflect evolutionary or ecological similarities. In fact, the aquatic members of these groups have relatively little in common, other than their low diversity (200 Ma (Winterton et al., 2010; Wang et al., 2012). An evolutionary origin for Sialidae in Eurasia is suggested by the locations of early fossils, although the common ancestor is presumed to have been globally distributed, with some generic diversification prior to the breakup of Pangaea (Liu et al., 2014). The Sialis lineage likely originated in the Nearctic, with subsequent dispersal and diversification in Eurasia. Several species of Sialis with North American distributions are very closely related to Eurasian species, suggesting that extensive faunal exchange via the Bering land bridge may have occurred over the last 100 million years (Liu et al., 2014). There are nine extant genera of Corydalinae, with the greatest diversity in Indomalaya (five genera, ∼88 species) and the Neotropics (three genera, ∼53 species). Unlike fishflies, which are only known from the Neotropics in Chile, Cuba, and Brazil, dobsonflies are widely distributed in Central and South America. Corydalinae is distributed throughout the Indomalayan region, with especially high diversity in China, India, and Southeast Asia. The distribution and diversity of dobsonflies outside of Indomalaya and the Neotropics are fairly limited. Several species of the primarily Neotropical genus Corydalus extend north into the Nearctic, and one species (Corydalus cornutus [Linnaeus, 1758]) is found exclusively in the United States and Canada. The monotypic genus Chloroniella is endemic to South Africa. Several species of the primarily Indomalayan genus Protohermes extend into the Palearctic in northern China and Japan. There are 18 extant genera of fishflies, with the greatest generic diversity in the Nearctic (six genera) and Indomalaya

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(five genera). At the species level, the greatest diversity is in Indomalaya (∼70 spp.), followed by Australasia (∼25 spp.). Neochauliodes (∼40 spp.) is the most speciose and widespread genus of fishflies, with a distribution throughout Indomalaya (including India, Southeast Asia, Indonesia, and China) and the eastern Palearctic (Japan, Russia, and Korea). In the Nearctic, two genera (Dysmicohermes and Orohermes) are endemic to the western United States, two genera (Nigronia and Chauliodes) are endemic to eastern and central North America, and one genus (Neohermes) is distributed across the region. Several genera have very disjunct distributions: Protochauliodes is found in Chile, Australia, and western North America, and Archichauliodes is quite speciose in Australasia (∼19 spp.) and is represented by two species in South America.

Life History, Ecology, and Behavior All Megaloptera have long-lived aquatic larvae and shortlived terrestrial pupae and adults (New and Theischinger, 1993). Final-instar larvae usually crawl out of the water to build pupal chambers under a stone or in moist soil or leaf litter (Figure 41.3). These chambers are often oval-shaped,

(a)

although some species create more tube-like chambers; chamber openings can either be open or closed. The prepupal stage is generally short (1–2 weeks), although Hamilton (1940) reported a 4-month-long prepupal period for Archichauliodes diversus (Walker, 1853) in a snow-fed New Zealand stream. The exarate pupae last for 1–4 weeks before emergence. Emergence times vary with species and climate. For example, Evans (1972) observed variation in the peak emergence of Sialis californica Banks, 1920 from April in coastal California to June in interior Oregon (United States). Adult Megaloptera live for 1–2 weeks and do not feed, although some have been reported to drink water or sugar solutions. Although sex ratios of adults have rarely been reported, males seem to be collected more frequently than females. A number of different types of sexual dimorphism have been reported in adults, including huge mandibles in males of Corydalus and Acanthacorydalis, differences in antennae in many fishflies, and body and wing coloration differences in Protohermes niger Yang and Yang, 1988 (Chang et al., 2013). Adult Megaloptera are almost always found near water sources, and are not believed to disperse long distances. Shortly after mating, females oviposit on vegetation or other structures that overhang the water

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FIGURE 41.3  Examples of the (a) larval, (b) prepupal, (c) pupal, and (d) adult life stages of Neohermes (Megaloptera: Corydalidae). Images by M.T. Bogan and M.R. Cover.

Chapter | 41  Minor Insect Orders

body. Upon hatching, larvae apparently fall or crawl into the water or dry streambed (Evans, 1972). Although there have been few successful attempts to rear larvae through their entire life cycle, some workers have reported 10–12 larval instars. Discrete cohorts are frequently observed in a given locality, with late-instar larvae overwintering prior to pupation and emergence of adults in spring or summer. Sialids typically have life cycles of 1–2 years, although 3-year life cycles have been observed in alpine areas (New and ­Theischinger, 1993). In California (United States), corydalid life cycles may last 4–5 years in intermittent (temporary) and cold perennial streams (Evans, 1972). In contrast, Corydalus cornutus was univoltine in warm streams of the southern United States. (Bowles, 1990). Larval sialids have been collected from both lotic and lentic habitats, although individual species seem to be specialists in one type of habitat. Larval corydalids are most commonly found in mountain streams, but have been found in a wide variety of aquatic habitats including large rivers, swamps, and water-filled vegetation. Several fishflies in the western United States, including Neohermes filicornis (Banks, 1903), Neohermes californicus (Walker, 1853), and several species of Protochauliodes, are specialists in temporary streams that may flow from 3 to 9 months per year (Evans, 1972; Bogan and Lytle, 2007). Immature larvae apparently burrow into the streambed as flow recedes, as they are not found in dry streambeds but reappear in the benthos within days of flow resumption. Final-instar larvae of Neohermes use stream drying as an environmental cue to begin pupation; they crawl under large stones to dig pupal chambers in moist sand and gravel as the streambed dries (Figure 41.3). Megalopteran larvae are all generalist predators of small invertebrates and will also scavenge for food and cannibalize smaller conspecifics, especially under laboratory conditions. Larvae of the dobsonfly Protohermes grandis (Thunberg, 1781) are presumably sit-and-wait predators, as a radio-tracking study revealed that larvae remained relatively stationary under stones in riffles for long periods of time (Hayashi and Nakane, 1989). Other hellgrammites have been observed to remain under stones during the day but actively crawl along the streambed in search of prey during the night.

Larval Morphology and Physiology Megaloptera larvae often have distinctly patterned head capsules and always have large, opposable mouthparts and a three-segmented thorax with three pairs of similar-sized, segmented legs, each with two tarsal claws. Larvae have a 10-segmented abdomen with paired lateral filaments (gills) on segments 1–7 (Sialidae) or 1–8 (Corydalidae) (Figures 41.1–41.3). A fossil from the upper Jurassic of a presumed larval sialid, Sharasialis, has a small pair of

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lateral filaments on the eighth abdominal segment (Ponomarenko, 2012), suggesting that this condition may be plesiomorphic for all Megaloptera. The tenth abdominal segment of sialids is composed of a single, long terminal filament, often with many attached hairs. In contrast, corydalids possess a pair of anal prologs and each proleg ends in a pair of claws. Corydalinae (dobsonfly) larvae are unique in having paired tufts of gills on the ventral side of abdominal segments 1–7. Larvae of many species of Chauliodinae possess modified abdominal spiracles on the eighth abdominal segment, which facilitate respiration. In some species of Parachauliodes and Chauliodes, these spiracles are modified to be long respiratory tubes that can be used as ‘snorkels’ to breathe subaerially (Takeuchi and Hoshiba, 2012).

Collecting, Rearing, and Specimen Preparation Larval megalopterans can be collected using many standard benthic sampling devices, including kick nets and Surber samplers in lotic habitats, and Ekman or Ponar grab samplers in lentic habitats. For targeted collecting of Megaloptera, hand-picking of larvae is a useful technique in shallow habitats with good water clarity. Although some species may actively crawl over the benthos in search of prey, most species seem to remain under cover during daylight hours. Corydalid larvae can often be found by turning over larger rocks in streams. Larvae should be preserved in 70% ethanol; higher concentrations of ethanol, although desirable for molecular analyses, can result in brittle specimens and rapid color loss. Larger larvae can persist for up to 30 min, despite being completely submerged in ethanol. Evans (1972) recommends injecting large larvae with a preservative before immersion in ethanol. Adult Megaloptera are most commonly collected during the day by using sweep nets in riparian vegetation, or at night by using black lights or other light traps. Eggs can be collected by scrapping egg masses off of the vegetation or rock substrate directly into containers. Megaloptera larvae can be reared in the laboratory in aquaria that mimic their natural habitats; important considerations include substrate, water temperature, and aeration. Larvae will consume a wide variety of food sources, including fresh or frozen animal tissue. Even when larvae are provided with sufficient food, cannibalism is a major concern. Ideally, individuals should be kept in separate containers during rearing. Final-instar larvae can often be encouraged to pupate by being placed in a container filled with damp soil or sand, preferably with the option of burrowing under a stone that is larger than the body length of the larvae. Once pupation begins, the substrate should be allowed to dry out to prevent fungal infections.

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NEUROPTERA Introduction Although the order Neuroptera contains 17 families and over 6000 species (Aspöck, 2002; Aspöck et al., 2012), only two neuropteran families are truly aquatic, and a third family includes some semiaquatic taxa. Sisyridae has aquatic larvae (Figure 41.4), whereas Nevrorthidae has aquatic larval and pupal stages. Larvae and pupae of three genera of Osmylidae are only reported from wetted habitats along the margins of water bodies. Although it is uncertain if these genera are completely dependent on aquatic habitats, they do occur in benthic samples collected from streams and springs (e.g., Ivković and Weissmair, 2011).

Phylogenetic Relationships As discussed previously in the Megaloptera section, phylogenetic relationships among taxa within the superorder Neuropterida and the order Neuroptera have been vigorously debated (e.g., Aspöck et al., 2012). Based on several different phylogenetic approaches, Nevrorthidae (suborder Nevrorthiformia) has been hypothesized to be a sister to all other Neuroptera (suborders Hemerobiiformia and Myrmeleontiformia: Beutel et al., 2010; Aspöck et al., 2012; Liu et al., 2012a), which suggests an aquatic ancestor for

FIGURE 41.4  Habitus of a larval spongillafly (Neuroptera: Sisyridae). Image by M.R. Cover.

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Neuroptera. Other analyses, however, have found a nonaquatic neuropteran to be the basal lineage, with a separate clade of Osmylidae and Sisyridae + Nevrorthidae (Winterton et al., 2010). Within the suborder Hemerobiiformia, Sisyridae and Osmylidae are generally considered to be more closely related than the other nine families in the suborder (Beutel et al., 2010; Aspöck et al., 2012).

Distribution and Diversity Of the three neuropteran families dealt with here, the Sisyridae is the most diverse (>70 species in four genera) and the most widely distributed (Cover and Resh, 2008). Sisyra (44 species) has a cosmopolitan distribution; Climacia (22 species) is limited to North and South America; Sisyrina (four species) is known from Africa, Asia, and Australia; and Sisyborina (two species) is endemic to Africa. The Osmylidae are fairly cosmopolitan, but most species are terrestrial (New, 2003), and only two subfamilies have representatives that may be dependent on aquatic habitats (Cover and Resh, 2008). The Kempyninae includes three genera, of which Kempynus (14 species) is known from Australia, New Zealand, and South America, whereas Australysmus (four species) and Clydosmylus (one species) are endemic to southeastern Australia. The subfamily Osmylinae includes the widely-distributed genus Osmylus (>20 species: Europe, Asia, and North Africa), as well as Grandosmylus (two species) and Lahulus (one species), which are both endemic to Asia (Oswald, 2013). Nevrorthidae (16 species in four genera) has a disjunct distribution that includes Europe, Africa, Australia, Japan, and China (Liu et al., 2012a). Nipponeurorthus (nine species) is known from Japan and China, Nevrorthus (four species) occurs in Europe and North Africa, Austroneurorthus (two species) is endemic to Australia, and Sinoneurorthus (one species) is endemic to China. Aquatic and semiaquatic Neuroptera are also well-represented in the fossil record. The Sisyridae and Nevrorthidae occur as fossils dating to the early and mid-Cretaceous (100 Ma and 140 Ma, respectively), whereas the earliest fossils of Osmylidae are from the late Permian (240 Ma: Jepson and Penney, 2007). Cretaceous (20–45 Ma) fossils of the three groups are commonly found in amber and lakebed shales, and these fossils suggest that the morphology and biology of these groups has not changed much since then. For example, Eocene (20 Ma) Osmylus are recorded from the wetted margins of paleolakes (Rasser et al., 2013). Also, Wichard et al. (2010) call extant Nevrorthidae ‘living fossils’ due to their similarities with fossil Nevrorthidae from Baltic amber (44 Ma). Although Nevrorthidae and Osmylidae currently do not occur in North America, fossil evidence indicates that they were present there in the Eocene (34 Ma: Cockerell, 1908; Wichard et al., 2010).

Chapter | 41  Minor Insect Orders

Life History, Ecology, and Behavior The Sisyridae, Nevrorthidae, and Osmylidae all have very distinct life histories, ecologies, and behaviors. Sisyridae (spongillaflies) are considered to be obligate predators of freshwater sponges, although they may occasionally prey on bryozoans as well (reviewed in Cover and Resh, 2008). Although freshwater sponges do not receive much scientific attention, they are widely distributed across the world and occur in ponds, lakes, rivers, and desert oases (Figure 41.5). Spongillafly larvae have specialized mouthparts for piercing and extracting food from sponge cells (Fig. 41.4). They preferentially feed upon ‘green’ sponges that have symbiotic zoochlorellae. One stable isotope study found that >98% of the carbon in populations of the spongillafly Climacia areolaris (Hagen, 1861) was derived from the symbiotic algae contained within its host sponge (Skelton and Strand, 2013). First-instar spongillafly larvae settle on an individual sponge and often do not leave that sponge until they reach their third and final instar. Mature third-instar larvae abandon their host and swim to shore or to emergent structures (e.g., vegetation, boat docks). At dusk, they emerge and crawl up to 30 m from the water to construct pupal cocoons on shore; they often spin net-like structures over these cocoons (Brown, 1952; Forteath and Osborn, 2012). Pupation duration is temperature dependent, and lasts 5–20 days. Adults generally emerge from pupal chambers between dusk and midnight, live for several days or weeks, and feed on pollen, mites, and aphids (Brown, 1952; Pupedis, 1987; Forteath and Osborn, 2012). Oviposition usually occurs

(a)

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between dusk and midnight, and eggs (usually two to five, but up to 20) are laid on overhanging vegetation along the margins of streams or lakes and covered with a silk mesh. A single female may lay several hundred eggs over the course of her life (Brown, 1952; Forteath and Osborn, 2012). After 10–20 days, eggs hatch and first-instar larvae wriggle or flip until they fall into the water, where they spend between an hour and 3 days swimming or drifting in search of a sponge host. Life cycle timing is variable within and across spongillafly populations and species, ranging from univoltine to bivoltine to multivoltine. For example, populations of Sisyra can be univoltine or bivoltine and often have one fast summer generation with a second generation that overwinters as mature larvae or prepupae in cocoons (Hölzel and Weissmair, 2002; Forteath and Osborn, 2012). In contrast, Climacia areolaris may have as many as three generations during the summer and then overwinter as third-instar larvae (Brown, 1952). Although spongillaflies have been relatively well studied, much less is known about the Nevrorthidae. Larvae of the family were undescribed until 1967, when Zwick published a study of Nevrorthus fallax (Rambur, 1842). Both larval and pupal Nevrorthidae are fully aquatic and appear to prefer clean, cold, montane streams with relatively high gradients and gravel to cobble substrate (Zwick, 1967; Aspöck and Aspöck, 2010; Liu et al., 2012a). Larvae are slender and agile predators of other benthic invertebrates. They likely pass through three larval instars before pupation. Life cycle timing is unknown; however, given the cool

(b)

FIGURE 41.5  An example of an aquatic environment that supports large populations of Sisyridae (Neuroptera): (a) desert oasis in Sonora, Mexico where (b) freshwater sponges (Spongillidae) grow on submerged palm roots and provide a habitat and food source for larvae of Climacia chapini. Images by M.T. Bogan.

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montane streams they inhabit, most populations are likely univoltine. Adult nevrorthids may emerge from streams in the spring (Ivković and Weissmair, 2011). They are often found under twigs and leaves that overhang fast-flowing sections of streams and have been observed during day and night (Liu et al., 2012a). Adult feeding, mating, and oviposition habits are still unknown, as are the details of egg incubation and hatching. Although most Osmylidae species are terrestrial, six genera in the subfamilies Osmylinae and Kempyninae are strongly associated with freshwater habitats. Of these genera, the most well-studied is Osmylus (Figure 41.6). This genus occurs widely throughout Europe, and all life stages occur only in riparian areas. The amphibious larvae of Osmylus fulvicephalus (Scopoli, 1763) live on wet mosses and mineral substrate along the margins of streams. First-instar larvae of O. fulvicephalus are known (Enders and Wagner, 1996) to consume eggs of Apatania (Trichoptera: Apataniidae). Second- and third-instar larvae prey upon Tipulidae, and all instars hunt Chironomidae by probing in mosses or mud or entering water to find their prey (Ward, 1965; Enders and Wagner, 1996; Kriska, 2013). Larval Osmylus have been observed consuming prey much larger than themselves; prey items are “quickly paralyzed and then leisurely sucked of its juices” (Withycombe, 1923). They are not known to be effective swimmers, but can crawl rapidly underwater (Kriska, 2013). They also have an air bubble in their larval digestive tube that eases submersion and prevents them from sinking too deep. O. fulvicephalus larvae can live on streamside mosses that are submerged for at least half the year (Nolte, 1991) and have been collected in benthic samples from both streams and springs (Zollhöfer et al., 2000; Ivković and Weissmair, 2011). Pupation occurs on the same wet mosses that larvae inhabit, and lasts about

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FIGURE 41.6  (a) Lateral and (b) dorsal views of a mature Osmylus fulvicephalus larva. Images by G. Kriska.

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10–14 days (Withycombe, 1923). Adult Osmylus are usually found in dense, overhanging vegetation along streams, and are often active between May and July in Europe. Both Osmylinae and Kempyninae adults consume mainly pollen and plant material, but also will prey upon moths, beetles, aphids, and mites (Devetak and Duelli, 2007). Adult females of Osmylus lay 2–12 eggs on hard surfaces along stream margins and generally avoid ovipositing on moss (Withycombe, 1923). The few studies of Osmylus life cycles indicate that populations are generally univoltine, and overwinter as second- or third-instar larvae.

Larval Morphology and Physiology Neuroptera larvae have distinct mouthparts, where the mandibles and maxillae are modified to form elongate, unsegmented stylets adapted for piercing and sucking. The exact shape of larval stylets and other aspects of body morphology vary among the three families of aquatic and semiaquatic Neuroptera. The stylets of Sisyridae originate centrally on the anterior portion of the head capsule, are generally held close together, and are slightly curved outward distally (Fig. 41.4). Each stylet can be used as a sucking tube, or both pairs can be joined together (Chandler, 1956). First-instar Sisyridae larvae are 0.2–0.6 mm long, whereas second and third instars are 0.7–3.0 and 2.7–8.5 mm long, respectively (Brown, 1952; Chandler, 1956; Forteath and Osborn, 2012). First-instar larvae obtain oxygen from the water through their cuticle, but second- and third-instar larvae have twoor three-segmented gills on abdominal segments 1–7. These abdominal gills can be vibrated rapidly to increase water current across gill surfaces. Third-instar larvae shed their gills prior to emergence from the aquatic environment. The thoracic and abdominal segments of Sisyridae larvae bear a variety of setae, tubercles, and sclerotized plates dorsally, which are used to distinguish genera and species within the family. The six-segmented antennae of Sisyridae larvae are long and slender, and often reach or exceed the tips of the stylets, but labial palps are lacking (Parfin and Gurney, 1956). Thoracic legs are five-segmented and slender, and each bears a single tarsal claw. Sisyridae larvae are fairly plump, being widest at the metathorax and narrow at the head and terminal abdominal segments, and the abdomen and thorax are roughly equal in length. Nevrorthidae larvae, in contrast, are very slender and elongate. Third-instar larvae can be 9.5–13 mm long, the abdomen is about 1.5 times as long as the thorax, and the narrow, dorsoventrally flattened head capsule is roughly 1.5 mm long (Zwick, 1967; Beutel et al., 2010). Maxillary stylets are inserted anterolaterally, are much more robust than those of Sisyridae, and are slightly upturned and curved inward distally. The maxillary stylets, antennae, and labial palps are subequal in length. Larval antennae appear to have as many as 18 segments, but this is the result of secondary

Chapter | 41  Minor Insect Orders

subdivisions of the penultimate antennomere (Beutel et al., 2010). Nevrorthid larvae have a strikingly long and sclerotized neck, which is an extension of the pronotum. The pronotum is fully sclerotized dorsally, and two dorsal plates adorn both the mesonotum and metanotum. Thoracic legs are slender, five-segmented, and have two tarsal claws and an empodium. The abdomen is pigmented dorsally, but not ventrally, and abdominal segment 10 forms a small pygopod. Each abdominal segment bears long setae dorsally and ventrally, with the longest setae on the terminal segment. Morphologies of the three genera with described larvae (Nevrorthus, Nipponeurorthus, and Austroneurorthus) are quite similar (Zwick, 1967). Larvae likely obtain oxygen from the water through their cuticle. Osmylidae larvae are the largest of the three aquatic and semiaquatic neuropteran families. First-instar Osmylus larvae can be as long as 4 mm, and mature third-instar larvae are 15 mm long (Withycombe, 1923; Kriska, 2013). The function, form, and insertion of the modified maxillae and mandibles are similar to those of the Nevrorthidae. The three-segmented antennae and the labial palps are ∼3/4 the length of the mandibles, and the head is slightly wider than long (Figure 41.6). The larval prothorax is fully sclerotized, whereas the mesothorax and metathorax have paired dorsal plates. The first eight abdominal segments bear spiracles and bristles, and all 10 segments have transverse rows of black setae. The 10th segment has a pair or eversible processes with recurved hooks and is used in locomotion and for grasping prey. Thoracic legs are slender, five-segmented, and have two tarsal claws and an empodium.

Collecting, Rearing, and Specimen Preparation Larval Sisyridae can be collected via a variety of methods, including picking them by hand from sponges, using benthic samplers in lotic habitats (e.g., Surber samplers, kick nets), and employing Ekman or Ponar grabs in lentic habitats. Most adult Sisyridae are drawn to light (e.g., UV, mercury vapor), but some species are not (e.g., Sisyra pedderensis Smithers, 2008: Forteath and Osborn, 2012). Larval and pupal Nevrorthidae can be collected from benthic substrate in streams by using Surber samplers or kick nets. The adults of some species of Nevrorthidae are attracted to light. Osmylidae larvae are best collected by picking through mosses and stones along the margins of streams, but they also have been captured from lotic habitats by using Surber samplers and Ekman grabs. Adults of all three families are most commonly collected by sweeping riparian vegetation with an aerial insect net. Rearing Sisyridae larvae in the lab can be complicated due to the difficulty in keeping their sponge host and food source alive. Few workers report rearing Nevrorthidae or Osmylidae in the lab. For morphological preservation, larvae and pupae of all three

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families should be fixed in Kahle’s solution for 24 h and then stored in 70% ethanol or preserved directly in 80% ethanol. Adult Neuroptera should be pinned or pointed, but can also be preserved in 70% ethanol.

BLATTODEA Blattodea (cockroaches and termites) is one of the oldest extant orders of insects, with fossils of ancestral cockroaches dating to the Paleozoic, and those of modern-type cockroaches dating to the Cretaceous (>140 Ma) (Bell et al., 2007). Although nearly 4600 extant cockroach species have been described and fossil aquatic cockroaches documented from the Mesozoic (>100 Ma), relatively few extant aquatic species are known (Nesemann et al., 2010). Most extant aquatic species belong to the subfamily Epilamprinae (Blaberidae) and occur at tropical latitudes. The highest levels of aquatic and amphibious cockroach species diversity occur in the Indomalaya and Neotropical regions. Although some workers have argued that aquatic cockroaches have special adaptations to aquatic life (e.g., using the abdominal tip as a snorkel or an air bubble as an accessory gill), most anatomical structures facilitating respiration in the aquatic environment are also found on terrestrial species (Bell et al., 2007). Aquatic and amphibious cockroaches generally occur in two types of habitat: (1) phytotelmata, and (2) streams and rivers. Many species occupying these habitat types tend to spend significant amounts of time in the terrestrial environment on the edges of aquatic habitat, but submerge themselves to find food or seek refuge from predators. Species occupying phytotelmata have been relatively well-documented in the Neotropics. Thirty-four aquatic or amphibious cockroach species (across 18 genera) are known from small pools in epiphytic bromeliads in Central and South America (Roche e Silva Albuquerque and Rodrigues-Lopes, 1976). In total, about 60 cockroach species have been collected from the leaf bases of bromeliads, but not all of these species are thought to be aquatic or amphibious (Bell et al., 2007). Little is known about the biology of cockroaches in phytotelmata, but some Costa Rican species are known to be nocturnal foragers, consuming mold and submerged rotting plant tissues in these habitats (Seifert and Seifert, 1976). Lotic habitats support aquatic and amphibious cockroach species in at least six genera of Epilamprinae, including Epilampra, Phlebonotus, Poeciloderrhis, Opisthoplatia, Rhabdoblatta, and Rhicnoda. Among Neotropical lotic cockroach species, individuals of Poeciloderrhis cribosa (Burmeister, 1838) have been recorded swimming against a current of 0.15 m/sec and can remain underwater for >3 min without replenishing their air supply (Roche e Silva Albuquerque et al., 1976). The Central American species Epilampra maya Rehn, 1902 readily enters aquatic habitats when disturbed, can remain submerged for >15 min, swims rapidly,

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and consumes aquatic plants (Crowell, 1946). This species has also been introduced into the United States, and has been observed along streams in Florida (Nickle and Sibson, 1984). Several natural history observations have also been made of lotic cockroach taxa in southern Asia and Japan. For example, the Japanese species Opisthoplatia orientalis (Burmeister, 1838) is not known to be a good swimmer, but it can rapidly move in lotic habitats by crawling along submerged substrate (Bell et al., 2007). In India, females of Phlebonotus pallens (Serville, 1831) provide maternal care for newly hatched nymphs (Pruthi, 1933). Nymphs cling to their mother’s abdomen and are shielded by her wings as she crawls along the stream bottom. Nymphs and apterous adult females of Rhicnoda natatrix Shelford, 1907 occupy montane (1600–2500 m) streams of India. They occur under submerged stones, are active swimmers and divers, occasionally run out of the water and along stream margins, and prefer aquatic microhabitats downstream of waterfalls and rapids (Nesemann et al., 2010). Although the eating habits of most lotic cockroach species are unknown, species in at least three genera are known to subsist largely on submerged leaf litter (Bell et al., 2007). Additional research should be focused on aquatic and amphibious cockroach species to better understand their ecologies, life histories, and distributions.

HYMENOPTERA Representatives from 11 families of Hymenoptera are known to be parasitoids of the aquatic stages of various arthropods. These 150 or so aquatic parasitoid species represent 50 aquatic and water-dependent orthopteran species occurs in Neotropical regions of South America (Amédégnato and Devriese, 2008). Little is known about the ecology and life history of most of these species, but recent studies have begun to address these knowledge gaps. The most widespread aquatic species is Cornops aquaticum (Bruner, 1906) (Acrididae), which occurs from southern Mexico to central Argentina. Water hyacinth is the primary habitat and food source for C. aquaticum. The first three nymphal instars consume water hyacinth exclusively, whereas nearly 80% of the diets of fourth and fifth instars and adults are hyacinth (Capello et al., 2011). Two other aquatic species of Acridiae in South America, Paulinia acuminata (De Geer, 1773) and Marellia remipes Uvarov, 1929, also exclusively consume aquatic plants (Capello et al., 2012) and have been used for biocontrol of aquatic weeds in Africa and Australia (Sands and Kassulke, 1986). All three of these species have fusiform bodies, strong hind femora, expanded hind tibiae for swimming, and modified ovipositor valves for egg insertion into aquatic plants (BentosPereira and Lorier, 1991). Six species of the genus Hydrolutos (Anostostomatidae) are endemic to streams and marshes of tepui (table-top mountains) in Venezuela (Issa and Jaffe, 1999; Derka and Fedor, 2010; Derka et al., 2013). Hydrolutos consume benthic algae and can stay submerged for over 20 min; this is made possible by a plastron formed by microsetation on their pleura and abdominal sternites. They can also swim rapidly, even against currents greater than 1 m per second. Given the relative paucity of entomological exploration in remote locations of South America, it is nearly certain that more aquatic orthopteran species remain to be found there. In addition to the Neotropics, many aquatic and water dependent species of Orthoptera are found in India and southwestern Asia. As many as 70 aquatic species may occur there, including members of the subfamilies Oxyinae, Tropidopolinae, and Hemiacridinae (Amédégnato and Devriese, 2008). The rest of Asia, Europe, Africa, and Australia all support very few known aquatic species. The phylogenetic relationships among all the different groups of aquatic and water-dependent Orthoptera are unclear, but it is likely that invasions of aquatic habitats have occurred independently multiple times in many of these families, subfamilies, and

genera (Amédégnato and Devriese, 2008). Much research is still needed on the ecology, life history, and phylogenetics of aquatic and water-dependent orthopteran groups.

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Chapter | 41  Minor Insect Orders

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Oswald, J.D., 2013. Neuropterida Species of the World: a Catalogue of the Species-Group Names of the Extant and Fossil Neuroptera, Megaloptera, Raphidioptera and Glosselytrodea (Insecta: Neuropterida) of the World. Web page: http://lacewing.tamu.edu/Species-Catalogue/index. html (accessed 25.02.14.). Parfin, S.I., Gurney, A.B., 1956. The spongilla-flies, with special reference to those of the Western Hemisphere (Sisyridae, Neuroptera). Proc. U. S. Natl. Mus. 105, 421–529. Pilgrim, R.L., 1972. The aquatic larva and the pupa of Choristella philpotti Tillyard, 1917 (Mecoptera: Nannochoristidae). Pac. Insects 14, 151–168. Ponomarenko, A.G., 2012. Supposed alderfly larva (Insecta, Megaloptera) from the Jurassic locality Shar-Teg, southwestern Mongolia. Paleontol. J. 46, 378–382. Pruthi, H.S., 1933. An interesting case of maternal care in an aquatic cockroach, Phlebonotus pallens Serv. (Epilamprinae). Curr. Sci. ­(Bangalore) 1, 273. Pupedis, R.J., 1987. Foraging behavior and food of adult spongila-flies (Neuroptera: Sisyridae). Ann. Entomol. Soc. Am. 80, 758–760. Rasser, M.W., Bechly, G., Bottcher, R., Ebner, M., Heizmann, E.P.J., Höltke, O., Joachim, C., Kern, A.K., Kovar-Eder, J., Nebelsick, J.H., Roth-Nebelsick, A., Schoch, R.R., Schweigert, G., Ziegler, R., 2013. The Randeck Maar: Palaeoenvironment and habitat differentiation of a Miocene lacustrian system. Palaeogeogr. Palaeoclimatol. Palaeoecol. 392, 426–453. Roble, S.M., 1985. Submergent capture of Dolomedes triton (Araneae, Pisauridae) by Anoplius depressipes (Hymenoptera, Pompilidae). J. Arachnol. 13, 391–392. Roche e Silva Albuquerque, I., Jose-Jurberg, R.T., Rebordões, A.M.P., 1976. Contribuição para o conhecimento ecológico de Poeciloderrhis cribrosa (Burmeister) e Poeciloderrhis verticalis (Bermeister) com un estudo sobre a genitália externa (Dictyoptera: Blattariae). Rev. Bras. Biol. 36, 239–250. Roche e Silva Albuquerque, I., Rodrigues-Lopes, S.M., 1976. Blattaria de bromélia (Dictyoptera). Rev. Bras. Biol. 36, 873–901. Rubinoff, D., Schmitz, P., 2010. Multiple aquatic invasions by an endemic, terrestrial Hawaiian moth radiation. Proc. Natl. Acad. Sci. 107, 5903–5906. Sands, D.P.A., Kassulke, R.C., 1984. Samea multiplicalis [Lep.: Pyralidae], for biological control of two water weeds, Salvinia molesta and Pistia stratiotes in Australia. Entomophaga 29, 267–273.

SECTION | VI  Phylum Arthropoda

Sands, D.P.A., Kassulke, R.C., 1986. Assessment of Paulinia acuminata (Orthoptera: Acridade) for the biological control of Salvinia molesta in Australia. Entomophaga 31, 11–17. Shimizu, A., 1992. Nesting behavior of the semi-aquatic spider wasp, Anoplius eous, which transports its prey on the surface film of water (Hymenoptera, Pompilidae). J. Ethol. 10, 85–102. Siefert, R., Seifert, F., 1976. Natural history of insects living in inflorescences of two species of Heliconia. J. N. Y. Entomol. Soc. 84, 233–242. Skelton, J., Strand, M., 2013. Trophic ecology of a freshwater sponge (Spongilla lacustris) revealed by stable isotope analysis. Hydrobiologia 709, 227–235. Takeuchi, Y., Hoshiba, H., 2012. The life histories of three species of Corydalidae (Megaloptera) from Japan. Aquat. Insects 34, 55–63. Wang, Y., Liu, X.Y., Winterton, S.L., Yang, D., 2012. The first mitochondrial genome for the fishfly subfamily Chauliodinae and implications for the higher phylogeny of Megaloptera. PLoS One 7, e47302. Ward, P.H., 1965. A contribution to the knowledge of the biology of Osmylus fulvicephalus (Scopoli 1763) (Neuroptera, Osmylidae). Entomol. Gaz. 16, 175–182. Winterton, S.L., Hardy, N.B., Wiegmann, B.M., 2010. On wings of lace: phylogeny and Bayesian divergence time estimates of Neuropterida (Insecta) based on morphological and molecular data. Syst. Entomol. 35, 349–378. Wichard, W., Buder, T., Caruso, C., 2010. Aquatic lacewings of family Nevrorthidae (Neuroptera) in Baltic amber. Denisia 29, 445–457. Wiegmann, B.M., Trautwein, M.D., Kim, J., Cassel, B.K., Bertone, M.A., Winterton, S.L., Yeates, D.K., 2009. Single-copy nuclear genes resolve the phylogeny of the holometabolous insect orders. BioMed. Central Biol. 7, 34. Withycombe, C.L., 1923. Notes on the biology of some British Neuroptera (Planipennia). Trans. R. Entom. Soc. Lond. 70, 501–594. Zollhöfer, J.M., Brunke, M., Gonser, T., 2000. A typology of springs in Switzerland by integrating habitat variables and fauna. Arch. für Hydrobiol. 121, 349–376. Zwick, P., 1967. Beschreibung der aquatischen Larve von Neurorthus fallax (Rambur) und Errichtung der neuen Planipennierfamilie Neurorthidae fam. nov. Gewässer Abwässer 44, 65–86.

Subject Index Note: Page numbers followed by “f” and “t” indicate figures and tables respectively

A

A1. See Antennules Abedus herberti (A. herberti), 951–953, 952f Acantharians, 120 Acanthobdella peledina (A. peledina), 567–568 anterior portion, 568f Acanthobdellida, 511, 565 collection, 583 crawling motions, 572f dispersal, 576–577 distribution and diversity, 566–567 ecological interactions behavior, 577–581 predators, 578–579 ecotoxicology, 582–583 external anatomy distinguishing characteristics, 567 external sensory structures, 568 segmentation, 567 suckers, 567–568, 569f internal anatomy alimentary canal, 568–570, 570f circulatory and respiratory systems, 571–572 excretory system, 570–571 locomotion, 572 reproductive system, 570 life history, 575–576 phylogenetic relationships, 566 physiology changes during winter, 573–574 ionic and osmotic regulation, 574 respiratory tolerance and adaptations, 572–573 population regulation, 581–582 preparation of specimens, 583–584 rearing, 583 reproduction, 574–575 and parental care, 571f systematics, 566 Acanthosentis dattai (A. dattai), 725–726 Acentropinae, 1068 Acidity, 40, 862–863 Acilius canaliculatus (A. canaliculatus), 1023f ACS. See Anisotropic crystalline structures Actinopod amebae, 119 Adephaga families, 1014. See also Coleoptera (Aquatic beetles); Myxophaga families; Polyphaga families. A. LeConte, 1020–1021 Aspidytidae, 1021 C. Latreille, 1022

D. Leach, 1022–1024 G. Latreille, 1024–1025 H. Aubé, 1025–1026 H. Régimbart, 1026–1027 Meruidae, 1027 Noteridae, 1027–1028 Adhesive warts, 322–323 Aeolosoma hemprichi (A. hemprichi), 521f Aeolosomatidae, 521 Aerial nets, 943 Aestivation, 985–986 AFA. See Alcohol–formalin–acetic acid Ahuauhtli, 959–960 Albertathyas montanus (A. montanus), 631f Alcohol–formalin–acetic acid (AFA), 560 Algophagidae, 612 Alien species, 83 Alkalinity, 862–863 Alluvial swamps, 46–47, 47f Alpine habitats, 24–25 Ameboid protozoa, 118–119. See also Ciliated protozoa. acantharians, 120 actinopod amebae, 119 foraminiferans, 119 heliozoans, 120 naked amebae, 119 Pelomyxa, 119 radiolarians, 119–120 and slime molds, 118f, 120 testate amebae, 119 Ameronothrus sp., 610f Amphibalanus amphitrite (A. amphitrite), 712 Amphibalanus improvisus (A. improvisus), 745f Amphicrossus japonicas (A. japonicas), 1036f Amphinemura palmeni (A. palmeni), 938f Amphipoda, 781–782, 782f, 784–785 cave-dwelling, 789f Epigean habitats, 788–789 Amphitrite genus, 522 Amphizoa sp., 1020–1021, 1021f Amphizoidae, 1004–1008 Amphizoidae LeConte (Trout-stream beetles), 1020–1021 Amphoteric rotifers, 251 Ampullarids, 394 Anactinotrichida. See Parasitiformes Anaerobic ciliates, 125 Anagenesis, 4 Ancylometes hewitsoni (A. hewitsoni), 603f Anhydrobiosis, 240–241, 361–362 Anisoptera, 894, 895f, 896

Anisotropic crystalline structures (ACS), 248–249, 249f Anisozygoptera, 895f, 896 Annelid groups, 12, 17f Annelida, inland water, 509. See also Habitats, inland water; Polychaeta, inland water. biology external anatomy, 514–515 internal anatomy, 515–516 physiology, 516 reproduction, 516–517 building plan, 509 collection, 519 distribution and diversity, 512 evolutionary origin, 512–513 ecology and behavior distribution and habitat selection, 517 feeding behavior, 518 locomotion and interspecific interactions, 518–519 physiological constraints, 517–518 predators and parasites, 518 phylogenetic relationships, 511–512 preparation for identification, 519 systematics, 510 Acanthobdellida, 511 Branchiobdellae, 511 Hirudinea, 511 Oligochaetes sensu stricto, 510 Annelids (Fresh and marine earthworms), 509 Anoplius depressipes (A. depressipes), 1068 Anoplius eous (A. eous), 1068 Anostraca (Fairy shrimp), 690 Anoxia. See Anoxybiosis Anoxybiosis, 361 Antennal glands, 675–676, 676f Antennules (A1), 765–766 Aorta, 853–854 Apertures, 440–441 Aphanoneura, 510, 521 Apharyn-geal pad, 537 Apicomplexans, 125 Apomorphies, 6 Apple snails. See Ampullarids Aquaculture, 93–94 Aquatic beetles. See Coleoptera Aquatic food webs, 72 lentic food webs, 75 lotic food webs, 74–75 techniques, 73–74

1073

1074

Aquatic habitats, global variations, 24 alpine habitats, 24–25 Australia, 28 oceanic islands, 26–28 polar zones, 24–25 temperate zones, 25 tropical zone, 25–26 Aquatic insects backswimmer Neoplea, 854f circulatory system, 853–854, 854f collecting, 868 ecology and behavior, 855–856 acidity, 862–863 and catastrophic drift, 858–859 feeding behavior, 863–864 habitat selection, 856–861 invertebrate communities, 862f ionic constraints, 861–862 osmotic constraints, 861–862 oxygen tension, 863 parasites, 864, 864f predators, 864 thermal maxima and minima, 862 ecology and behavior, 855–864 evolutionary diversification, 851 excretion, 854–855 generation time, 855 hemimetabolous and holometabolous development, 856f internal tracheal system, 853f literature sources, 851 metamorphosis, 855 osmotic balance, 854–855 plate-like tracheal gills, 854f regions of insect anatomy, 852f reproduction and development generation time, 855 metamorphosis, 855 respiratory system, 852–853 tagmatization, 852 Aquatic light trap, 59 Aquatic sowbugs. See Isopoda Aquatic–terrestrial ecotones, 70–71 Aquiferous system, 142 Aquifers, 775 Arachnida, 600, 601t–602t Arachnids, 600. See also Hydrachnidiae (Water mites). collecting and extracting techniques, 649–654 collecting concentrated sample, 650–652 deutonymphs and adults, 650–652 extraction, 652 larvae, 652–653 rearing, 653–654 freshwater Sarcoptiformes, 609 Astigmatina, 611–613 Oribatida, 609–611 mites, 600–601 parasitiformes, 607 Mesostigmata, 607–609 preservation and preparation, 654–655 Prostigmata, 613 Halacaroidea, 613–614 Parasitengonina, 615–617

  Subject Index

Raphignathoidea, 614–615 Araneae (Spiders), 601 Ctenidae, 603 Cybaeidae, 603 feeding behavior, 601–602 Lycosidae, 603–604 Pisauridae, 604–606 Thomisidae, 606 Trechaleidae, 606–607 Archichauliodes clade, 1061 Architaenioglossa, 389 Areoles, 305–306 Argulids collection and culturing, 749–750 specimen preparation, 750 Argulus coregoni (A. coregoni), 741 Argulus foliaceus (A. foliaceus), 741 Argulus japonicus (A. japonicus), 741 Argyroneta aquatica (A. aquatica), 602–603 Arrenurus fissicornis (A. fissicornis), 632f Arrenurus kitchingi (A. kitchingi), 646 Arthropod subphyla, 17f Crustacean classes and Malacostracan orders, 12–13 C. centrura, 18f E. nevadensis, 18f freshwater branchiopod crustaceans, 18f freshwater crustaceans, 19f Hexapoda classes, 13, 19f Insecta orders, 13–21 A. abnormis, 20f Acilius, 20f Diptera, 19f H. diabasia, 20f Hemiptera, 20f Odonata, 20f R. fuscula, 20f Sialis larva, 20f Arthropoda, 593f collection, 597 culturing, 597 distribution and diversity, 592 ecology and behavior feeding behavior, 596 habitat selection, 596 physiological constraints, 596 predators and parasites, 596–597 external anatomy, 592–594 internal anatomy and physiology, 594 body cavities, 594 circulatory and respiratory systems, 595 excretory and osmoregulatory systems, 595 neural system, 595 reproduction and development, 595–596 specimens preparation, 597 Arthropods, 591 phylogenetic relationships within Arthropoda, 592 relationships among related phyla, 591–592 Asellus aquaticus isopods, 786, 788, 791 Aspidytes wrasei (A. wrasei), 1021f Aspidytidae (Cliff water beetles), 1021 Asplanchna sieboldii (A. sieboldii), 235f Astacid crayfish, 801. See also Crayfish.

Astacopsiphagus parasiticus Viets, 613–614 Astigmata. See Astigmatina Astigmatina, 611–612 diversity and biology, 613 morphology, 612 Atria, 538–539 Atya genus, 803 Atyidae, 800 Australia, 28 Australian crayfish. See Cherax Axopodia, 119

B

Baetis fuscatus (B. fuscatus), 879f Balanocochlis glandiformis (B. glandiformis), 389f Barbicambarus cornutus, 814f Barnacles collection, 750 culturing, 750 specimen preparation, 751 Basommatophora, 391 Batracobdella, 569f Bay barnacle, 712 Bdelloid anhydrobiosis, 241 Bdelloid rotifers, 229f Beating net. See Beating sheet Beating sheet, 60 Beetle mites. See Oribatida Benthic invertebrates, 59. See also Planktonic invertebrates. aquatic light trap, 59 beating sheet, 60 decapods, 59 dip net, 59 flight intercept traps, 60 hard surfaces, 59 interactions, 484 kick net, 59 littoral benthic substrate, 59 malaise traps, 60 Berlese collector, 667 Biochemical oxygen demand (BOD), 478–479 Bioindicators, 175 Biological species concept, 5. See also Cohesion species concept. drawback, 5 gene exchange, 5 gene flow and reproductive behavior, 6 limitation, 5–6 units of conservation, 5 Bithyniidae, 394 Bivalve shells, 384–385. See also Gastropod shells. Bivalvia, 424 age and growth, 467–468 benthic invertebrate interactions, 484 biotic interactions diseases, 471 parasites, 470–471 predation, 468–470 changes in hydrology channelization, 477–478 floods, 476–477

Subject Index

impoundments, 477 collection data recording, 485–486 season, permits, and gear, 485 survey methods, 484–485 consumptive and commercial impacts, 482–483 Corbicula diversity, 434 culture, propagation, and rearing, 486–487 curation, 486 Dreissena diversity, 434 ecosystem processes, 471–472 endangered species and conservation, 481–482 evolution and phylogenetics, 425–426 external and internal anatomy, 434 Ctenidia, 441 external shell structure, 438–440 internal shell structure, 436–438 mantle cavity and apertures, 440–441 shell and mantle, 434–435 shell growth and microstructure, 435–436 feeding structures and mechanisms, 441–442 assimilation, 445–448 clearance rates, 442–444 deposit feeding, 448–449 digestion, 445–448 ingestion, 445–448 labial palps, 445 particle sorting, 445 retention efficiencies, 442–444 suspension feeding mechanics, 442 foot and locomotion, 449–451 freshwater mussel genera, 489f habitat and abiotic interactions, 472 air exposure, 475–476 depth effects, 475 effects of pH, 474–475 macro-and microhabitat characteristics, 472–473 oxygen, 475–476 substrate, sediment, and stream flow preferences, 473–474 temperature, 476 invasive bivalves, 483–484 larval morphology, 459–460 life-cycles, 460–461 Sphaeriids and Corbicula, 466–467 unionid, 461–466 Nearctic Unionoidea diversity, 426 family-level relationships, 426–428 molecular evolution, 431 phylogeography, 431 population genetics, 431 species diversity and distributions, 428–431 North American Sphaeriidae diversity, 431–432 diversity and distributions, 433 polyploidization, 433 supra-specific relationships, 432–433 phylogenetic relationships, 424 physiology circulation, 451–452 excretion, 452–453

1075

neural systems, 453–454 osmoregulation, 452–453 respiration, 451–452 sense organs, 453–454 pollution and freshwater bivalves, 478 essential elements, 480 metals, 479 non-essential metals, 479–480 pesticides and PCBs, 481 sewage, 478–479 population demography, 468 reproductive anatomy, 454 brooding, 457–459 gonads, gametes, and fertilization, 454–457 zebra mussels, 484 Blattodea, 1067–1068 BOD. See Biochemical oxygen demand Body and appendage morphology, 764–767 Body bauplan, 138 freshwater sponge individual/population, 140f habitus of freshwater sponges, 138–139, 138f L. baicalensis, 140 meta-individual, 139–140 sexual and asexual elements, 139f Boeckella, 710–711 Bottom-up control, 76 Brachionid rotifers, 240 male mating behavior in, 252f Brachionus calyciflorus (B. calyciflorus), 235f Brachionus manjavacas (B. manjavacas), 253f female and male survival, 253f Brachionus species, 227f Brachycerans, 1043–1044 Brachyura (True crabs), 798. See also Crabs. global distribution, 799t infraorder, 805f–806f primary freshwater crabs, 806 Branchinecta raptor (B. raptor), 691f Branchinella ondonaguae (B. ondonaguae), 692f Branchiobdellae, 511 Branchiobdellida, 551 C. cirratus, 552f crustacean hosts collection, 560 culturing, 560 distribution and diversity, 552–553 ecology and behavior branchiobdellidan–host relationship, 559 habitat selection, 557–558 interspecific competition, 559 population abundance, 558 predators and parasites, 559 sympatric crayfish ectosymbionts, 559 external anatomy, 553, 554f head of oligochaete, 556f internal anatomy digestive system, 554 excretory system, 555 muscular system, 553–554 neural system, 554 reproductive system, 555

secretory glands, 554–555 vascular system, 554 locomotion, 556 nutrition, 556 phylogenetic relationships, 552 physiology, 555–556 preparation for identification, 560–561 reproduction, 556–557 sampling, 560 systematics, 551–552 X. victoriensis, 552f Branchiobdellidan–host relationship, 559 Branchiopoda (Water flea), 678f, 687 culture methods, 701 distribution and diversity, 689–690 families and number of extant genera, 688t field collection, 700 fossil record, 688–689 life history and ecology behavior, 697–698 foraging, 698–699 population regulation, 699–700 predators and parasites, 699 morphology, 690 Anostraca, 690 Diplostraca, 693–694 Laevicaudata, 691–692 Notostraca, 690–691 phylogenetic relationships, 687–688 physiology, 694–696 reproduction and development, 696–697 sample preparation, 700–701 Branchiura, 711. See also Copepoda; Thecostraca. argulid branchiurans, 736f Argulus sp., 735f control, 742 diversity and distribution, 740–741 external anatomy, 734–737 host, 741–742 infection, 741–742 internal anatomy, 737–739 reproduction and life history, 739 life history, 739–740 mating, 739 Broadcasting, 462 Brood care, 769 Brood selection, 769 Bryozoa. See Ectoprocta Bryozoans, 327–328 collection, 341–342 culturing, 342 lophophores, 330f specimen preparation, 342–343 Buccal–pharyngeal apparatus, 354–355, 355f–356f Bulbous pharynx, 193 Burrowing water beetles. See Noteridae

C

Caddisflies. See Trichoptera Caddisfly larva (Glossoma nigrior), 864 Caenis horaria (C. horaria), 880f Caenogastropoda, 389, 398

1076

Caenogastropods, 393 ampullarids, 394 Bithyniidae, 394 Hydrobiidae, 395 Littorinidae, 395 Neritidae, 394 pomatiopsids, 395 Stenothyridae, 395 Thiaridae family, 395 valvatids, 394–395 viviparids, 393–394 Calanoida, 710–711 Calopterygoidea, 895 Calopteryx splendens (C. splendens), 899f Calyptostoma, 616f Calyptostomatidae, 615–616 Calyptostomatids, 616 Cambarus dubius (C. dubius), 814f Caminella peraphora (C. peraphora), 608–609 Campanulates, 242 Candona elliptica (C. elliptica), 764f Cannibalism, 1063 Canopy fogging, 944–945 Canthocamptidae, 711 Capsules, 196 Carabidae Latreille (Ground beetles), 1022 Carapace. See Hinged calcite shell Caridea (Shrimps), 798, 800f. See also Decapoda. collecting and culturing, 837 diets, 830 distribution and global diversity, 803–804 habitat selection, 824 lateral view, 808f preservation, 839 as prey, 832 reproduction and life history, 821–822 systematic and phylogenetic relationships, 800 “Carina”, 385 Central nervous system (CNS), 191–192, 191f, 812–813 Cephalion, 214–216 Cercopagis pengoi (C. pengoi), 691f Cerithoidea, 390 Chaetonotida, 211–212, 212f–213f, 215f Chaetonotus, 218 Chaetonotus hstrix (C. hstrix), 215f Channelization, 41 Chaoborus (phantom midges), 859f Chemoreception, 677–678, 785 Cherax (Australian crayfish), 805, 826 Chironomidae, 618–619 Choanocytes, 142, 143f Choanoflagellates, 120–121 Chronogaster troglodytes (C. troglodytes), 282f Chrysomelidae Latreille (Leaf beetles), 1028–1029 Chrysomonads, 121 Chytridiomycetes, 125 Cicindis horni (C. horni), 1022f Cilia, 187–188, 188f Ciliated protozoa, 122–123. See also Flagellated protozoa. ciliate species, 124

  Subject Index

anaerobic, 125 in soil, 125 form and function in, 124f free-living species, 123 microbial loop, 123–124 Cingulum, 232 Circulatory system aquatic insects, 853–854, 854f Arthropoda, 595 Copepoda, 717 Decapoda, 810–812 Hirudinida, 571–572 Ostracoda, 767 Cirripedia, 711–712 A. improvisus, 745f balanid cirriped, 743f distribution, 747 ecology, 746–748 external anatomy, 742–743 impacts and control, 747–748 internal anatomy, 743–744 reproduction and life history, 744–746 CITES. See Convention on International Trade in Endangered Species of Wild Fauna and Flora Cladocera (Water-fleas), 689, 694 Cladoceran eggs, 697 Clam shrimps. See Conchostraca Claws, 351 eutardigrade, 353f morphologies, 352f reduction, 353–354 tardigrade, 352f Clearance rates. See Filtration rates Cliff water beetles. See Aspidytidae Clitellata, 510 Clitellum, 529, 536 Clithon, 395f Cneoglossa edsoni (C. edsoni), 1028–1029 Cneoglossidae Champion, 1029 Cnidaria, 159–161 biology external anatomy, 164–166 internal anatomy, 166 physiology, 166–167 reproduction and life history, 168–171 body forms, 160f collection, 175–176 culturing, 176 distribution and diversity, 162–164 ecological aspects bioindicators, 175 competition, 173–174 dispersal, 174 ecological impacts, 174–175 invasive species, 175 locomotion, 174 ecology and behavior feeding behavior, 172–173 habitat selection, 171–172 physiological constraints, 172 predators, 173 features, 159 global distribution, 164t Hydra, 164f

phylogenetic relationships, 161–162 specimen preparation, 176–177 systematics, 159–161 taxonomy and nematocyst types, 163t Cnidarians, 159, 160f nematocyst, 161f Cnidocysts, 159 Cnidome, 159 CNS. See Central nervous system Coarse particulate organic matter (CPOM), 990 Cockroaches. See Blattodea Coelom, 329 Coelomocytes, 537 Coelomoducts. See Gonoducts Cohesion species concept, 6–7. See also Evolutionary species. ecological factors, 7 evolutionary lineages, 7 genetic exchangeability, 7 speciation and generation process, 7 COI. See Cytochrome c oxidase subunit I Coleoptera (Aquatic beetles), 1004 collecting, 1017–1018 community patterns, 1015 conservation and environmental susceptibility, 1016 culturing, 1018 dispersal, 1014–1015 diversity and global distribution, 1004–1008 external anatomy, 1009 families, 1005t–1007t feeding behavior, 1015 herbivory, 1016 predation, 1016 habitat associations, 1013–1014 internal anatomy, 1010 digestive system, 1010 neural system, 1010–1011 reproductive system, 1011 respiratory and muscular systems, 1010 life cycles and reproduction, 1013 morphological adaptations, 1009–1010 physiological adaptations, 1011 to combat microorganisms, 1012 osmotic and ionic adaptations, 1012 respiratory adaptations, 1011–1012 sensory adaptations, 1012 preservation, 1018 systematic and phylogenetic relationships, 1008 variation in size, 1009 Collectors, 72, 885 Collembola (Springtails), 850. See also Hexapoda (Insects). aquatic, 865 basal piece and arms, 865f collecting, 868 Collophore, 864 courting, 850f culturing, 868–869 development, 866 ecology, 867 constraints, 867–868 feeding behavior, 868 habitat, 867

Subject Index

parasites, 868 physiological adaptations, 867–868 predators, 868 external anatomy, 865–866 paddle-shaped structure, 866f phylogeny, 865 reproduction, 866 species diversity, 865 specimen preparation, 869 Collophore, 864 Collothecacea, 233f, 234–235 Colonial rotifers, 244, 245f. See also Sessile rotifers. C. unicornis, 245 colony formation, 246f, 247t allorecruitive, 245 autorecruitive, 245 geminative, 246 F. ringens, 244 hypotheses, 246 juveniles, 245 Coloration, 904, 905f Comb-clawed cascade beetles. See Meruidae Competition, 76. See also Predation. Branchiobdellida, 559 Cnidaria, 173–174 Decapoda, 831 exploitative, 76 gastropod, 405 Porifera, 148–149 Rotifera, 261 Competitive exclusion, 76 Conchology, 434–435 Conchostraca (Clam shrimps), 687–688 Conglutinates, 462–463, 465f Contulma paluguillensis (C. paluguillensis), 974f Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES), 57 Cool springs, 34 Copepoda, 710–711. See also Branchiura; Thecostraca. adaptations, 720–722 behavior, 722 collection, 748 conservation, 728–732 culturing, 748 distributional patterns, 727–728 diversity, 726–727 external anatomy, 712–717 cyclopid, 715f, 719f diaptomid calanoid, 714f harpacticoid, 716f SEM-colored specimens, 713f feeding, 722–723 habitats of freshwater, 729f–732f importance for humans food, biological indicators, and source of chitin, 733–734 mosquito larvae biological control, 732–733 pathogenic agents intermediate hosts, 733 internal anatomy circulatory system, 717

1077

digestive system, 717 excretory system, 717–718 neural system, 718 reproductive system, 718 population regulation food density and quality, 724–725 heavy metals, 725 larval mortality, 725 impact of parasites, 725–726 pH, 724 predator control, 725 temperature, 723–724 reproduction and life history, 718–720 specimen preparation, 748–749 Corbicula, 425–426, 466–467. See also Dreissena. diversity, 434 in North America, 433 Cordylophora, 165f, 167f asexual reproduction in, 169–170 colonies, 164–165, 167 distribution, 162, 164t dormancy stage, 172 dreissenid mussels with, 172f hydranths, 173, 173f nematocysts in, 162f polyps, 171 prevalence, 162 species, 161 Cornops aquaticum (C. aquaticum), 1070 Corona caps, 226 Corvospongilla mesopotamica (C. mesopotamica), 137–138, 137f Corydalinae (Dobsonfly), 1063 lineages, 1061 Costae, 385 CPOM. See Coarse particulate organic matter Crabs, 798 collecting and culturing, 838–839 diets, 830–831 distribution and global diversity, 805–807 dorsal view, 810f–811f habitat selection, 829 infraorder Anomura, 807f preservation, 839 as prey, 832 reproduction and life history, 823 systematic and phylogenetic relationships, 802–803 Craspedacusta sowerbii (C. sowerbii), 168f life cycle, 168f medusae budding stages in, 169f Crawling water beetles. See Haliplidae Aubé Crayfish, 100, 798 behavior agonistic, 831f cambarid copulatory, 827f burrows, 827f chelate in, 807–808 collecting and culturing, 837–838 diets, 830 differences in color pattern, 818f distribution and global diversity, 804–805 dorsolateral view, 809f habitat selection, 824–829

infraorder Astacidea, 801f pool habitat, 823f preservation, 839 as prey, 832 reproduction and life history, 822–823 stages of molting, 815f systematic and phylogenetic relationships, 800–802 thoracic region, 812f ventral view, 816f Cretacimermis protus (C. protus), 274f Creutzeria (Histiostomatidae), 613 Crustacea, 671, 798f collection, 682 culturing, 682 development, 678 distribution and diversity, 672–673 external anatomy, 673, 673f appendages, 673–675, 674f exoskeleton, 673 shrimp, 674f feeding behavior, 681 growth, 678–679 habitat selection, 679–680 internal anatomy and physiology circulation and respiration, 675 crayfish chelae, 675f digestive system, 675 fluid and solute balance, 675–676 neural system and receptors, 676–677 Murray River crayfish, 682f Opossum shrimp, 678f ostracod, 672f physiological constraints Argulus fish lice, 680f hardness and pH, 681 salinity, 680 temperature, 680–681 predators and parasites, 682 reproduction, 678 specimen preparation, 682 systematic and phylogenetic relationships, 671–672 taxonomic groups, 672t Cryptobiosis, 147 anhydrobiosis, 361–362 anoxybiosis, 361 cryobiosis, 361 osmobiosis, 361 Cryptomonads, 122 Cryptosporidium hominis (C. hominis), 125 Cryptosporidium parvum (C. parvum), 125 Ctenidae (Wandering spiders), 603 Ctenidia, 441 Ctenostomata, 328 Cumacea, 782–783 Cupelopagis vorax (C. vorax), 233f Curculionidae Latreille (Weevils/snout beetles), 1029 Cuticle, 276 Cuticula, 904 Cyathobodonids, 122 Cybaeidae, 603 Cyclestheria hislopi (C. hislopi), 693f Cyclestherida (Tropical clam shrimp), 693–694

1078

Cyclomorphosis, 242, 369 Cylindroiulus punctatus (C. punctatus), 662f Cyphon larvae, 1012 Cyprididae, 772, 774–775 Cyrtonia tiba (C. tiba), 234f Cytochrome c oxidase subunit I (COI), 759–760, 995–996, 1058 Cyzicus sp., 691f

D

Daily vertical migration (DVM), 243–244 Daphnia pulicaria (D. pulicaria), 695f Daphnia sp., 691f Darwinula stevensoni (D. stevensoni), 766f Decapoda, 798, 800f collecting and culturing crabs, 838–839 crayfish, 837–838 shrimps, 837 diet crabs, 830–831 crayfish, 830 shrimps, 830 dispersal, 829–830 distribution and global diversity crabs, 805–807 crayfish, 804–805 shrimps, 803–804 ecological importance, 832–833 commensals, 833–834 crayfish impacts, 837 interactions in Missouri streams, 835–836 interactions in upper Midwest lakes, 836–837 parasites, 833–834 pathogens, 833–834 population regulation, 834–835 external morphology, 807 habitat selection, 823 crabs, 805–807 crayfish, 824–829 shrimps, 824 internal anatomy biramous appendages, 811f circulatory system, 810–812 digestive system, 809 excretory system, 810–812 neural system, 812–813 respiratory system, 809–810 interspecific and intraspecific competition, 831 physiology aspects circadian rhythms, 815 ecdysis, 815–816 to fresh water, 819–821 life history, 821–823 pigmentation color, 817–819 reproduction, 816–817, 821–823 reproductive hormones, 817 respiration, 813–814 thermal control, 814–815 tolerances, 814–815 predation, 831–832 crabs, 832 crayfish, 832 shrimps, 832

  Subject Index

preservation, 839 species of, 798t systematic and phylogenetic relationships, 798–800 crabs, 802–803 crayfish, 800–802 Crustacea, 798f global distribution, 799t Decapods, 59, 798 Demanian system, 278 Demographic exchangeability, 7 Density-dependent control competition, 76 predation, 76–77 Density-independent control, 77–78 Deposit feeders. See Collectors Deposit feeding, 448–449 Deutonymph, 642–643 Deutonymphal astigmatan mite, 612f Diapause, 985–986 diapausing embryos, 237 “diapausing” eggs, 697 DIC. See Differential interference contrast Differential interference contrast (DIC), 221, 304, 348–349, 560 Diffusion feeding, 127 Digestive system Branchiobdellida, 554 Coleoptera, 1010 Copepoda, 717 Crustacea, 675 Decapoda, 809 Nemata, 278–283 Ostracoda, 767 Peracarida, 784 Syncarida, 784 Tardigrada, 359 Dilution methods, 130 Dimictic process, 45 Dinoflagellates, 122 Dip net, 59, 942–943 Diphascon sp., 359f Diplopoda (Millipedes), 661 Anadenobolus sp., 662f anatomy external morphology, 662 internal anatomy, 662–663 biogeography and diversity, 662 C. punctatus, 662f culturing, 668 ecology and behavior of freshwater aquatic Australian millipedes, 666 O. gracilis, 665f P. angustus, 665f physiological problems, 664–665 S. semiaquaticum, 667f subterranean habitats of Europe, 666–667 swamps and rivers, 666 tropical islands and Amazonian floodplains, 665–666, 665f evolution, classification, and phylogenetic relationships, 661–662 extraction, 667 fixing and mounting, 667–668 N. polydesmoides, 663f

physiology defensive glands, 664 nervous system, 663–664 neurosecretion, 663–664 osmoregulation and water balance, 663 respiration, 663 toxic substances, 664 sampling methods, 667 Diplostraca, 687–688, 693 Diptera (True flies), 98–100, 1043 aquatic/semiaquatic, 1045t–1047t collecting, 1057 distribution and diversity, 1044 ecology and behavior, 1053 exemplar pupae of, 1051f feeding behavior, 1056 habitat selection flowing waters, 1053–1054 special habitats, 1054–1056, 1055f standing waters, 1054 larval morphology, 1044–1050 life cycles, 1050–1052, 1051f phenology, 1053 phylogenetic relationships, 1043–1044, 1044f predators, parasites, and parasitoids, 1056–1057 preserving specimens for identification, 1058 pupal morphology, 1050 rearing Diptera larvae, 1057 means for associating life stages, 1058 pupae, 1057–1058 reproduction oviposition, 1052–1053 swarming and mating, 1052 Dipterans, 857 Displayers, 463–465 Dissolved organic carbon (DOC), 252–253 Dissolved organic matter (DOM), 70 Dissolved oxygen, 770–771 Diving-bell spider. See Argyroneta aquatica (A. aquatica) Dobsonfly. See Corydalinae DOC. See Dissolved organic carbon Dolomedes, 604 Dolomedes aquaticus Goyen, 604–605 Dolomedes tenebrosus (D. tenebrosus), 606f Dolomedes triton (D. triton), 604–605 Dolomedes vittatus Walckenaer, 604–605 DOM. See Dissolved organic matter Donacia cf. subtilis, 1028f Dormant egg. See Opsiblastic egg Dorsoventral musculature (DVM), 436–437 Double-stranded breaks (DSB), 241 Doubly uniparental inheritance (DUI), 431 Dragonflies. See Odonata Dreissena, 425–426, 438f, 440, 457, 466–467 diversity, 434 in North America, 483–484 Dreissenids, 102 Drift net sampler, 943 Drifting, 1009–1010 Drumming. See Vibrational communication Dryopidae Billberg (Long-toed water beetles), 1029–1030 DSB. See Double-stranded breaks

Subject Index

Dugesia benazzi (D. benazzi), 191f DUI. See Doubly uniparental inheritance DVM. See Daily vertical migration; Dorsoventral musculature Dwarf males, 237 Dyar’s Law, 1050–1052 Dyar’s rule, 958–959 Dysmicohermes clade, 1061 Dytiscidae Leach (Predaceous diving beetles), 1022–1024

E

Earthworms. See Oligochaeta Ecdysis, 368, 855 Decapoda, 815–816 ECM. See Extracellular matrix Ectoprocta (Bryozoa), 327–328 dispersal, 337 ecological interactions biotic interactions, 338–339 parasites, 338 predators, 338 ecology of species Hislopidae family, 340 Lophopodidae family, 340 Pectinatellidae family, 339 Plumatellidae family, 339–340 external and internal anatomy, 328–329 body wall, 329–331 coelom, 329 neural system, 329 food, feeding, and digestion, 331 habitats, 336–337 life history temperate regions, 337–338 tropical regions, 338 origin and evolutionary development, 328 ctenostomata, 328 phylactolaemata, 328 reproduction asexual dormant bodies, 332–335 colony fragmentation and fission, 332 sexual reproduction and larvae, 335–336 swimming zooids, 332 zooid budding, 332 Ectosymbiosis, 198–199 Electromigration method, 130–131 Elimia, 393, 397–398, 409 Elmidae Curtis (Riffle beetles), 1030–1031 Elmis sp., 1030f Elodes, 1012 Emergence traps, 944 Encystment, 362 Endocyst, 329 Enrichment, 130 Entoprocta, 340–341 Ephemera danica (E. danica), 877 male imago, 878f male subimago, 878f Ephemera lineata (E. lineate), 879f Ephemeral streams. See Intermittent streams Ephemeroptera (Mayflies), 873 behavior of winged stages Baetis, 884f

1079

Ecdyonurus sp., 884f emergence, 881–882 flight activity, 883–884, 883f H. lauta, 882f subimaginal stage, 884–885 collection, 888 distribution, 876–877 diversity, 876–877 ecology and behavior, 885–886 E. virgo, 888f environmental changes, 886–887 feeding behavior, 885 habitat selection, 884–885 human effects, 886–887 interactions with humans, 887–888 parasitic and commensal relationships, 886 physiological constraints, 885 predators, 886 endemism, 876–877 extant families, 875t–876t history and paleontology, 873 imagos external anatomy A. albinatii, 878f abdomen, 878 head, 877 thorax, 877–878 winged stages, 877 internal anatomy of nymphs, 880 life cycle, 880–881 nymphs external anatomy, 878 abdomen, 879–880 B. fuscatus, 879f C. horaria, 880f E. lineate, 879f head, 878–879 P. submarginata, 880f R. semicolorata, 879f S. ignita, 880f thorax, 879 phylogenetic relationships, 874–876, 877f rearing, 888 reproduction, 884 specimen preparation, 888 speciose, 876t systematics, 874 Ephemeroptera, 98 Ephemeroptera, Plecoptera, and Trichoptera (EPT), 995–996 Ephoron virgo (E. virgo), 888f Epigean habitats Amphipoda, 788–789 Isopoda, 789 Mysidacea, 789 Epimetopidae Zaitzev (E. Zaitzev), 1031 Epipleura, 211–212 Epistome, 328–329 EPT. See Ephemeroptera, Plecoptera, and Trichoptera Equivalve, 435 Erpobdellidae body shape, 568f transverse sections, 567f Esophagus, 359–360 EST. See Expressed sequence tag Ethanol (ETOH), 793, 869

Eubranchipus bundyi (E. bundyi), 690f Eucypris virens (E. virens), 772–773, 777 Euglena, 113, 115–116 Euglenids, 121 Euhrychiopsis lecontei (E. lecontei), 1030f Eulichadidae Crowson (Forest stream beetles), 1031 Eustigmaeus frigida (E. frigida), 615f Eutardigrade. See also Tardigrada (Water bears). buccal–pharyngeal apparatus, 357f morphology, 348f musculature and ventral ganglia, 348f reproductive apparatus, 363f in vivo micrographs, 357f Eutely, 232 Evolutionary scandal, 249 Evolutionary species, 6 Excretory system Arthropoda, 595 Branchiobdellida, 555 Copepoda, 717–718 Decapoda, 810–812 Hirudinida, 570–571 Nemata, 283–284 Rotifera, 236–237 Tardigrada, 360 water mites, 633–634 Exoskeleton eclosion and sclerotization, 643 evolution, 632–633 Exploitative competition, 76 Expressed sequence tag (EST), 759–760 Extracellular matrix (ECM), 188–189

F

“Facultative Water” beetles, 1013–1014 FADA. See Freshwater Animal Diversity Assessment False rhabdites, 188 “False Water” beetles, 1013–1014, 1037–1038 Fashing. See Hormosianoetus mallotae Faunal differences, 857–858 Faunus ater (F. ater), 391f Feeding behavior aquatic insects, 863–864 Cnidaria, 172–173 Coleoptera, 1015 herbivory, 1016 predation, 1016 Collembola, 868 Crustacea, 681 Diptera, 1056 Ephemeroptera, 885 Gastrotricha, 220 Hirudinida, 577–578 inland water Annelida, 518 inland water Polychaeta, 525–526 Nemata, 288 Nematomorpha, 319 Oligochaeta, 543 Ostracoda, 773–774 Peracarida, 790–791 Plecoptera, 939–940

1080

Porifera, 146–147 spiders, 601–602 Female reproductive organs, 768 FFG. See Functional feeding group Filterers, 72 Filtration rates, 260, 442–444 Fine particulate organic matter (FPOM), 72 Fingernail clams, 472 Fishing spiders. See Pisauridae (Nursery-web spiders) Flagellated protozoa, 120 choanoflagellates, 120–121 chrysomonads, 121 cryptomonads, 122 cyathobodonids, 122 dinoflagellates, 122 euglenids, 121 form and function in, 121f freshwater ciliated protozoa, 123f haptomonads, 122 heteromitids, 122 heterotrophic flagellates, 120 I. chabelardi, 122 Flatworms. See Platyhelminthes Flight intercept traps, 60 Floscularia ringens (F. ringens), 244 Flow cytometry, 130–131 Foraminiferans, 119 Forest stream beetles. See Eulichadidae Crowson Fossil record, 760 FPOM. See Fine particulate organic matter FPZ. See Functional process zone Fredericella species, 328 Free living, 286 Free-living protozoa, 117f biological diversity ameboid protozoa, 118–120 ciliated protozoa, 122–125 flagellated protozoa, 120–122 functional groups, 116–118 collection, 129–130 culturing, 130–131 diversity assessment, 129–130 ecology functional roles, 126–127 groundwater protozoa, 128–129 life in low-oxygen environments, 128 protozoa and ecosystem function, 127–128 symbiotic associations, 127 nature as group, 113–114 protozoan species, 114–115 protozoan cell, 115–116 Fresh and marine earthworms. See Annelids Freshwater Animal Diversity Assessment (FADA), 760–761 Freshwater invertebrates, 93. See also Inland water invertebrates. A. abnormis, 20f Acilius, 20f acquiring energy aquatic food webs, 72–75 functional feeding groups, 71–72 habitat traits, 71 traits in general, 71

  Subject Index

aquatic–terrestrial ecotones, 70–71 B. zetlandicus, 17f biomonitoring, 97, 97t C. centrura, 18f C. spinifer, 16f Clitellata, 17f Clithon and H. altilis, 16f Coleps sp., 14f Collembola, 19f in commerce aquaculture, 93–94 freshwater pearls, 95–97, 95f–96f insect jewelry, 95–97, 95f–96f invertebrate–fish connection, 94–95 density-dependent control competition, 76 predation, 76–77 density-independent control, 77–78 Diptera, 19f direct market economic benefits, 100–101 damages, 101–102 diseases Crustaceans, 107 habitats of human disease vectors, 105–106 incidence, 104–105 insect vectors of human disease, 106 snails, 106–107 E. muelleri, 15f E. nevadensis, 18f Epiphanes brachionus spinosa, 15f freshwater branchiopod crustaceans, 18f crustaceans, 19f mites, 17f functional relationships, 65 G. difficilis, 17f green hydra and C. jsowerbyi, 14f H. diabasia, 20f Hemiptera, 20f indirect nonmarket economic benefits, 102 damages, 102–103 lentic ecosystems, 68 physical and ecological characteristics of, 68 spatially distributed habitat features, 69–70 temporal factors, 68–69 lotic ecosystems billabong murray river, 67f hydrogeomorphic patterns, 67–68 lateral riverscape patterns of organisms, 68 longitudinal patterns in habitat and organisms, 66–67 physical and ecological characteristics of, 66 riverine landscape, 67f medicinal leeches, 103–104, 103f nuisance aquatic insects, 98 diptera, 98–100 ephemeroptera, 98 trichoptera, 98 Odonata, 20f P. pachysoma, 17f P. paludosus, 19f Prostoma, 16f

R. fuscula, 20f S. mediterranea, 16f S. semiaquaticum, 17f Sialis larva, 20f stresses to livestock, 104 U. gracilis, 15f wildlife from biting flies, 104 Freshwater Mesostigmata, 608–609 Freshwater mussel lures, 427f Freshwater Sarcoptiformes, 609. See also Parasitiformes; Prostigmata. Astigmatina, 611–613 Oribatida, 609–611 Freshwater snails, 100 Functional feeding group (FFG), 863 Functional process zone (FPZ), 32 Furca, 764–765

G

Gambusia spp. See Mosquito fishes Gammarus minus (G. minus), 788–789 Gastrodermis, 159 Gastromermis anisotis (G. anisotis), 284f egg clusters, 284f infective-stage juvenile, 284f slightly greenish juvenile, 296f Gastropoda (Snails), 388–389 B. glandiformis, 389f collection, 412–413 conservation ecology, 408 channeled apple snail, 412f conservation and propagation, 410–411 hydrobiidae ecology, 410 invading New Zealand mud snail, 412f invasive species, 411–412 leptoxis foreman oviposition, 411f pleuroceridae ecology, 409–410 culturing, 413 diets, 399–400 dispersal, 402 distribution and diversity, 392 caenogastropods, 393–395 hotspots, 393f pulmonates, 396 ecological determinants, 404 biogeographic factors, 404–405 competition, 405 flexibility in shell architecture, 408 flow and hydroperiod, 405 predation, 405 snail response to predators, 405–408, 407t watershed connections and chemical composition, 404 evolutionary pathways, 392 feeding effects, 401–402, 401t gastropod shells, 385 grazers, 401–402 habitat choice, 399 as intermediate hosts of human disease, 106–107 North American, 390t pachychilidae, 392f phylogenetic relationships, 389–392 population regulation

Subject Index

food quality, 402 parasitism, 402–403 production ecology, 403–404, 403t reproduction and life history, 397 caenogastropoda, 398 life-cycles in freshwater snails, 397f pulmonata, 398–399 specimen preparation and identification, 413 systematic studies, 391–392 systematics, 389–392 T. winteri, 388f in United States, 410t Gastropods. See Gastropoda (Snails) Gastrotricha, 211 biology external anatomy, 214–216 internal anatomy and physiology, 216–217 reproduction, 217–218 collection, 220–221 culturing, 220–221 distribution and diversity, 214 ecology and behavior density and biomass, 219t feeding behavior, 220 habitat selection, 218, 219t physiological constraints, 218–220 predators and parasites, 220 phylogenetic relationships, 212–214 specimen preservation, 220–221 systematics, 211–212 Gatherers. See Collectors Genital pores, 536–537 Genitalia, 905, 908f. See also Reproductive system. Geographical Information Systems (GIS), 472–473, 481 Georissidae Latreille (Minute mudloving beetles), 1031–1032 Georissus crenulatus (G. crenulatus), 1032f Gerris water striders, 958, 958f GIS. See Geographical Information Systems Glandularia, 634–635 Global Positioning Systems (GPS), 485–486 Glochidia, 424 Glomerin, 664 Glossiphoniidae body shape, 568f transverse sections, 567f Gonoducts, 523 Gonopods, 808–809 Gordian worms, 303 Gordiid larvae, 314 GPS. See Global Positioning Systems GR. See Guild Ratio Graphoderus sp., 1023f Green glands. See Antennal glands Groundwater protozoa, 128–129 Guild Ratio (GR), 259 Gyrinidae Latreille (Whirligig beetles), 1024–1025

H

Habitat selection, 788 aquatic insects, 856 brine fly, 857f

1081

caddisfly net, 858f cranefly larvae, 861f deep-water habitats, 860–861 ephemeral vs. permanent, 856–857 epigean vs. hypogean, 861 fairy shrimp, 857f infauna vs. epifauna, 861 littoral zone, 860–861 lotic vs. lentic, 857–859 mosquito larvae, 860f oceans and saline lakes vs. freshwater, 857 phantom midge, 859f planktonic vs. benthic, 859–860 Playa Lake, 856f surface, 860–861 water strider on pleuston, 860f whirligig beetle, 860f Arthropoda, 596 Cnidaria, 171–172 Gastrotricha, 218, 219t inland water Annelida, 517 inland water Polychaeta, 525 Nemata, 286 Nematomorpha, 318–319, 318f Oligochaeta, 542 Ostracoda, 769–770 Peracarida epigean habitats, 788–789 Hyalella amphipod species, 788f hypogean habitats, 789–790 Platyhelminthes, 197, 197f–198f Porifera, 145–146 Shrimps, 824 Habitats, inland water. See also Polychaeta, inland water. global variations in, 24 alpine habitats, 24–25 Australia, 28 oceanic islands, 26–28 polar zones, 24–25 temperate zones, 25 tropical zone, 25–26 lentic ecosystems, 43–50 lotic ecosystems, 28–41 subterranean habitats, 41–43 Habitus of Corydalidae larvae, 1060f Habitus of larval spongillafly, 1064f Habrophlebia lauta (H. lauta), 882f Hairworms. See Nematomorpha (Horsehair worms) Halacaridae, 613–614 Halacaroidea, 613 Halacaridae, 613–614 Pezidae, 613–614 Haliplidae, 1004–1008 Haliplidae Aubé (Crawling water beetles), 1025–1026 Haliplids, 1025 Hand picking, 943 Haptomonads, 122 Haptonema, 122 Harpacticoids, 723 Head arrester system, 904–905 Headwaters, 34 Heat shock proteins (HSP), 242

Helioflagellates, 121 Heliozoans, 120 Hellgrammites, 1060 Helobdella stagnalis (H. stagnalis), 576 Helophoridae Leach, 1032 Helophorus sp., 1032f Hemiptera (True bugs), 951 A. herberti, 951–953, 952f collection, 960–961 conservation, 960 culturing, 960–961 distribution, 951–953 diversity, 951–953, 952f ecology and behavior body size, 958–959 evolution of paternal care, 958–959, 959f feeding, 957 flood survival, 959 Gerris water striders, 958f importance to humans, 959–960 L. medius, 957f mating, 958 external anatomy, 953 global species richness, 954t life cycle, 956–957 locomotion aquatic, 953–955 flight, 955–956 Notonecta, 955f phylogenetic relationships, 951–953 physiology osmoregulation, 956 predators, 956 respiration, 956 reproduction, 957 specimens preparation, 960–961 Hemocyanin, 853 Hemoglobin in hemolymph of larval Chironomidae, 1050 in larval Chironomidae, 1050 “Herbivorous” cladocerans, 698 Herbivory, 1015–1016 Hess sampler, 943 Heteroceridae Mac Leay (Variegated mudloving beetles), 1032–1033 Heteromitids, 122 Heterotrophic flagellates, 120 Hexapoda (Insects), 849. See also Collembola (Springtails). aquatic insects backswimmer Neoplea, 854f circulatory system, 853–854, 854f collecting, 868 culturing, 868–869 ecology and behavior, 855–864 evolutionary diversification, 851 excretion, 854–855 generation time, 855 hemimetabolous and holometabolous development, 856f internal tracheal system, 853f literature sources, 851 metamorphosis, 855 osmotic balance, 854–855 plate-like tracheal gills, 854f

1082

regions of insect anatomy, 852f respiratory system, 852–853 specimen preparation, 869 tagmatization, 852 components, 850 phylogenetic relationships, 850–851 Hexapods. See Hexapoda Hexarthra, 235, 236f, 238 Hibernaculae, 334f Hindgut, 359–360 Hinged calcite shell, 763 Hirudinea (True leeches), 511 Hirudinida (Leeches), 565 collection, 583 crawling motions, 572f dispersal, 576–577 distribution and diversity, 566–567 ecological interactions behavior, 579–581 feeding behavior, 577–578 predators, 578–579 ecotoxicology, 582–583 external anatomy distinguishing characteristics, 567 external sensory structures, 568 segmentation, 567 suckers, 567–568, 569f internal anatomy alimentary canal, 568–570, 570f circulatory and respiratory systems, 571–572 excretory system, 570–571 locomotion, 572 reproductive system, 570 life history, 575–576 phylogenetic relationships, 566 physiology changes during winter, 573–574 ionic and osmotic regulation, 574 respiratory tolerance and adaptations, 572–573 population regulation, 581–582 preparation of specimens, 583–584 rearing, 583 reproduction, 574–575 and parental care, 571f systematics, 566 Hirudiniformes, 567f–568f Hislopia, 328, 329f egg release in, 336 Hibernaculae in, 335 mixing cilia in, 331 zooid in, 335 Hislopia malayensis (H. malayensis), 334f, 336f Hislopia natans (H. natans), 332f Hislopidae family, 340 Histiostomatid mite, 612f Homocaligidae, 614f Homocaligids, 614 Hormosianoetus mallotae (Fashing), 613 Horsehair worms. See Nematomorpha Host trapping, 430f, 465 HSP. See Heat shock proteins Human disease vectors, 106t

  Subject Index

habitats of, 105–106 insect vectors, 106 Hydra species, 175, 175f Hydra viridissima (H. viridissima), 167, 170f Hydrachnidae, 628–629 Hydrachnidiae (Water mites), 617–626. See also Arachnids. biogeographic and ecological characteristics, 620t–625t, 626 classification, 619 diversity, 619 ecology, 646 external morphology, 626 A. montanus, 631f adult exoskeleton evolution, 632–633 Chelicerae of Hydrachna sp., 631f Deutonymphs and adults, 628–632 extendible gnathosoma, 631f Hydryphantoidea, 626–628 of larvae, 626 palps of Trichothyas sp., 631f soft-bodied condition, 631–632 general relationships, 618 habitats and assemblages, 645 interstitial habitats, 645 lakes, permanent ponds, marshes, swamps, and bogs, 645–646 phytotelmata, 646 riffle habitats, 645 springs, 645 stenothermic pools, 645 temporary pools, 646 impact as parasites effect on other mites sharing same host, 647 effects on individual hosts, 646–647 variation in mite load on hosts, 647 impact as predators, 647–648 importance as prey, 648–649 internal anatomy, 633 Arrenurus species, 634f digestive system, 633 excretory system, 633–634 H. despiciens, 628f Limnesia sp., 630f neural system, 634–635 reproductive system, 635 respiratory system, 634 S. ecphyma, 627f S. ecphyma, 629f life history, 635–645 attaching to host and engorgement, 641–642 detaching from host, 641–642 deutonymph, 642–643 direct spermatophore transfer, 643–645 eclosion and sclerotization of exoskeleton, 643 eggs, 636–637 indirect spermatophore transfer, 643 larvae, 637–640 larvae of Eylais sp., 639f phenology of larvae, 642 protonymph, 642 raft of larval Eylais, 636f

searching for hosts, 637–638 seeking and locating hosts, 639–640 selecting feeding site on host, 640 tritonymph, 643 variations among taxa, 638–639 origin and phylogeny, 618–619 potential as environmental quality indicators, 649 Hydraenidae Mulsant (Minute moss beetles), 1033 Hydraulics, 36–37 Hydrobiidae, 395, 410 Hydrocanthus sp., 1028f Hydrochidae, 1033, 1034f Hydrochus brevis (H. brevis), 1033 Hydrodroma despiciens (H. despiciens), 628f Hydrogeomorphic patches, 30–32 Danforth Spring, 31f Mississippi River, 31f ozarks of southeastern Missouri, 31f Tagliamento River, 31f Hydrology, 36–37 Bivalvia changes channelization, 477–478 floods, 476–477 impoundments, 477 Ostracoda, 772–773 Hydrolutos, 1070 Hydrophilidae Latreille (Water scavenger beetles), 1034 Hydroscapha natans (H. natans), 1018f Hydroscapha redfordi (H. redfordi), 1018f Hydroscaphidae LeConte (Skiff beetles), 1018 Hydrozetes spp., 610f, 611 Hydryphantoidea, 626–628 Hygrobia davidii (H. davidii), 1026–1027 Hygrobia hermanni (H. hermanni), 1026f Hygrobia tarda (H. tarda), 1026f Hygrobiidae Régimbart (Squeak beetles), 1026–1027 Hygrotus salinarius (H. salinarius), 1012 Hymenoptera, 1068 Hyphochytriomycetes, 125 Hypodermis, 278 Hypogean habitats, 789–790. See also Epigean habitats. Hypopleura, 211–212 Hyporheic zones, 41–42, 776 Hyposmocoma, 1069, 1069f Hypostome, 766

I

Ichthyodinium chabelardi (I. chabelardi), 122 ICL. See Intracytoplasmic lamina ICZN. See International Code of Zoological Nomenclature Ilyobius lineage, 1060–1061 Imago external features, 900–902 tracheal system, 909 Imagochrysalis, 643 Immature life stages, 1059 Infundibulum, 232 Inland water invertebrates, 3

Subject Index

phylogenies, 4–9 species, 4–9 using systematics and taxonomy, 9 ICZN, 9–10 identification and consistent approaches, 10 IUBS, 10 taxonomic keys, 10 annelid groups, 12 arthropod subphyla, 12–21 freshwater invertebrates key, 11 mollusca classes, 11–12 Insecta aquatic insects, diversification of, 851 literature sources, 851 Insects. See Hexapoda Instars, 855 Interference competition. See Exploitative competition Intermittent streams, 32–33 headwater prairie stream and upper three runs creek, 33f hydroperiod length, 33 Ozarks of southern Missouri, 33f resident invertebrates, 33–34 International Code of Zoological Nomenclature (ICZN), 9, 428 International Union of Biological Sciences (IUBS), 10 Intertropical Convergence Zone (ITCZ), 918 Intracytoplasmic lamina (ICL), 230–231 Invasive Alien invertebrates accumulation, 85f ecological impacts, 86–88 prediction, 88 freshwater invertebrate invasions, 83–84, 84t human vectors of dispersal, 84–86 hypothesized attributes, 86t invasive aquatic invertebrates management, 88–89 traits conferring invasion success, 86 Invasive aquatic invertebrates management, 88–89 Invertebrates. See also Freshwater invertebrates; Inland water invertebrates. collection conservatively, 58 legally, 57–58 safely, 58 culturing, 62 fixing specimens, 61–62 preservation, 61–62 record-keeping, 61 sampling, 58 benthic invertebrates, 59–60 myriad ways, 58–59 planktonic invertebrates, 60–61 Ion composition, 770 Ionic content, 39–40 Ionization radiation (IR), 241 IR. See Ionization radiation Isolation methods, 130 Isopoda, 782, 782f, 784–785 appendages in freshwater, 784f Epigean habitats, 788–789 ITCZ. See Intertropical Convergence Zone

1083

Iteroparity, 575 IUBS. See International Union of Biological Sciences

J

Jellyfish (Medusozoa), 159–161 Cnidaria classes in, 161–162 fresh-water, 166f inclusion of class Polypodiozoa, 162 “pulsing” behavior, 167 Johnstonianidae, 615–616, 616f Jumps, 238

K

Kairomone, 262–263 Kempyninae, 1064–1065 Kentrophoros, 124, 127 Kick net, 59 Kick screen, 943 Kidneys, 387 Killer shrimp, 87–88 Koenikea wolcotti Viets, 633f Kurzia longirostris (K. longirostris), 693f

L

Labial palps, 445 Lacinia mobilis, 783 Laevicaudata (Smooth clam shrimp), 691–692 Lakes, 43 biotic zonation, 45, 45f geomorphology and physicochemical zonation, 43–45 Lake Ohrid, 793 Ostracoda, 775 turnover, 45 Lansing effect, 252 Lara sp., 1031f Larva, 855 external features, 899–900 labium of odonate, 900f Larval Carabidae, 1022f Larval corydalids, 1060 Larval gill systems, 908–909 Larval sialids, 1060, 1063 Late Embryo Abundant genes (LEA genes), 242 LEA genes. See Late Embryo Abundant genes Leaf beetles. See Chrysomelidae Latreille Leeches. See Hirudinida Leiodidae Fleming (Round fungus beetles), 1035 Lentic ecosystems, 43, 68. See also Lotic ecosystems. human impacts on, 49–50 hypersaline lakes and pools, 48–49 lakes, 43 biotic zonation, 45, 45f geomorphology and physicochemical zonation, 43–45 physical and ecological characteristics of, 68 phytotelmata, 49 reservoirs, 46 spatially distributed habitat features, 69 littoral zone, 69–70 neuston, 70 zooplankton, 70

temporal factors, 68–69 wetlands, 46, 47f alluvial swamps, 46–47, 47f invertebrates of seasonal, 46 Lentic food webs, 75 Lepiceridae, 1019 Lepicerus pichilingue (L. pichilingue), 1019f Lepidodermella squamata (L. squamata), 214 Lepidoptera, 1068–1069 Lepidurus bilobatus (L. bilobatus), 691f Leptestheria compleximanus (L. compleximanus), 693f Leptodora kindti (L. kindti), 694f Lesteva longoelyrata (L. longoelyrata), 1038f Lethocerus medius (L. medius), 957f Light microscopy (LM), 142–143 Light traps, 944 Limnadia lenticularis (L. lenticularis), 693f Limnichidae Erichson (Minute marshloving beetles), 1035–1036 Limnichites sp., 1036f Limnocharid mites, 647–648 Limnocythere, 770 Limnocythere sanctipatricii (L. sanctipatricii), 766f Limnoterrestrial habitat, 226–227 Limnoterrestrial heterotardigrades, 355 Linderiella occidentalis (L. occidentalis), 690f Liocanthydrus bicolor (L. bicolor), 1027f Littoral benthic substrate, 59 Littoral zone, 69–70 plankton, 61 Littorinidae, 395 LM. See Light microscopy Long-jawed orb-weavers. See Tetragnathidae Long-toed water beetles. See Dryopidae Billberg Lophophorates, 328 Lophophore, 331 Lophopodella carteri (L. carteri), 340, 340f Lophopodidae family, 340 Lotic ecosystems, 28–29 acidity, 40 Billabong Murray river, 67f chemical environment, 39 headwaters, 34 human impacts, 40–41 hydraulics, 36–37 hydrogeomorphic patterns, 67–68 hydrology, 36–37 intermittent streams, 32–33 headwater prairie stream, 33f hydroperiod length, 33 ozarks of southern Missouri, 33f resident invertebrates, 33–34 ionic content, 39–40 lateral riverscape patterns of organisms, 68 longitudinal patterns in habitat and organisms, 66–67 oxygen requirements, 39 physical and ecological characteristics of, 66 riverine landscape, 67f rivers, 35–36 springs cool, 34 thermal, 34

1084

structure of rivers, 29 hydrogeomorphic patches, 30–32 macrosystem, 29 Mianus River, 30f network patterns, 29 Pecos River, 30f reaches, 32 stream order, 29–30 substrates, 37–38 thermal environment, 38–39 Lotic food webs, 74–75 Low-dissolved oxygen, 813–814 Lubomirskia baicalensis (L. baicalensis), 140 Lumbricoidea, 512–513 Lutrochidae (Travertine beetles), 1036 Lycosidae (Wolf spiders), 603–604 Lymnaea peregra Müller, 398 Lymnaeidae, 391 Lymnaeids, 400 Lynceus brachyurus (L. brachyurus), 692f Lyrifissures, 635

M

MAA. See Mycosporine-like amino acid Macrodasyida, 211–212, 212f Macrohectopus branickii (M. branickii), 792–793 Macroinvertebrates, 583 Macroplea mutica (M. mutica), 1028f Macrosystem, 29 Macrothrix rosea (M. rosea), 694f Majoidea, 802 Malaise traps, 60, 944 Mandibles (Md), 765–766 Mantle cavity, 440–441 Marsh beetles. See Scirtidae Marsupium, 457 Mating Branchiura, 739 Copepoda, 720 Diptera, 1052 Hemiptera, 958 Odonata, 911 Trichoptera, 987 wheel, 911 Mature nymphs, 879 Mature Sialis larva, 1060f Maxillipeds, 807 Maxillopoda, 710 Branchiura, 711 Copepoda, 710–711 Thecostraca, 711–712 Maxillulae (Mx1), 765–766 Mayflies. See Ephemeroptera Md. See Mandibles Mechanoreception, 677 Mecoptera (Scorpionflies), 1069 Medicinal leeches, 103–104, 103f Medusae, 166, 166f Medusozoa. See Jellyfish Megadriles, 510, 529 Megaloptera, 1060 distribution and diversity, 1061–1062 larval morphology and physiology, 1063 life history, ecology, and behavior, 1062–1063

  Subject Index

phylogenetic relationships, 1060–1061 Megalopteran larvae, 1063 Megareoles, 305–306 Menonts, 167 Mermithidae, 294 Meru phyllisae (M. phyllisae), 1027f Meruidae (Comb-clawed cascade beetles), 1027 Mesostigmata, 601 biology, 608–609 morphology, 607–608 Mesostigmatan mite, 607f terrestrial, 608f Metamorphosis, aquatic insects, 855 Metamorphotype, 995 Metanephridia, 541 Metaniidae, 137 Microbotrophic nutrition, 286 Microchorista philpotti (M. philpotti), 1069 Microfossils, 689 Midgut, 359–360 Millipedes. See Diplopoda Minor insect orders Blattodea, 1067–1068 Hymenoptera, 1068 Lepidoptera, 1068–1069 Mecoptera, 1069 Megaloptera, 1060–1063 Neuroptera, 1064–1067 Orthoptera, 1069–1070 Minute bog beetles. See Sphaeriusidae Erichson Minute moss beetles. See Hydraenidae Mulsant Minute mudloving beetles. See Georissidae Latreille MIP. See Mixis induction protein(s) Misumenops nepenthicola (M. nepenthicola), 606 Mites, 600, 607 Mixis induction protein(s) (MIP), 250 Molluscs (Mollusca), 383–384 classes, 11–12, 16f digestive system, 386–387 diversity, 383–384 environmental physiology, 388 excretory and neural systems, 387–388 phylogenetic relationships, 384 reproductive system and larval development, 388 respiratory and circulatory systems, 387 shell morphology, 384–385, 385f soft anatomy, 385–386, 386f systematics, 384 Mollusks. See Molluscs (Mollusca) Molting. See Ecdysis Monogenea, 181 Monogonont rotifers, 229f diapausing eggs of, 250f life cycle for, 249f Monomictic process, 45 Morphotype, 242 Mosquito fishes (Gambusia spp.), 1056–1057 Mucronothrus nasalis (M. nasalis), 611 Murray River crayfish, 682f Muscular system Branchiobdellida, 553–554 Coleoptera, 1010

Nemata, 278 Rotifera, 235–236 Mx1. See Maxillulae Mycosporine-like amino acid (MAA), 252–253 Myoretronectes paranaensis (M. paranaensis), 186 Mysidacea (Opossum shrimp), 678f, 782, 782f Epigean habitats, 788–789 Mysis diluviana (M. diluviana), 103 Myxophaga families, 1008, 1014. See also Adephaga families; Coleoptera (Aquatic beetles); Polyphaga families. H. LeConte, 1018 Lepiceridae, 1019 S. Erichson, 1019 T. Steffan, 1019–1020 Myxozoans, 125–126

N

NACODe. See North American Combined Ostracode Database Naked amebae, 118–119 NANODe. See North American Nonmarine Ostracode Database Narcotization, 749 Nautizooids, 332 Nearctic Unionoidea diversity, 426 family-level relationships, 426–428 molecular evolution, 431 phylogeography, 431 population genetics, 431 species diversity and distributions, 428–431 Nemata (Nematoda), 273 biogeography and diversity, 274–275 culturing, 300 ecology and behavior dispersal, 290f dune lakes, 290f extreme and specialized habitats, 288–291 feeding behavior and sites, 288 habitat selection, 286 invertebrate and vertebrate parasitic nematodes, 292–295 physiological constraints, 286–288 predators and parasites, 291–292 Sagehen Creek in California’s Sierra Nevada, 290f evolution and phylogenetic relationships, 273–274 external anatomy, 275 cuticle, 276 sense organs, 276–278 extraction, 298–299 fixing and mounting, 299–300 growth and reproduction, 284–286 internal anatomy digestive system, 278–283 excretory system, 283–284 hypodermis, 278 muscular system, 278 nervous system, 284 respiration, 284 sampling methods, 297 Nematocysts, 159 Nematomorpha (Horsehair worms), 303

Subject Index

collection, 321 culturing, 321–322 distribution and diversity, 304–305 ecology and behavior effects of pesticides, 319 feeding behavior, 319 habitat selection, 318–319, 318f host specificity, 316–317 physiological constraints, 319 predation, parasitism, and commensals, 319–321 sex ratios, 317 external anatomy adult free-living gordiids, 306f, 308f P. varius, 307f fossil hairworms, 305f Gordian knot, 304f internal anatomy body wall and musculature, 306 neural system, 306–308 reproductive organs, 308 phylogenetic relationships, 304 physiology, 308 reproduction and life cycle, 308, 309f egg laying, 311, 313f fertilization, 309–311 hosts and emergence, 308–309 larvae, cysts, and hosts, 312–316, 314f–315f specimen preparation, 322–323, 322f systematics, 304 Nemertea (Ribbon-worms), 205, 207t biology external anatomy, 207 internal anatomy and physiology, 207–208 reproduction and development, 208 collection and culturing, 209 distribution and diversity, 206–207 ecology and behavior, 208–209 morphological feature, 205, 206t phylogenetic relationships, 206 specimen preparation, 209 systematics, 205–206 Neoblasts, 189–190 Neochauliodes, 1061–1062 Neodermata, 181 Neohermes, 1062f Neohermes filicornis (N. filicornis), 1060f Neotoma, 761–763 Nephridia, 523 Neritidae, 394 Nervous system Diplopoda, 663–664 Nemata, 284 Tardigrada, 360 Neural systems, 453–454 Branchiobdellida, 554 Coleoptera, 1010–1011 Copepoda, 718 Crustacea, 676–677 Decapoda, 812–813 Peracarida, 785 phylum Arthropoda, 595 phylum Nematomorpha, 306–308 phylum Rotifera, 236

1085

Syncarida, 785 water mites, 634–635 Neuroptera, 1064 collecting, rearing, and specimen preparation, 1067 distribution and diversity, 1064 larval morphology and physiology, 1066–1067 life history, ecology, and behavior, 1065–1066 phylogenetic relationships, 1064 Neuropterida, 1060 Neurosecretion, 663–664 Neuston, 70 Nevrorthidae larvae, 1066–1067 Nilus curtus (N. curtus), 605–606 Niphograpta albiguttalis (N. albiguttalis), 1068–1069 Nitidulidae Latreille (Sap beetle), 1036 NODE. See Nonmarine Ostracod Distribution in Europe non-Clitellata, 510 Nonmarine Ostracod Distribution in Europe (NODE), 761–763 Noroviruses, 471 North American Combined Ostracode Database (NACODe), 761–763 North American marbled crayfish. See Procambarus “Marmorkrebs” North American Nonmarine Ostracode Database (NANODe), 761–763 North American Sphaeriidae diversity, 431–432 diversity and distributions, 433 polyploidization, 433 supra-specific relationships, 432–433 Noteridae (Burrowing water beetles), 1027–1028 Notostraca (Tadpole shrimp), 689–691 Notostracan thorax, 690 Notum, 852 Nuisance aquatic insects, 98 diptera, 98–100 ephemeroptera, 98 trichoptera, 98 Nursery-web spiders. See Pisauridae Nymphochrysalis, 642

O

Oceanic islands, 26–28 Ocellus, 278 Ochthebius corrugatus Rosenhauer (Hydraenidae), 1014, 1017 Octomyomermis troglodytis (O. troglodytis), 288–289 Odonata (Dragonflies), 894 biotic interactions, 922–923 abiotic limitations and, 924–925 intraspecific interactions, 924 parasitism and interactions, 925 predation, 923–924 body shapes and microhabitat types, 909f impact of climate change, 926–927 collection and sampling, 928–929 conservation status and biotic indicators, 927–928 culturing, 929

dispersal and migration, 918 diversity patterns, 925–926, 927f egg structure, 902–903, 903f external features imago, 900–902, 901f larva, 899–900, 899f flight, 910–911 foraging adult, 918 Anax imperator Leach, 919f larval, 917–918 habitats, 920f acidic ponds and lakes, 922 forest and shade, 922 generalists vs. specialists, 918–919 lotic waters, 921 microhabitat occupancy, 919–921 saline waters, 922 selection, 919 temporary, 921–922 terrestrial, 922 life cycle, 913, 914f adult life span, 915–916 egg development, 913 larval development, 913–914, 915f metamorphosis and emergence, 914–915, 917f prereproductive period, 915 seasonal patterns, 915 types and voltinism, 916–917 perception, 905–906 compound eyes and Ocelli, 906–907 tactile sensory organs, 907–908 phase relationships, 912f preservation, 929–930 reproduction, 911 mating systems, 911 oviposition, 911–912 sexual dimorphism, 911 respiration, 910f larval gill systems, 908–909 oxygen demands, 909 physical-gill respiration, 909–910 tracheal system of imago, 909 size, 902 systematic and phylogenetic relationships, 894 Anisoptera, 896 Anisozygoptera, 896 species numbers, 897–899, 897t–898t Zygoptera, 894–896 thermoregulation, 910 ultrastructures, 903 coloration, 904 cuticula, 904 genitalia, 905 head arrester system, 904–905 wing structures, 903–904, 904f Oligochaeta (Earthworms), 529, 530f biology E. velutinus, 539f external anatomy, 535–537 internal anatomy, 537–540 L. variegatus, 538f physiology, 540–541

1086

reproduction and life history, 541–542 spermatozeugmata in spermathecae, 540f T. tubifex, 537f chaetae types in, 536f collection, 544–545 distribution and diversity, 533–535 ecology and behavior ecological aspects, 544 feeding behavior, 543 macrohabitat distribution, 542 microhabitat selection, 542 physiological constraints, 542–543 predators and parasites, 543–544 location of reproductive organs, 532f male duct types in, 539f phylogenetic relationships, 531–533 preparation for identification, 545–548 rearing, 545 systematics, 529–531 tubificid worm dissection, 547f Oligochaetes sensu stricto (Sludge and earthworms), 510 chaetae, 514, 515f digestive track, 515f megadrile, 510, 512 microdrile, 512–513 faunal similarity of, 512f nephridial system, 516f wave activity, 518f OMEGA. See Ostracod Metadatabase of Environmental and Geographical Attributes Ontological species, 5 Oogenesis-flight syndrome, 955 Oomycetes, 125 Opalinids, 125 Open tracheal system, 1011 Operational species, 5 Opossum shrimp. See Mysidacea Opsiblastic egg, 217–218 Oral pore, 192–193 Orconectes australis (O. australis), 814f Orconectes rusticus (O. rusticus), 826f Orconectes virilis (O. virilis), 817 Oribatida (Beetle mites), 609 diversity and biology, 611 morphology, 609–611 Orohermes crepusculus (O. crepusculus), 1060f Orthoptera, 1069–1070 Osmobiosis, 361 Osmoregulation in aquatic beetles, 1012 Arthropoda, 595 Hemiptera, 956 Order Trichoptera, 984–985 Peracarida, 785 Syncarida, 785 Osmotic balance, 854–855 Osmylidae larvae, 1067 Osmylinae, 1064–1065 Osmylus fulvicephalus larva (O. fulvicephalus larva), 1066 Osphradia, 453–454 Ostracod Metadatabase of Environmental and Geographical Attributes (OMEGA), 761–763

  Subject Index

Ostracoda, 757–758 biogeographic database, 761–763 biology, 763 appendages, 764–765, 766t brood care, 769 circulatory system, 767 D. stevensoni, 766f digestive system, 767 egg, 769 female reproductive organs, 768 gender identification, 767 head region, 765–766 internal morphology, 765f L. sanctipatricii, 766f male reproductive structure, 768–769 under microscope, 763f podocopid ostracodes, 764f reproduction, 768–769 respiratory system, 767 shell and dermal layers, 763f shell morphology, 763–764 thoracic region, 766 visual system, 767–768 distribution, 760–761 diversity, 760–761 ecology and behavior environmental constraints on distribution, 770–773 feeding behavior, 773–774 habitat selection, 769–770 ion hydrochemistry, 771f parasites, 774 predators, 774 environmental constraints, 770 dissolved oxygen, 770–771 hydrology, 772–773 ion composition, 770 ion hydrochemistry, 771f water temperature, 771–772, 773t field collection, 774–775 aquifers, 775 hyporheic zone, 776 lakes, 775 for nonmarine ostracodes, 775f springs, 775 streams, 775 wetlands, 775 fossil record, 760 general systematics, 758–759 hydrology, 772–773 hyporheic zone, 776 lakes, 775 male reproductive structure, 768–769 microscopic crustaceans, 757–758 nonmarine ostracodes, 762f, 767t podocopid lineages, 761f predators, 774 rearing techniques, 777 sample preparation, 776–777 sampling localities in databases, 762f springs, 775 streams, 775 superfamilies with nonmarine taxa, 759t taxonomic database, 761–763 valve morphology, 758f

wetlands, 775 Ostracodes, 757–758 Oviferon, 237–238 Oviposition aquatic beetles, 1013 Diptera, 1052–1053 Odonata, 911–912, 913f Trichoptera, 987–988 Oxygen tension, 863

P

“pa kh-gnong”, 734 Paddles, 238 PAH. See Polyaromatic hydrocarbon Palaemonetes genus, 803–804 Palaemonetes paludosus (P. paludosus), 819f Palaemonidae, 803 Palaeonema phyticus (P. phyticus), 274, 274f Palmen body, 880 Pan traps, 944–945 Pancrustacea. See Tetraconata Papillae, 568 PAR. See Photosynthetically active radiation Parabasalians, 120 Paracles genus, 1069 Paraleptophlebia submarginata (P. submarginata), 880f Paramacrobiotus richtersi (P. richtersi), 348f Parasimulium (Black fly), 1053 Parasitengone mite, 618f Parasitengonina, 615 non-hydrachnidian Calyptostoma, 616f Calyptostomatidae, 615–616 Johnstonianidae, 616f Johnstonianidae, 615–616 Stygothrombidiidae, 615–617 Stygothrombium, 617f Parasites, 774 Parasitiformes, 601, 607. See also Freshwater Sarcoptiformes; Prostigmata. Mesostigmata biology, 608–609 morphology, 607–608 Parasitoids, 1056 Particle sorting, 445 PCBs. See Polychlorinated biphenyls PCR. See Polymerase chain reaction Pectinatella magnifica (P. magnifica), 330f, 339 Pectinatellidae family, 339 Pedal feeding, 449, 455f Pedinellid stramenopiles. See Helioflagellates Peltodytes dispersus (P. dispersus), 1025f Penial chaetae, 536–537 PER. See Photoenzymatic repair Peracarida. See also Syncarida. Amphipoda, 781–782, 782f collecting, 793–794 community ecology, 791–792 culturing, 793–794 Cumacea, 782–783 diet, 790–791 dispersal, 790 environmental physiology calcium, 786

Subject Index

pH, 786 salinity, 786–787 temperature, 785–786 external morphology, 783–784 feeding behavior, 790–791 habitat selection, 788 epigean habitats, 788–789 Hyalella amphipod species, 788f hypogean habitats, 789–790 internal anatomy circulation, 784–785 digestive system, 784 excretion, 785 neural system, 785 osmoregulation, 785 reproductive anatomy, 785 respiration, 784–785 sensory system, 785 invasive species ecological impacts, 792 Isopoda, 782, 782f life history, 787–788 Mysidacea, 782, 782f notable radiations, 792–793 population, 791–792 reproduction, 787 specimen preparation, 793–794 Spelaeogriphacea, 782–783 Stygiomysida, 782–783 Tanaidacea, 782–783 Thermosbaenacea, 782–783 Pereiopods, 807 Peterson grab sampler, 943 Pezidae, 613–614 PGM. See Platinum group metals Pharynx, 193–194, 193f Phenetics, 8 Photoenzymatic repair (PER), 252–253 Photoreception, 676–677 Photosynthetically active radiation (PAR), 69–70 Phreatic zones, 41–42 Phylactolaemata, 328 Phylactolaemate larva, 335f Phylogenetic species, 6 asexual and sexual species, 6 disadvantage, 6 identification, 6 Phylogenetic trees, 8 using allozyme markers and mtDNA haplotypes, 8 computer packages, 8–9 phenetics and cladistics disciples, 8 shared derived and primitive character, 8 Phylogenies ecology and behavior studies, 7–8 phylogenetic trees, 8–9 Physa acuta (P. acuta), 406 Physa genus, 396 morphological responses, 408 reducing algal biomass, 401 reproductive system, 398f Physa gyrina (P. gyrina), 396 Physella vernalis (P. vernalis), 387f Physical-gill respiration, 909–910 Physid gastropods, 406 Physidae, 391

1087

Phytophagous groups, 1056 Phytotelmata, 49 Pirata piraticus (P. piraticus), 603–604 Pirata spp., 603–604 Pisauridae (Nursery-web spiders), 603–604 Piscicolidae, 566 body of leeches, 567 body shape, 568f cocoons of, 574–575 transverse sections, 567f Pit traps, 944–945 PKD. See Proliferative kidney disease Planktonic invertebrates, 60 littoral zone plankton, 61 rotifers, 60–61 zooplankton, 60–61 Plastron, 853 Platinum group metals (PGM), 480 Plationus patulus (P. patulus), 236f Platyhelminthes (Flatworms), 181 body wall, epidermis, and sensory structures basal membrane, 186–187 cell connections, 186–187 cilia, 187–188 epidermal structures, 188 external epithelial, 186–187 musculature, 188 collection, 199–200 culturing, 200 digestive tract, 192 intestine, 193 oral, 192–193 pharynx, 193–194, 193f ecology and behavior ectosymbiosis, 198–199 food web role, 197–198 habitat selection, 197, 197f–198f physiological constraints, 199 excretory systems, 194, 194f geographical distribution, 184–186 neural system central nervous system, 191–192, 191f sensory elements, 192 osmoregulatory systems, 194 parenchyma cell types and musculature, 189–190 functions of, 190 organization and structure, 188–189 phylogenetic relationships, 184, 185f regeneration, 190–191 reproduction development, 196–197 organs and gametes, 194–196, 195f types, 196 species diversity and abundance, 186 specimen preparation, 200 systematic, 181–183 turbellaria groups and position, 182f Platynothrus peltifer (P. peltifer), 611 Platypsyllus castoris (P. castoris), 1035f Plecoptera (Stoneflies), 933 adults collection, 943 aerial nets, 943 beating sheet, 943–944 canopy fogging, 944–945

emergence traps, 944 hand picking, 943 light traps, 944 Malaise traps, 944 pan traps, 944–945 pit traps, 944–945 sticky traps, 944–945 sweep nets, 943 conservation, 940 eggs of, 939f external anatomy, 936f adult, 936 female subgenital plates, 938f immature, 935 mandibles and maxillae, 937f feeding behavior, 939–940 genitalia of, 937f labeling specimens, 946 life history, 937–938 macro-and micro-habitat usage, 939 nymphs collection dip net, 942–943 drift net sampler, 943 hess sampler, 943 kick screen, 943 Peterson grab sampler, 943 ponar sampler, 943 stovepipe sampler, 943 surber sampler, 943 parasites and symbionts, 940 phylogeny and biogeography, 933–935, 934t physiological constraints, 939 preservation techniques, 945–946 rearing, 946–947 vibrational communication, 940–942 Plecoptera Species File (PlecSF), 934 PlecSF. See Plecoptera Species File Plesiomorphies, 6 Pleuroceridae ecology, 409–410 Plumatella casmiana (P. casmiana), 334f Plumatella reticulata (P. reticulata), 332f Plumatella vaihiriae (P. vaihiriae), 339f Plumatellidae family, 339–340 Pneumocystis, 125 Podura aquatica (P. aquatica), 866 floating spermatophore, 866f Polar zones, 24–25. See also Tropical zone. Polyaromatic hydrocarbon (PAH), 478 Polyarthra species, 242 Polychaeta, 520–521, 522f Polychaeta, inland water, 519 biology external anatomy, 522 internal anatomy, 522–524 physiology, 524 reproduction, 524–525 collection and specimen preparation, 526 distribution and diversity, 521–522 ecology and behavior, 525 distribution and habitat selection, 525 feeding behavior, 525–526 invasive tendencies, 526 locomotion, 526 physiological constraints, 525 predators and parasites, 526

1088

internal building plan, 523f phylogenetic relationships, 521 systematics Aphanoneura, 521 Namanereis and Stratiodrilus, 520f Polychaeta, 520–521 Polychlorinated biphenyls (PCBs), 481, 583 Polymerase chain reaction (PCR), 375 Polyp anatomy, 164–166 C. sowerbii, 165f Calpasoma, 165f Cordylophora, 165f, 167f P. hydriforme, 166f Polyphaga families, 1008. See also Adephaga families; Coleoptera (Aquatic beetles); Myxophaga families. C. Champion, 1029 C. Latreille, 1028–1029 C. Latreille, 1029 D. Billberg, 1029–1030 E. Crowson, 1031 E. Curtis, 1030–1031 E. Zaitzev, 1031 G. Latreille, 1031–1032 H. Latreille, 1034 H. Leach, 1032 H. Mac Leay, 1032–1033 H. Mulsant, 1033 Hydrochidae, 1033 L. Erichson, 1035–1036 L. Fleming, 1035 Lutrochidae, 1036 N. Latreille, 1036 P. Lacordaire, 1036–1037 P. Laporte, 1037 S. Erichson, 1038 S. Fleming, 1037–1038 S. Latreille, 1038–1039 Polyploidization, 433 Polypodium hydriforme (P. hydriforme), 161, 170–171, 171f Polyzoans. See Bryozoans Polyzonimine, 664 Pomacea spp., 101 Pomatiopsids, 395 Ponar sampler, 943 Ponto-Caspian basin, 793 Ponto-Caspian hydroid, 175 Population demography, 468 Porifera (Sponges), 133 in active phase, 138f biogeography and diversity, 135, 138 Corvospongilla, 137–138 endemic taxa sensu stricto, 138 metaniidae, 137 sponge species, 135–136 spongillina, 136–137 biology anatomy and physiology, 140–143 Badiaga genus, 141f body bauplan, 138–140 life history, life cycle, and reproduction, 143–144, 146f life history changes, 140 body architecture and aquiferous system, 142f

  Subject Index

collection, 149–151 ecology and behavior behavioral adaptive traits, 147–148 competition and cooperation, 148–149 feeding behavior, 146–147 habitat selection, 145–146 sponges as natural resource, 149 evolution and phylogenetics, 134–135 evolutionary history, 148f freshwater sponges, 134f geographic distribution, 136f identification, 151f as natural resource, 149 preparation, 149–151 rearing, 149–151 spongillaflies, 150f systematics, 133–134 Potamocypris unicaudata, 774, 777 Predaceous diving beetles. See Dytiscidae Leach Predation, 76–77 aquatic beetles, 1016 pressure, 831–832 Predator–prey interactions, 261–264 Predators, 774 Prionocyphon limbata (P. limbata), 1037f Prionocyphon sp., 1038f Proales sordida (P. sordid), 252 Procambarus “Marmorkrebs”, 100 Procambarus clarkii (P. clarkii), 100, 814f Proliferative kidney disease (PKD), 338 Pronotum, 1066–1067 Prostatic glands, 538–539 Prostigmata, 613. See also Freshwater Sarcoptiformes; Parasitiformes. Halacaroidea, 613–614 Parasitengonina, 615–617 Raphignathoidea, 614–615 Protochauliodes clade, 1061 Protochauliodes spencer (P. spencer), 1060f Protonephridia, 217 Protonymph, 642 Protosialis, 1060–1061 Protozoan cell characteristics, 115 food intake, 116 locomotion, 115–116 reproduction, 116 structural elements, 116 structure and function, 115 Pruinescence, 904 Pruinosity. See Pruinescence Psammon, 70 Psephenidae, 1008, 1017 Psephenidae Lacordaire (Water pennies), 1036–1037 Psephenus herricki (P. herricki), 1037f Pseudocellus. See Ocellus Pseudocoelom, 232 Pseudopenis, 538–539 Pseudopodia, 115–116 Pseudosuccinea columella (P. columella), 387f Pteronarcys scotti Ricker, 937f Ptilodactylidae Laporte (Toe-winged beetles), 1037

Pulmonata, 398–399 Pulmonate gastropods. See Basommatophora Pulmonates, 391f, 396

Q

Quorum sensing (QS), 250

R

Radiolarians, 119–120 Raft spiders. See Pisauridae (Nursery-web spiders) Random amplified polymorphic DNA methods (RAPD methods), 740–741 RAPD methods. See Random amplified polymorphic DNA methods Raphignathoidea, 614 Homocaligids, 614 Stigmaeidae, 615 RCC. See River Continuum Concept RCO. See Retrocerebral organ Reaches, 32 Rearing techniques, 777 “Red queen” hypothesis, 403 Reproductive system Branchiobdellida, 555 Coleoptera, 1011 Copepoda, 718 Hirudinida, 570 Peracarida, 785 Rotifera, 237–238 Syncarida, 785 water mites, 635 “RESonate”, 32 Respiratory system aquatic insects, 852–853 Arthropoda, 595 Coleoptera, 1010 Decapoda, 809–810 Hirudinida, 571–572 Ostracoda, 767 Trichoptera, 984 water mites, 634 Resting eggs, 362–363 Resurrection ecology, 689 Retention efficiencies, 442–444 Retrocerebral organ (RCO), 236 Rhabdites, 188 Rhithrogena semicolorata (R. semicolorata), 879f Rhizopod amebae, 118–119 Rhynchocoel, 208 Rhyncholimnochares, 638 Rhynchomonas nasuta (R. nasuta), 127 Ribbon-worms. See Nemertea Riffle beetles. See Elmidae Curtis River Continuum Concept (RCC), 74 River crabs, 829 Rivers, 35 Amazon River basin, 36 Colorado River, 35f ecological characteristics, 35 mountain stream in Glacier National Park, 35f nature, 35–36

Subject Index

Rotifera (Wheel animals), 226f bdelloid rotifers, 229f biogeography, 228–230, 230f Brachionus species, 227f characteristics, 225–228 collection, 265–266 culturing aquaculture, 267–268 laboratory culture, 266–267 diversity and distribution colonial rotifers, 244–246 Filinia species, 243f phenotypic variation, 242–243 and population movements, 243–244 sessile rotifers, 246–249 ecological interactions competition with zooplankton, 261 foraging behavior, 258–259 functional role in ecosystem, 259–261 parasitism on rotifers, 264 predator–prey interactions, 261–264 rotifers as parasites, 265 environmental physiology anhydrobiosis, 240–241 environmental toxicology, 240 locomotion, 238 physiological ecology, 238–240 stress responses generalization, 242 evolutionary relationships, 230 external morphology, 231–232 little-known habitats, 228f male, 237f monogonont rotifers, 229f organ system structure and function, 232 C. vorax, 233f collothecacea, 233f corona, 232 excretory system, 236–237 muscular system, 235–236 neural system, 236 reproductive system, 237–238 trophi and gut, 232–235 preparation for identification, 268–269 reproduction and life history, 249 aging, 252–253 cyclical parthenogenesis and diapause, 249–251 diapausing embryos in sediments, 251 dynamics of field populations, 255–257 embryogenesis, 251 genetic variation, 258 life tables, 253–255 monogonont rotifers, 249f population dynamics, 253–258 population dynamics in chemostats, 257–258 reproductive behavior, 251–252 sessile species, 231f trophi types, 234f Round fungus beetles. See Leiodidae Fleming Rove beetles. See Staphylinidae Latreille

S

s.s. See sensu strict Sap beetle. See Nitidulidae Latreille Saprobic Index, 240

1089

Saprophagy, 1056 Sars’ Method, 700 Satonius stysi (S. stysi), 1020f Scanning electron microscope (SEM), 151, 304, 348–349, 749 Schwiebea, 612f Scirtidae (Marsh beetles), 1012, 1037–1038 Sclerites, 904 Scorpionflies. See Mecoptera Scrapers, 885 Screen barrier net. See Kick screen Scuds. See Amphipoda SDPOM. See Sewage-derived particulate organic matter Segmental organs, 523 SEM. See Scanning electron microscope Semelparity, 575 Semi-aquatic Cheiroseius, 608f Semisulcospira, 393 Sense organs, 453–454 Sensory elements, 192 Sensory pore X-organ, 718 Sensory system Peracarida, 785 Syncarida, 785 sensu strict (s.s), 510 Septal glands, 537 Serradium semiaquaticum (S. semiaquaticum), 666–667, 667f Serratella ignita (S. ignita), 880f Sessile rotifers, 246, 247f ACS, 248–249, 249f F. conifera, 246–247, 248f hydrophyte species, 247 juvenile motile stages, 246 substrate selection behaviors, 247 tube construction, 248, 248f Sewage-derived particulate organic matter (SDPOM), 478–479 Sexual dimorphism, 364, 364f SG. See Stress Granules Shared derived character, 8 Shared primitive character, 8 Shell external structure, 438–440 growth and microstructure, 435–436 internal structure, 436–438 and mantle, 434–435 morphology, 763–764 Shore beetles, 1013–1014, 1032–1033 Shrimps. See Caridea Sialidae, 1061 Sialis lineage, 1060–1061 Sideswimmers. See Amphipoda Simuliid pupae, 1057–1058 Siphons, 440 Sisyridae, 1064–1065 aquatic environment, 1065f Skiff beetles. See Hydroscaphidae LeConte Smooth clam shrimp. See Laevicaudata Snails. See Gastropoda Space competition. See Exploitative competition Species, 4 biological species, 5–6 categories, 5

cohesion species, 6–7 evolutionary species, 6 influential concepts, 5 interpretation of variation, 4 phylogenetic species, 6 Spelaeogriphacea, 782–783 Spercheidae Erichson (Filterfeeding water scavenger beetles), 1038 Spercheus emarginatus (S. emarginatus), 1038f Sperchonopsis ecphyma (S. ecphyma), 627f, 629f Sperm drop. See Spermatophore Sperm-morulae, 455–456 Spermathecal pores, 536–537 Spermatophore, 309–310 transfer direct, 643–645 indirect, 643 Sphaeriids, 466–467 Sphaerius sp., 1019f–1020f Sphaeriusidae Erichson (Minute bog beetles), 1019 Spiders. See Araneae Spinicaudata (Spiny clam shrimp), 689, 693–694 Spiny clam shrimp. See Spinicaudata Spirurid nematodes, 292–294 Sponges. See Porifera Spongillina gemmules, 141f geographic distribution and taxonomic richness, 137 K and r phases, 145f monophyletic status, 135, 135f Spring viremia of carp virus (SVCV), 742 Springs cool, 34 Ostracoda, 775 thermal, 34 Springtails. See Collembola Spumalin, 971 Squeak beetles. See Hygrobiidae Régimbart Staphylinidae Latreille (Rove beetles), 1038–1039 Statoblasts, 332–333 bryozoan, 333f Statocysts, 192, 192f, 785 Stenocolus scutellaris, (S. scutellaris), 1031f Stenosialis lineage, 1060–1061 Stenostomum sp., 189f–190f Stenothyridae, 395 Sticky traps, 944–945 Stigmaeidae, 615 Stigmata, 607–608 Stoneflies. See Plecoptera Stovepipe sampler, 943 Stream order, 29–30 Streams, 775 Streptocephalus moorei (S. moorei), 692f Stress Granules (SG), 242 Stygiomysida, 782–783 Stygofauna, 42–43 Stygothrombidiidae, 615–617 Stygothrombium, 617f Subaquatic spider. See Argyroneta aquatica (A. aquatica)

1090

Subimaginal stage, 884–885 “Subitaneous” eggs, 697 Substrates, 37 limitations, 38 Murray River of Australia, 38f nature and provision, 37–38 Subterranean habitats aquatic habitats, 42–43, 42f blind cave crayfish, 43f hyporheic zones, 41–42 karst system topography, 42f phreatic zones, 41–42 Suckers, 567–568, 569f Sulci, 763–764 Superareoles, 305–306 Superconglutinates, 463, 483f Supernumerary muscles, 553–554, 554f Supplements, 276–278 Surber sampler, 943 Surface-dwelling larvae, 637 Surface-swimming Gyrinidae, 1017 Suspension feeding mechanics, 442 SVCV. See Spring viremia of carp virus Swarming adult dipterans, 1057 Diptera, 1052 Sweep nets, 943 Swimmerets, 784 Swimming zooids, 332 Symplesiorphies. See Shared primitive character Synapomorphies. See Shared derived character Syncarid, 783f Syncarida, 783 Systellognatha, 935

T

Tachyblastic egg, 217–218 Tactile cues, 648 Tactile sensory organs, 907–908 Tadpole shrimp. See Notostraca Taenidia, 904 Tagmatization, 852 abdomen, 852 head, 852 thorax, 852 Tanaidacea, 782–783 Tardigrada (Water bears), 347–348 biogeography, 349–350 collection and extraction, 376 culturing, 377 cyclomorphosis, 369 ecology and behavior dispersal, 375–376 habitats, 369–370 North American distribution and substrates, 371t–374t population dynamics, 370–374 trophic relationships, 375 features, 347–349 with habitat references, 350t internal anatomy buccal–pharyngeal apparatus, 359 digestive system, 359 esophagus, 359–360 excretory system, 360

  Subject Index

hindgut, 359–360 midgut, 359–360 musculature, 360, 360f nervous system, 360 respiration and circulation, 360 latent states, 360–361 cryptobiosis, 361–362 diapause, 362–363 life history, 368–369 light and electron microscopy, 376–377 molting, 368 morphological characters, 351 buccal–pharyngeal apparatus, 354–355 claws, 351–354 cuticle, 355–357 eggs, 357–359, 358f phylogenies and species identification molecular analyses, 377–378 taxonomic keys, 378 reproduction and development, 367 eggs and parental care, 366 gametogenesis and gametes, 364–366 mating and fertilization, 366 postembryonic development, 367 reproductive apparatus, 363–364 reproductive modes, 366 sexual conditions, 363 sexual dimorphism, 364, 364f SEM, 349f systematics and phylogenetic relationships, 350–351, 351f Tarsus, 852 Taxonomic database, 761–763 TCA. See Thiazolidine-4-carboxylic acid TDS. See Total dissolved salts TEM. See Transmission electron microscopy Temperate zones, 25 Termites. See Blattodea Territoriality. See Exploitative competition Testate amebae, 119 Tetraconata, 592 Tetragenous, 457 Tetragnathidae (Long-jawed orb-weavers), 602–603 Tetramermis fissispina (T. fissispina), 292 Thecostraca, 711–712. See also Branchiura; Copepoda. Thermal springs, 34 Thermoreception, 677–678 Thermosbaenacea, 782–783 Thesocytes, 143 Thiara winteri (T. winteri), 388f Thiaridae family, 395 Thiazolidine-4-carboxylic acid (TCA), 252 Thomisidae, 606 Thomisus nepenthiphilus (T. nepenthiphilus), 606 Thoracic region, 766 Thoracopods, 673–675 Toe-winged beetles. See Ptilodactylidae Laporte Top-down control. See Predation Torrent beetles. See Torridincolidae Steffan Torridincolidae Steffan (Torrent beetles), 1019–1020 Total dissolved salts (TDS), 574

Transient species, 470–471 Transmission electron microscopy (TEM), 348–349 Travertine beetles. See Lutrochidae Trechalea tirimbina (T. tirimbina), 606f Trechaleidae, 606–607 Trichodina diaptomi (T. diaptomi), 726 Trichoptera (Caddisflies), 965 biogeographic regions, 970f classification, 966t–967t distribution, 969–970 ecology and behavior case-and retreat-making behavior, 988–989 climate change, 993–994 conservation, 993–994 disturbance effects, 992–993 drift, 989–990 food and feeding, 990 habitats and aquatic adaptations, 991–992 human impacts, 993–994 parasitism, 988 predation, 988 secondary production, 990–991 external morphology adults, 979–982, 980f–982f C. paluguillensis, 974f eggs, 971 larvae, 971–978, 972f–973f, 975f Neoatriplectides sp., 976f primary setal pattern, 974f pupae, 978–979, 979f field collection capturing adult caddisflies, 995 larval caddisflies, 994–995 rearing and association, 995–996 fossil Trichoptera case, 970f larval cases, 968f phylogenetic relationships, 965–969, 969f physiological adaptations aestivation, 985–986 diapause, 985–986 osmoregulation, 984–985 respiration, 984 reproduction and life history, 986 emergence and seasonality, 986–987 flight and dispersal, 987 mating behavior, 987 oviposition, 987–988 specimen preparation, 996 Trichoptera, 98 Tritonymph, 643 Trochus, 232 Troglocambarus maclanei (T. maclanei), 819f Tropical clam shrimp. See Cyclestherida Tropical zone, 25–26 True bugs. See Hemiptera True crabs. See Brachyura True flies. See Diptera True leeches. See Hirudinea True Water beetles, 1013–1014 “True” parenchymal cells, 190 Tubercles, 568 Tubificidae sexual organs, 516f Turbellaria, 183f Two-winged flies. See Diptera (True flies)

Subject Index

Tylomelania sulawesi (T. sulawesi), 391f Typhlocarididae, 804

U

Ultraviolet radiation (UVR), 252–253 Umbilicus, 385 Uncus, 233 Unionicola intermedia (U. intermedia), 644 Unionid life-cycles, 461 acquired immunity, 465–466 broadcasting, 462 conglutinates, 462–463 displayers, 463–465 host specificity, 465–466 host trapping, 465 superconglutinates, 463 Urnatella gracilis (U. gracilis), 341, 341f

V

Valvatids, 394–395 Variegated mud-loving beetles. See Heteroceridae Mac Leay Varuna litterata (V. litterata), 806–807 Veliger, 454

1091

Vibrational communication, 940–942, 940t, 941f Viviparids, 393–394 Viviparity, 144 Viviparus georgianus (V. georgianus), 400

W

Wandering spiders. See Ctenidae Wandesia, 638 Water bears. See Tardigrada Water boatmen eggs. See Ahuauhtli Water core, 265 Water flea. See Branchiopoda Water hyacinth, 100–101 Water mite parasitism, 646–647 Water mites. See Hydrachnidiae Water pennies. See Psephenidae Lacordaire Water scavenger beetles. See Hydrophilidae Latreille Water-fleas. See Cladocera Waterfowl, 337 Wetlands, 46, 47f alluvial swamps, 46–47, 47f invertebrates of seasonal, 46 Ostracoda, 775

Wheel animals. See Rotifera Whirligig beetles. See Gyrinidae Latreille White spot syndrome virus (WSSV), 834 Winter stoneflies, 937 Wolbachia bacteria, 774 Wolf spiders. See Lycosidae Wound closure, 190–191 WSSV. See White spot syndrome virus

X

x-organ, 217–218 Xerosomes, 241, 241f

Z

Zebra mussels, 484 Zetomimus francisi (Z. francisi), 610f Zooids, 328 bryozoans, 329f budding, 332 phylactolaemate, 329f Zooplankton, 70 Zoosporic fungi, 125 Zwickia species, 613 Zygoptera, 894–896, 895f, 901f, 902

Taxonomy Index Note: Page numbers followed by “f” and “t” indicate figures and tables respectively

A Abedus, 955–957, 959, 961 A. herberti, 951, 952f, 955–956 Abtrichia, 979–981, 981f Abyssidrilus, 543 Acalyptonotidae, 601t–602t, 620t–625t Acanthacorydalis, 1061–1063 Acanthametropodidae, 874–876, 875t–876t, 878–879 Acanthamoeba, 118–119 Acanthobdella A. livanowi, 511, 566 A. peledina, 511, 552, 566, 568f, 570, 572, 576, 583 Acanthobdellida, 510–511, 529, 533, 565, 567 Acanthobdellidae, 12, 566–567 Acanthocephala, 212, 230 Acanthocyclops, 718, 721, 723, 726–728 Acanthodrilidae, 512–513 Acanthogammaridae, 792–793 Acanthogammarus, 792–793 Acantholeberidae, 688t Acanthosentis dattai, 725–726 Acantobdellidae, 566 Acari, 375f, 600, 601t–602t, 834 Acaridae, 601t–602t, 612f, 613 Acariformes, 600–601, 601t–602t, 609–617, 609f Acarina, 600 Acaroidea, 601t–602t Acartia, 731–732 Acella, 396 A. haldemani, 396 Acentropinae, 1068–1069 Achaetobdellae, 512 Acherontacaridae, 601t–602t, 620t–625t Acherontacarus, 637 Acheta domesticus, 307f Achromadora, 292 Aciculata, 520–521 Acilius, 20f A. canaliculatus, 1022–1024, 1023f Acineta, 117f Acipenser fulvescens, 319–320 Acipenseridae, 170–171 Aclitellata, 510–511 Acochlidiida, 389, 401 Acoela, 186–189, 188f, 191–196, 198 Acoelomorpha, 181, 184, 186–188, 190, 192–194 Acrasis, 120 Acrididae, 1069–1070 Acroloxidae, 390t, 391, 394t

Acroloxoidea, 390t Acroloxus, 396 Acroneuria A. abnormis, 20f A. internata, 936, 937f A. lycorias, 936, 938f Acrothoracica, 711–712 Actinomonas, 121 Actinomyxidia, 518 Actinonaias, 424, 489f A. ligamentina, 466 A. pectorosa, 480 Actinophrys, 118f, 120 Actinosphaerium, 120 Acutuncus, 350t, 357–359, 369 Acyclus inquietus, 248–249, 249f Adamietta, 395 Adephaga, 1004, 1005t–1007t, 1008–1012, 1014–1016, 1020–1028 Adineta, 264 A. ricciae, 241 A. vaga, 241 Adorybiotus, 350t Aedes, 654, 1043–1044, 1045t–1047t, 1053 A. aegypti, 732–733 A. albopictus, 732 A. cinereus, 640 A. communis, 642 A. excrucians, 640 A. punctor, 642 Aegla, 802–803, 807–808, 823, 829–832 A. laevis, 823 A. rostrata, 823 Aeglidae, 798, 799t, 802–803, 807, 824, 825t Aegloidea, 807 Aeolosoma, 521 Aeolosoma, 172–173, 526 A. hemprichi, 521f, 523–525 A. maritimum, 525 A. variegatum, 525 Aeolosomatidae, 12, 519–526, 529, 536 Aepophilidae, 951–953, 954t Aeromonas A. hydrophila, 570 A. salmonicida, 471 Aeshna, 913, 922 A. cyanea, 906–907, 910–911, 912f, 922 A. eremita, 677 A. grandis, 911–912, 913f, 922 A. isoceles, 923, 923f A. juncea, 902–903, 903f A. viridis, 918–919

Aeshnidae, 896–900, 897t–898t, 900f, 902–903, 908, 909f, 910–912, 917–920, 922, 926, 928f, 929 Aeshnoidea, 896–899, 897t–898t Aetideidae, 726 Africasiinae, 620t–625t Afrithelphusa monodosa, 807–808, 810f, 814–816, 816f, 822, 828f Afrocyclops, 729 Agapetus A. bifidus, 985 A. fuscipes, 988 A. illini, 985 A. occidentis, 985 Agnetina flavescens, 936, 938f Agriocnemis, 902 Agriodrilus, 543 Agriotypus armatus, 988 Aktedrilus, 543 Alainites albinatii, 877, 878f Alasmidonta, 424, 460, 489f A. heterodon, 477 A. mccordi, 429, 429t A. robusta, 429, 429t A. undulate, 468 A. wrightiana, 429, 429t Albertathyas montanus, 631f Albertia, 543 Albiinae, 620t–625t Alexandrovia ringulata, 542–543 Algivores, 400t Algophagidae, 601t–602t, 612–613 Algophagus pennsylvanicus, 613 Algophilus, 1025–1026 Alisotrichia, 972–974, 973f Allocapnia, 939 A. granulata, 936, 937f Allocrangonyx pellucidus, 789f Allocyclops, 730 Allodero, 542–543 Allodiaptomus, 734 Allomyia scotti, 972–974, 973f Alluroididae, 517, 533 Alma, 512 Almidae, 510, 512 Alnus, 62 Alona, 694 Alpheidae, 798, 799t, 804 Alpheus, 804 Alticinae, 1028–1029 Amara alpina, 310f Amazonatolica hamadae, 979–982, 983f

1093

1094

Amblema, 424, 461–462, 489f A. plicata, 455, 459f, 482–483 Ambleminae, 424, 427, 457–458, 460, 462, 489f Amblemini, 424, 489f Amblypygi, 600 Ambrysus, 953, 955–956, 959 A. amargosus, 960 Ambystoma dumerilii, 741 Ameiridae, 711, 727–728 Ameletidae, 874–876, 875t–876t Ameletopsidae, 874–877, 875t–876t, 879 Ameronothridae, 601t–602t, 611 Ameronothroidea, 601t–602t Ameronothrus, 610f Ametropodidae, 874–876, 875t–876t Amniclineus, 207t A. zhujiangensis, 207t Amnicola, 395, 400, 401t, 403–404, 406 Amnicolidae, 390t Amoeba, 118–119, 118f A. proteus, 117f Amoebophrya ceratii, 122 Amoenacaridae, 601t–602t, 620t–625t Amphibalaninae, 712 Amphibalanus, 712, 745–746 A. amphitrite, 712, 746–748, 750 A. improvisus, 49, 712, 742, 744–748, 745f, 751 Amphicrossus japonicus, 1036 Amphidinium, 122 Amphiesmenoptera, 965 Amphilinidea, 181 Amphinaias pustulosa, 455 Amphinemura palmeni, 936, 938f Amphinomidae, 520–521 Amphipoda, 13, 41, 66, 68, 75, 578, 672–675, 672t, 678–680, 682, 781–794, 788f–789f Amphipterygidae, 895–899, 897t–898t, 909 Amphitrite, 522 Amphizoa, 1020–1021, 1021f Amphizoidae, 1004–1014, 1005t–1007t, 1020–1021, 1021f Ampullariidae, 100, 389, 390t, 391, 394, 394t, 400, 412f Ampullarioidea, 390t Anabaena, 260 Anabolia A. bimaculata, 965–969, 968f, 972–974, 973f A. furcata, 985 A. nervosa, 985 Anabrus simplex, 316 Anacharis, 62 Anacroneuria, 940 Anactinotrichida, 607 Anadenobolus, 662f Anaspidacea, 13, 674f, 783–785 Anax, 899–900, 900f, 902, 908, 909f, 920, 923–925, 924t A. ephippiger, 918–919 A. imperator, 917–918, 919f, 923, 923f, 926–927 A. junius, 918, 922 Anchitrichia, 992 Anchycteis, 1037 Anchytarsus, 1037

Taxonomy Index

Ancylidae, 387, 390t, 391, 394t, 396, 400t Ancylometes, 603–604 A. hewitsoni, 603f Ancylus, 396 A. fluviatilis, 400, 405 A. planus, 644 Androprosopa, 1043–1044, 1045t–1047t Anepeorus, 885 Angiostrongylus cantonensis, 101 Anisitsiellidae, 601t–602t, 620t–625t, 637–638, 646 Anisitsiellinae, 620t–625t Anisops, 853 Anisoptera, 894–902, 895f, 897t–898t, 899f–901f, 905, 908–912, 912f, 914, 918, 923–924, 924t, 926, 929 Anisozygoptera, 894–899, 895f, 897t–898t, 905, 908f Annelida, 509–513, 519–521, 571–572 Annerossella, 614, 614f A.knorri, 614 Annulipalpia, 965–972, 966t–967t, 974–979, 981–982, 984, 988–989, 991–992, 995 Anodonta, 86, 424, 443–445, 445f, 447, 450, 489f A. anatina, 474, 480 A. cygnea, 446–447, 452f, 474–476, 747 Anodontites trapesialis, 474 Anodontoides, 424, 462, 489f A. ferussacianus, 475 A. ferussacinaus, 476 Anomalopsychidae, 965, 966t–967t, 971–972, 974f, 976–978 Anomopoda, 687 Anomura, 798, 798t–799t, 807, 807f, 824, 825t Anopheles, 106, 603–604, 654, 732–733, 1043– 1044, 1045t–1047t A.crucians, 640 A. gambiae, 105f Anopla, 205–206, 206t–207t Anoplius, 1068 A. depressipes, 1068 A. eous, 1068 Anostostomatidae, 1070 Anostraca, 673–675, 674f, 687–690, 688t, 694–697, 700–701 Anostracta, 855, 857f Antarctoperlaria, 934–935, 934t, 940–941 Antechiniscus, 364f Antimelania, 395 Antipodoeciidae, 965–969, 966t–967t Antocha, 1043–1044, 1045t–1047t Anuraeopsis fissa, 237f, 238, 256f, 262 Anurida, 867 A. maritime, 867 Anystides, 601t–602t Anystoidea, 618, 626 Apatania, 1066 Apataniidae, 965, 966t–967t, 972–974, 973f, 985, 990, 992 Apatronemertes, 207t A. albimaculosa, 207t Apeltosperchontinae, 620t–625t Aphanomyces astaci, 101, 834 Aphanoneura, 510–511, 519–521, 529

Aphanoneura, 520 Aphelenchoides, 280f, 288 Aphelenchoididae, 280f Aphelocheiridae, 951–953, 954t, 956 Aphelocheirus, 956 Apheviderulicidae, 601t–602t, 620t–625t Aplexa, 397 A. hypnorum, 400 Aplodinotus grunniens, 469–470 Apochauliodes, 1061 Apochela, 350t, 352, 354–357, 359, 366 Apocorophium, 673f Apocyclops, 721 Apodera vas, 127 Apodibius, 350t, 353–354 A. confusus, 370 Aporodesminus wallacei, 665–666 Apteraliplus, 1025–1026 Apteropanorpidae, 1069 Apusomonas, 122 Aquarius, 951–953, 952f, 959 Arachnida, 12, 60, 592, 600–601, 601t–602t, 988 Araeolaimida, 273 Araneae, 600–607, 601t–602t Araneomorphae, 601, 601t–602t Aranimermis, 294 Arcella, 117f, 118–119, 231–232 Archaeochlus, 1043–1044, 1045t–1047t Archichauliodes, 1061–1062 A. diversus, 1062–1063 Archigetes, 543–544 Archilestes grandis, 920 Archimonotresis limnophila, 184–185 Architaenioglossa, 389, 390t, 391 Archostemmata, 1004, 1008 Arcidens, 424, 489f Arctiinae, 1068–1069 Arctodiaptomus, 727–728 A. rectispinosus, 721 A. salinus, 721 Arctoperlaria, 934–936, 934t Arctopsyche, 990 Arctopsychinae, 990 Arenicola, 522 A. marina, 524 Arenohydracaridae, 601t–602t, 620t–625t Arenotus, 216, 218 Argiolestidae, 896–899, 897t–898t Argiope bruennichi, 923, 923f Argulidae, 711, 735f, 737–738, 740, 740f, 742, 749–750 Argulus, 593f, 680, 680f, 711, 734–742, 735f, 750 A. ambystoma, 734–735 A. catostomi, 741–742 A. chinensis, 735–736 A. coregoni, 739–742 A. foliaceus, 711, 738–742 A. japonicus, 734, 737–741 A. multicolor, 737 A. personatus, 734 A. siamensis, 740–742 Argyrodiaptomus cavernicolax, 726 Argyroneta, 604 A.aquatica, 602–603

Taxonomy Index

Arhynchobdellida, 511–512, 566 Arius assimilis, 741–742 Arkansia, 424, 489f Arrenuridae, 601t–602t, 620t–625t, 631f–632f, 637–638, 639f, 646 Arrenuroidea, 601t–602t, 618–619, 620t–625t, 632–633, 632f, 635, 637–639, 641–644, 647–648, 653–654 Arrenurus, 631f, 634f, 635–636, 638–648, 639f, 653–654, 925 A. angustilimbatus, 642 A. birgei, 634f A. bleptopetiolatus, 634f A. cuspidator, 641–642 A. danbyensis, 642, 647 A. delawarensis, 642 A. fissicornis, 632f A. kenki, 640, 642 A. kitchingi, 646 A. madaraszi, 641 A, manubriator, 644–645, 649 A. megalurus, 634f A. papillator, 641 A. planus, 641 A. pseudocylindratus, 632f A. pseudosuperior, 636–637, 647 A. rotundus, 634f A. ventropetiolatus, 642 A. wardi, 17f Arrhopalites, 867 Artemia, 175–176, 596, 694–696, 699, 701, 721, 748, 750 A. franciscana, 676 A. monica, 48 A. salina L., 679 Artemiidae, 688t Artemiopsis, 696, 698 Arthropoda, 351, 591–592, 601t–602t, 661– 662, 850 Arthrotardigrada, 350t, 351, 369 Asajirella, 331 Asajirella gelatinosa, 84–85 Ascomorpha, 234f A. eucaudis, 231–232 Ascothoracida, 711–712 Asellus, 559, 792–793 A. aquaticus, 604, 677, 785–786, 788–791, 793 Asellus species, 294 Aspidiophorus, 212 A. oculifer, 216–217 Aspidisca, 122–123 Aspidogastrea, 181 Aspidytes A. niobe, 1021 A. wrasei, 1021 Aspidytidae, 1004, 1005t–1007t, 1008–1010, 1014, 1021, 1021f Asplanchna, 173, 229f, 232, 234–238, 234f, 242, 246, 250–251, 253, 258–264, 262f A. brightwellii, 238, 242, 251, 262 A. girodi, 237f, 250f, 256f, 262–263 A. intermedia, 242 A. priodonta, 244, 257 A. sieboldii, 235f, 236–237, 242, 262

1095

Asplanchnidae, 234–235 Asplanchnopus, 232, 234–235, 259, 262–263 Assiminea pecos, 410t Assimineidae, 390, 390t Assulina, 118f Astacidae, 798–801, 799t, 801f, 804–805, 824, 825t, 826 Astacidea, 672t, 798–801, 798t, 801f, 824, 825t Astacocrotonidae, 601t–602t, 620t–625t Astacoidea, 798–802, 799t, 801f, 805, 824, 825t Astacopsidrilus, 542–543 A. naceri, 534 Astacopsiphagus parasiticus, 613–614 Astacopsis, 805, 824, 825t A. gouldi, 824, 825t Astacus, 614, 801, 805, 824, 825t A. astacus, 94, 553 A. leptodactylus, 101, 553, 824 A. pachypus, 824 Astasia, 121 Astatumen, 350t, 354–355, 356f Astigmata, 834 Astigmatina, 601t–602t, 609–613 Astrohydra, 159–161, 164–166, 164t, 168–169, 172–173 A. japonica, 163t, 168 Astrosclera, 147 Asynarchus nigriculus, 988 Atanatolica, 991–992 Athearnia, 394t, 409 Athericidae, 1043–1044, 1045t–1047t, 1049– 1050, 1049f, 1052–1054 Atherix, 1043–1044, 1045t–1047t, 1049, 1049f Athernia A. anthoni, 410t Athienemanniidae, 601t–602t, 620t–625t Atlantoastacus, 805 Atopsyche, 971–972, 971f, 974–976, 975f Atractides, 631f, 640 Atriplectididae, 965, 966t–967t, 974–976, 990 Atrochidae, 232, 233f, 246, 258–259 Attheyella, 711, 730 Aturidae, 620t–625t, 632f, 639f Aturinae, 620t–625t Aturus, 632f, 639f Atya, 803 A. lanipes, 681 A. scabra, 824 Atyaephyra, 803, 824 Atyella, 803 Atyidae, 59, 798, 799t, 800, 803, 824, 825t Aulodrilus, 533–534, 540–542, 545 Aulophorus, 534–535, 542 Austeruseus, 350t Australiothyadinae, 620t–625t Australocypris insularis, 773–774 Australysmus, 1064 Austremerellidae, 874–876, 875t–876t Austridotea lacustris, 790 Austroneurorthus, 1064, 1066–1067 Austroperlidae, 934–935, 934t Austropetaliidae, 896–899, 897t–898t Austropotamobius, 801, 805, 824–826, 825t A. italicus, 678 A. pallipes, 557–558

Austrosialis, 1060–1061 Austrothelphusa transversa, 829 Austrotinodes, 971–972 A. texensis, 972–974, 973f Axonopsinae, 620t–625t, 632 Axonopsis, 632, 639, 645–646 Axymyia, 1043–1044, 1045t–1047t, 1048f, 1049 Axymyiidae, 1043–1044, 1045t–1047t, 1048f, 1049 Azolla, 62, 1068–1069

B

Badiaga, 141f Baetidae, 874–878, 875t–876t, 878f–879f, 881–882, 885–886 Baetis, 76, 881, 883–887, 884f B. fuscatus, 878, 879f B. rhodani, 881 Baetiscidae, 874–877, 875t–876t, 879–882 Bagoinae, 1029 Baicalarctiidae, 191–192, 195 Baicalasellus, 792–793 Baicalellia evelinae, 196 Baikalodrilus, 534, 542 Baikaloperla elongata, 859 Baikalospongia bacillifera, 138f Balanidae, 712 Balanocochlis glandiformis, 389f Balanomorpha, 711–712 Balantidium coli, 123 Balanus, 742–745 B. amphitrite, 744 B. amphitrite saltonensis, 49 B. improvisus, 746 Balatro, 543 Banksiola dossuaria, 972–974, 973f Barbarochthonidae, 965–969, 966t–967t Barbicambarus, 825 B. cornutus, 813–814, 814f Barbronia weberi, 574–576 Barnacles, 712 Basommatophora, 390t, 391–392 Bathynella, 790 B. baicalensis, 783 Bathynellacea, 13, 672t, 783–784 Batracobdella, 568f–569f Bdellerogatus plumbeus, 580–581 Bdellodrilinae, 511, 551–552 Bdellodrilus illuminatus, 554, 558 Bdelloidea, 227, 234–235, 237, 249, 614 Bdellonemertea, 205–206 Beauchampia, 248 B. crucigera, 248f Behningiidae, 874–876, 875t–876t, 878–879 Bellamya, 83–84, 393–394 B. angularis, 389, 389f, 393–394 B. japonica, 411 B. purificata, 389, 389f Belostomatidae, 639f, 951–959, 952f, 954t, 957f, 959f Belostomatinae, 958 Bennelongia, 760, 764, 774 Beornidae, 350t Beorn leggi, 350t, 351 Beraea gorteba, 974–976, 975f

1096

Beraeidae, 965, 966t–967t, 974–976, 975f, 986–987 Bergtrollus, 350t Beringia, 969 Berosus, 1034 Bertolanius, 369 Bertolanius, 350t, 352f, 354, 357–359, 362 B. nebulosus, 362–363, 371t–374t B. smreczyneskii, 371t–374t B. volubilis, 361 B. weglarskae, 366, 371t–374t Bezzia, 647, 1044, 1048f, 1049 Bibionidae, 1052 Bicosoeca, 121 Bigelowiella natans, 783 Bindius, 350t Biocellata, 606 Biomphalaria, 58, 400 B. alexandrina, 773–774 B. glabrata, 400, 773–774 Biomyxa, 119 Biserovus, 350t, 356f Bithynia, 400 B. tentaculata, 400, 404 Bithyniidae, 394, 394t, 399–401, 400t Bittacomorpha, 1043–1044, 1045t–1047t, 1048f, 1049 Bivalvia, 12, 384–388, 925 Biwadrilidae, 510 Blastodinium, 122, 725–726 Blatta lateralis, 695–696 Blattidae, 308 Blattisociidae, 607–608, 608f Blattodea, 1059, 1067–1068 Blepharicera, 1043–1044, 1045t–1047t, 1048f, 1049–1050, 1051f Blephariceridae, 1043–1044, 1045t–1047t, 1048f, 1049–1054, 1051f, 1055f, 1056, 1058 Bodo, 121 Boeckella, 710–711, 726, 728, 731–732 B. triarticulata, 731 Bogatiidae, 601t–602t, 620t–625t Bombina bombina, 868 Borealibius, 350t, 354, 369 B. zetlandicus, 17f, 366, 367f, 371t–374t Bosmina, 45, 74–75, 173, 688t, 698, 701, 724 B. freyi, 10 B. longirostris, 260 Bostrichiformia, 1008 Bothriocephalus, 725–726 Bothrioneurum, 533–534, 540–542, 545 B. grandisetosum, 543 Bothrioplana semperi, 183f, 186, 199 Bothrioplanidae, 183f, 186, 191–192, 196–197 Bouchardina, 825 B. robison, 817–819, 818f Bougainvillidae, 163t Boyeria, 920 Brachionus, 227f, 230, 234–238, 250–253, 255, 258–259, 261f, 263–266, 265f, 721 B. angularis, 243 B. calyciflorus, 235, 235f, 237f, 238, 240, 243, 245, 250f, 251, 253–254, 254f–257f, 257, 259, 261–264, 261f, 263f, 266 B. caudatus, 242

Taxonomy Index

B. ibericus, 258 B. manjavacas, 253, 253f B. plicatilis, 48–49, 230–231, 236, 238–241, 250–252, 254–255, 258, 264, 266 B. rubens, 260, 265–266 B. urceolaris, 262–263 Brachiopoda, 328 Brachious B. plicatilis, 237f B. rotundiformis, 258 Brachycentridae, 965, 966t–967t, 974–976, 989–990 Brachycentrus, 990, 992 Brachycera, 1043–1044, 1045t–1047t, 1050, 1056 Brachypoda, 644 Brachypylina, 601t–602t Brachythemis, 902–903, 908, 909f B. lacustris, 902–903, 919 Brachytron pratense, 922 Brachyura, 672t, 798, 798t–799t, 805–806, 805f–806f, 824, 825t Bradleystrandesia, 768, 773t Bradytictia, 458–459 Branchinecta, 695–696, 698 B. campestris, 695–696 B. gigas, 690, 698 B. mackini, 695–696 B. paludosa, 672–673 B. raptor, 690, 691f, 698 Branchinectidae, 688t Branchinella B. hardingi, 9–10 B. nana, 690 B. occidentalis, 690 B. ondonaguae, 692f Branchinellites, 9–10 Branchiobdella, 552–553, 555, 557 B. astaci, 556–558 B. balcanica, 557–558 B. hexodonta, 557–559 B. italica, 557–558 B. kozarovi, 555–557 B. parasita, 556–558 B. pentodonta, 557–558 Branchiobdellae, 511 Branchiobdellida, 509–511, 529, 533, 551–552, 559, 565 Branchiobdellidae, 12, 511, 551–552 Branchiobdellids, 72 Branchiobdellinae, 551–552 Branchiodrilus, 535 Branchiopoda, 12, 66–67, 70, 592, 672–675, 672t, 678–679, 678f, 687–690, 688t, 697, 699–700, 850, 857 Branchipodidae, 688t Branchiura, 672t, 680, 710–711, 734–742, 740f B. sowerbyi, 533–535, 542–543 Brevicaudaturus, 635–636, 641 Brevitentoria, 965, 966t–967t, 969 Bromeliaceae, 218, 1054 Brotia, 390f, 391–392, 392f, 395 Brychius, 1025 B. hungerfordi, 1016–1017 Bryocyclops, 730

Bryospilus, 694 Bryozoa, 11, 327–329, 330f, 331–332, 337–343, 340f Buccinidae, 401 Buddenbrockia plumatellae, 338 Buenoa macrotibialis, 725 Buenos, 853 Bulimnea, 396 B. megasoma, 396 Bulinus, 400 B. globosus, 398–399 Byrrocryptus, 1037 Byrroidea, 1008 Byrsopteryx, 992 B. mirifica, 971–972, 971f Byssanodonta, 432–433 Bythinella, 395 Bythinia B. tentaculata, 402–403 Bythotrephes, 103, 694, 698–699 B. longimanus, 87–88, 103, 245, 254

C

Caecidotea, 782f, 784f, 792–793 C. r. racovitzai, 789 Caenidae, 874–885, 875t–876t, 880f Caenis, 880–883, 888 C. horaria, 879–880, 880f Caenogastropoda, 385–395, 390t, 394t, 397–398, 400, 401t, 402, 403t, 404 Caenomorpha, 125 Caenorhabditis briggsae, 288 Cafeteria, 121 Calamoceratidae, 965–969, 966t–967t, 968f, 974–976, 975f, 979–981, 981f–982f, 986–987 Calamoecia, 726 Calanoida, 710–713, 713f, 717–734 Calanoids, 70 Calasellus, 19f Calathaemon holthuisi, 804 Calcarea, 133 Calcarobiotus, 350t Caligohomus, 615 Calineuria californica, 942 Callibaetis, 881 Callinectes C. bocourti, 820 C. sapidus, 806–807, 820 Callinectes sapidus, 557 Calliobdella vivida, 579 Callipodida, 664 Callistocypridinae, 759t Caloca saneva, 992 Calocidae, 965, 966t–967t Calohypsibiidae, 350t, 353–354, 366 Calohypsibius, 350t, 352f, 353–354, 357 Calopterygidae, 897–899, 897t–898t, 908–911, 909f, 912f, 926 Calopterygoidea, 895–899, 897t–898t Calopteryx, 904, 910–911, 921 C. haemorrhoidalis, 909–910 C. maculata, 911 C. splendens, 899–900, 899f, 904–907, 905f–907f

Taxonomy Index

Calosopsyche, 971–972, 971f Calpasoma, 159–161, 164–165, 164t, 165f, 168–169, 172–173, 175–176 C. dactylopterum, 163t, 168 Calyptostase, 615 Calyptostoma, 616f Calyptostomatidae, 601t–602t, 615–617, 616f, 632 Calyptostomatoidea, 601t–602t, 618 Camallanida, 725–726 Camallanus, 294, 725–726 C. cotti, 292–294 Cambalidae, 308 Cambarellus, 805, 822, 825 C. puer, 838 Cambaridae, 798–801, 799t, 801f, 804–805, 824, 825t, 826 Cambarincola, 552–553 C. fallax, 555–556 C. gracilis, 553 C. ingens, 555–556, 558 C. mesochoreus, 553, 555–558 C. okadai, 553 C. pamelae, 555–556 Cambarincolinae, 511, 551–552 Cambaroides, 801, 804–805 Cambarus, 805, 812f, 813, 824–828 C. bartonii, 824, 825t C. brachydactylus, 828 C. diogenes, 838 C. distans, 813–814, 813f C. dubius, 813–814, 814f C. friaufi, 828 C. gentryi, 813–814, 813f C. jonesi, 828 C. laevis, 828 C. limosus, 824, 825t C. ludovicianus, 817–819, 818f C. robustus, 824, 825t, 830 C. strigosus, 807, 811f Caminella peraphora, 608–609 Camisiidae, 611 Campbellonemertes, 207t C. johnsi, 207t Campeloma, 393–394, 398, 400, 403, 406 C. decampi, 410t C. decisum, 399, 404, 484 C. demersum, 401–402 C. rufrum, 399 Campostoma ornatum, 957, 957f Canace, 1043–1044, 1045t–1047t Canacidae, 1043–1044, 1045t–1047t, 1056 Canalipalpata, 520–521 Candona, 758f, 773t C. acutula, 771–772 C. candida, 771–772 C. caudata, 770–771 C. elliptica, 764f C. subtriangulata, 770–771 C. suburbana, 765f Candonidae, 758–761, 758f, 759t, 764f, 768 Candoninae, 767–768 Candonocypris novaezelandiae, 773–774 Canthocamptidae, 711, 720, 727 Canthocamptus staphylinus, 720

1097

Caobangia, 520 Capilloventridae, 531–534, 536 Capnia lacustra, 859, 939 Capniidae, 859, 934–936, 934t, 937f, 940t, 942–944 Carabidae, 308, 310f, 1004, 1005t–1007t, 1009, 1013–1014, 1016, 1020–1022 Carabus, 1022 C. clathratus, 1022 C. menetriesi, 1022 C. variolosus, 1022 Carapacea, 691–692, 874–876 Carcinus maenas, 748 Caridea, 672t, 798, 798t–799t, 800, 800f, 824, 825t Caridella, 803 Caridina, 803, 824 C. steineri, 824 Caridinides, 803 Caridinophila, 556–557 Caridinophilinae, 551–552 Caridinophilus unidens, 551–552, 557 Carphania, 350t, 351, 352f, 355, 356f, 359, 363–364 C. fluviatilis, 355, 369 Carphaniidae, 350t, 351, 356–357 Caryophyllaeidae, 543–544 Caspiastacus, 805 Castor C. canadensis, 1035 C. fiber, 1035 Castoroides ohioensis, 771–772 Castrada, 184–185 Castrella C. truncate, 199 Castricollis, 688 Catenula, 200 C. lemnae, 183f, 200 Catenulida, 181, 183f, 184, 186–188, 188f, 190–198, 200 Catenulidae, 183f, 192 Catocala communis, 789 Catocala vidua, 772–774, 777 Catostomidae, 291 Caudofoveata, 384 Cavostelium, 120 Cenocorixa bifida, 642 Centrocypris, 758f Centropagidae, 710–711, 728 Centroptiloides, 885 Centroptilum triangulifer, 884 Cepangopaludina C. chinensis, 100 C. japonica, 100 Cephaliophora, 265f Cephalobidae, 278 Cephalocarida, 672, 710 Cephalodella, 229f, 231, 234f, 238–240 C. forficula, 231–232 C. hoodii, 239 Cephalopods, 384 Cephalothrix, 205 Ceralcea, 149, 990–992, 995 C. transvera, 986–987 Cerambycidae, 1028–1029

Ceratium, 116, 122 Ceratomyxa shasta, 526 Ceratophylum demersum, 400, 1019 Ceratopogonidae, 58, 106, 274, 274f, 375f, 608, 637–638, 925, 1043–1044, 1045t–1047t, 1048f, 1049, 1053–1054, 1056–1058 Ceratozetoidea, 601t–602t Cercariae, 773–774 Cercomonas, 122 Cercopagidae, 688t Cercopagis, 103, 172–173, 694, 698–699 C. pengoi, 87–88, 103, 691f Ceriagrion, 894–895 Ceriodaphnia, 654, 694 C. quadrangula, 693f Cerithioidea, 390–392, 390t Cernotina, 990 Cestoda, 181, 518, 925 Ceutorhynchinae, 1009–1010, 1029 Chaetae, 514, 515f, 535 Chaetoceros, 750 Chaetogammarus ischnus, 782f Chaetogaster, 518, 534, 537, 543 C. limnaei, 542 C. limnaei limnaei, 544 C. limnaei waghini, 543 Chaetonotida, 211–218 Chaetonotidae, 211–214, 213f, 218, 219t Chaetonotus, 212–214, 216, 218 C. hystrix, 218 C. maximus, 218 C. oculifer, 216–217 C. spinifer, 16f C. spinulosus, 214 Chaetopterus variopedatus, 58 Chaetopteryx villosa, 985 Chalcolestes viridis, 913–914, 919 Chaoboridae, 637–638, 1043–1044, 1045t–1047t, 1048f, 1049, 1054, 1057–1058 Chaoborus, 45, 238, 699, 725, 859, 859f, 1043–1044, 1045t–1047t Chaoborus astictopus, 98–99 Chappuisides, 632 Chappuisididae, 601t–602t, 620t–625t, 633f Chappuisidinae, 620t–625t Chara, 773–774 Chara zeylanica, 295 Chathamiidae, 965, 966t–967t, 985, 992 Chauliodes, 1061–1063 Chauliodinae, 1060–1061, 1063 Cheirogenesia, 883–884 Cheiroseiulus, 608 C. reniformis, 608 Cheiroseius, 607–608, 608f Chelicerae, 606–608, 612 Chelicerata, 12, 592, 600, 850 Chelicorophium curvispinum, 86–87 Chelifera, 1043–1044, 1045t–1047t Chelonibia manati, 712 Cherax, 26, 94, 805, 825–826 C. destructor, 100, 676–677, 824 C. quadricarinatus, 28, 824 C. tenuimanus, 824 Cheylostigmaeus, 615

1098

Chilinidae, 391 Chilodonella, 122–123, 124f Chilomonas, 122 Chilopoda, 661 Chilostigma itasca, 987 Chimarra, 971–972, 971f, 974–976, 975f Chinonelasmatiodea, 712 Chirocephalidae, 688t, 855, 857f Chironomidae, 149, 200, 274–276, 288f, 292, 310f, 375f, 578, 618–619, 637–638, 639f, 647–648, 654, 853, 869, 917–918, 940, 956, 1043–1044, 1045t–1047t, 1049–1050, 1052f, 1053–1058, 1055f, 1066, 1069 Chironomus, 19f, 642, 654, 853, 863, 1054–1056 C. plumosus, 544 C. riparius, 580 Chlamydomonas, 748 C. acidophilia, 240 C. reinhardtii, 342 Chlamydotheca, 773t C. arcuata, 772 Chlorella, 117f, 124, 167, 172–173, 220, 254, 377 Chlorocyphidae, 895, 897–900, 897t–898t, 908–909 Chlorogomphidae, 896–899, 897t–898t, 921, 928 Chloronia, 1061 Chloroniella, 1061 Chloroperlidae, 934–935, 934t, 940t, 941–944 Chone, 525 Chonopeltis, 711, 734, 735f, 736–740, 750 C. australis, 737–738 Chordariidae, 191–192 Chordeumatida, 663f, 665 Chordodes, 304–306, 309–312, 315, 315f, 317–318, 321 C. brasiliensis, 317 C. japonensis, 315–317 C. kenyaensis, 308, 315–317, 319 C. morgani, 306f, 308f, 311f, 313–314, 313f–314f, 316 C. nobilii, 319 C. nobilli, 320–321 Chrissia, 773t Chromadora, 292 Chromadorida, 273, 290–291 Chronogaster, 289–290 C. africana, 289–290 C. troglodytes, 282f, 285, 289–290 Chrysogaster, 1043–1044, 1045t–1047t Chrysomelidae, 1004, 1005t–1007t, 1009–1011, 1013–1014, 1028–1029, 1028f Chrysomelinae, 1028–1029 Chrysops, 1043–1044, 1045t–1047t Chthamaloidea, 712 Chthamalus, 742 Chydorid, 679 Chydoridae, 688t, 697–698, 701 Cichlids, 410 Cicindis horni, 1022 Ciliophrys, 121 Cipangopaludina, 83–84 Cirripedia, 672t, 710–712, 742–748, 750–751 Cirrodrilus

Taxonomy Index

C. cirratus, 17f, 552f C. ezoensis, 558 Cladocera, 41, 69f, 70, 75, 77, 200, 375f, 578, 687–689, 688t, 693–699, 701, 734, 768, 1026 Cladocerans, 678–679 Cladophora, 408, 789 Cladotanytarsus mancus, 648 Clappia, 394t Clathrosperchoninae, 620t–625t Claudiella, 1019–1020 Clavidae, 163t Cletocamptus, 713f Clibanarus, 807–808 C. fonticola, 807 Climacea, 149 Climacia, 1064 C. areolaris, 1065 C. chapini, 1065, 1065f Clionaidae, 147 Clitellata, 12, 17f, 509–512, 521–523, 529, 531, 533, 536, 551–552, 565–566 Clitellate, 565 Clithon, 395f Cloeon, 877, 881, 883–885 C. dipterum, 885 Clogmia, 1043–1044, 1045t–1047t Clostridium botulinum, 102 Clunio, 1056 Clydosmylus, 1064 Cneoglossa edsoni, 1029, 1029f Cneoglossidae, 1004, 1005t–1007t, 1014, 1029, 1029f Cnesia, 1044, 1048f, 1049 Cnidaria, 40–41, 122, 191–192 Cochlacocyclops, 727 Cochliopidae, 390t Coeangrionidae, 909–910 Coelogynopora biarmata, 184–185 Coelom, 329 Coelomomyces, 725–726 Coelopa, 1043–1044, 1045t–1047t Coelopidae, 1043–1044, 1045t–1047t, 1056 Coelopina, 1043–1044, 1045t–1047t Coenagriocnemis reuniensis, 902–903 Coenagrion, 894–895, 899–900, 900f C. mercuriale, 918 C. puella, 904–905, 905f, 915–916, 923–924 Coenagrionidae, 894–895, 897–901, 897t–898t, 901f, 908, 909f, 922, 926, 928f Coenagrionoidea, 894–895, 897–899, 897t–898t Coenagrion puella, 641–642 Cognettia, 540 Coleoptera, 13–14, 20f, 100–101, 149, 375f, 579, 635–637, 851, 853, 855, 860–861, 860f, 863, 869, 970, 994, 1004, 1005t–1007t, 1008–1009, 1011, 1014, 1016–1017, 1025–1026, 1060 Coleps, 14f, 124f Collembola, 13, 19f, 70–71, 850–851, 853, 864–868, 1038–1039 Collotheca, 233f–234f, 248, 258–259 C. campanulata, 247, 247f C. ferox, 233f

C. libera, 233f, 238 C. trilobata, 233f Collothecacea, 233f, 234–235 Collothecidae, 231f, 232, 246 Collozoum longiforme, 119–120 Coloburiscidae, 874–877, 875t–876t, 879–880 Colpoda, 124f, 125 Concavinae, 712 Conchophthirus, 470 Conchostraca, 687 Conochilidae, 244–245, 245f Conochiloides C. natans, 250f Conochilopsis C. causeyae, 243–244 Conochilus, 237, 244, 246f, 247t, 251, 263–264, 698–699 C. dossuarius, 244, 245f C. hippocrepis, 244, 246, 255, 257f C. unicornis, 244–246, 254 Conoesucidae, 965, 966t–967t, 992 Continenticola, 183f Contulma, 971–972, 976–978, 993 C. paluguillensis, 974f, 986–987 Copepoda, 578, 672–673, 672t, 677–679, 681, 1026 Copepodid, 722, 724–725 Copepodite, 713f Copepods, 24, 41, 66–67, 69f, 70, 72, 75, 710–734, 713f, 715f–716f, 719f, 729f–732f, 748–750 Copestylum, 1043–1044, 1045t–1047t Coptotomus, 1022–1024 Coquillettidia, 1050 Coquillettidia perturbans, 642, 647 Corallomyxa, 119 Corbicula, 86–87, 424–426, 433–434, 434t, 438, 440, 444–445, 449, 449f, 454, 457, 466–469, 471–472, 474, 478, 481, 483–485, 747 C. fluminea, 83–84, 87, 433–434, 434t, 452–453, 466, 469, 474, 478–481 C. leana, 433–434, 434t, 481 Corbiculidae, 425–426, 432–433, 434t, 435, 438, 466 Corbiculoidea, 432 Cordulegaster bidentata, 922 Cordulegasteridae, 921 Cordulegastridae, 896–900, 897t–898t, 900f, 919 Cordulegastroidea, 896–900, 897t–898t Cordulia aenea, 914, 916f Corduliidae, 896–900, 897t–898t, 902–903, 912, 920–921, 928 Cordylophora, 159–161, 162f, 164–165, 164t, 165f, 167, 167f, 169–176, 172f–173f C. caspia, 173, 175 C. caspia japonica, 163t C. mashikoi solangiae, 163t Cordylophoridae, 163t Coregonus C. clupeaformis, 102–103, 319–320 C. nasus, 469 C. oidschian, 469 Corethrella, 1043–1044, 1045t–1047t

Taxonomy Index

Corethrellidae, 1043–1044, 1045t–1047t Corixidae, 639f, 951–957, 952f, 954t Cornops aquaticum, 1070 Coronuloidea, 712 Corophiidae, 793 Corophium, 208 Corophium curvispinum, 786–787 Corvospongilla, 137–138 C. mesopotamica, 137–138, 137f Corydalidae, 988, 1060–1063, 1060f, 1062f, 1069 Corydalinae, 1060–1061, 1063 Corydalus, 1061–1063 C. cornutus, 1061–1063 Coryphoridae, 874–877, 875t–876t Cosmopterigidae, 1069, 1069f Cosmopteriginae, 1068 Cougourdella, 864 Coxoplectoptera, 873 Crambidae, 1068–1069 Crangonyx, 790–791 C. pseudogracilis, 787 Crangonyx gracilis, 677 Craspacusta, 164–165 Craspedacusta, 159–169, 164t, 171–176 C. chuxiogensis iseana, 163t C. jsowerbyi, 14f C. podocysts, 167 C. sowerbii, 83–84, 165f–166f, 168f–169f, 171–175 C. sowerbii sinensis, 163t C. vovasi, 163t Craspedosomatidae, 663f Crassiclitellata, 510, 531, 533, 536, 538–540 Crassostrea, 445 Crassostrea gigas, 747 Cratenemertea, 206 Cretachordodes burmitis, 304–305, 305f Cretacimermis C. burmitis, 274 C. libani, 274 C. protus, 274f Creutzeria, 613 Criconema, 281f Cricotopus, 48–49, 288f, 289, 296f, 1043–1044, 1045t–1047t C. nostocicola, 295f Criodrilidae, 512 Criodrilus, 541 C. lacuum, 540–541, 545 Cristatella, 332 C. mucedo, 336, 338 Cristatellidae, 332 Crocothemis C. divisa, 917 C. erythraea, 914, 915f, 918–919, 921–922 C. servilia, 918–919 Crotoniidae, 601t–602t Crotonioidea, 601t–602t Crustacea, 12, 24, 28, 31f, 38–39, 671–682, 672t, 711, 759–760, 786, 798, 798f, 798t, 850, 852, 855, 857, 857f, 1026–1027 Crustaceans, 70, 73, 75 Cryptocorypha ornata, 665 Cryptomonas, 122, 239–240, 748

1099

Cryptophytes, 723 Cryptopygus C. antarcticus, 867 C. sverdrupi, 867 Cryptosporidium, 125, 264 C. hominis, 125 C. parvum, 125, 471 Ctenidae, 601t–602t, 602–604, 603f Ctenidia, 444 Ctenochauliodes, 1061 Ctenocheilocharis, 710 Ctenopoda, 687 Ctenostomata, 327–328, 334f, 340 Ctenothyadidae, 601t–602t, 620t–625t Cucujiformia, 1008 Culex, 19f, 603, 654, 732–733, 1043–1044, 1045t–1047t Culex pipiens quinquefasciatus, 300 Culicidae, 58, 106, 608, 637–638, 859–860, 1015, 1043–1044, 1045t–1047t, 1048f, 1049–1050, 1051f, 1053–1054, 1056–1057 Culicoides, 1043–1044, 1045t–1047t Culicomorpha, 1050, 1052 Culoptila, 979–981 C. moselyi, 965–969, 968f, 979–982, 983f Cumacea, 13, 672t, 782–784 Cumberlandia, 424, 489f Cupelopagis, 237–238, 259 C. vorax, 233f, 248–249, 249f, 258–259, 266–267 Curculionidae, 100–101, 1004, 1005t–1007t, 1009–1014, 1016, 1029, 1029f Curicta, 959 Cyanophthalma obscura, 206, 208 Cyathobodo, 122 Cybaeidae, 601t–602t, 602–604 Cyclestheria, 696 C. hislopi, 693f Cyclestherida, 687–689, 688t, 691–694, 696–697 Cyclestheriidae, 688t Cyclidium, 122–123, 124f, 125 Cycliophora, 212 Cyclocypris, 773t C. ampla, 771–772 C. ovum, 768–769 C. sharpei, 771–772 Cyclominae, 1029 Cyclomorphosis, 719 Cyclonaias, 424, 489f C. tuberculata, 454 Cycloneuralia, 212 Cyclophyllidea, 725–726 Cyclopidae, 717, 720, 726–729 Cyclopoida, 375f, 710, 712–727, 713f, 719f, 730–734 Cyclopoids, 70 Cyclops, 107, 291, 294 C. abyssorum, 725 C. furcifer, 724 C. scutifer, 724 C. singularis, 722 C. strenuus, 723 C. vicinus, 721, 724, 726 Cyclopsiella, 979–981

Cyclorrhapha, 1043–1044, 1050 Cyclothyadinae, 620t–625t Cylindroiuluspunctatus, 662f Cylindrothelphusa steniops, 829 Cymatia bonsdorfi, 646 Cyphon, 1012 Cyphonautes, 336f Cypretta, 773t C. kawatai, 773–774 Cypria, 773t C. ophtalmica, 772 C. ophthalmica, 771–772 C. turneri, 773 Cypricercinae, 768 Cypricercus centrura, 18f Cypriconcha, 773t Cyprideis, 769 Cyprididae, 758–761, 758f, 759t, 765f, 767–768, 772, 774–775 Cypridoidea, 757–760, 758f, 759t, 761f, 764, 764f, 766, 769 Cypridopsinae, 759t Cypridopsis, 765–766, 773t C. hartwigi, 773–774 C. vidua, 769 Cypridopsis vidua, 654 Cyprids, 745–746, 751 Cyprinidae, 768 Cyprinodon pecosensis, 410 Cyprinotinae, 759t Cyprinotus, 758f Cyprinus carpio, 465 Cypris, 743f, 745–746, 745f, 750 Cypris, 261, 773t C. balnearia, 772 C. pubera, 262f Cyprogenia, 424, 462–463, 489f C. aberti, 463, 465f Cyproidinae, 759t Cyrtobagous salviniae, 100–101 Cyrtonia tiba, 234f Cyrtophorids, 123f Cystobranchus C. meyeri, 572f C. salmositicus, 569f C. verrilli, 572f C. virginicus, 577f Cytheridae, 766f Cytherideidae, 758f, 759t Cytheridella ilosvayi, 769 Cytherissa, 758f C. lacustris, 770–771 Cytheroidea, 757–760, 758f, 759t, 764f, 766, 768–769 Cytheruridae, 759t Cyzcidae, 688t Cyzicus, 691f, 694–695

D

Dactylobiotus, 350t, 352–354, 352f–353f, 357–359, 357f, 362, 369–374 D. ambiguus, 371t–374t D. dispar, 368–369, 371t–374t D. grandipes, 371t–374t D. octavi, 371t–374t

1100

D. parthenogeneticus, 361, 362f D. selenicus, 354f, 371t–374t Dactylocephala, 199 Dactylopodola D. baltica, 216–217 Dactylopodolidae, 216 Dalyelliidae, 183f, 184–185, 193, 195, 197 Dalyellioida, 183f, 184, 186, 188 Daphnia, 173, 238, 261, 261f, 265–266, 265f, 268, 292, 654, 678–679, 688–689, 691f, 694–695, 697–700 D. laevis, 648 D. pulex, 261f D. pulicaria, 695f D. similis, 48–49 Daphniidae, 688t Daphniopsis, 697 Darlingtonia, 1054, 1055f Darwinula, 758f, 769 D. stevensoni, 766f, 768–769 Darwinulidae, 758f, 759t, 766f, 768, 774 Darwinuloidea, 757–760, 758f, 759t, 761f, 764f, 766, 768–769 Dasydtidae, 214–216 Dasydytes ornatus, 220f Dasydytidae, 211–212, 213f, 214–216, 218, 219t, 220–221 Dasytricha, 125 Daubaylia, 281f, 285, 291, 295 D. olseni, 295f–296f Daubayliidae, 281f Decapoda, 672t, 673–675, 674f, 678, 681–682, 798–800, 798t–799t, 800f Delevea, 1019–1020 Delphacidae, 100–101 Demanietta, 829 Demeijerea, 149 Demospongiae, 133, 135f, 142–143 Dendrobranchiata, 675f, 798–800 Dendrocephalus alachua, 690 Dendrocoelum lacteum, 186 Derocheilocaris, 710 Desmocarididae, 798, 799t, 804, 824 Desmocaris, 804 D. bislineata, 824 D. trispinosa, 808, 808f, 824 Desmodorida, 273 Desmognathus quadramaculatus, 577f Desmolaimus, 292 Desmoscolecida, 273 Deuterophebiidae, 1050 Deuterophlebia, 1043–1044, 1045t–1047t, 1048f, 1049 Deuterophlebiidae, 1043–1044, 1045t–1047t, 1048f, 1049–1050, 1052–1054, 1056, 1058 Deuterostoma, 184 Deutonymphs, 616f, 642–643 Devadattidae, 895–899, 897t–898t Diacyclops, 719–721, 723, 726–728 D. biceri, 720–721 D. languidus, 724 D. nanus, 724 Diacypris compacta, 773–774 Diamesa, 1043–1044, 1045t–1047t, 1054–1056 Diamesinae, 1054–1056

Taxonomy Index

Diamphidaxona, 632 Diamphipnoidae, 934–935, 934t Dianous, 1038–1039 D. coerulescens, 1038–1039 Diapause, 719–720 Diaphanosoma, 694 D. birgei, 648 Diaptomidae, 710, 713f, 724–729, 731–732 Diaptominae, 728 Diaptomus connexus, 48–49 Dibusa angata, 985 Dicercomyzidae, 874–877, 875t–876t Dichaeturidae, 213f, 214–216, 218 Dicranophoridae, 234–235 Dicranophorus, 234f Dicteriadidae, 895, 897–899, 897t–898t Dictynoidea, 601t–602t Dictyostelium, 120 Didacna, 747 Didymops, 896 Diestrammena, 318–320 Difflugia, 119, 231–232 Digenea, 181 Dikerogammarus villosus, 87–88, 87f, 788, 790–792 Dileptus, 124f Dinema, 121f, 725–726 Dineutes, 1010–1011 Dineutus, 1024–1025 Dinobryon, 121 Dioctophyme renale, 559 Diogenidae, 798, 799t, 807 Diosaccidae, 728 Diphascon, 350t, 356f–357f, 359f, 369 D. alpinum, 371t–374t D. carolae, 371t–374t D. granifer, 371t–374t D. higginsi, 371t–374t D. nobilei, 371t–374t D. patanei, 371t–374t D. recamieri, 371t–374t D. scoticum, 357f, 361, 368–369, 371t–374t Diplacodes, 908, 909f Diplectrona modesta, 972–974, 973f Diplopoda, 661–664, 662f–663f, 665f, 667, 667f Diplostraca, 687–688, 688t, 693–694 Diplura, 850–851 Diporeia, 102–103 Dipseudopsidae, 965, 966t–967t, 972–976, 975f, 991–992 Diptera, 13, 19f, 98–100, 106, 148–149, 150f, 274, 274f, 375f, 637–639, 639f, 641, 851, 853–855, 854f, 859–863, 859f, 869, 988, 991, 1004, 1013–1014, 1043–1044, 1044f, 1045t–1047t, 1049–1050, 1052–1058 Dipteromimidae, 874–877, 875t–876t Dipteropeltis, 711, 734, 735f, 736–738, 740 Dipteropeltis hirundo, 734, 740 Disparoneurinae, 894–895 Distocambarus, 825–826 D. crockeri, 817–819, 818f Distromatonemertea, 206 Dithinozerconidae, 608 Dixa, 1043–1044, 1045t–1047t

Dixella, 1043–1044, 1045t–1047t Dixidae, 1043–1044, 1045t–1047t, 1054, 1057–1058 Dolania, 884–885 D. americana, 883 Dolerocypris, 773t sinensis, 763f Dolichopodidae, 1043–1044, 1045t–1047t, 1054 Dolomedes, 604–606, 1068 D. aquaticus, 604–605 D. plantarius, 605 D. scriptus, 604–605 D. striatus, 604–605 D. tenebrosus, 593f, 604–605, 606f D. triton, 604–606, 868 D. vittatus, 604–605 Dolophilodes distinctus, 979–981, 981f Dolops, 711, 735–738, 735f, 740 D. discoidalis, 734 D. longicauda, 734 D. ranarum, 737–741 Donacia subtilis, 1028f Donaciinae, 1009, 1028–1029, 1028f Dorosoma petenense, 98–99 Dorvilleidae, 514 Dorydrilidae, 533–534 Dorylaimida, 273, 291 Dorylaimidae, 277f, 281f Dorylaimus, 276f–277f, 281f, 292 D. stagnalis, 286 Doryphoribius, 350t, 354, 369 D. doryphorus, 371t–374t D. evelinae, 371t–374t D. longistipes, 371t–374t D. macrodon, 371t–374t D. parthenogeneticus, 361, 368–369 D. scoticum, 361 D. tergumrudis, 371t–374t Dracunculus, 725–726 D. insignis, 733 D. medinensis, 733 Dranunculus D. medinensis, 107, 107f Dreissena, 86–87, 388, 424–426, 433–434, 440, 442–444, 445f, 449, 449f, 454, 457, 466–467, 469–471, 474–475, 483–484 D. bugensis, 102–103, 432f, 433–434, 434t, 472, 483–484 D. polymorpha, 83–84, 101–103, 432f, 433–434, 434t, 466–467, 472, 479–480, 483–484, 830, 925 D. rostriformis, 434 D. rostriformis bugensis, 83–84 Dreissenidae, 433, 434t, 435, 449 Drepanosticta, 894 Drepanotenia, 725–726 Drilophaga, 543 Dromus, 424, 489f Drosophila, 768–769 Drusus, 993 Dryomyzidae, 1043–1044, 1045t–1047t, 1056 Dryopidae, 1004, 1005t–1007t, 1008–1009, 1011–1014, 1016, 1029–1030 Dryopoidea, 1008

Taxonomy Index

Dryops, 1029–1030 Drypoidae, 1036 Dugastella, 803 Dugesia, 183f, 200, 291 D. benazzi, 191f D. tigrina, 186 Dugesiidae, 183f Dumontidae, 688t Dysmicohermes, 1061–1062 Dyticidae, 1014 Dytiscidae, 1004, 1005t–1007t, 1008–1017, 1020–1028, 1023f Dytiscoidea, 1008, 1021 Dytiscus, 1010–1011, 1013, 1022–1024 D. alaskanus, 1014–1015 D. dauricus, 1009 D. latissimus, 1016–1017

E

Ecclisocosmoecus, 991–992 Ecdyonurus, 883–884, 884f Ecdysozoa, 212, 591–592 Eceritotarsus catarinensis, 100–101 Echiniscidae, 350t, 351, 355 Echiniscoidea, 350t, 351, 369 Echiniscoides sigismundi, 361 Echiniscoididae, 350t Echiniscus, 350t, 352f, 355, 356f, 364f, 369–370, 377 E. blum, 371t–374t E. duboisi, 349f E. granulatus, 375 E. testudo, 368–369 E. wendti, 371t–374t Echinocotyle, 725–726 Echiura, 512, 522 Ecnomidae, 965, 966t–967t, 971–976, 973f, 990 Ectinosomatidae, 727–728 Ectobidae, 308 Ectocyclops, 721 Ectoprocta, 11, 327–336 Edoneus, 824 Eichhornia crassipes, 100–101, 614, 771, 1068–1069 Eimeria, 125 Elaphoidella, 711, 713f, 730 E. bidens, 720 Elaphoidellopsis, 730 Elassoneuria, 887–888 Elateridae, 1013–1014 Elateriformia, 1008 Electrogena, 881–882 Eleutherengonides, 601t–602t Elimia, 76, 393, 394t, 397–398, 400, 401t, 405, 409–410 E. cahawbensis, 404 E. clara, 404 E. crenatella, 410t E. livescens, 408, 410, 484 E. potosiensis, 408 Ellipsaria, 424, 489f Elliptio, 424, 454, 457–458, 462, 484, 489f E. arca, 455 E. complanata, 454, 461–462, 467–468, 475–476, 478–481, 484

1101

E. dilatata, 457–458, 462f E. nigella, 429, 429t Elliptoideus, 424, 461–462, 489f Ellisodrilus, 556–557 Elmidae, 1004, 1005t–1007t, 1008–1017, 1029–1031, 1030f–1031f, 1036 Elminae, 1030–1031 Elmis, 1030–1031, 1030f Elodea, 175–176, 247, 265–266, 576, 838 E. canadensis, 247, 247f Elodes, 1012 Elophila interruptalis, 1068 Elosa worallii, 239–240 Elpidium, 773, 773t Embolocephalus velutinus, 539f Emiliania, 122 Empedimermis cozii, 294, 294f Empididae, 988, 1050, 1052, 1056 Empidoidea, 1043–1044, 1045t–1047t, 1054 Enallagma, 894–895, 904, 923 E. ambiguum, 905, 907f E. clausum, 48–49 E. cyathigerum, 903–904, 904f, 909–910 E. ebrium, 646 E. risi, 905, 907f Encentridophorinae, 620t–625t Enchytraeida, 533 Enchytraeidae, 510, 517, 531, 533–534, 538–543, 545, 548 Endopterygota, 1060 Engaeus, 614, 805, 825–826 E. fultoni, 614 Engaewa, 805 Enoicyla, 992 E. pusilla, 992 Enopla, 205–206, 206t–207t Enoplida, 273, 278, 291 Enoplidae, 278 Enterococcus faecalis, 733 Entocytheridae, 759t, 763, 774 Entodinium, 125 Entognatha, 13, 19f, 850–851 Entoprocta, 11, 327, 340–341, 341f Entosiphon, 121 Eohypsibiidae, 350t, 353–355 Eohypsibioidea, 350t Eohypsibius, 350t, 353, 356f, 357–359, 369 E. nadjae, 371t–374t Eophreatoicus, 789 Eospongilla morrisonensis, 134 Eothinia, 234f Epactophanes richardi, 720 Epallage fatime, 909 Epallagopodinae, 620t–625t Epalxella, 125 Ephemera E. danica, 877, 878f, 881 E. lineata, 878–879, 879f Ephemerella, 886 Ephemerellidae, 874–877, 875t–876t, 879–884, 880f Ephemeridae, 873–879, 875t–876t, 879f, 881–882, 885 Ephemeroidea, 874–876

Ephemeroptera, 14, 98, 149, 292, 851, 853, 854f, 855, 859, 863, 873–876, 880–881, 887–888, 953, 959, 995–996, 1004, 1013–1014 Ephemerythidae, 874–876, 875t–876t Ephoron, 882, 887–888 E. shigae, 882 E. virgo, 887–888, 888f Ephydatia, 147 E. fluviatilis, 138–139, 138f–139f, 144, 146–148 E. muelleri, 15f Ephydra, 1056 E. cinerea, 857 E. gracilia, 48 E. hians, 596, 854–855, 854f, 857 Ephydridae, 1043–1044, 1045t–1047t, 1050, 1051f, 1054, 1056 Ephyridae, 596, 854–855, 854f Epigean, 781–782 Epilampra, 1067–1068 E. maya, 1067–1068 Epilamprinae, 1067–1068 Epimetopidae, 1004, 1005t–1007t, 1008, 1013–1014, 1031, 1031f Epimetopus, 1031 Epioblasma, 424, 430f, 438–439, 465, 482, 489f E. arcaeformis, 429, 429t E. biemarginata, 429, 429t E. brevidens, 430f, 465 E. capsaeformis, 438–439, 443f, 480 E. flexuosa, 429, 429t E. florentina florentina, 429, 429t E. haysiana, 429, 429t E. lenior, 429, 429t E. lewisii, 429, 429t E. personata, 429, 429t E. propinqua, 429, 429t E. sampsonii, 429, 429t E. stewardsonii, 429, 429t E. torulosa gubernaculum, 429, 429t E. torulosa rangiana, 430f, 465 E. torulosa torulsa, 429, 429t E. triquetra, 430f, 465 E. turgidula, 429, 429t Epiophlebia, 896, 910–911, 923–924, 924t Epiophlebiidae, 894–895, 895f Epiophlebioidea, 897–899, 897t–898t Epiphanes, 229f, 230, 234f, 250, 264 E. senta, 232, 251–252 Epiphanes brachionus spinosa, 15f Epiphragma, 1056 Epiprocta, 896 Epischura, 262, 711, 726–728 Epischura nevadensis, 18f Epistylis, 559 Epistylis niagarae, 726 Epitheca, 902–903 Epitrachys rugosus, 513 Epoicocladius, 886 Epophthalmia, 896 Eremobiotus, 350t Eriocheir sinensis, 94, 680, 806–807, 820, 830–831, 831f

1102

Erirhininae, 1029 Eristalis, 1043–1044, 1045t–1047t, 1049, 1049f Erpobdella, 581 E. fervida, 576–577 E. microstoma, 580–581 E. obscura, 570, 572–576, 578–582 E. octoculata, 577–581 E. punctata, 569f, 572–573, 575–582, 577f Erpobdellidae, 511, 566–572, 567f–568f, 571f, 574, 578 Erpobdelliformes, 511–513 Erpodiaceae, 375 Errantia, 531 Erythemis simplicicollis, 647, 918 Erythraeidae, 988 Erythraeoidea, 618 Erythraiae, 601t–602t Erythrodiplax berenice, 922 Erythromma lindenii, 906–907, 906f Esanthelphusa, 829 Escherichia coli, 58, 444 Esox lucius L., 677 Etherioidea, 428, 459–460 Ethmolaimus, 286, 288 Euastacus, 805, 824–826, 825t E. spinifer, 614 Eubasilissa, 979–981 Eubranchipus, 695–696 E. bundyi, 690f E. moorei, 695 Eucarida, 13 Eucestoda, 181 Euchlanis, 229f, 237–238, 262f E. dilatata, 238, 241–242, 244 Euchordodes, 318 E. nigromaculatus, 317 Eucorethra, 1043–1044, 1045t–1047t, 1048f, 1049 Eucyclopinae, 727 Eucyclops, 713f, 723, 726–728 E. bathanalicola, 721 E. serrulatus, 723 Eucypridinae, 759t Eucyprinotus, 773t Eucypris, 773t E. virens, 760, 768–769, 772–774, 777 Eudiaptomus gracilis, 734 Eudorylaimus E. andrassyi, 286–288 Eudrilidae, 512–513 Eudriloidea, 512–513 Euglena, 121 Euglenoids, 725–726 Euglypha, 118–119 Euhirudinea, 511–512 Euholognatha, 934–935, 934t Eulichadidae, 1004, 1005t–1007t, 1009, 1013–1015, 1031, 1031f Eulichas, 1031 Eulimnadia diversa, 695–696 Eulimnogammaridae, 792–793 Eumalacostraca, 672t, 784 Eunapius, 137 E. carteri, 84–85, 141f, 149 E. fragilis, 484

Taxonomy Index

Eunicida, 514, 520–521 Euparyphus, 1043–1044, 1045t–1047t Eupera, 432–433, 438–439, 441, 459 E. cubensis, 432–433 Euperinae, 432–433, 438 Euphaeidae, 895, 897–899, 897t–898t, 908–909 Euphrosinidae, 521 Euplotes, 122–123, 125 E. daidaleos, 117f Eupodides, 601t–602t Euproctus platycephalus, 149 Eurotatoria, 227, 237 Eurypterida, 592, 600 Euryrhynchidae, 798, 799t Euryrhynchina, 804 Euryrhynchoides, 804 Euryrhynchus, 803–804 Eurytemora, 726, 733 Eurytemora affinis, 83–84 Eustheniidae, 934–935, 934t Eustigmaeus, 615 E. frigida, 615f Eutardigrada, 347–348, 348f–349f, 350–357, 350t, 352f, 356f–357f, 359–360, 363–364, 364f, 366, 367f, 369–370 Euthyadinae, 620t–625t Euthyplociidae, 874–876, 875t–876t, 879 Eutrochozoa, 184, 196 Eylaidae, 601t–602t, 620t–625t, 628–629, 637–638, 639f Eylaioidea, 635–637, 653 Eylais, 635–644, 636f, 639f, 646–648, 653 E. discreta, 636 E. euryhalina, 636–637 Eylaoidea, 601t–602t, 618–619, 620t–625t, 628, 632–634, 637, 641–643

F

Fabaeformiscandona, 773t Fabria inornata, 965–969, 968f Facetotecta, 711–712 Falcataria, 62 Fallicambarus, 805, 825–826 F. devastator, 826, 838 F. fodiens, 826–827 F. gordoni, 826 F. strawni, 817–819, 818f Famelobiotus, 350t Fasciola hepatica, 101 Faunus ater, 391–392, 391f, 398 Faxonella, 825 F. clypeata, 838 Fecampiida, 182, 196–197 Feltria, 639f, 644 Feltriidae, 601t–602t, 620t–625t, 639f Ferradasiidae, 601t–602t, 620t–625t Filinia, 235, 243, 243f, 263–264, 263f F. longiseta, 255, 256f, 262–263 F. terminalis, 256f Fimbriaria, 725–726 Fimbricyclops, 730 Fletcherimyia, 1043–1044, 1045t–1047t, 1049, 1049f, 1054 Floscularia, 245, 247t, 248, 258–259 F. conifera, 244, 246–248, 246f, 248f–249f, 253

F. janus, 248 F. ringens, 244 Flosculariacea, 232, 234–235 Flosculariidae, 231f, 244, 246, 258–259 Fluminicola, 394t Fontigens, 395 Fontinalis, 677 Forcipomyia, 1043–1044, 1045t–1047t Forelia, 632, 644 Foreliinae, 620t–625t, 644–645 Fossaria, 391, 396 Fractonotus, 350t, 357–359 Francisella tularensis, 101 Fredericella, 328–329, 330f, 332–333, 337–338 F. indica, 336–337 F. sultana, 333f, 336–337 Frontipodopsidae, 601t–602t, 620t–625t Frontipodopsis, 632 Frontonia, 262f Frullania eboracensis, 231–232 Furcatergalia, 874–876 Fusconaia, 424, 461–462, 489f F. ebena, 457, 461f, 467–468, 471 F. flava, 438–439, 443f, 463, 465f F. subrotunda, 457–458, 462f

G

Galatheoidea, 807 Galba, 396 Galerucella nymphaeae, 1028–1029 Galerucinae, 1028–1029 Gallocaris, 824 Gambusia, 1056–1057 G. affinis, 410, 868 Gammaridae, 791, 793 Gammarinema, 295 Gammarus, 173, 292, 673f, 790–791, 793 G. acherondytes, 790 G. duebeni, 786–787, 792 G. fasciatus, 792 G. fossarum, 786–788 G. lacustris, 677, 786–787, 792 G. limnaeus, 785–786 G. minus, 677, 681, 786, 788–791 G. pseudolimnaeus, 787 G. pulex, 677, 788–789, 791–792 G. roeseli, 787–788 G. tigrinus, 792 Gastromermis anisotis, 284f, 286, 292, 293f, 294–295, 295f–296f Gastropoda, 383–400, 389f, 401t, 402f, 404–410, 411f, 413, 578 Gastrotricha, 11, 211–214 Gecarcinucidae, 798, 799t, 802, 805–806, 824, 825t Gelastocoridae, 951–953, 952f, 954t, 955–956 Geleia, 122–123 Gelyella, 710 G. droguei, 717 Gelyelloida, 710, 712–717, 728–729 Geocentrophora G. baltica, 199 G. sphyrocephala, 199 Geocharax, 805

Taxonomy Index

Georissidae, 1004, 1005t–1007t, 1008, 1013–1014, 1031–1032, 1032f Georissus, 1031–1032 G. crenulatus, 1031–1032, 1032f Geosesarma notophorum, 806–807, 820 Gerridae, 70, 605, 637, 646, 647f, 859–860, 860f, 951–955, 952f, 954t, 957–958 Gerris, 637, 955, 958, 958f G. incognitus, 958, 958f G. thoracicus, 958, 958f Gerromorpha, 951–955, 952f, 954t Giardia, 58, 264 G. intestinalis, 120 G. lamblia, 471 Gieysztoria, 184–185 G. rubra, 183f Glacidorbidae, 389, 391, 401 Glacidorboidea, 389 Glebula, 424, 489f Globigerinoides ruber, 119 Globonautes macropus, 814, 823, 838–839 Glochidia, 459–461, 466 Glomerida, 664 Glossinidae, 58, 106 Glossiphonia complanata, 572, 574–581 Glossiphoniidae, 511–512, 516, 566–572, 567f–568f, 570f–571f, 574–575, 577–578 Glossosomatidae, 965–969, 968f, 971–972, 971f, 974–982, 981f, 983f, 985–986, 989–990 Glossosomatoidea, 965, 966t–967t Glutops, 1043–1044, 1045t–1047t Glyphidrilidae, 512 Glyphopsyche sequatchie, 993 Glyphotaelius pellucidus, 985–986, 992 Glypotendipes paripes, 580 Glyptograpsidae, 798, 799t Gmelinoides fasciatus, 785–786 Gnathopods, 784 Gnathostomulida, 184, 187, 230 Gnosonesimidae, 195 Gobio gobio, 923–924 Gobius alcockii, 149 Goeracea genota, 965–969, 968f, 974–976, 975f Goeridae, 965–969, 966t–967t, 968f, 974–976, 975f, 985, 990–992 Gomphidae, 896–902, 897t–898t, 901f, 908, 909f, 914, 917–921, 926, 928–929, 928f Gomphidia, 908, 909f Gomphocythere, 769 Gomphoidea, 896–899, 897t–898t Gomphus G. flavipes, 915, 917f G. vulgatissimus, 915, 921 Goneplacidae, 798, 799t Gonidea, 424, 489f G. angulata, 427 Gonideini, 424, 489f Goniomonas, 121f, 122 Gonopods, 668 Gonyaulax catenella, 122 Gordiida, 304–311, 309f–312f, 313–317, 314f, 320–322 Gordionus, 308f, 310f, 322–323 chinensis, 318–319 lokaaus, 308f

1103

Gordius, 305–306, 309–310, 312, 314f–315f, 315, 317–318, 320–321 G. aquaticus, 306, 312–313 G. difficilis, 306f, 308f–309f, 309–311, 312f–313f, 317–318 G. dimorphus, 317 G. robustus, 305, 306f, 311f, 312–314, 313f, 315f, 316–317, 320 G. robustus, 304f G. tolosanus, 315–316 Gordius difficilis, 17f Gramastacus, 805 Grandosmylus, 1064 Graphiurus monardi, 717 Graphoderus, 1022–1024, 1023f G. bilineatus, 1016–1017 Graptemys G. geographica, 470 G. versa, 469 Gretacarinae, 620t–625t Gripopterygidae, 934–935, 934t Grumicha grumicha, 965–969, 968f Grumichella, 992 Gryllidae, 308, 1069–1070 Gryllotalpidae, 1069–1070 Gryllus, 310f G. firmus, 307f Grypopterygidae, 945 Guineaxonopsinae, 620t–625t Guloptiloides, 885 Gymnodinium, 122 G. simplex, 122 Gymnolaemata, 327–329, 336 Gymnoplea, 710, 712–713 Gymnostomes, 123f Gyratrix, 184, 197–198 G. hermaphroditus, 183f, 184, 197–198 Gyraulus albus, 404 Gyrinidae, 860, 860f, 1004, 1005t–1007t, 1008–1014, 1016–1017, 1024–1025, 1024f Gyrinus, 1024 G. marinus, 1024–1025 Gyrocotylidea, 181 Gyrotoma, 393, 394t, 409

H

Haber, 536–537 Habrophlebia lauta, 881–882, 882f Habrotrocha, 231–232 H. rosa, 266–267 Hadodiaptomus dumonti, 726 Hadromerida, 135f Haemadipsidae, 511 Haementaria H. ghilianii, 568–570, 578–579 Haementeria ghilianii, 514 Haemopidae, 511, 566, 568–570, 578 Haemopis H. grandis, 569f H. marmorata, 569f, 576–577, 580–581 H. sanguisuga, 579 H. terrestris, 577f Halacaridae, 601t–602t, 609f, 613–614 Halacaroidea, 601t–602t, 613–614 Halesus

H. digitatus, 985 H. radiatus, 985 Haliclonidae, 135f, 147 Halicyclops thermophilus, 721 Haliplidae, 1004–1011, 1005t–1007t, 1013–1017, 1025–1026, 1025f Haliplus, 1025–1026 H. ruficollis, 1026 Halobates, 857, 953, 955, 957 Halobiotus, 350t H. crispae, 360–361, 368–369 Halomonhystera parasitica, 295, 297f Halteria, 124f Haltidytes festinans, 215f Hamiota, 463 H. altilis, 427f, 463–465 Hamiota altilis, 16f Haplohexapodibius, 350t, 353–354 Haplomacrobiotus, 350t, 353–354 Haplopharyngida, 191–193, 195 Haplopoda, 687 Haplosclerida, 133, 135, 135f Haplosclerina, 135 Haplosialis, 1060–1061 Haplotaxida, 533 Haplotaxidae, 531, 533, 536–539, 543 Haplotaxis H. aedeochaeta, 535 H. gordioides, 543 Haplozetidae, 601t–602t Haptorids, 123f Harnischia, 1053 Harpacticidae, 711 Harpacticoida, 375f, 710–723, 713f, 725, 727–730, 733, 748 Harpacticoids, 70 Harpagopalpidae, 601t–602t, 620t–625t Hebesuncus, 350t, 357–359 Hebridae, 951–953, 954t Hedriodiscus, 1056 Helcomyzidae, 1056 Helichus, 1013, 1029–1030 Helicophidae, 965, 966t–967t Helicopsyche, 992 H. borealis, 965–969, 968f Helicopsychidae, 965–969, 966t–967t, 968f, 976–978, 990–991 Heliosphaera, 118f Heliozoa, 121 Helisoma, 100, 396, 400, 401t, 830 H. anceps, 396, 399 Helobdella, 17f, 568f H. papillata, 576–577 H. robusta, 566 H. stagnalis, 572, 574, 576–581 Helophoridae, 1004, 1005t–1007t, 1008, 1011–1014, 1032, 1032f Helophorus, 1032 H. brevipalpis, 1032 H. micans, 1032 H. nubilus, 1032 H. orientalis, 1013, 1032 H. rufipes, 1032 Helotrephidae, 951–953, 954t Hemerobiiformia, 1064

1104

Hemiacridinae, 1070 Hemiboeckella, 726 Hemicordulia, 896 Hemicycliophora, 280f Hemicycliophoridae, 280f Hemicytheridae, 759t Hemimysis, 94 Hemiphlebiidae, 894, 897–899, 897t–898t Hemiptera, 13, 20f, 100–101, 149, 595–596, 635–638, 851, 853, 855, 857, 859–860, 860f, 863–864, 864f, 951–961, 952f, 957f Hemisarcoptoidea, 601t–602t Hemistena, 424, 489f Hemocyanin, 853 Heptagenia diabasia, 20f Heptageniidae, 874–876, 875t–876t, 878–885, 879f, 884f Heptagenioidea, 874–876 Herbrossus elouardi, 885 Hermaphrodites, 364–366, 394–395, 696 Hermatobatidae, 951–953, 954t Herpetocypris, 773t Hesperocorixa, 951–953, 952f Hesperodiaptomus, 711, 727–728 H. shoshone, 720 Hesperus kovaci, 1038–1039 Heteragrionidae, 896–899, 897t–898t Heterobranchia, 389, 390t, 394t Heterocapsa, 122 Heteroceridae, 1004, 1005t–1007t, 1008, 1013–1014, 1032–1033, 1033f Heterocheilidae, 1056 Heterocope, 711, 726–728 Heterocypris, 292, 765–766, 773t H. carolinensis, 773 H. incongruens, 769, 772–774, 777 Heterodonta, 425–426, 450–451 Heterolepidoderma, 212–214, 220f H. macrops, 215f H. ocellatum, 215f, 216–217 Heteromita, 122 Heteronemertea, 205–207, 206t–207t Heteroplectron americanum, 965–969, 968f Heteroptera, 605, 639f, 853, 854f, 951–953 Heterotardigrada, 347–348, 349f, 350–351, 350t, 352f, 355–357, 356f, 359–360, 363–364, 364f, 369–370, 371t–374t Heterotrichs, 123f Hexactinellida, 133 Hexagenia, 881–882, 885–886 H. limbata, 98 H. rigida, 98, 99f, 881 Hexamita, 120, 121f Hexapoda, 12, 759, 849–851, 853 Hexapodibius, 350t Hexarthra, 235, 236f, 238, 243, 250, 263–264 H. fennica, 48–49, 250f, 255 H. mira, 243–244 Hexatoma, 1043–1044, 1045t–1047t Hibernaculae, 332–335 Himalopsyche, 976 H. phryganea, 976 Hirudinaria bpling, 569f Hirudinea, 12, 509–514, 516–517, 521, 529, 533, 565, 834

Taxonomy Index

Hirudinea, 511 Hirudinida, 512, 565–567 Hirudinidae, 566–570, 569f, 577 Hirudiniformes, 511–513, 567–570, 567f–568f, 570f, 572 Hirudo H. medicinalis, 104, 568–570, 569f, 578–581, 583 H. verbana, 568–570, 578, 583 Hislopia, 329f, 331, 335–336, 340 H. malayensis, 334f, 336f H. natans, 332f Hislopidae, 340 Hispinae, 1028–1029 Histeroidea, 1008 Histiogaster, 612f Histiostomatidae, 601t–602t, 612–613, 612f Histiostomatoidea, 601t–602t Histriobdellidae, 519–520, 520f Hobbseus, 825 Hofstenidae, 192 Holometabola, 1060 Holopediidae, 688t Holothyrida, 607 Holtodrilus truncatus, 553, 557, 559 Hominidae, 8 Homocaligidae, 601t–602t, 614–615, 614f Homocaligids, 614 Homocaligus, 614, 614f Homoptera, 100–101, 951–953 Homoscleromorpha, 133 Hoplonemertea, 205–206, 206t–207t, 208 Horelophinae, 1034 Horelophopsinae, 1034 Hormosianoetus mallotae, 613 Huitfeldtia rectipes, 642 Huitfeldtiinae, 620t–625t Hungarohydracaridae, 601t–602t, 620t–625t Huntemanniidae, 711 Hyalella, 677, 680, 784f, 786–791, 788f, 793 H. azteca, 680, 790 H. montezuma, 578, 791 Hyalinella H. orbisperma, 336 H. punctata, 336–338 Hyallela, 19f Hybomitra, 1043–1044, 1045t–1047t Hydatophylax, 991 Hydra, 159–162, 160f, 163t–164t, 164–167, 164f, 170–177, 172f, 175f H. littoralis, 172 H. viridissima, 164f, 167, 170–173, 170f H. visidissima, 172 Hydracarina, 925 Hydrachna, 631f, 635–642, 639f, 646–649, 653–654 H. conjecta, 638, 646 Hydrachnidae, 601t–602t, 620t–625t, 628–629, 631f, 637–638, 639f, 642–643 Hydrachnidiae, 601, 601t–602t, 604, 613–650, 618f, 620t–625t Hydrachnoidea, 601t–602t, 618–619, 620t–625t, 632–633, 635–637, 641, 653 Hydradephaga, 1008, 1034

Hydraenidae, 1004, 1005t–1007t, 1008–1009, 1011–1014, 1017, 1033, 1033f Hydrellia, 1043–1044, 1045t–1047t Hydridae, 161, 163t Hydrilla H. azteca, 581 H. verticillata, 576 Hydrobiidae, 389–390, 390t, 392, 394t, 395, 398, 400, 409–412, 409f, 412f, 1009–1010 Hydrobiosidae, 965–969, 966t–967t, 971–972, 971f, 974–979, 975f, 989–992 Hydrochidae, 1004, 1005t–1007t, 1008, 1011–1014, 1033 Hydrochoreutinae, 620t–625t Hydrochus, 1033, 1034f Hydrocyphon, 1037–1038 Hydrodroma, 645–648, 651, 654 H. despiciens, 628f, 635, 637, 648 Hydrodromidae, 601t–602t, 620t–625t, 628f Hydrolutos, 1070 Hydrometra, 951–953, 952f, 957 Hydrometridae, 637, 951–953, 952f, 954t, 957 Hydropharus, 1043–1044, 1045t–1047t Hydrophilidae, 1004, 1005t–1007t, 1008–1009, 1011–1014, 1016, 1022–1025, 1033–1034, 1034f Hydrophilinae, 1034 Hydrophiloidea, 1008, 1011–1012, 1033–1034 Hydroporinae, 1022–1024 Hydropsyche, 986–987, 990 Hydropsychidae, 863–864, 965, 966t–967t, 971–981, 971f, 973f, 975f, 982f, 986, 989–992 Hydropsychoidea, 965, 966t–967t Hydroptila, 979–981, 982f Hydroptilidae, 965–969, 966t–967t, 971–982, 971f, 973f, 981f–983f, 985–986, 988–992, 994 Hydroptiloidea, 965, 966t–967t Hydropysche, 992 Hydrosalpingidae, 965, 966t–967t Hydroscapha, 1018 H. natans, 1018, 1018f H. redfordi, 1018, 1018f Hydroscaphidae, 1004, 1005t–1007t, 1008–1009, 1011, 1013–1015, 1018, 1018f Hydrous piceus, 1009, 1016 Hydrovolzia, 637–638 Hydrovolziidae, 601t–602t, 620t–625t Hydrovolzioidea, 601t–602t, 618–619, 620t–625t, 637 Hydrovolzoidea, 653 Hydrozetes, 609–611, 610f Hydrozetes lacustris, 611 Hydrozetidae, 601t–602t, 609–611, 610f Hydrozetoidea, 601t–602t Hydryphanidae, 631f Hydryphantes, 628–629, 638–639, 646–648, 654 H. tenuabilis, 635–636, 638–639, 641 Hydryphantidae, 601t–602t, 620t–625t, 631f, 637, 639–640, 642–643, 646, 651–652, 654 Hydryphantinae, 620t–625t Hydryphantoidea, 601t–602t, 618–619, 620t–625t, 626–628, 632–634, 637, 639, 641, 645, 653

Taxonomy Index

Hygobiidae, 1008, 1013 Hygrobates, 631f, 636, 639–640, 642–643, 645–646, 648 H. fluviatilis, 649 H. nigromaculatus, 648 Hygrobatidae, 601t–602t, 620t–625t, 631f, 988 Hygrobatoidea, 601t–602t, 618–619, 620t–625t, 632–633, 635, 637–639, 641–644, 647–648, 653–654 Hygrobia H. austalasieae, 1026–1027 H. davidii, 1026–1027 H. hermanni, 1026–1027, 1026f H. nigra, 1026–1027 H. tarda, 1026–1027, 1026f H. wattsi, 1026–1027 Hygrobiidae, 1004, 1005t–1007t, 1008, 1011, 1013–1016, 1021, 1026–1027, 1026f Hygrophila, 389, 390t, 391 Hygrotus salinarius, 1012 Hymenoptera, 13, 851, 855, 863, 988, 1057, 1059–1060, 1068 Hymenosomatidae, 798, 799t Hymenostomes, 123f Hypania invalida, 526 Hypechiniscus, 350t, 355, 369 Hypochilus petrunkevitchi, 611 Hypogastruridae, 867 Hypolestidae, 896–899, 897t–898t Hypopodes, 612 Hyposmocoma, 1069, 1069f Hypotrichinidae, 191–192 Hypotrichs, 123f Hypsibiidae, 350t, 353–355, 366 Hypsibioidea, 350t Hypsibius, 350t, 352f, 354, 357f, 359, 366, 369, 377 H. arcticus, 371t–374t H. augusti, 371t–374t H. convergens, 368–369, 371t–374t H. dujardini, 357f, 361, 366–369, 371t–374t, 377 H. janetscheki, 369–370 H. klebelsbergi, 369–370, 371t–374t H. microps, 371t–374t H. roanensis, 359, 371t–374t H. scabropygus, 359 H. thaleri, 369–370 Hypsiibiidae, 366 Hypsogastropoda, 389–390, 390t Hyridella depressa, 480 Hyriidae, 428, 440, 460 Hyriopsis schlegeli, 95–96, 95f

I

Iberobathynella andalusica, 783f Ichneumonidae, 988 Ichthybotidae, 874–876, 875t–876t Ichthydium, 212–214, 216, 218 I. podura, 218 I. skandicum, 215f I. squamigerum, 215f Ichthyobodo, 121 Ichthyodinium chabelardi, 122 Ichthyophthirius, 123

1105

Ictinogomphus I. ferox, 915 I. rapax, 902–903 Illinobdella, 567 Illyocryptus, 18f Ilybius, 1013 I. montanus, 1017 Ilyobius, 1060–1061 Ilyocryptidae, 688t Ilyocyprididae, 758–759, 758f, 759t Ilyocypris, 758f, 773t Ilyodrilus templetoni, 530f, 541, 543 Incertae Sedis, 350t Incoltorrida, 1019–1020 Indosialis, 1060–1061 Insecta, 13, 850–851 Insuetifurca, 350t, 356f Integripalpia, 965–972, 966t–967t, 976–979, 989–992, 995 Io, 410–411 I. fluvialis, 389f, 390 Iridinidae, 440, 457–458 Ironus, 292 Ischnura, 894–895, 908, 909f, 911, 918–919 I. aurora, 918 I. elegans, 902–903, 903f, 906–907, 906f, 910–911, 912f, 913–914, 918–919 I. hastata, 911 I. heterosticta, 922 I. senegalensis, 911, 918–919 I. verticalis, 647 Isobactrus uniscutatus, 613–614 Isocapnia, 859 Isochaetides, 534 Isocypridinae, 759t Isocypris, 773t Isohypsibiidae, 350t, 352, 354, 357, 366 Isohypsibioidea, 350t Isohypsibius, 350t, 352f, 353–354, 356f–357f, 366, 369, 371t–374t I. annulatus, 366 I. augusti, 371t–374t I. baldii, 371t–374t I. basalovoi, 371t–374t I. canadensis, 371t–374t I. deflexus, 352 I. elegans, 371t–374t I. granulifer, 364, 366, 371t–374t I. kenodontis, 371t–374t I. lunulatus, 349f, 354f, 371t–374t I. monoicus, 363f, 364–366, 365f, 368–369 I. monoicus, 363f I. nodosus, 366 I. papillifer, 371t–374t I. sattleri, 371t–374t I. schaudinni, 371t–374t I. tetradactyloides, 371t–374t I. tuberculatus, 371t–374t Isohysibius I. deconincki, 371t–374t Isomermis benevolus, 294, 294f Isonychiidae, 874–876, 875t–876t, 879, 885 Isopoda, 13, 672–673, 672t, 675, 678–680, 682, 782–787, 789–794 Isoptera, 863

Isostictidae, 894–895, 897–899, 897t–898t Isotomurus palustris, 867 Itaipusa graefei, 184–185 Itaquascon, 350t, 355 I. trinacriae, 363f Itaspiella parana, 184–185 Itauara julia, 979–981, 981f Ithytrichia, 976–978 Itura, 234f

J

Johnstoniana rapax, 616 Johnstonianidae, 601t–602t, 615–617, 616f Juga, 390, 400, 401t Julidae, 662f, 664–665 Jutumia kosteri, 410t

K

Kabatarina pattersoni, 711 Kakaducarididae, 798, 799t, 824, 825t Kakaducaris glabra, 804 Kalyptorhynchia, 183f, 184, 186–188, 197–198 Kambaitipsychidae, 965, 966t–967t, 969, 969f Kantacaridae, 601t–602t, 620t–625t Karyorelictids, 123f Kathablepharis, 122 Katodinium, 122 Kawamuracarinae, 620t–625t Kazacharthra, 687 Kellicottia bostoniensis, 250f Kempyninae, 1066 Kempynus, 1064 Kentrophoros, 124, 127 Keratella, 229–230, 234–235, 238, 261, 263–264, 331 K. americana, 238 K. cochlearis, 239–240, 243–244, 255, 256f–257f, 257, 260–263 K. quadrata, 242, 255, 256f K. slacki, 262–263 K. taurocephala, 239 K. testudo, 238, 255, 262–263 K. tropica, 262f Kinorhyncha, 212, 591–592 Koenikea wolcotti, 633f Koinoporus, 207t K. mapochi, 207t Kokiriidae, 965, 966t–967t, 974–976, 975f, 990 Kongsbergia, 632f Krendowskiidae, 601t–602t, 620t–625t, 637–638 Kurzia longirostris, 693f

L

Labyrinthula, 125 Lacinia mobilis, 783 Lacinularia, 244, 247t, 263 L. elliptica, 244 L. flosculosa, 231f, 245f, 246 Lacrymaria, 122–123 Laevicaudata, 687–688, 688t, 691–692, 696–697 Lagenidium, 725–726 Lamprodrilus, 534–535 L. isoporus, 542–543 L. mrazeki, 541–542

1106

Lampsilini, 489f, 424, 458–460, 462 Lampsilis, 489f, 424, 438–439, 444, 447f, 453–454, 460, 463–465 L. abrupta, 466 L. binominata, 429, 429t L. cardium, 438–439, 443f, 455, 459f, 460, 463, 463f, 465f, 466 L. cariosa, 468 L. fasciola, 427f, 455, 463–465 L. ornata, 455 L. radiata, 87f L. reeviana, 466 L. siliquoidea, 444, 460, 466, 476, 480–481 Lampyridae, 1004, 1005t–1007t, 1009–1010, 1013–1014, 1035, 1035f Lancaris, 803 Lanx, 410t Laophontidae, 711, 727 Lara, 1016, 1031f Larainae, 1030–1031 Larval L. aquatica, 638–639 Lasaea, 433 Lasidia, 459–460 Lasiocephala basalis, 987–988 Lasmigona, 424, 427, 460, 489f L. complanata, 457, 461f, 465 L. compressa, 454 L. subviridis, 454 Latona, 694 Latrodectus, 605 Laversia, 637 Laversiidae, 601t–602t, 620t–625t Lavigeriinae, 393 Lebertia, 638–639 Lebertiidae, 601t–602t, 620t–625t Lebertioidea, 601t–602t, 618–619, 620t–625t, 632–633, 635, 637–639, 641–643, 647–648, 653–654 Lecane, 229f, 234–235, 262f, 266, 268 L. quadridentata, 237f, 251 Lecithoepitheliata, 183f, 184, 186, 192–198 Ledermuellaria, 615 Leeches, 511 Legionella, 58 Leiodidae, 1004, 1005t–1007t, 1013–1015, 1035, 1035f Lemiox, 424, 489f Lemna, 62, 218, 247, 867 Lepiceridae, 1004, 1005t–1007t, 1008–1009, 1013–1015, 1019, 1019f Lepicerus L. bufo, 1019 L. inaequalis, 1019 L. pichilingue, 1019, 1019f Lepidocaris, 688 Lepidochaetus zelinkai, 215f Lepidodasyidae, 216 Lepidodermella, 213–214, 218 L. squamata, 214, 215f, 216–218, 217f, 220 L. triloba, 220f Lepidoptera, 13, 100–101, 149, 851, 855, 863, 965, 970, 978–979, 1059, 1068–1069, 1069f Lepidostoma, 992 L. basale, 987–988 L. podagerum, 991

Taxonomy Index

Lepidostomatidae, 965, 966t–967t, 974–976 Lepidurus, 689–690, 696 L. bilobatus, 691f L. lemmoni, 695 Lepomis, 319–320 L. cyanellus, 677 L. gibbosus L., 406 L. microlophus, 406 Leptestheria compleximanus, 693f Leptestheriidae, 688t Leptinillus validus, 1035 Leptoceridae, 148–149, 965–969, 966t–967t, 968f, 971–976, 971f, 973f, 975f, 978–982, 979f, 981f–983f, 989–992, 995 Leptocerus, 991 L. americanus, 965–969, 968f Leptocytheridae, 759t Leptodea, 424, 489f L. fragilis, 452, 456f Leptodiaptomus, 713f L. minutus, 724 Leptodora, 245, 694, 696, 698–699 L. kindti, 694f Leptodoridae, 688t Leptohyphidae, 874–878, 875t–876t, 883–884 Leptomyxa, 119 Leptonema, 974–976, 975f Leptophlebia, 882, 887 L. cupida, 881 Leptophlebiidae, 874–885, 875t–876t, 880f, 882f Leptopodidae, 951–953, 954t Leptopodomorpha, 951–953, 952f, 954t Leptosialis, 1060–1061 Leptoxis, 394t, 400, 409–411, 411f L. ampla, 410t L. plicata, 410t L. taeniata, 410t Lestes, 894, 899–900, 913, 921, 924–925, 926f L. barbarus, 906–907, 906f L. congener, 913, 922, 924–925, 926f L. disjunctus, 640, 924–925, 926f L. dryas, 924–925 L. eurinius, 924–925 L. forcipatus, 924–925, 926f L. rectangularis, 924–925, 926f L. sponsa, 923–924 L. vigilax, 924–925 L. virens, 911–912, 913f Lesteva, 1038–1039 Lestidae, 897–899, 897t–898t, 911, 926 Lestoidea, 894, 897–899, 897t–898t Lestoideidae, 897–899, 897t–898t Lethaxonidae, 601t–602t, 620t–625t Lethocerus, 951–953, 952f, 955–956, 959, 961 L. maximus, 951, 953 L. medius, 957, 957f Leucorrhinia, 922 L. dubia, 914, 915f L. intacta, 918 L. pectoralis, 929 Leucotrichia, 76, 991–992 L. fairchildi, 979–982, 983f Leucotrichiinae, 989 Leuctra, 936, 938f

Leuctridae, 934–935, 934t, 937, 940t, 942–944 Lexingtonia, 489f, 424 Leydigia, 694 Libellula, 899, 918 L. depressa, 907–908 L. luctuosa, 911 L. quadrimaculata, 911, 918, 922 Libellulidae, 858, 858f, 896–904, 897t–898t, 901f, 908, 909f, 910–912, 914, 915f, 917–922, 926, 928f Libelluloidea, 896–900, 897t–898t, 902 Liberonautes, 832, 833f L. latidactylus, 807–808, 810f, 822–823, 828f L. paludicolis, 829 Liberonautes latidactylus, 19f Ligula, 725–726 Ligumia, 424, 463–465, 489f L. nasuta, 484 L. recta, 457, 461f L. subrostrata, 452–453, 457, 458f, 461f, 475 Limmenius, 350t Limnadia lenticularis, 693f Limnadiidae, 688t Limnemertes, 207t L. poyangensis, 207t Limnephilidae, 965–969, 966t–967t, 968f, 971–981, 971f–973f, 980f, 982f, 985–987, 990–991, 993 Limnephilus, 971–972, 971f L. ademus, 992 L. affinis, 985 L. assimilis, 985–986 L. centralis, 985–986 L. externus, 988 L. lunatus, 985–986 Limnesia, 630f, 635, 639, 645–648 L. maculata, 641 Limnesiidae, 601t–602t, 620t–625t, 638 Limnesiinae, 620t–625t Limnias, 231–232, 244–245, 247t, 248, 265 L. melicerta, 245f, 248, 248f Limnichidae, 1004, 1005t–1007t, 1008–1009, 1011, 1013–1014, 1035–1036, 1036f Limnoasellus, 792–793 Limnobium, 62 Limnocalanus, 710–711, 719 L. macrurus, 719–720, 734 Limnocaridina, 803 L. iridinae, 824 Limnocentropodidae, 965, 966t–967t, 974–976, 975f Limnocentropus, 990 L. mergatus, 974–976, 975f Limnochares, 635–640, 643, 646, 653–654 L. americana, 631f, 637 L. aquatica, 637 Limnocharidae, 601t–602t, 620t–625t, 628–629, 635, 647f, 654 Limnocharinae, 620t–625t Limnocnida, 159–167, 164t, 169, 171–174, 176 L. indica nepalensis, 163t L. tanganicae, 163t Limnocythere, 773t Limnocythere, 758f, 770

Taxonomy Index

L. sanctipatricii, 766f L. staplini, 771–772 Limnocytheridae, 758–759, 758f, 759t, 767–768 Limnodriloidinae, 539–540 Limnodrilus, 533–534 L. hoffmeisteri, 518–519, 533–534, 541–545 L. silvani, 296f Limnohalacarus wackeri, 614 Limnoithona, 731–732 Limnomedusae, 159–161 Limnomysis, 94 Limnoperna, 87 L. fortunei, 86–87 Limnophora, 1043–1044, 1045t–1047t, 1054 Limnoposthia polonica, 186 Limnozetidae, 601t–602t, 611 Limonia, 1043–1044, 1045t–1047t Limoniidae, 375f Limpets, 400 Lindeniinae, 896 Linderiella occidentalis, 690f Lindiidae, 234–235, 234f Lineus longissimus, 205 Liocypridinae, 772 Lioplax, 393–394 L. cyclostomatiformis, 410t Lipostraca, 687 Lipsothrix, 1043–1044, 1045t–1047t, 1056 Lirceus, 789 L. fontinalis, 677, 785–786, 789, 791 Lissorhoptrus simplex, 1029 Listeria aquatica, 640–641, 646–647, 654 Lithasia, 410–411 L. obovata, 409–410 Lithoglyphidae, 390t Littorinidae, 395 Loa loa, 106 Lobopodia, 591–592 Longitarsus nigerrimus, 1028–1029 Lontra L. canadensis, 1035 L. provocax, 832 Lophogastrida, 13 Lophophore, 328 Lophopodella, 330f, 337, 340 L. capensis, 333f L. carteri, 15f, 84–85, 332–333, 336–338, 340, 340f, 342, 484 Lophopodidae, 332–334, 340 Lophopus crystallinus, 336, 341–342 Lophotrochozoa, 184 Loricifera, 304, 591–592 Loxoconchidae, 759t Loxodes rex, 127 Loxosomatoides sirindhornae, 340–341 Lubomirskia L. baicalensis, 140 L. baikalensis, 151 Lubomirskiidae, 135, 135f, 137–138, 147 Luciola, 1035 Lumbricidae, 519, 539–540 Lumbricoidea, 512–513 Lumbriculida, 533, 565 Lumbriculidae, 519, 529, 530f, 531, 533–535, 538–543, 552

1107

Lumbriculus, 540, 542, 545 L. variegatus, 530f, 534, 538f, 543, 545 Lurchibates, 642–643 Luridae, 192 Lurus, 192 Lutodrilidae, 510 Lutrochidae, 1004, 1005t–1007t, 1008, 1013–1014, 1016, 1036, 1036f Lutrochus, 1016, 1036 Lycastoides, 521–522 Lycosidae, 601t–602t, 602–604, 604f, 988, 1068 Lycosoidea, 601t–602t, 602–603 Lymnaea, 396–397, 401t L. elodes, 397–398, 401–403, 406–408, 407t L. emarginata, 399 L. obovata, 409–410 L. peregra, 398, 400, 404 L. stagnalis, 48–49, 391, 396–398 Lymnaeidae, 100, 385, 387, 387f, 390t, 391, 394t, 396, 400, 400t Lymnaeoidea, 390t Lynceidae, 688t Lynceus, 691–692 L. brachyurus, 691f–692f

M

Maccaffertium vicarium, 885 Machadorythidae, 874–876, 875t–876t Macrobdella decora, 568–570, 569f Macrobdellidae, 566–570, 568f, 577 Macrobiotidae, 350t, 352–355, 366, 369 Macrobiotus, 351–352, 352f, 354, 354f, 356f–357f, 357–360, 364–366, 377 M. echinogenitus, 371t–374t M. harmsworthi, 371t–374t M. hufeland, 369 M. hufelandi, 361, 368–369, 371t–374t M. joannae, 366, 368–369, 377 M. lazzaroi, 371t–374t M. martini, 371t–374t M. nelsonae, 371t–374t M. palarii, 371t–374t M. persimilis, 375 M. sapiens, 368–369, 375 M. tardigradum, 377 Macrobrachium, 93–94, 596, 747, 798, 800, 803–804, 821 M. carcinus, 681 M. nipponense, 93 M. rosenbergi, 93 M. rosenbergii, 817–819 M. vollenhoveni, 803 Macrocyclops, 723, 732–733 M. albidus, 723–724 Macrodasyida, 211–214, 217–218 Macrohectopidae, 792–793 Macrohectopus, 792–793 M. branickii, 792–793 Macromia, 896 M. illinoiensis, 925 Macromiidae, 896–901, 897t–898t, 900f–901f, 908, 909f, 920–921, 923–924, 924t Macronema, 976–978 M. hageni, 979–981, 982f

Macronematinae, 976, 979–981, 990 Macronychus, 1016 Macrophytes, 402f Macroplea, 1011, 1013 M. appendiculata, 1028–1029 M. mutica, 1028–1029, 1028f Macroseius biscutatus, 608 Macrostemum, 979–981 M. arcuatum, 979–981, 982f Macrostomida, 183f, 184, 186–187, 191–196 Macrostomidae, 183f, 197–198 Macrostomorpha, 196–197 Macrostomum, 183f, 184, 186, 191, 197–198, 468–469 M. lignano, 200 M. platensis, 189f–190f Macrothricidae, 688t, 701 Macrothrix rosea, 694f Macrotrachela quadricornifera, 239–241, 251 Macroveliidae, 951–953, 954t, 957 Macroversum, 350t, 352–353, 369 Madachauliodes, 1061 Maharashtracarinae, 620t–625t Majoidea, 802 Malacobdella, 206 Malaconothridae, 601t–602t, 611 Malacostraca, 12, 672–675, 672t, 783–785, 798 Malawispongiidae, 135, 135f, 137–138, 147 Malcostraca, 850 Mallomonas, 121 Manayunkia speciosa, 525–526 Manophylax, 992 Maphrodites, 696 Marellia remipes, 1070 Marenzelleria wireni, 525 Margaritifera, 424, 431, 450, 489f M. falcata, 454, 467–468, 471 M. margaritifera, 95, 95f, 455, 460, 467–469, 474–475 Margaritiferidae, 489f, 424–426, 428, 440–441, 444f, 458–460, 462 Marginellidae, 401 Marilia, 990–991 Marinellina, 214–216 M. flagellate, 211–212, 212f Marisa, 394 Marisa cornuarietis, 100–102 Marstonia, 394t, 395 Marthasterias glacialis, 208–209 Maruina, 1043–1044, 1045t–1047t, 1048f, 1049 Marvinmeyeria, 568f Mastigamoeba, 119 Mastigella, 119 Mastigodiaptomus, 727–728 Maxillopoda, 12, 18f, 672, 672t, 710–711, 717 Mayorella, 119 Mecistogaster, 902 Mecoptera, 1059, 1069 Medionidus, 424, 463–465, 489f M. mcglameriae, 429, 429t Megabalaninae, 712 Megadasys pacificus, 214 Megadytes ducalis, 1009 Megalagrion amaurodytum, 922

1108

Megalocypridinae, 759t, 772 Megalocypris princeps, 772 Megalonaias, 424, 461–462, 489f M. nervosa, 455, 467, 482–483 Megaloprepus caerulatus, 902, 917 Megaloptera, 14, 20f, 149, 851, 855, 988, 1059–1064, 1060f, 1062f Megaluracarus, 632f Megamelus scutellaris, 100–101 Meganeuropsis permiana, 894 Megapodagrionidae, 895–899, 897t–898t Megascolecidae, 512–513 Megascolecoidea, 512–513 Melampus, 392 Melanemerellidae, 874–877, 875t–876t Melanoides, 395, 403 M. tuberculata, 411 Melanopsidae, 390 Melosira, 341 Menidia beryllina, 98–99 Menzeliella, 727 Mermithidae, 273, 275, 278, 283f, 285–286, 294, 1057 Meropeidae, 1069 Merops apiaster, 923 Merostomata, 600 Meruidae, 1004, 1005t–1007t, 1009–1010, 1014, 1027, 1027f Meru phylissae, 1027 Mesamphisopus, 789 M. capensis, 790 Mesoasellus, 792–793 Mesobelostomum deperditum, 958–959 Mesocrista, 350t Mesocyclops, 262, 723, 725–729, 731–733 M. aspericornis, 723–724 M. edax, 720, 723 M. leuckarti, 724 M. longisetus, 723, 733 M. ogunnus, 731 M. pehpeiensis, 731 M. thermocyclopoides, 731–733 Mesophylax aspersus, 992 Mesostigmata, 601t–602t, 607–609, 608f–609f Mesostoma, 183f M. ehrenbergii, 189f–190f, 197–198, 200 M. lingua, 197–198, 200 Mesostomidae, 183f, 195 Mesostominae, 188, 195, 200 Mesotardigrada, 350t Mesothelae, 601 Mesoveliidae, 951–953, 954t, 957 Metacyclops, 727, 733 M. gracilis, 724 M. mendocinus, 734 M. minutus, 719, 724 Metacypris, 758f Metania reticulata, 138f Metaniidae, 135f, 137, 147 Metapterygota, 874 Metazoa, 128–129, 187–188 Metopus, 125 Metretopodidae, 874–876, 875t–876t, 879 Metrichia, 320–321

Taxonomy Index

Metriocnemus, 1054 Metschnikowia, 725–726 Metschnikowiidae, 135, 135f, 137–138, 147 Micracanthia, 951–953, 952f Micractinium, 260 Microcerberus, 790 Microchorista philpotti, 1069 Microcyclops, 713f, 723 Microdalyellia tennesseensis, 199 Microdarwinula, 769 Microdiaptominae, 726 Microdiaptomus, 726 M. cokeri, 726, 728–729 Micrognathozoa, 212, 230 Microhypsibiidae, 350t, 366 Microhypsibius, 350t, 352f, 353, 369 M. bertolanii, 371t–374t M. minimus, 371t–374t Micromelaniidae, 394t Micronecta scholtzi, 958 Micropterus M. dolomieu, 832, 833f M. salmoides, 460, 463f, 480 Microsporidae, 1019 Microsporidia, 338 Microsporidium, 293f Microsporidium (Plistophora), 264 Microsporus, 1019 Microstomum, 197–198, 291 M. lineare, 192–193 Microtrombidium, 616f Microvelia, 960 M. reticulata, 868 Micruropodidae, 792–793 Mictrotrichiae, 608f Midea, 644 M. expansa, 636 Mideidae, 601t–602t, 620t–625t, 638 Mideopsellinae, 620t–625t Mideopsidae, 601t–602t, 620t–625t Mideopsis, 632, 645–646 Millipedes, 661–667, 667f Milnesiidae, 348, 350t, 352, 354–355 Milnesioides, 350t M. exsertum, 364 Milnesium, 351, 352f, 354–355, 369–370, 371t–374t, 377 M. tardigradum, 364, 369, 375 Minibiotus, 350t, 354, 357–359 M. pustulatus, 371t–374t Minilentus, 350t, 354 Miracidia, 773–774 Miraciidae, 727 Miridae, 100–101 Mischoderus, 1043–1044, 1045t–1047t Misophrioida, 710, 728–729 Misthodotidae, 873 Misumenops nepenthicola, 606 Mixibius, 350t, 369 M. saracenus, 371t–374t Mixolimnesiinae, 620t–625t Mniobia, 231–232 Mochlonyx, 1043–1044, 1045t–1047t Moina, 694, 697

Moinidae, 688t Molanna, 991–992 Molannidae, 965, 966t–967t, 985, 990 Mollusca, 11, 383–388, 384f Momoniidae, 601t–602t, 620t–625t, 638 Monhystera, 286, 288, 292 Monhysteridae, 273, 295 Monhystrella plectoides, 295 Moniligastridae, 533 Monoculus foliaceus, 711 Monocystis, 125 Monogenea, 181 Monogononta, 227, 231, 237, 249 Mononchida, 273, 291 Mononchus, 286 Mononema, 725–726 Monopisthocotylea, 181 Monoporeia, 791 Monosacanthes, 725–726 Monosiga, 120–121, 121f Monospilus, 694 Monostilifera, 205–206, 207t Monotoplanidae, 191–192 Monstrilloida, 710 Mopsechiniscus, 364f, 367 Morimotacarinae, 620t–625t Mormonilloida, 710 Motobdella, 570, 572, 581 M. montezuma, 566, 578–579, 581 Mucronothrus, 611 M. nasalis, 611 Multipeniatidae, 191–192, 195 Multitubulatina, 213–214 Murphyella, 879–880 Murrayidae, 350t, 352–354, 366 Murrayon, 350t, 352f, 354, 357–359, 366, 369 M. dianeae, 371t–374t M. hastatus, 371t–374t M. pullari, 371t–374t Muscidae, 1043–1044, 1045t–1047t Muscocyclops, 730 Musculium, 432–433, 439, 441, 449–450, 459, 466, 474 Mutelocloeon, 886 Mya arenaria L., 747 Mycetopodidae, 457–458 Mycetozoa, 120 Mycobacterium ulcerans, 960 Mycobatidae, 601t–602t Mygalomorphae, 601 Mylopharyngodon pisceus, 469 Myodocopa, 759–760 Myoretronectes paranaensis, 186, 197 Myriapoda, 12, 592, 661–662, 666–667, 850 Myriophyllum, 247, 265–266 Myrmecodesmus M. adisi, 665 M. duodecimlobatus, 665 Myrmeleontiformia, 1064 Mysida, 672, 672t, 678 Mysidacea, 782–785, 789–793 Mysis, 94, 789, 793 M. diluviana, 88, 103, 782f, 789–790, 792–793 M. relicta, 88, 103, 787–788

Taxonomy Index

Mystacides M. azurea, 987–988 M. sepulchralis, 979–981, 982f Mystacocarida, 710–711 Mytilocypridinae, 759t, 772 Mytilocypris henricae, 774 Mytilus, 747 M. edulis, 450–451, 747 Myxobolus cerebralis, 125–126 Myxophaga, 1004, 1005t–1007t, 1008, 1014, 1018–1020 Myxozoan, 338 Myzobdella, 567

N

Naegleria, 118–119, 122 Naiadacarus arboricola, 613 Naididae, 529–535, 530f, 537–540, 542–545 Nais N. barbata, 543 N. elinguis, 544 Najadicola ingens, 631f, 647–648 Najadicolinae, 620t–625t Namalycastis, 521–522 Namanereidae, 525 Namanereidinae, 521–522 Namanereinae, 519–520 Namanereis, 520–522 N. araps, 521–522 N. catarractarum, 521–522 N. cavernicola, 521–522 N. quadraticeps, 525 Nannochloris oculata, 261 Nannochloropsis oculata, 254 Nannochorista, 1069 Nannochoristidae, 1069 Nannophya pygmaea, 911 Nannopus palustris, 719 Nanocladius, 940, 1056 Nanogona polydesmoides, 663f Narapa bonettoi, 535 Narapidae, 533 Nassula tumida, 117f Nasturtium officinale, 867 Natantia, 798–800 Naucoridae, 864, 951–957, 954t Nauplii, 696, 722–725, 745f, 748 Naupliicola, 725–726 Nebela, 118–119 Necopinatidae, 350t, 366 Nectonema, 304 N. munidae, 306 Nectonematida, 304 Nectopsyche, 974–976, 975f, 979–981, 981f, 993 N. aureofasciata, 979–981, 981f N. gemmoides, 971–972, 971f, 991 N. punctata, 979–981, 981f Necturus maculosus, 469 Nehalennia speciosa, 646 Nelumbo lutea, 337 Nemata, 11, 273 Nematocera, 638, 1043–1044 Nematoda, 212, 273, 375f, 591–592 Nematoidea, 273

1109

Nematomorpha, 212, 303–305, 308, 316, 319–322, 591–592, 864 Nematoplana nigrocapitula, 199 Nemertea, 11, 205 Nemertodermatida, 188–189, 191–196 Nemertoplanidae, 192 Nemoura arctica, 939 Nemouridae, 934–935, 934t, 937, 940t, 942–944 Nemurella pictetii, 937–938 Neoacaridae, 601t–602t, 620t–625t, 633f Neoatractidinae, 620t–625t Neocaridina, 553, 559, 821–822 Neochauliodes, 1061–1062 Neochetina N. bruchi, 100–101 N. eichhorniae, 100–101 Neochordodes, 308f, 309–310, 311f, 318, 320–321 N. occidentalis, 304, 308f, 311–312, 313f–315f, 314–315 Neodasyidae, 211–212, 216 Neodermata, 181 Neoephemeridae, 874–876, 875t–876t Neogossea antennigera, 215f Neogosseidae, 211–212, 213f, 214–216, 218, 219t Neohelice granulatus, 676–677 Neohermes, 1061–1063, 1062f N. californicus, 1063 N. filicornis, 1060, 1060f, 1063 Neohydronomus affinis, 1029 Neolimnochares, 637 Neomamersa, 632 Neomamersinae, 620t–625t Neomysis, 94, 678f Neoneuromus, 1061 Neoophora, 182 Neoperla N. sabang, 936, 939f N. stewarti, 940 Neopetalia, 896 Neopetaliidae, 896–899, 897t–898t Neophylax, 76, 979–981, 982f, 987 N. nacatus, 987 N. ornatus, 987 N. rickeri, 992–993 Neoplea, 853, 854f Neoptera, 874, 933–934 Neoradina, 395 Neosarmatium, 820 Neoshoenobia, 1068 Neotelmatoscopus, 1043–1044, 1045t–1047t Neotenotrocha, 514 Neothricidae, 688t Neotorrenticolinae, 620t–625t Neotrichia, 989 Neotrichiinae, 989 Neotrigonia, 425–426 Nepenthaceae, 1054 Nepenthes, 606, 1054 Nephelopsis obscura, 407 Nephridia, 387 Nephropoidea, 800–801 Nepidae, 951–959, 952f, 954t, 959f

Nepomorpha, 951–955, 952f, 954t, 958–959 Nereidae, 519–520 Nereididae, 519–522, 520f, 524–525 Neritidae, 388, 390t, 394, 394t, 395f, 400t, 402 Neritimorpha, 390t Neritina reclivata, 394 Neritoidea, 390t Neritomorpha, 394t Nerthra, 951–953, 952f Nesameletidae, 874–879, 875t–876t Neumania, 647–648 N. papillator, 643–644, 649 Neurhermes, 1061 Neurogomphus, 908, 909f Neuroptera, 13, 148–149, 851, 855, 863, 1059–1060, 1064–1067, 1064f–1065f Neuropterida, 1060, 1064 Nevromus, 1061 Nevrorthidae, 1064–1067 Nevrorthiformia, 1064 Nevrorthus, 1064, 1066–1067 N. fallax, 1065–1066 Nicalimnesiinae, 620t–625t Nigronia, 1061–1062 Nilaparvata, 960 Nilotoniinae, 620t–625t Nilus, 604 N. curtus, 605 Niphargus, 789–790 N. rhenorhodanensis, 789–790 N. virei, 790 Niphograpta albiguttalis, 100–101, 1068–1069 Niponosialis, 1061 Nipponacaridae, 601t–602t, 620t–625t Nipponeurorthus, 1064, 1066–1067 Nipponocyphoninae, 1037–1038 Nitidulidae, 1004, 1013–1014, 1016, 1036, 1036f Nocticanace, 1043–1044, 1045t–1047t Noctiluca, 122 Nostoc, 296f Noteochorododes N. cymatium, 317 N. talensis, 317 Noteridae, 1004, 1005t–1007t, 1008–1011, 1013–1014, 1016, 1022–1024, 1026–1028, 1027f–1028f Noterus, 1027–1028 Nothochauliodes, 1061 Notholca, 229–230, 237–238 N. acuminata, 242, 256f Nothopsyche ruficollis, 987–988 Nothrina, 601t–602t Notidobiella brasiliana, 979–982, 983f Notoaeschna sagittata, 919 Notoaturinae, 620t–625t Notodromadidae, 758–759, 758f, 759t Notodromadinae, 759t Notodromas monacha, 767–768 Notogomphus, 908, 909f Notommata, 234–235, 234f, 250 N. copeus, 231–232, 258–259 Notomudamellinae, 620t–625t Notonecta, 953–955, 955f

1110

Notonectidae, 605, 646, 853, 859, 864, 864f, 951–957, 954t Notonemouridae, 934–935, 934t, 945 Notosolenus, 121 Notostraca, 687–692, 688t, 695–698 Nudomideopsidae, 601t–602t, 620t–625t, 633f Nudomideopsis, 644 Nuphar, 337 Nymphaea, 247, 337 Nymphicula, 1068 Nymphochrysalis, 642 Nymphomyia, 1043–1044, 1045t–1047t Nymphomyiidae, 1043–1044, 1045t–1047t, 1050, 1053–1054 Nymphulinae, 1068

O

Obliquaria, 424, 489f O. reflexa, 457, 461f Obovaria, 489f, 424 Ochlerotatus sierrensis, 49 Ochridacyclops, 721 Ochrotrichia, 992 Ochteridae, 951–953, 954t Ochthebius, 1014 O. caucasius, 1033 O. corrugatus, 1014 Ocnerodrilidae, 512–513 Octochaetidae, 512–513 Octomyomermis troglodytis, 288–289 Odonata, 13, 20f, 149, 637–638, 639f, 851, 853, 855, 869, 874, 886, 894, 897–899, 899f, 902, 904–906, 908, 909f, 910–914, 913f–914f, 916f, 917–919, 922–923, 925–928, 988, 1004 Odonatoptera, 894 Odontoceridae, 965–969, 966t–967t, 968f, 979–981, 986–987, 990 Odontolinus, 1038–1039 Odontomyia, 1043–1044, 1045t–1047t, 1049, 1049f Odontostomes, 123f Oecetis, 986–987, 990, 992 Oeconesidae, 965, 966t–967t Oedoparena, 1043–1044, 1045t–1047t, 1056 Oestridae, 956 Oligochaeta, 12, 375f, 509–511, 514–517, 515f, 521, 523, 529, 530f, 531, 532f, 533–537, 536f, 539–540, 539f, 543–545, 565, 578, 917–918 Oligochaeta, 535 Oligochaete, 553, 556, 556f Oligochoerus, 186 O. limnophilus, 186 Oligoneuriidae, 874–876, 875t–876t, 878–879, 883–885, 887–888 Oligotrichs, 123f Olindiidae, 159–161, 163t Olinga feredayi, 992 Omaliinae, 1038–1039 Omaniidae, 951–953, 954t Omartacaridae, 601t–602t, 620t–625t Omartacarinae, 620t–625t Ombrastacoides, 801–802

Taxonomy Index

Onchidiidae, 391 Onchocerca volvulus, 106, 107f, 834 Oncholaiidae, 278 Oncomelania, 403 Oncorhynchus O. mykiss, 319–320, 741, 832 O. nerka, 88, 103 Oncosclera, 137 Oniscigastridae, 874–877, 875t–876t Onychodiaptomus sanguineus, 725 Onychogomphus, 908, 909f O. forcipatus, 909 O. uncatus, 911, 912f, 914–915, 916f Onychophora, 304, 591–592 Onychopoda, 687, 698–699 Oodes helopioides, 1022 Ophiogomphus cecilia, 921 Ophryoxidae, 688t Opilioacarida, 601, 607 Opiliones, 600 Opisthoplatia, 1067–1068 O. orientalis, 1067–1068 Opisthopora, 533 Opistocystidae, 529–535, 540, 542 Orconectes, 470, 553, 557–558, 805, 815–816, 822, 824–828, 828f, 835–836 O. australis, 813–814, 814f, 823f, 828–829 O. compressus, 828 O. eupunctus, 835–836 O. hylas, 835–836 O. immunis, 813, 824 O. inermis, 42–43, 815, 822–823, 828 O. inermis testii, 43f O. limosus, 557–558, 837 O. luteus, 835 O. nais, 813 O. neglectus chaenodactylus, 835–836 O. palmeri longimanus, 817–819, 818f O. pellucidus, 815 O. peruncus, 835–836 O. propinquis, 173 O. propinquus, 817–819, 830, 836–837, 836f O. punctimanus, 835 O. quadruncus, 835–836 O. rusticus, 100, 102, 173, 407, 407t, 824, 825t, 826f, 832, 833f, 835–836, 836f, 838 O. sheltae, 828 O. virilis, 681, 815–817, 816f, 824, 825t, 836–837, 836f Oreella, 350t, 357–359 Oreellidae, 350t, 351, 356–357 Oreogeton, 1043–1044, 1045t–1047t Oreoleptidae, 1043–1044, 1045t–1047t, 1050, 1054 Oreoleptis torrenticola, 1043–1044, 1045t–1047t Oribatida, 601t–602t, 609–611 Oripodoidea, 601t–602t Ornamentula, 218 O. paraënsis, 215f Orohermes, 1061–1062 O. crepusculus, 1060, 1060f Orthetrum, 908, 909f, 921 O. cancellatum, 902–903, 903f, 922 O. coerulescens, 923, 923f

Orthocladius, 1043–1044, 1045t–1047t Orthogalumna terebrantia, 100–101 Orthoptera, 13, 851, 855, 1059, 1069–1070 Orthorrhapha, 1043–1044 Orthotrichia, 988 Osmylidae, 1064–1067 Osmylinae, 1064, 1066 Osmylus, 1064, 1066–1067 O. fulvicephalus, 1066, 1066f Osphranticum, 710–711 Ostracoda, 375f, 672–673, 672f, 672t, 678–680, 711 Ostracods, 41, 757–777, 758f, 759t, 762f, 764f, 771f, 773t, 775f Otomesostoma O. arovi, 186 O. auditivum, 186 Otomesostomidae, 191–192 Otonemertes, 207t O. denisi, 207t Otoplana rauli, 197 Otoplanidae, 186 Ototyphlonemertes, 205 Oukuriella, 149 Oxidae, 601t–602t, 620t–625t Oxidus gracilis, 664–665, 665f Oxus, 638 Oxycera, 1043–1044, 1045t–1047t Oxyinae, 1070 Oxytricha, 122–123 Ozobranchidae, 511, 566

P

Pachychilidae, 391–392, 392f Pachylasmatoidea, 712 Pachyschesidae, 792 Pachyschesis, 681 Pacifastacus, 801, 805, 824–826, 825t P. fortis, 101 P. leniusculus, 94–95, 101–102, 291, 552f, 553, 813, 817–819, 818f, 835, 837 P. leniusculus trowbridgii, 822 Paguroidea, 807 Palaeagepetus, 976–978 Palaemon, 800, 803 P. concinnus, 803 Palaemonetes, 800, 803–804, 821 P. alabamae, 807, 811f, 821, 824, 830 P. antrorum, 821 P. cummingi, 821 P. ganteri, 808, 808f, 830 P. kadakiensis, 803 P. kadiakensis, 814–815, 821 P. paludosus, 819, 819f, 821, 824, 830 Palaemonias, 824 P. ganteri, 821, 824 Palaemonidae, 798, 799t, 800, 803, 824, 825t Palaemoninae, 800, 803 Palaemon paludosus, 19f Palaeoheterodonta, 425–426 Palaeohirudo eichstaettensis, 513 Palaeonema, 274 P. phyticus, 274f Palaeonemertea, 205–207, 206t

Taxonomy Index

Palaeospongilla chubutensis, 134, 147 Paleochordodes protus, 304–305, 305f Paleodictyopterida, 851 Paleoptera, 874 Palingenia, 882–884 P. longicauda, 887–888 Palingeniidae, 874–876, 875t–876t, 879, 882–884 Palpata, 521 Palpigradi, 600 Palpomyia, 1044, 1048f, 1049 Paludicella, 331–332, 335–336 P. articulata, 334f Panarthropoda, 350, 360 Pancrustacea, 592, 671, 759–760, 850 Pandirodesmus disparipes, 665–666 Pangaea, 800–801 Pannota, 874–876, 879–880 Pantala, 908, 909f P. flavescens, 917–918, 920–922, 924 Parachauliodes, 1061, 1063 Parachela, 350t, 352–355, 366 Parachironomus longiforceps, 338–339 Parachordodes, 308f Paracles, 1069 Paracrobiotus richtersi, 357f Paracrostoma, 395 Paracyclops, 721, 730 P. fimbriatus, 724 Paradiaptominae, 726–728 Paradiphascon, 350t Paradoxosomatidae, 665f, 667 Paragnetina media, 939 Paragomphus, 921, 908, 909f Paragonimus, 833–834 Paragordius, 304, 308f, 311–312, 315, 315f, 317–319, 321 P. obamai, 308, 315–317, 320–321 P. tricuspidatus, 316–318, 320f P. varius, 306f–307f, 307–309, 310f, 313, 313f–314f, 315–317 P. varius, 314 Parajapyx, 850–851, 850f Paraleptophlebia, 885 P. submarginata, 879–880, 880f Paralichas, 1037 Paralimnetis, 691–692 Paramacrobiotus, 350t, 357–359, 369 P. areolatus, 361 P. halei, 371t–374t P. megalonyx, 364–366 P. richtersi, 348f, 360f, 362–364, 364f, 368– 369, 368f, 371t–374t, 375, 377 P. tonollii, 368–369, 371t–374t Paramecium, 114–115, 122–123, 769, 773–774 Paramelita, 788–789 P. nigroculus, 785–786 Paramideopsis susanae, 633f Paramphisopus palustris, 790 Paramysis, 793 Paranadrilus, 541 Paranephrops, 801–802, 805, 824, 825t Paranophrys, 125 Paraphanolaimus, 292

1111

Paraphrynoveliidae, 951–953, 954t Paraphysomonas, 121 Parapsyche cardis, 972–974, 973f Pararotatoria, 227, 234–235, 237, 249 Parartemia, 596, 695–696 Parartemiidae, 688t Parascon, 350t, 355, 356f Paraseison, 234–235 Parasimulium, 1053 Parasitengona, 617f Parasitengones, 615–617 Parasitengonina, 601t–602t, 615–649, 616f, 618f Parasitiformes, 600–601, 601t–602t, 607–609 Parastacidae, 798–802, 799t, 801f, 805, 824, 825t, 826 Parastacoidea, 798–801, 801f Parastacus, 801–802, 805, 824, 825t, 826 P. defossus, 826 P. nicoleti, 826 P. pugnax, 826 P. zonangulus, 824, 838 Parastenocarididae, 711, 720, 727 Parastenocaris, 711 Parathelphusa, 829–830 Paratrechalea, 606–607 P. azul, 606–607 Paratya, 803, 824 Pardosa, 603–604, 1068 P. messingerae, 604 Parhexapodibius, 350t P. kathmanae, 370 P. pilatoi, 366 Parisia, 803, 824 Parvicapsula minibicornis, 526 Parvicorbicula, 120–121 Parvidrilidae, 531–536 Parydra, 1043–1044, 1045t–1047t Paucitubulatina, 213–214 Paulinia acuminata, 1070 Pauropoda, 661 Pectinatella magnifica, 330, 330f, 333, 335–339 Pectinatellidae, 339 Pectispongilla, 141f Pediastrum duplex, 341 Pedomoecus sierra, 965–969, 968f Pedunculata, 711–712 Pegias, 424, 489f Pelecorhynchidae, 1043–1044, 1045t–1047t Pelecypoda, 449–450 Pellioditis, 277f, 280f Pelomyxa, 119 Peltodytes, 1025–1026 Peltoperlidae, 934–935, 934t, 940t, 941–944 Penelia, 694 Peneoidea, 798 Peniculids, 123f Pentaphlebiidae, 895–899, 897t–898t Pentastomida, 710–711 Penthesilenula brasiliensis, 760, 769 Peracarida, 13, 672, 680, 781–788, 791–792 Perca fluviatilis, 923–924 Percidae, 465

Percina caprodes, 430f, 465 Percolomonas, 122 Pereiopods, 673–675, 784 Perenethis fascigera, 606–607 Pericoma, 1043–1044, 1045t–1047t Peridiniopsis berolinensis, 122 Peridinium, 122 Perilestidae, 894, 897–899, 897t–898t Periphyton, 399, 402f Peritricha, 518 Perlidae, 934–935, 934t, 937, 937f, 939, 940t, 941–942, 944, 988 Perlodidae, 934–935, 934t, 937, 940t, 942–944 Permoplectoptera, 873 Petalomonas, 121 Petalura, 902 Petaluridae, 896–899, 897t–898t, 902–903, 922 Petaluroidea, 896–899, 897t–898t Petrobiona, 147 Petrolisthes, 807–808 P. armatus, 807 Petronica macrocephala, 320 Petrothrincidae, 965, 966t–967t Pezidae, 601t–602t, 613–614 Phaenocora, 199 P. typhlops, 199 P. unipunctata, 199 Phaenocorinae, 193 Phagocata P. vitta, 186 P. woodworthi, 85–86 Phagodrilus, 537, 543 Phallobaikalus gladiiseta, 534 Phallocryptus, 596, 695–696 Phallodrilinae, 534, 539–540, 543 Pharyngostomoides procyonis, 559 Phenes raptor, 922 Pheromermis, 294 P. pachysoma, 17f, 283f, 284–285 Philocasca alba, 986–987, 991 Philodina, 230, 264 P. citrina, 252 P. roseola, 241 Philogangidae, 895–899, 897t–898t Philogeniidae, 896–899, 897t–898t Philometra, 725–726 Philopotamidae, 965, 966t–967t, 971–976, 971f, 975f, 978–981, 981f, 986, 988–991 Philorheithridae, 965, 966t–967t, 979–982, 983f Philosinidae, 897–899, 897t–898t Philosyrtis rauli, 184–185 Phlebonotus, 1067–1068 P. pallens, 1067–1068 Phlebotomidae, 58 Phoreticovelia, 958 Phoronida, 328 Phreatodyte, 1027–1028 Phreodrilidae, 531–535, 538–539, 542–543 Phreodrilus branchiatus, 535 Phricothelphusa, 829 Phryganea, 992 Phryganeidae, 965–969, 966t–967t, 968f, 971–976, 973f, 978–981, 980f, 990, 993 Phryganopsychidae, 965, 966t–967t

1112

Phyganeidae, 978–979 Phylactolaemata, 327–331, 329f, 335–336, 335f, 338, 340, 342 Phyllobranchiates, 675f Phyllodocida, 519–521 Phyllognathopus viguieri, 723, 730 Phylloicus, 991–992 P. abdominalis, 979–981, 981f P. aeneus, 965–969, 968f, 974–976, 975f P. bromeliarum, 992 P. mexicanus, 979–981, 982f Phyllomacromia, 896, 908, 909f Phylocentropus, 974–976, 975f, 991–992 Physa, 321, 396–397, 398f, 401, 406–408 P. acuta, 396, 403, 405–409 P. ancillaria, 405 P. gyrina, 396, 400–401, 406 P. heterostropha, 396 P. integra, 396, 399 P. natricina, 410t P. vernalis, 400 P. virgata, 396, 407 Physarum, 118f, 120 Physella, 83–84, 387f, 401t, 830 P. heterostropha, 398–399 P. vernalis, 387f, 405 P. virgata, 791 Physidae, 100, 385, 387, 390t, 391, 394t, 396, 400, 400t, 406 Physocypria, 773, 773t Phytophthora infestans, 125 Phytotelmata, 602–603, 1054 Piersigia, 637, 646–648 Piersigiidae, 601t–602t, 620t–625t Piguetiella blanci, 542 Pila, 394, 400 P. luzonica, 101 Pilidae, 394 Piona, 632, 635–636, 639, 642–649, 654 P. limnetica, 648 P. napio, 641 Pionatacinae, 620t–625t Pionidae, 601t–602t, 620t–625t, 631f, 642–644 Pioninae, 620t–625t Pirata, 603–604, 604f P. piraticus, 603–604 Pirata piscatorius, 603–604 Pisaura P. lama, 606–607 P. mirabilis, 605 Pisauradae, 1068 Pisauridae, 593f, 601t–602t, 602–607, 606f Piscicola P. geometra, 569f, 579 P. milnera, 569f P. punctata, 569f Piscicolidae, 511–512, 516, 566–572, 567f–568f, 574–575, 577 Pisciforma, 874–876 Pisidiinae, 432 Pisidium, 432–433, 438–439, 441, 449, 459, 466, 474, 483–484, 488f–489f P. amnicum, 433, 434t P. casertanum, 431–432, 474

Taxonomy Index

P. henslowanum, 433, 434t P. moitessierianum, 433, 434t P. punctiferum, 483 P. supinum, 433, 434t Pistia, 62, 1068–1069 P. stratiotes, 100–101, 614 Pisuliidae, 965, 966t–967t Placobdella, 568f P. costata, 579 P. hollensis, 569f, 572 P. ornata, 572 P. papillifera, 576, 578, 583 P. parasitica, 572 Plagiopyla, 124f, 125 Plagiostomidae, 191–192, 195 Plagiostomum evelinae, 196 Planolineus, 207t P. exsul, 207t Planorbella, 406–407 P. trivolvis, 400, 406 Planorbidae, 100, 386f, 387, 390t, 391, 394t, 396, 400, 400t Planorbis, 100 P. contortus, 405 P. planorbis, 404 P. vortex, 400 Planorboidea, 390t Plasmodiophora brassicae, 125 Plasmodium, 125 Platicrista, 350t Plationus patulus, 236f Platychauliodes, 1061 Platychirograpsus spectabilis, 820 Platycnemididae, 894–895, 897–899, 897t–898t, 926, 928f Platycnemis pennipes, 919, 920f, 923–924 Platycopioida, 710, 728–729 Platyhelminthes, 181–182, 182f, 184–189, 191–199, 197f–198f Platyias quadricornis, 237f, 251 Platyneuromus, 1061 Platynothrus peltifer, 611 Platypsyllinae, 1035 Platypsyllus castoris, 1035 Platyseius, 608 P. italicus, 608 Platystictidae, 894, 897–899, 897t–898t Platystictoidea, 894, 897–899, 897t–898t Platythelphusa, 829–830 P. armata, 830–831 Platyzoa, 212 Plaumanniinae, 620t–625t Plecoptera, 14, 20f, 149, 375f, 638, 851, 853, 855, 859, 863, 933–934, 934t, 944, 953, 988, 995–996, 1004, 1016 Plecopterocoluthus, 940 Plectidae, 273, 275, 275f Plectomerus, 424, 489f Plectrotarsidae, 965, 966t–967t Plectus, 275, 292 Pleidae, 853, 854f, 951–956, 954t Pleistoannelida, 531 Pleistocene, 395 Plenitentoria, 965, 966t–967t, 969

Pleocyemata, 672t, 798–800 Pleopods, 678, 784 Plesiogammarus, 788–789 P. brevis, 788–789 P. mussauensis, 782–783 P. zienkowiczii, 788–789 Plethobasus, 424, 461–462, 489f Pleureta lineata, 234f Pleurobema, 424, 462–463, 489f P. altum, 429, 429t P. avellanum, 429, 429t P. bournianum, 429, 429t P. chattanoogaense, 429, 429t P. flavidulum, 429, 429t P. hagleri, 429, 429t P. hanleyianum, 429, 429t P. johannis, 429, 429t P. murrayense, 429, 429t P. nucleopsis, 429, 429t P. rubellum, 429, 429t P. sintoxia, 475 P. troschelianum, 429, 429t P. verum, 429, 429t Pleurobemini, 489f, 424, 462 Pleurocera, 410–411 P. canaliculum, 409–410 P. unciale, 478 Pleuroceridae, 385, 386f, 389f–390f, 390, 390t, 392–393, 394t, 398–400, 400t, 408–411, 409f Pleuronema, 122–123 Pleurotrocha, 265 Pleuroxus, 694 Plumatella, 173–174, 329f, 333, 335, 337 P. casmiana, 329–330, 334f, 337–338 P. emarginata, 333f, 334–338 P. fruticosa, 331–332, 336 P. fungosa, 331, 336–337 P. mukaii, 334–335 P. nitens, 336–337 P. recluse, 336 P. repens, 331, 336–339, 342 P. reticulata, 332f P. rugosa, 333f P. vaihiriae, 336–340, 339f Plumatellidae, 335, 338 Pneumocystis, 125 P. carinii, 125 P. jiroveci, 125 Podiceps nigricollis, 48 Podocopida, 757–760, 759t, 764f, 766t, 767–769 Podocytes, 717–718 Podon, 694 Podonidae, 688t Podonominae, 1054–1056 Podophrya, 122–123, 124f Podoplea, 710 Podostemaceae, 1019–1020 Podura, 604 P. aquatica, 865–868, 866f Poduridae, 865, 865f Poeciloderrhis, 1067–1068 P. cribosa, 1067–1068

Taxonomy Index

Poecilostomatoida, 710 Poecilotheria ornata, 606–607 Pogonophora, 520–521 Pollicipalpinae, 620t–625t Polyartemia, 690 Polyartemiella, 690 Polyarthra, 234–235, 238, 242, 256f, 258–259, 263–264, 263f, 725 P. dolichoptera, 253, 256f P. vulgaris, 257, 260 Polycelis nigra, 186 Polycentropodidae, 965, 966t–967t, 974–981, 986, 990 Polycentropus, 990 Polychaeta, 12, 509–511, 519–521, 525–526, 565–566 Polycladida, 182, 188–197 Polycystididae, 183f Polydesmida, 664–667, 665f, 667f Polydesmidae, 665f, 667f Polydesmus angustus, 665f Polykrikos, 122 Polymerurus, 218 P. nodicaudus, 215f Polymitarcyidae, 874–878, 875t–876t, 882, 885, 887–888, 888f Polyodontidae, 170–171 Polyopisthocotylea, 181 Polyphaga, 1004, 1005t–1007t, 1008–1012, 1014–1016, 1028–1039 Polyphemidae, 688t Polyphemus, 694, 698–699 Polyplaston, 125 Polyplectropus, 976–978 Polypodiidae, 163t Polypodium, 167, 174 P. hydriforme, 161–162, 163t–164t, 166, 166f, 170–171, 171f, 173 Polysphondylium, 118f Polystilifera, 205–206 Polythoridae, 895, 897–899, 897t–898t, 909 Polyzoniida, 664–665 Pomacea, 26–27, 83–84, 100–101, 394, 400 P. canaliculata, 100–102, 394, 411–412, 412f P. insularum, 101–102 P. paludosa, 411–412 Pomatiopsidae, 390, 390t, 394t, 395, 398 Pompilidae, 1068 Pontarachnidae, 601t–602t, 620t–625t Pontastacus, 805 Pontogammaridae, 793 Pontomyia, 1056 Popenaias, 461–462 P. popeii, 455 Populus, 62 Porcellanidae, 798, 799t, 807 Porifera, 11, 41, 133, 139–143, 143f, 147 Portunidae, 798, 799t, 806–807 Potamalpheops, 804 Potamanthidae, 874–876, 875t–876t Potamarcha congener, 921 Potamidae, 798, 799t, 802, 805–806, 824, 825t Potamilus, 489f, 424, 460 P. alatus, 455, 459f, 460

1113

Potamocoridae, 951–953, 954t Potamocypris, 680, 758f, 772, 773t P. smaragdina, 769 P. unicaudata, 774, 777 Potamodrilidae, 519–526, 529 Potamodrilus, 521, 523–524 P. fluviatilis, 525 Potamogeton, 579, 838 Potamoidea, 802, 805–806 Potamolepidae, 135f, 137, 147–148 Potamonautes, 829 P. raybouldi, 838–839 Potamonautidae, 798, 799t, 802, 805–806, 824, 825t Potamonemertes, 207t P. gibsoni, 207t P. percivali, 207t Potamopyrgus, 400, 403 P. antipodarum, 83–84, 101–102, 398, 411–412, 412f P. jenkinsi, 400 Potamothrix, 533–535 P. bedoti, 541–542 P. hammoniensis, 540f, 541, 543–544 Pottiaceae, 375 Pottsiella, 335–336 Povilla, 887–888 P. adusta, 882, 887 Priapulida, 212, 591–592 Pristina, 514–515, 543 Pristinidae, 529–535, 540, 542, 544–545 Pristolycus, 1035 Proales, 234f, 265 P. gigantea, 234f P. sordida, 252 Proasellus, 793 P. coxalis, 791 Probopyrus pandalicola, 681 Procamallanus, 725–726 Procambarus, 804–805, 822, 825–828 P. acutus, 817–819, 822, 826–827, 828f P. alleni, 815–816, 815f, 822, 827f, 831, 831f P. clarkia, 822–823 P. clarkii, 9–10, 27, 84–85, 94, 100–102, 553, 557–558, 673f, 804–805, 812–815, 813f–814f, 824, 830–831, 835, 837–838 P. enoplosternum, 813–814, 813f P. fallax virginalis, 822 P. hagenianus, 823 P. lunzi, 807, 811f, 813–814, 813f P. paeninsulanus, 817–819 P. pygmaeus, 817–819, 818f, 826–827, 838 P. simulans, 813 P. spiculifer, 817–819, 818f P. tenuis, 828 P. versutus, 812f, 813 P. zonangulus, 822–823 Prodesmodora, 292 Progne subis, 1057 Progymnoplea, 710 Proichthydidae, 213f Proichthydioides, 218 Proichthydium, 218 Prolecithophora, 182, 184–186, 191–197

Prolecithoplana lutheri, 194 Proleptus, 294 Promenetus, 401t Propappidae, 533–534 Prorhynchidae, 183f Prorhynchus stagnalis, 183f, 192–193, 197–199 Prorodon, 122–123 Proschizorhynchus triductibus, 199 Proseriata, 184–186, 188, 191–193, 195–197 Prosimulium, 1043–1044, 1045t–1047t, 1052–1053 Prosopistoma, 885 Prosopistomatidae, 874–876, 875t–876t, 879–880 Prostigmata, 601, 601t–602t, 613–649 Prostoma, 207t, 291 P. Asensoriatum, 207t P. canadiensis, 207t P. communopore, 207t P. eilhardi, 206, 207t P. eilhardi macradenum, 207t P. graecense, 206, 207t, 208 P. hercegovinense, 207t P. jenningsi, 207t P. kolasai, 207t P. ohmiense, 207t P. puteale, 207t Protanthea simplex, 341 Protanyderus, 1043–1044, 1045t–1047t Protereismatidae, 873 Proterometra macrostoma, 408 Proteus mirabilis, 606–607 Protista, 11 Protochauliodes, 1061–1063 P. spenceri, 1060, 1060f Protogonyaulax, 122 Protohermes, 1061 P. grandis, 1063 P. niger, 1062–1063 Protolimnesiinae, 620t–625t Protomonotresidae, 191–192 Protoneuridae, 894–895 Protonymph, 642 Protoperidinium, 121f, 122 Protoplanellinae, 197 Protoplasa, 1043–1044, 1045t–1047t Protoptila, 971–972, 971f Protoptilinae, 989 Protosialis, 1060–1061 Protosticta, 894 Protozoa, 69f, 70, 113–131 Protura, 850–851 Protzia, 638 Protziinae, 620t–625t Psalteriomonas, 122 Psammolimnesiinae, 620t–625t Psammoryctides, 534–535 P. barbatus, 543 Psephenidae, 1004, 1005t–1007t, 1008–1009, 1011, 1013–1015, 1017, 1036–1037, 1037f Pseudechiniscus, 350t, 355, 356f, 369 P. suillus, 371t–374t Pseudiron, 885

1114

Pseudobiotus, 350t, 354, 356f, 364, 366, 367f, 369–370 P. augusti, 371t–374t P. kathmanae, 366, 367f, 371t–374t P. megalonyx, 364 Pseudobodo, 121 Pseudobranchus axanthus, 741 Pseudocandona marchica, 769 Pseudochordodes P. bedriagae, 306 P. dugesi, 317 Pseudodiaptomidae, 710–711, 728 Pseudodiaptomus, 710–711, 728, 731–732 Pseudohalmyrapseudes aquadulcis, 782–783 Pseudohexapodibius, 350t Pseudohydryphantes, 628–629 Pseudohydryphantinae, 620t–625t Pseudolestidae, 896–899, 897t–898t Pseudomonas hirudinis, 570 Pseudoneureclipsidae, 965, 966t–967t, 969, 969f Pseudopalaemon, 803 Pseudophyllidea, 725–726 Pseudorhynchelmis, 534 Pseudoscorpiones, 600 Pseudostigma accedens, 913 Pseudostigmatidae, 922 Pseudostomum quadrioculatum, 195–196 Pseudosuccinea, 396 P. columella, 387f, 396, 400, 405 Pseudosyrtis P. fluviatilis, 184–185 P. neiswestnovae, 184–185 Pseudothelphusidae, 798, 799t, 802, 805–806, 824, 825t Pseudotrichomonas, 125 Psilotreta, 965–969, 968f Psychodidae, 1043–1044, 1045t–1047t, 1048f, 1049–1050, 1054, 1056–1058 Psychomyiidae, 965, 966t–967t, 972–976, 990 Pterastericolidae, 197 Pteridomonas, 121, 121f Pteriomorphia, 450–451 Pteronarcyidae, 934–935, 934t, 940t, 942 Pteronarcys P. biloba, 940 P. californica, 938 P. scotti, 936, 937f Ptilocolepidae, 965–969, 966t–967t, 974–979, 989 Ptilodactylidae, 1004, 1005t–1007t, 1008–1009, 1013–1014, 1037 Ptychobranchus, 424, 489f P. fasciolaris, 457, 461f P. subtentum, 463, 465f Ptychoptera, 1043–1044, 1045t–1047t Ptychopteridae, 1043–1044, 1045t–1047t, 1048f, 1049–1050, 1053–1054 Ptygura, 234f, 248, 258–259, 265 P. beauchampi, 238, 247 P. libera, 238 P. pilula, 244, 248, 248f Pulmonata, 385–388, 390t, 391f, 392, 396–399, 401, 401t, 403t

Taxonomy Index

Puri aleca, 1061 Pycneus, 803, 824 Pycnisia, 803 Pycnogonida, 592, 600 Pycnopsyche, 979–981, 982f, 987 Pyganodon, 489f, 424, 460–462, 466 P. cataracta, 461–462 P. grandis, 467, 469, 475–476, 480, 484 P. implicata, 468 Pyralidae, 100–101 Pyraustinae, 1068–1069 Pyrgodesmidae, 665f, 666 Pyrgulopsis, 394t, 395 P. bruneauensis, 410t P. neomexicana, 410t P. ogmoraphe, 410t P. pachyta, 410t P. roswellensis, 410t

Q

Quadrula, 424, 454, 489f Quadrulini, 424, 462, 489f Quincuncina, 424, 489f Quistadrilus multisetosus, 533–534

R

Radiospongilla, 137 Radix, 396, 406 R. auricularia, 100 R. balthica, 408 Rallidentidae, 874–877, 875t–876t Ramajendas, 350t, 352–353, 369 Ramazzottidae, 350t, 353 Ramazzottius, 350t, 353, 357–359, 370 R. oberhaeuseri, 353f, 361 Ramiheithrus virgatus, 979–982, 983f Rana, 294, 741 R. clamitans, 319–320 R. erythraea, 320f R. temporaria, 868 Ranatra, 951–953, 952f, 959 Randiellidae, 531, 533 Raphidioptera, 1060 Raphignathina, 601t–602t Raphignathoidea, 601t–602t, 614–615 Raptobaetopus, 885 Rechtracheata, 874–876 Redudasyidae, 211–212 Redudasys, 214–216 R. fornerise, 211–212, 212f Regina alleni, 832 Rehbachiella, 688 Remanella, 124f Remipedia, 672, 710 Reticulomyxa, 119 Retortamonas, 120 Retronectidae, 186, 194 Rhabditida, 273, 290–291, 295 Rhabditidae, 277f, 278, 280f Rhabditoidea, 290–291 Rhabditophora, 181, 184, 186–188, 188f, 190–192, 195 Rhabditrina, 273 Rhabdoblatta, 1067–1068

Rhabdocela, 186 Rhabdochona pracox, 294 Rhabdocoela, 182, 183f, 184–186, 188, 192–193, 195–200, 834 Rhabdocoelida, 182, 197 Rhantus, 1013, 1022–1024 Rhaphidophoridae, 308 Rhaycophilidae, 971 Rheumatobates, 958 Rhicnoda, 1067–1068 R. natatrix, 1067–1068 Rhinoglena, 258–259 R. fertoeensis, 255, 256f R. frontalis, 243, 250, 253 Rhithrogena semicolorata, 878, 879f Rhithropanopeus harrisii, 680, 820 Rhizocephala, 711–712 Rhyacodrilinae, 533, 539–540 Rhyacodriloides, 534 Rhyacodrilus, 534 R. falciformis, 543 Rhyacophila, 976, 990, 992 R. angelieri, 993 R. fuscula, 20f Rhyacophilidae, 965–969, 966t–967t, 974–979, 989–992, 995 Rhyacopsyche, 992 Rhynchelmis, 534, 540, 542 R. brachycephala, 517 Rhynchobdellida, 511–512, 566 Rhynchocoela, 205–206 Rhynchohydracaridae, 601t–602t, 620t–625t Rhyncholimnochares, 628, 637–638, 645 R. kittatinniana, 17f, 631f, 653 Rhyncholimnocharinae, 620t–625t Rhynchomonas, 121, 121f R. nasuta, 120, 127 Rhynchoscolex simplex, 196 Rhyniognatha hirsti, 849 Riccia, 62 Richardsonianus australiensis, 583 Richtersius, 350t R. coronifer, 353f, 357f, 375 R. oberhaeuseri, 364 Ricinulei, 600 Riekoperla darlingtoni, 939 Rimanellidae, 895–899, 897t–898t Ripistes, 518, 542 Riseriellus occultus, 208–209 Rissooidea, 390, 390t Romanomermis R. culicivorax, 283f, 300 Rossianidae, 965, 966t–967t Rotalia, 118f Rotaria, 231–232 Rotifera, 11, 212, 225–226, 230, 249, 258, 375f Rotifers, 24, 69f, 70 Rutripalpidae, 601t–602t, 620t–625t

S

Sabellidae, 519–521 Saldidae, 951–953, 952f, 954t Salenthydrobia, 395 Salifidae, 566, 578

Taxonomy Index

Salmo S. trutta, 319–320, 577 S. trutta fario, 832 Salpingoeca, 120–121 Salticidae, 602–603 Salvelinus S. fontinalis, 319–320, 471 S. leucomaenis japonicus, 320 Salvinia, 62, 1068–1069 S. molesta, 100–101 Samastacus, 801–802, 805 Samea multiplicalis, 1068–1069 Sander vitreus, 87 Santiagocarinae, 620t–625t Saprodinium, 125 Sararcinia purpurea, 49f Sarcophaga, 1043–1044, 1045t–1047t Sarcophagidae, 1043–1044, 1045t–1047t, 1049, 1049f Sarcoptiformes, 601t–602t, 609–613 Sarracenia, 608, 1054, 1055f S. purpurea, 228f Sarraceniopus hughesi, 608 Sathodrilus, 552–553 S. attenuatus, 553 Satonius, 1019–1020 S. stysi, 1019–1020, 1020f Saturnospongilla carvalhoi, 141f Scapholeberis, 70 S. kingi, 261 Scaphydra, 1018 Scarabaeiformia, 1008 Scardinius erythrophthalmus, 923–924 Scathophagidae, 988 Scenedesmaceae, 264 Scenedesmus, 260, 748 S. obliquus, 264 Schistocephallus, 725–726 Schistonota, 874–876 Schistosoma S. haematobium, 107f S. mansoni, 773–774 Schizodrilus, 543 Schizomida, 600 Schmardaella, 517 Schmidtea, 200 S. mediterranea, 16f Schminkeinae, 620t–625t Schusterius, 350t Schwiebea, 612f Scilla siberica, 726 Sciomyzidae, 1043–1044, 1045t–1047t, 1056 Scirtidae, 1004, 1005t–1007t, 1009, 1013–1014, 1037–1038, 1037f–1038f Scirtinae, 1037–1038 Scleorchious rubecula, 320 Scolecida, 520–521 Scopuridae, 934–935, 934t Scorpiones, 600 Scutariella japonica, 559 Scuticociliates, 123f Scylla serrata, 806–807 Sedentaria, 531 Seison, 234–235

1115

Seisonidae, 227 Selminosoma chapmani, 667 Semblis, 979–981 Semibalanus, 742 Semisulcospira, 393, 403 Semisulcospiridae, 390, 390f Semotilus atromaculatus, 319–320 Senecella, 719, 726 S. calanoides, 734 Sepedon, 1043–1044, 1045t–1047t Sericostoma personatum, 986–987 Sericostomatidae, 965–969, 966t–967t, 968f, 979–982, 983f Sermyla, 395 Serradium S. hirsutipes, 666–667 S. semiaquaticum, 17f, 666–667, 667f Serratella ignita, 879–880, 880f Sesarma, 820 S. bidentatum, 820 S. verleyi, 820 S. windsor, 820 Sesarmidae, 798, 799t, 820 Sessilia, 711–712 Setisura, 874–876 Setodes, 991–992 Setopus, 220f Sharasialis, 1063 Sialia mexicana, 320 Sialida, 1060–1061 Sialidae, 1060–1061, 1060f, 1063 Sialis, 20f, 375f, 1060–1061, 1060f S. californica, 1062–1063 Sida, 694 Sididae, 688t Siettitia, 1017 Sigara selecta, 956–957 Silphopsyllus desmanae, 1035 Silvius, 1043–1044, 1045t–1047t Simpsonaias, 424, 489f S. ambigua, 469 Simulidae, 463, 465f Simuliidae, 58, 104, 106, 294, 375f, 925, 1043–1044, 1045t–1047t, 1048f, 1049–1054, 1056–1057 Simulium, 39, 834, 1043–1044, 1045t–1047t S. arcticum, 104 S. damnosum, 105f, 106 S. metallicum, 294f S. vittatum, 104 Sinanodonta woodiana, 86, 430 Sinantherina, 251, 258–259, 263–264 S. semibullata, 238, 245, 247t S. socialis, 237–238, 244, 246–249, 246f, 247t, 249f–250f, 251, 258, 264, 264f S. spinosa, 263–264 Sinocalanus, 731–732 Sinochauliodes, 1061 Sinoneurorthus, 1064 Siolineus, 207t S. turbidus, 207t Siphlaenigmatidae, 874–876, 875t–876t Siphlonuridae, 874–878, 875t–876t, 883 Siphlonuroidea, 874–876, 878

Siphlonurus occidentalis, 885 Siphluriscidae, 874–876, 875t–876t, 879 Siphluriscus chinensis, 874–876 Siphonostomatoida, 710 Siphonuridae, 881–882 Sipuncula, 512 Sisyborina, 1064 Sisyra, 149, 1064–1065 S. pedderensis, 1067 Sisyridae, 148–149, 150f, 1064–1067, 1064f–1065f Skeletonema, 750 S. costatum, 254 Skistodiaptomus, 727–728 Smicridea, 992 S. thermophila, 992 Sminthuridae, 865, 865f Sminthurides, 604, 865 S. aquaticus, 866–867 S. lepus, 850–851, 850f Solenogastres, 384 Solifugida, 600 Somatochlora, 913, 922 S. alpestris, 922 S. flavomaculata, 923, 925f S. metallica, 902–903, 903f Somatogyrus, 394t, 395 Sommaniathelphusa, 829 Sorbeoconcha, 389, 391 Spanglerogyrus, 1024–1025 Sparganophilidae, 510, 519 Spelaeodiaptomus rouchi, 726 Spelaeogriphacea, 13, 782–784 Speodiaptominae, 726 Speodiaptomus birsteini, 726 Speonoterus, 1027–1028 Spercheidae, 1004, 1005t–1007t, 1008–1009, 1013–1014, 1038 Spercheus, 1038 Sperchon, 638, 645–646 Sperchonopsis ecphyma, 627f, 629f Sperchontidae, 601t–602t, 620t–625t Spermathecae, 539–540, 548 Spermatozeugmata, 539–540, 540f Sphaeridiinae, 1034 Sphaeriidae, 384, 388, 425–426, 431–435, 434t, 437f, 438–439, 441, 454–455, 457–460, 466 Sphaeriinae, 432–433 Sphaerium, 432–433, 438–439, 441, 443–445, 445f, 449–450, 459, 466, 471, 474, 476, 483–484, 488f–489f S. corneum, 433, 434t, 481 S. occidentale, 476 S. partumeium, 476 S. rhomboideum, 433 S. securis, 476 S. striatinum, 443–444, 446f Sphaerius, 1019, 1019f–1020f S. africanus, 1019 Sphaeriusidae, 1004, 1005t–1007t, 1009, 1011, 1013–1014, 1019f–1020f, 1019 Sphaeroceridae, 1050, 1051f Sphaerodactylus parvus, 534–535, 543

1116

Sphaeroeca, 117f, 120–121 Sphagnophylax meiops, 965–969, 968f Sphagnum, 119, 218, 220, 228f, 231–232, 922, 1019 Spharuiusidae, 1008 Spicipalpia, 965–969, 971–972, 989 Spinadis, 885 Spinalona, 694 Spinastacoides, 801–802 Spinicaudata, 687–689, 688t, 691–695, 697–698, 757–758 Spinochordodes tellinii, 316–317 Spionidae, 519–521 Spirobolida, 662f, 664 Spirobolidae, 308 Spirosperma ferox, 541 Spirostreptida, 664 Spirulina, 777, 794 Spirurina, 273 Spiruroidea, 294 Spongia palustris, 149 Spongilla S. fluviatilis, 149 S. lacustris, 138f–139f, 140, 141f, 144, 147, 149 Spongillidae, 135f, 137, 147, 1065, 1065f Spongillina, 134–135, 134f–135f, 137–138, 138f, 141f, 145f, 147 Sporozoa, 518 Spumella, 121, 121f Stagnicola, 396 S. caperata, 391 Staphylinidae, 1004, 1005t–1007t, 1009, 1013– 1014, 1038–1039 Staphyliniformia, 1008 Staphylininae, 1038–1039 Staphylinoidea, 1008 Stegoelmis, 1016 Stenasellus virei, 790 Steninae, 1038–1039 Stenochironomus, 1056 Stenocolus, 1031 S. scutellaris, 1031 Stenocyphoninae, 1037–1038 Stenocypris, 773t Stenomelania, 395, 398 Stenonema, 885 Stenopelmatidae, 308 Stenophylax, 992 Stenopodidea, 674f Stenopsychidae, 965, 966t–967t, 990 Stenosialis, 1060–1061 Stenostomidae, 183f, 192 Stenostomum, 172–173, 183f, 189f–190f, 192, 200 Stenothyra, 395 Stenothyridae, 395 Stentor, 122–123 Stenus, 1038–1039 S. comma, 868 S. fornicatus, 1038–1039 Stephanella hina, 336 Stephanoceros, 248, 258–259 Stephanoeca, 120–121 Sterna paradisea, 292

Taxonomy Index

Stictocladius, 1053 Stigmaeidae, 601t–602t, 614–615 Stratiodrilus, 520 Stratiomyidae, 1043–1044, 1045t–1047t, 1049, 1049f, 1052–1054, 1056 Stratiomys, 1056 Stratiotes aloides, 918–919 Streptocephalidae, 688t Streptocephalus, 695–696 S. mackini, 694–695 S. moorei, 692f S. sealii, 695 Strombidium, 122–123, 125 Strophitus, 424, 466, 489f S. undulates, 457–458, 462f Stygameracarinae, 620t–625t Stygatoniidae, 620t–625t Stygiocaris, 803 Stygiomysida, 672t, 782–783 Stygobromus, 789–790 Stygodiaptomus kieferi, 726 Stygolimnesiinae, 620t–625t Stygoparnus, 1013–1014, 1029–1030 S. comalensis, 1016–1017, 1029–1030 Stygotantulus stocki, 710 Stygothrombiae, 601t–602t Stygothrombidiidae, 601t–602t, 615–617, 617f Stygothrombidiidae, 615–617 Stygothrombidioidea, 601t–602t, 618 Stygothrombium, 617f Stygotoniidae, 601t–602t Stylaria, 542 S. lacustris, 530f, 534 Stylochus ellipticus, 748 Stylodrilus, 514–515, 534 S. cernosvitovi, 534–535, 543 S. heringianus, 538–539, 544–545 Stylommatophora, 392 Styloperlidae, 934–935, 934t Styraconyx hallasi, 369 Suberitidae, 147 Suctoria, 123f Sudanonautes S. monodi, 823 S. monodosus, 814 Sulcarius biannulatus, 988 Sulcospira, 395 Suragina, 1043–1044, 1045t–1047t Symbiocladius, 886, 1056 Symbiocloeon, 886 Symbiodinium, 119–120, 122 Sympecma, 917 Sympetrum, 913, 916, 918, 922 S. internum, 641 S. obtrusum, 641 S. sanguineum, 902–903, 906–907, 921 S. striolatum, 914, 915f, 917, 923–924 Symsagittifera roscoffensis, 188 Syncarida, 13, 781, 783–785, 783f, 788, 790 Syncaris, 803, 824 S. pacifica, 824 Synchaeta, 226, 229–230, 234–235, 234f, 256f, 257, 263–264 S. grandis, 253 S. oblonga, 256f, 257

S. pectinata, 244, 251, 257, 264 S. tremula, 256f Syndermata, 230 Syndinium, 725–726 Syngenodrilidae, 533 Synlestidae, 894, 897–899, 897t–898t Synthemistidae, 897–899, 897t–898t, 929 Synura, 121 Syrphidae, 1043–1044, 1045t–1047t, 1049–1050, 1049f, 1052, 1054, 1056 Systellognatha, 934–935, 934t

T

Tabanidae, 58, 106, 1043–1044, 1045t–1047t, 1049–1050, 1049f, 1051f, 1054, 1056–1058 Tabanus, 1043–1044, 1045t–1047t, 1049–1050, 1049f, 1051f Tachidiidae, 711 Tachytrechus, 1043–1044, 1045t–1047t Taeniopterygidae, 934–935, 934t, 937f, 940t, 942–944 Taeniopteryx, 946 T. burksi, 936, 939f Taiochauliodes, 1061 Talorchestria brito, 295 Tanaidacea, 13, 672, 672t, 782–783 Tanjistomella verna, 974–976, 975f Tantulocarida, 710–712, 727–728 Tanycypris, 773t Tanyderidae, 1043–1044, 1045t–1047t, 1054, 1056 Tanymastigitidae, 688t Tanypodinae, 1056 Tanypteryx pryeri, 922 Tanytarsus, 1043–1044, 1045t–1047t Tarachodes, 923, 923f Taraxitrichia, 971–974, 973f Tardigrada, 11, 348–351, 349f, 350t, 351f–352f, 354–355, 355f, 357, 360–364, 362f, 367– 370, 375–378, 375f, 591–592 Tartarothyadinae, 620t–625t Tartarothyas, 628–629 Tasimiidae, 965, 966t–967t Taxiphyllum, 62 Taylorconcha serpenticola, 410t, 411–412 Tegeocranellidae, 601t–602t Teinobasis, 894–895 Telephlebia brevicauda, 921 Teloganellidae, 874–877, 875t–876t Teloganodidae, 874–877, 875t–876t Temnocephala, 183f, 199 Temnocephalidae, 183f, 184, 186, 197–199, 834 Temnostoma, 1043–1044, 1045t–1047t, 1056 Temoridae, 727–728 Tenuibiotus, 350t, 352f T. willardi, 371t–374t Tenuibranchiurus, 805 Teratocyclops, 727 Teratothyadidae, 601t–602t, 620t–625t Terebellida, 520–521 Terrestricytheroidea, 758–759 Terrimegadrili, 510 Testechiniscus spitzbergensis, 371t–374t Testudacarinae, 620t–625t

Taxonomy Index

Testudinella, 229f Tetanocera, 1043–1044, 1045t–1047t Tetracanthagyna plagiata, 902 Tetracapsuloides T. bryozalmonae, 338 T. bryozoides, 338 Tetraclitoidea, 712 Tetraconata, 12–21, 592, 850 Tetragnathida, 602–603 Tetrahymena, 114–115, 122–123, 124f Tetramermis fissispina, 292 Tetrastemma, 208 Tetrigidae, 1069–1070 Tettigoniidae, 308, 1069–1070 Teutonia, 637 Teutoniidae, 601t–602t, 620t–625t, 637–638 Thalassarachna basteri, 613–614 Thalassius spinosissimus, 605 Thalassodrilus hallae, 543 Thalerius, 350t Thamnocephalidae, 688t Thamnocephalus salinarum, 695–696 Thaumalea, 1043–1044, 1045t–1047t Thaumaleidae, 1043–1044, 1045t–1047t, 1050, 1053, 1057 Thaumatoneuridae, 896–899, 897t–898t Thaumatopsylloida, 710 Thecostraca, 710–712 Theliderma, 424, 437, 463 T. intermedia, 438, 439f T. metanevra, 454 T. tuberosa, 429, 429t Theodoxus, 400, 401t Theridiidae, 605 Theristus, 292 Thermacaridae, 601t–602t, 620t–625t Thermacarus, 645 Thermobathynella adami, 680, 785–786 Thermocyclops, 727–729, 731, 733 Thermomesochra reducta, 721 Thermonectus T. marmoratus, 1010–1011 T. nigrofasciatus ornaticollis, 9–10 Thermopsis thermophila, 772 Thermosbaena T. mirabilis, 680, 785–786 Thermosbaenacea, 13, 672t, 782–783, 785–786 Thermosphaeroma T. subequalum, 680 T. thermophilum, 680, 785–786 Thermozodiidae, 350t Thermozodium, 350t T. esakii, 350 Theromyzon, 578 T. tessulatum, 574–575, 579 T. trizonare, 574–576, 578–580, 583 Thiara, 395 T. winteri, 388f Thiaraidae, 391 Thiaridae, 100, 389–390, 394t, 398 Thomisidae, 601t–602t, 602–603, 606 Thomisoidea, 601t–602t Thomisus nepenthiphilus, 606 Thoracica, 711–712 Thoracopods, 673–675

1117

Thremmatidae, 965, 966t–967t, 979–981, 980f, 982f Thulinius, 350t, 352f–353f, 354, 366–367, 369 T. augusti, 370, 371t–374t T. ruffoi, 371t–374t T. saltursus, 371t–374t T. stephaniae, 371t–374t, 377 Tiguassuidae, 533 Timiriaseviinae, 759t, 767, 769 Tinodes, 992 T. waerneri, 988–989 Tintinnids, 123f Tintinnopsis, 122–123, 124f Tiphyinae, 620t–625t, 644–645 Tiphys, 643–644 T. americanus, 641 Tipula, 1050, 1051f Tipulidae, 608, 861, 861f, 1043–1044, 1050, 1054, 1056, 1066 Tipuloidea, 1043–1044, 1045t–1047t, 1049–1050, 1051f, 1053–1054, 1056–1058 Tobrilidae, 281f, 286 Tobrilus, 281f, 292 T. gracilus, 288 T. grandipapellatus, 288 Todothyas, 636, 639, 647–648 T. barbigera, 637 Tolhuaca T. brasiliensis, 979–982, 983f T. cupulifera, 979–982, 983f Tolypothrix, 777 T. tenuis, 777 Tonnacypris, 773t Tonnoirocladius, 1054, 1055f Torrenticola, 618f, 630f, 638, 645–646 Torrenticolidae, 601t–602t, 620t–625t Torrenticolinae, 620t–625t Torridincola, 1019–1020 Torridincolidae, 1004, 1005t–1007t, 1008–1011, 1013–1014, 1019–1020, 1020f Toxolasma, 424, 463–465, 489f T. parvus, 454 T. pullus, 454 T. texasiensis, 444, 449f Toxoplasma gondii, 125 Toxorhynchites, 1044, 1048f, 1049–1050, 1051f Tracheloraphis, 122–123 Trajancypris, 773t Tramea basilaris, 911–912, 913f, 914, 915f Trebouxia, 375 Trechalea, 604, 606–607 T. tirimbina, 606f Trechaleidae, 601t–602t, 602–604, 606–607, 606f Trechaleoides, 606 Trechus triamicorum, 1022 Trematoda, 181, 403, 411–412, 925 Trepomonas, 120 Trhypochthoniidae, 601t–602t, 611 Triaenodes, 974–976, 979–981, 982f, 991 T. perna, 972–974, 973f T. tardus, 965–969, 968f Trichechus, 712 Trichobothria, 609–610, 616–617 Trichocerca

T. pusilla, 251 T. rattus, 234f, 258–259 Trichocorixa verticalis, 956 Trichodactylidae, 798, 799t, 802, 805–806, 824, 825t Trichodactyloidea, 805–806 Trichodina, 725–726 T. diaptomi, 726 Trichodrilus pauper, 534–535, 543 Trichomitopsis, 120 Trichomonas, 125 T. vaginalis, 120 Trichoptera, 13, 98–99, 148–149, 285f, 375f, 637–638, 851, 855, 858, 858f, 863–864, 953, 959, 965–976, 966t–967t, 969f–971f, 973f–975f, 978–982, 979f–981f, 983f, 984–993, 995–996, 1004, 1069 Trichothyas, 631f Tricladida, 183f, 186, 188–193, 195–197, 200 Tricorixia verticalis, 857, 858f Tricorythidae, 874–877, 875t–876t, 883–884 Tricorythodes, 885 T. minutus, 881 Tricula, 403 Tridactylidae, 1069–1070 Tridentiger bifasciatus, 173 Trigonioida, 425–426 Trilobita, 592, 850 Trilobus, 292 Trimyema, 125 Triops, 18f, 689, 694–695 Triopsidae, 688t Triplonchida, 273 Tripyla, 292 Trithemis kirbyi, 923, 923f Tritogonia, 424, 489f T. verrucosa, 457–458, 462f Tritonymph, 643 Trochophore, 388 Trochosa, 603–604 Trochospongilla horrida, 141f Trochospongilla leidii, 84–85 Troglocambarus, 827–828 T. maclanei, 673f, 819, 819f, 828, 830 Troglocaris, 803, 824 Troglocubanus, 824 Troglodiaptomus, 726 Troglomexicanus, 824 Trombiculoidea, 601t–602t Trombidiae, 601t–602t Trombidiformes, 601, 601t–602t Trombidioidea, 618 Tropidopolinae, 1070 Tropocyclops, 262, 721–722, 725, 730 T. extensus, 725 T. prasinus, 723–724 Truncilla, 424, 489f Tryonia, 395 Trypanosoma brucei, 121 Tubifex, 542, 576, 1026–1027 T. pomoricus, 541 T. tubifex, 518–519, 533–534, 537f, 541, 543–545 Tubificidae, 516f, 529–536, 530f, 538–543, 545 Tubificinae, 539–542

1118

Tulotoma, 393–394 T. magnifica, 410t Turbanella hyalina, 220 Turbellaria, 182–184, 182f–183f, 187, 196, 375f, 834 Tylenchidae, 288 Tylenchina, 273 Tylomelania, 395 T. sulawesi, 391–392, 391f Typhlatya, 803, 824 Typhlocarididae, 798, 799t, 804, 824, 825t Typhlocaris galilea, 804 Typhloplanidae, 184–185, 193, 197, 199–200 Typhloplanoida, 183f, 184, 186, 191 Tyronia alamosae, 410t Tyrrellia, 645–646 Tyrrellinae, 620t–625t

U

Uchidastygacarinae, 620t–625t Uchidastygacarus, 645 U. acadiensis, 633f Uenoidae, 965, 966t–967t, 985, 990 Uglukodrilus hemophagus, 558 Ula, 1043–1044, 1045t–1047t Ulianinidae, 195 Umagillidae, 197 Umborotula bogorensis, 141f Umma, 908, 909f, 920 Uncinais uncinata, 542–543 Undula, 218 Unio, 427 Uniomerus, 424, 475, 489f U. tetralasmus, 454 Unionacea, 424 Unionicola, 634–637, 640, 643, 645–648, 651, 654 U. crassipes, 641 U. foili, 641–642, 649 U. intermedia, 644 Unionicolidae, 470–471, 601t–602t, 620t–625t, 633f, 988 Unionicolinae, 620t–625t Unionidae, 95, 424–428, 440–441, 444f, 457–460, 461f, 489f Unioninae, 489f, 424, 427, 457–458, 460, 462 Unionoida, 424–428, 431–432, 434, 453–454, 458 Unionoidea, 489f, 424, 425f, 426–431, 438–439, 455, 457, 460 Urastomidae, 182, 191–192 Urnatella, 340–341 U. gracilis, 15f, 340–341, 341f

Taxonomy Index

Uroglena, 121 Uropygi, 600 Utaxatax, 637 Utricularia, 265–266 Utterbackia, 489f, 424, 461–462, 466 U. imbecillis, 454, 466–467, 481 U. suborbiculata, 456, 460f

V

Vallisneria, 33f Valvata, 400, 403–404 V. utahensis, 410t Valvatidae, 389, 390t, 394t, 398, 400 Vampyrella, 118–119 Vannella, 118–119 Variechaetadrilus harmani, 533–534 Varuna, 820 V. litterata, 806–807 Varunidae, 798, 799t, 806–807, 820 Vejdovskyella, 518 V. comata, 543 Velesunio ambiguus, 480 Veliidae, 951–955, 954t, 957–958 Velkovrhia, 164–165 V. enigmatica, 162, 163t–164t, 167, 169–170 Veneroida, 425–426, 432, 434 Venustaconcha, 424, 463–465, 489f Vesicularia, 62 Vibrio cholerae, 733 Victorella, 330f V. pavida, 174, 334f Vietnamellidae, 874–876, 875t–876t Villosa, 424, 463–465, 489f V. iris, 449, 455f, 466, 478, 480 V. vanuxemi, 455 Virilastacus, 801–802, 805 V. araucanius, 826 V. rucapihuelensis, 826 Viviparidae, 100, 389, 389f, 390t, 391, 394t, 399–401, 400t Viviparoidea, 389, 390t Viviparus, 393–394, 400 V. georgianus, 397, 400, 402–403 V. subpurpureus, 404 Volsellacarus sabulonus, 633f Vorticella, 117f, 122–123, 245f Vriesea, 1054, 1055f

W

Wandesia, 637–638, 645 Wandesia (Partnuniella) thermalis, 637 Wandesiinae, 620t–625t

Wettinidae, 601t–602t, 620t–625t Wolbachia, 320–321, 774, 940 Wolffia, 867 Wormaldia, 986–987 Wyeomyia, 1054

X

Xanthidae, 798, 799t Xanthocnemis zealandica, 924 Xenochironomus, 149 Xerobiotus, 350t Xestoleberidae, 759t Xiphocaridae, 59 Xiphocarididae, 798, 799t, 804 Xiphocaris, 804 X. elongata, 677, 804 X. gomezi, 804 Xiphocentronidae, 965, 966t–967t, 972–974, 978–979, 990 Xiphosura, 592, 600 Xironodrilinae, 511, 551–552 Xironodrilus formosus, 555–556 Xironogiton, 552–553 X. instabilis, 555–558 X. victoriensis, 552f, 553, 555–558, 556f Xubida infusella, 100–101 Xylotopus, 1056

Y

Yara, 1018 Yinia, 207t Y. pratensis, 207t Ytu, 1019–1020 Y. zeus, 1019–1020

Z

Zeacumantus subcarinatus, 408 Zelandothyadidae, 601t–602t, 620t–625t Zelandothyadinae, 620t–625t Zetomimidae, 601t–602t, 611 Zetomimus, 609–610 Z. francisi, 610f Zwickia, 613 Zygonoides occidentis, 923, 925f Zygonyx, 908, 909f, 920–921 Zygoptera, 639f, 853, 894–903, 895f, 897t–898t, 899f–901f, 905, 908–912, 908f, 912f, 914, 917–920, 922, 926, 929

Thorp and Covich’s Freshwater Invertebrates

A Global Series of Books on the Identification, Ecology, and General Biology of Inland Water Invertebrates by Experts from Around the World Fourth Edition Series Editor: James H. Thorp Volume I: Ecology and General Biology Edited by James H. Thorp and D. Christopher Rogers Published 2015 Volume II: Keys to Nearctic Fauna Edited by James H. Thorp and D. Christopher Rogers Published 2016 Volume III: Keys to Palaearctic Fauna Edited by D. Christopher Rogers and James H. Thorp Expected Publication Date: 2017 Volumes in Preparation and Under Contract Keys to Neotropical and Antarctic Fauna Keys to Neotropical Hexapoda Keys to Fauna of the Australian Bioregion Possible Future Volumes of the Fourth Edition Keys to Oriental and Oceana Fauna Keys to Oriental and Oceana Hexapoda Keys to Palaearctic Hexapoda Keys to Afrotropical Fauna Keys to Afrotropical Hexapoda Related Publications Ecology and Classification of North American Freshwater Invertebrates Edited by J.H. Thorp and A.P. Covich First (1991), Second (2001), and Third (2010) Editions Field Guide to Freshwater Invertebrates of North America by J.H. Thorp and D.C. Rogers

Keys to Nearctic Fauna Thorp and Covich’s Freshwater Invertebrates - Volume II

Fourth Edition

Edited by

James H. Thorp D. Christopher Rogers

AMSTERDAM • BOSTON • HEIDELBERG • LONDON • NEW YORK • OXFORD • PARIS SAN DIEGO • SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

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Dedications from the Editors

To Henry B. Ward, George C. Whipple, W. Thomas Edmondson, and Robert W. Pennak—pioneers who blazed a publishing trail with books on the ecology and identification of North American freshwater invertebrates. To Alan P. Covich, a longtime friend and valued colleague, who not only helped develop the first three editions but also made possible the fourth edition’s improved taxonomy and worldwide coverage by introducing the current editors to each other. James H. Thorp and D. Christopher Rogers

Contributors to Volume II

Fernando Álvarez  [Chapter 16] Departamento de Zoología, Instituto de Biología, U.N.A.M., Circuito exterior s/n, Ciudad Universitaria, Copilco, Coyoacán, A.P. 70-153, México, Distrito Federal. C.P. 04510, México; email: [email protected] Bonnie A. Bain [Chapter 12] Department of Biological Sciences, Southern Utah University, Cedar City, Utah 84720, USA; email: [email protected] llse Bartsch [Chapter 16] Forschungsinstitut Senckenberg, c/o DESY, Gebaeude 3, Raum 316, Notkestr. 85, 22607, Hamburg, Germany; email: [email protected] Valerie Behan-Pelletier [Chapter 16] Agriculture and Agri-Food Canada, K.W. Neatby Building, 960 Carling Avenue, Ottawa, Ontario K1A 0C6, Canada; email: ­[email protected] Matthew G. Bolek  [Chapter 10] Department of Zoology, Oklahoma State University, 501 Life Sciences West, Stillwater, Oklahoma 74078, USA; email: bolek@ okstate.edu Ralph O. Brinkhurst  [Chapter 12] 205 Cameron Court, Hermitage, Tennessee 37076, USA Francisco Brusa [Chapter 5] División Zoologia Inverte­ brados, Museo de La Plata, FCNyM-UNLP, 1900 La Plata, Argentina; email: [email protected] Richard D. Campbell  [Chapter 4] Department of Develop­ mental and Cell Biology, University of California, Irvine, CA, USA; post mail: 2561 Irvine Ave., Costa Mesa, California, 92627 USA; email: [email protected] Joo-lae Cho [Chapter 16] Invertebrate Research Division, National Institute of Biological Resources, Environmental Research Complex, Gyoungseo-dong, Incheon, 404-170, South Korea; email: [email protected] David R. Cook [Chapter 16] 7725 North Foothill Drive South, Paradise Valley, Arizona 85253, USA; email: [email protected] Kevin S. Cummings  [Chapter 11] Illinois Natural History Survey, Center for Biodiversity, 607 East Peabody Drive, Champaign, Illinois 61820, USA; email: ksc@ inhs.uiuc.edu

Cristina Damborenea [Chapter 5] División Zoología Invertebrados, Museo de La Plata, FCNyM-UNLP, Paseo del Bosque, 1900 La Plata, Argentina; email: [email protected] R. Edward DeWalt  [Chapter 16] Illinois Natural History Survey, Center for Biodiversity, 607 East Peabody Drive, Champaign, Illinois 61820, USA; email: edewalt@inhs. illinois.edu Genoveva F. Esteban  [Chapter 2] Conservation Ecology and Environmental Sciences Group, Faculty of Science and Technology, Bournemouth University, Dorset, United Kingdom; email: gesteban@bournemouth. ac.uk James W. Fetzner Jr.  [Chapter 16] Biodiversity Services Facility, Section of Invertebrate Zoology, Carnegie Museum of Natural History, 4400 Forbes Avenue, Pittsburgh, Pennsylvania 15213-4080, USA; email: [email protected] Bland J. Finlay [Chapter 2] School of Biological and Chemical Sciences, Queen Mary University of London, The River Laboratory, Wareham, Dorset, BH20 6BB, United Kingdom; email: [email protected] Stuart R. Gelder  [Chapter 12] Department of Science and Math, University of Maine at Presque Isle, Presque Isle, Maine 04769, USA; email: [email protected] Fredric R. Govedich  [Chapter 12] Department of Biological Sciences, Southern Utah University, 351 West University Blvd, Cedar City, Utah 84720, USA; email: govedich@ suu.edu Daniel L. Graf [Chapter 11] The Academy of Natural Sciences, 1900 Benjamin Franklin Parkway, Philadelphia, Pennsylvania 19103, USA; email: [email protected] Roberto Guidetti  [Chapter 15] Department of Biology, University of Modena and Reggio Emilia, via Campi 213/D, 41125, Modena, Italy; email: roberto.guidetti@ unimore.it Ben Hanelt  [Chapter 10] Department of Biology, Univer­ sity of New Mexico, 163 Castetter Hall, Albuquerque, New Mexico 87131, USA; email: [email protected]

xi

xii

Brenda J. Hann  [Chapter 16] Department of Biological Sciences, W463 Duff Roblin, University of Manitoba, Winnipeg, Manitoba R3T 2N2, Canada; email: hann@ cc.umanitoba.ca Tom Hansknecht [Chapter 16] Barry A. Vittor and Associates, Inc., 8060 Cottage Hill Rd., Mobile, Alabama 36695, USA; email: [email protected] David J. Horne [Chapter 16] School of Geography, Queen Mary University of London, Mile End Road, London E1 4NS, United Kingdom; email: d.j.horne@ qmul.ac.uk Julian J. Lewis [Chapter 16] Lewis & Associates LLC, 17903 State Road 60, Borden, Indiana 47106-8608, USA; email: [email protected] Lawrence L. Lovell [Chapter 12] Research Associate, Polychaetous Annelids, Research & Collections, Natural History Museum of Los Angeles County, 900 Exposition Blvd., Los Angeles, California 90007, USA; email: ­[email protected] Tobias Kånneby [Chapter 7] Department of Zoology, Swedish Museum of Natural History, 10405, Stockholm, Sweden; email: [email protected] Renata Manconi [Chapter 3] Dipartimento di Scienze della Natura e del Territorio (DIPNET), Università di Sassari, Muroni 25, I-07100, Sassari, Italy; email: [email protected] William E. Moser  [Chapter 12] Smithsonian Institution, National Museum of Natural History, Department of Invertebrate Zoology, Museum Support Center, 4210 Silver Hill Road, Suitland, Maryland 20746, USA; email: [email protected]

Contributors to Volume II

Anna J. Phillips [Chapter 12] Smithsonian Institution, National Museum of Natural History, Department of Invertebrate Zoology, 10th and Constitution Ave, NW, Washington, DC 20560-0163, USA; email: phillipsaj@ si.edu George O. Poinar Jr.  [Chapter 9] Department of Zoology, Oregon State University, Corvallis, Oregon 97331, USA; email: [email protected] Wayne Price [Chapter 16] Department of Biology, University of Tampa, 401 W. Kennedy Blvd., Tampa, Florida 33606, USA; email: [email protected] Roberto Pronzato [Chapter 3] Dipartimento di Scienze della Terra, dell’Ambiente e della Vita (DISTAV), Università di Genova, Area Scientifico-Disciplinare 05 (Scienze biologiche), Settore BIO/05, Genova, Italy; email: [email protected] Lorena Rebecchi [Chapter 15] Department of Biology, University of Modena and Reggio Emilia, via Campi 213/D, 41125, Modena, Italy; email: lorena.rebecchi@ unimore.it Janet W. Reid  [Chapter 16] Virginia Museum of Natural History, 1001 Douglas Avenue, Martinsville, Virginia 24112, USA; email: [email protected] Vincent H. Resh  [Chapter 16] Department of Environmental Science, Policy, and Management, University of California, 305 Wellman Hall, Berkeley, California 94720, USA; email: [email protected] Dennis J. Richardson  [Chapter 12] School of Biological Sciences, Quinnipiac University, 275 Mt. Carmel Avenue, Hamden, CT 06518, USA; email: Dennis. [email protected]

Diane R. Nelson  [Chapter 15] Department of Biological Sciences, East Tennessee State University, Johnson City, Tennessee 37614-1710, USA; email: [email protected]

D. Christopher Rogers  [Chapters 1, 11, 16] Kansas Bio­ logical Survey and Biodiversity Institute, Higuchi Hall, University of Kansas, 2101 Constant Avenue, Lawrence, Kansas 66047, USA; email: [email protected]

Carolina Noreña [Chapter 5] Departamento Biodiversidad y Biología Evolutiva, Museo Nacional de Ciencias Naturales (CSIC), Madrid, España; email: norena@mncn. csic.es

S.S.S. Sarma [Chapter 8] Laboratorio de Zoología Acuática, Unidad de Morfología y Función, Facultad de Estudios Superiores, Universidad Nacional Autónoma de México, Av. de lo Barrios, no. 1, Los Reyes, Tlalnepantla, Edo. de Méx. C.P. 54090, México; email: [email protected]

Roy A. Norton  [Chapter 16] SUNY College of Environ­ mental Science and Forestry, 134 Illick Hall, 1 Forestry Drive, Syracuse, New York 13210, USA; email: ranorton@ esf.edu Alejandro Oceguera-Figueroa [Chapter 12] Laboratorio de Helmintologiá, Instituto de Biologiá, Universidad Nacional Autoñoma de México, Tercer circuito s/n, Ciudad Universitaria, Copilco, Coyoacán. A.P. 70-153, Distrito Federal, C. P. 04510, México; email: aoceguera@ ib.unam.mx

Andreas Schmidt-Rhaesa  [Chapter 10] Zoological Museum, University Hamburg, Martin Luther-King. Platz 3, 20146 Hamburg, Germany; email: [email protected] Hendrik Segers  [Chapter 8] School of Freshwater Biology, Belgian Biodiversity Platform, Royal Belgian Institute of Natural Sciences, Vautierstraat 29, B-1000, Brussels, Belgium; email: [email protected]

Contributors to Volume II

Alison J. Smith [Chapter 16] Department of Geology, Kent State University, Kent, Ohio 44242, USA; email: [email protected] Ian M. Smith  [Chapter 16] Systematic Acarology, Environ­ mental Health Program, Agriculture and Agri-Food Canada, K.W. Neatby Building, 960 Carling Ave., Ottawa, Ontario K1A 0C6, Canada; email: [email protected] T.W. Snell  [Chapter 8] School of Biology, Georgia Institute of Technology, 310 Ferst Drive, Atlanta, Georgia 30332, USA; email: [email protected] Malin Strand  [Chapter 6] The Swedish Species Information Centre, Swedish University of Agricultural Sciences, Uppsala, Sweden; email: [email protected] Per Sundberg  [Chapter 6] Department of Zoology, University of Gothenburg, P.O. Box 463, SE-405 30 Gothenburg, Sweden; email: [email protected] Christopher A. Taylor [Chapter 16] Curator of Fishes and Crustaceans, Prairie Research Institute, Illinois Natural History Survey, University of Illinois at UrbanaChampaign, 1816 S. Oak, Champaign, Illinois 61820, USA; email: [email protected] Roger F. Thoma  [Chapter 16] Midwest Biodiversity Insti­ tute, 4673 Northwest Parkway, Hilliard, Ohio 43026, USA; email: [email protected] James H. Thorp  [Chapters 1, 11, 12] Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, 2101 Constant Avenue, Lawrence, Kansas 66047, USA; email: [email protected]

xiii

Robert J. Van Syoc  [Chapter 16] California Academy of Sciences, Department of Invertebrate Zoology and Geology, 55 Music Concourse Drive, San Francisco, California 94118, USA; email: Bvansyoc@ calacademy.org L. Cristina de Villalobos  [Chapter 10] Facultad de Ciencias Naturales y Museo, Departamento de Invertebrados, Paseo del Bosque S/N 1900 La Plata, Argentina; email: [email protected] Robert L. Wallace [Chapter 8] Department of Biology, Ripon College, 300 Seward Street, Ripon, Wisconsin 54791, USA; email: [email protected] Elizabeth J. Walsh  [Chapter 8] Department of Biological Science, University of Texas at El Paso, 500 W. University Avenue, El Paso, Texas 79968, USA; email: [email protected] Alan Warren [Chapter 2] Department of Life Sciences, Natural History Museum, Cromwell Road, London SW7 5BD, United Kingdom; email: [email protected] Timothy S. Wood [Chapters 13, 14] Department of Biological Sciences, Wright State University, 3640 Colonel Glen Highway, Dayton, Ohio 45435, USA; email: [email protected] Fernanda Zanca [Chapter 10] Facultad de Ciencias Naturales y Museo, Departamento de Invertebrados, Paseo del Bosque S/N 1900 La Plata, Argentina; email: [email protected]

About the Editors

field and lab. While his research emphasizes aquatic invertebrates, he also studies fish ecology, especially as related to food webs. He has published more than hundred refereed journal articles, books, and chapters, including three singlevolume editions of Ecology and Classification of North American Freshwater Invertebrates (edited by J.H. Thorp and A.P. Covich) and the first volume (Ecology and General Biology) in the current fourth edition of Thorp and Covich’s Freshwater Invertebrates.

Dr. James H. Thorp has been a Professor in the Department of Ecology and Evolutionary Biology at the University of Kansas (Lawrence, KS, USA) and a Senior Scientist in the Kansas Biological Survey since 2001. Prior to returning to his alma mater, Prof. Thorp was a Distinguished Professor and Dean at Clarkson University, Department Chair and Professor at the University of Louisville, Associate Professor and Director of the Calder Ecology Center of Fordham University, Visiting Associate Professor at Cornell, and Research Ecologist at the University of Georgia’s Savannah River Ecology Laboratory. He received his Baccalaureate from the University of Kansas (KU) and both Masters and PhD degrees from North Carolina State. Those degrees focused on zoology, ecology, and marine biology with an emphasis on the ecology of freshwater and marine invertebrates. Dr. Thorp has been on the editorial board of three freshwater journals and is a former President of the International Society for River Science. He teaches freshwater, marine, and general ecological courses at KU, and his master’s and doctoral graduate students work on various aspects of the ecology of organisms, communities, and ecosystems in rivers, reservoirs, and wetlands. Prof. Thorp’s research interests and background are highly diverse and span the gamut from organismal biology to community, ecosystem, and macrosystem ecology. He works on both fundamental and applied research topics using descriptive, experimental, and modeling approaches in the

Dr. D. Christopher Rogers is a research zoologist at the University of Kansas with the Kansas Biological Survey and is affiliated with the Biodiversity Institute. He received his PhD degree from the University of New England in Armidale, NSW, Australia. Christopher specializes in freshwater crustaceans (particularly Branchiopoda and Decapoda) and the invertebrate fauna of seasonally astatic wetlands on a global scale. He has numerous peer reviewed publications in crustacean taxonomy and invertebrate ecology, as well as published popular and scientific field guides and identification manuals to freshwater invertebrates. Christopher is an Associate Editor for the Journal of Crustacean Biology and a founding member of the Southwest Association of Freshwater Invertebrate Taxonomists. He has been involved in aquatic invertebrate conservation efforts all over the world.

xv

Preface to the Fourth Edition

Those readers familiar with the first three editions of our invertebrate book (Ecology and Classification of North American Freshwater Invertebrates, edited by J.H. Thorp and A.P. Covich) will note that the fourth edition has expanded from a North American focus to worldwide coverage of inland water invertebrates. We gave our book series on inland water invertebrates the name Thorp and Covich’s Freshwater Invertebrates to: (1) associate present with past editions, unite current volumes, and link to future editions; (2) establish a connection between the ecological and general biology coverage in Volume I with the taxonomic keys in the remaining volumes; and (3) give credit to Professor Alan Covich for his work on the first three editions. For the sake of brevity, we refer to the current edition as T&C IV. Whether the fifth edition of T&C will ever appear is certainly problematic, but who knows! At present we are considering producing up to 11 volumes in the fourth edition. While I am the sole editor of the book series at this point, Christopher has been a major and highly valued partner in developing ideas for the fourth edition and is thus far an editor on the first three volumes (senior editor on the third). He will also play a major role in many of the remaining volumes because of his diverse and global knowledge of freshwater invertebrates, especially in the area of taxonomy. As we made significant progress on the first three volumes, we began contacting some potential coeditors and authors to develop volumes for other zoogeographic regions and negotiations with a few of those volumes are now underway. However, we are still seeking experts in fields of invertebrate taxonomy for various zoogeographic regions to serve as highly dependable coeditors, especially those who both work and live in the zoogeographic regions covered by the various future volumes.

Our concept for T&C IV included producing one book (Volume I, published in late 2014 with a 2015 copyright date) with 6 chapters on general environmental issues applicable to many invertebrates, followed by 35 chapters devoted to individual taxa at various levels (order to phylum, or even multiple phyla in the case of the protozoa). Volume I was designed both as an independent book on ecology and general biology of various invertebrate taxa and as a companion volume for users of the keys in the regional taxonomic volumes, thereby reducing the amount of information duplicated in the taxonomic volumes. The perhaps 10 taxonomic volumes to be published in the next decade or so will contain both keys for identifying invertebrates in specific zoogeographic regions and descriptions of detailed anatomical features needed to employ those keys. While the vast majority of authors in T&C editions I–III were from the United States or Canada, we attempted in T&C IV to attract authors from many additional countries in six continents. Although we largely succeeded in this goal, we expect the fifth edition of T&C—if it is ever published—to continue increasing the proportion of authors from outside North America as our books become better known internationally. Our goals for T&C IV are to improve the state of taxonomic and ecological knowledge of inland water invertebrates, help protect our aquatic biodiversity, and encourage more students to devote their careers to working with these fascinating organisms. These goals are especially important because the verified and probable losses of species in wetlands, ponds, lakes, creeks, and rivers around the globe exceed those in most terrestrial habitats. James H. Thorp

xvii

Preface to Volume II

This is the second volume of the fourth edition of Thorp and Covich’s Freshwater Invertebrates (T&C IV) and the first to focus almost exclusively on taxonomy. Information on the ecology and general biology of the groups can be found in Volume I (Ecology and General Biology, edited by Thorp & Rogers, 2015), the companion text for the current and all remaining books in this series. All taxonomic volumes (other than those focused exclusively on Hexapoda) are expected to consist of an introductory chapter, a chapter on protozoa (multiple kingdoms), and 14 chapters on individual phyla from Cnidaria to Arthropoda. Some of the chapters are very small (e.g., Chapter 14 on Entoprocta), whereas others are huge, especially Chapter 16 on Arthropoda. A typical chapter includes a short introduction, a brief discussion of limits to identification of taxa in that chapter, important information on terminology and morphology that is needed to use the keys, techniques for preparing and preserving material for identification (also covered in Volume I), the taxonomic keys, and a few references. In the large chapters on Mollusca (11), Annelida (12), and Arthropoda (16), different individuals have contributed separate sections, and thus there are multiple sections on introduction through keys and references. While this may confuse some readers, it has allowed us to gain contributions from an increased number of experts around the world. The multilevel keys are formatted to enable users to work easily at the level of their taxonomic expertise and the needs of their project. For that reason, we separated keys by major taxonomic divisions. For example, a student in a college course might work through one or more of the initial crustacean keys to determine the family in which a freshwater shrimp belongs. In contrast, someone working on an environmental monitoring project might need to identify a crayfish or crab to genus or even species, and thus would use the relevant, detailed keys that require more background experience. We also designed the keys, where possible, to proceed from a general to a specific character within a couplet.

We have asked authors to include only taxa that are recognized internationally by publication in reputable scientific journals that follow the International Code of Zoological Nomenclature. Thus, no taxa that have merely been proposed should be included even if they have been identified by the world’s expert on that group. “Common” species are not designated because a common species in one area may not be common in another, and this designation can lead to overly frequent and false identifications. Authors have been encouraged to end the keys at the point where further identification without genetic analysis is not practical or when it is clear that too many of the extant fauna have yet to be described in scientific publications. Users of these keys need to realize that taxonomy is a growing and vibrant field in which new taxa are being described and previously accepted relationships reevaluated. For some users, this volume may be sufficient for their needs, but for others, a companion text listing known species in a smaller geographic region may also be helpful. This edition is strongly focused on species found in fresh through saline inland waters, with a nonexclusive emphasis on surface waters, thereby reflecting the bias of existing scientific literature. Again, most estuarine and parasitic species are not covered in this book, but we do discuss species whose life cycle includes a free-living stage (e.g., Nematomorpha) and species that live in hard freshwaters through to brackish waters even though they may be normally associated with estuarine or marine habitats in some parts of their life cycles (e.g., some shrimp and crabs). It is our hope that scientists and students from around the world will benefit from this volume. Suggestions for improving future volumes are welcome. Editors James H. Thorp D. Christopher Rogers

xix

Acknowledgments for Volume II

Many people contributed to this volume in addition to the chapter authors and those acknowledged in individual chapters. We greatly appreciate all our colleagues who have contributed information, figures, or reviews to Volume II, and also thank those who provided similar services for the earlier editions, upon which the present book partially relies. We are again grateful to the highly competent people at Academic Press/Elsevier who helped in many aspects of the book’s

production from the original concept to the final marketing. In particular, we appreciate our association with Elsevier editors and production team including Candace Janco, Rowena Prasad, Laura Kelleher, and the entire United States and overseas production teams, especially Julia Haynes. James H. Thorp D. Christopher Rogers

xxi

INTRODUCTION

Chapter 1

Introduction1 James H. Thorp Kansas Biological Survey and Department of Ecology and Evolutionary Biology, University of Kansas, Lawrence, KS, USA

D. Christopher Rogers Kansas Biological Survey and Biodiversity Institute, University of Kansas, Lawrence, KS, USA

Chapter Outline Introduction to This Volume and Chapter 1 Components of Taxonomic Chapters How to Use This Volume

1 1 2

INTRODUCTION TO THIS VOLUME AND CHAPTER 1 This is the second volume in the fourth edition of Thorp and Covich’s Freshwater Invertebrates. Unlike the first three editions of Ecology and Classification of North American Freshwater Invertebrates (edited by Thorp and Covich in 1991, 2001, and 2010), the fourth edition has been split into multiple texts, with Volume I (Thorp & Rogers, 2015) providing global coverage of the ecology, general biology, phylogeny, and collection techniques for inland water invertebrates. Subsequent volumes provide keys to identify fauna in specific zoogeographic regions. This division of volumes enabled us to produce reasonable sized volumes at relatively moderate prices instead of publishing one massive, high priced tome. While some labs may have multiple copies of the “Keys to Fauna” in their region, we also recommend that they have at least one copy of Volume I, in order to obtain useful background information on each invertebrate group. The current chapter is organized into an introduction, a section explaining the organization of most taxonomic chapters, and a key to larger taxonomic groups. This chapter’s key is designed to help the reader locate the most pertinent chapter (important probably only for students and beginning taxonomists) and begin identifying organisms in their samples. Readers will note that chapters within and

Key to Kingdoms and Phyla in This Volume 3 References4

among volumes vary in specificity of their taxonomic keys. This reflects both the likely percent of the fauna that has been named and how easily taxa can be separated by alpha taxonomic methods and associated keys.

COMPONENTS OF TAXONOMIC CHAPTERS This volume is an identification manual to the inland water invertebrates of the Nearctic Region where we present information needed to diagnose and determine these organisms to various taxonomic levels. Other information concerning ecology, morphology, physiology, phylogeny, and both collecting and culturing techniques can be found in Volume I of this series. Each of the remaining 15 chapters in the current volume is limited to a single phylum, except Chapter 2’s coverage of multiple phyla of unicellular protists. Chapter 2 is designed for readers who only need general information about protists. We have attempted to include the following five sections in those chapters: (1) a brief introduction to the broader taxon; (2) a description of identification limitations for each taxon; (3) details of pertinent terminology and morphology; (4) information on preparing and preserving specimens for identification; and (5) taxonomic keys (separated by level of identification). A restricted number of especially pertinent references are given in each chapter following appropriate taxonomic sections. Readers can find a much more extensive list of references to their group in

1. This chapter was written to be a useful starting point for taxonomic volumes (II, III, etc.) in all zoogeographic regions. Consequently, there will be only minor differences among volumes. Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385028-7.00001-9 Copyright © 2016 Elsevier Inc. All rights reserved.

1

INTRODUCTION

2

Volume I (Chapters 3 and 7–41) along with more details on collecting, preparation, and preserving major taxa. Figures in each chapter are limited to those needed for effective use of the keys. For additional anatomical information, including figures, see the relevant chapter in Volume I.

HOW TO USE THIS VOLUME There is an old maxim that says “keys are written by people who do not need them for people who cannot use them.” We have made every effort to make these keys as user friendly as publication limitations would permit. Each section begins with a basic introduction to the morphology and terminology used in diagnosing the taxa of that section. Limitations to the current state of taxonomic knowledge are also presented so that the reader may gauge the reliability of the information presented. Only the established, peer reviewed scientific literature was used to define the taxonomic categories and epithets included. All names, as far as we are aware, conform to the International Code of Zoological Nomenclature (ICZN). All nomina and taxonomic arrangements used, as well as the rejection of old names was based on peer reviewed scientific literature. Names from unpublished manuscripts, dissertations, “in house” designations, or records that have not been validated are not acceptable. Provisional names and species designated “taxon 1” or “species 1” were not used unless they were previously recognized and accepted in the peer reviewed scientific literature (Richards & Rogers, 2011). No new species descriptions or previously unpublished taxonomic arrangements are presented. The keys are dichotomus (no triplets or quadruplets are used) and are hierarchical. Thus, for a given group, the first keys are to the highest taxonomic category. The second set of keys is to the next level, the third set to the level below that one, and so on, down to the lowest justifiable taxonomic level based on current knowledge of that group. This level is different for different groups depending upon the state of resolution in the scientific literature. Organisms not identifiable beyond a particular taxonomic level are left at that level. Properly prepared keys typically employ specific, primary, diagnostic characters. Older keys often use different characters than the more recent keys. This shift in primary characters results from systematists and taxonomists testing the importance of characters. The ultimate goal of the systematist is to ensure that the interpretation of which characters are important will converge with biological reality. To a non-taxonomist, this process may seem merely to be “lumping and splitting,” rather than the result of employing the scientific method to reveal natural relationships. Surprisingly, many users do not know how to interpret a dichotomus key, making the fundamental assumption that a

Thorp and Covich’s Freshwater Invertebrates

correct identification answer is always present in the key. This assumption generally takes one of the following three forms: 1. A  ll species are identifiable using a given key. Many new species have yet to be described, let alone discovered. Generalized geographic ranges are provided for most taxa presented herein, yet species ranges shrink, swell, and change elevation constantly, particularly as weather and climate patterns shift. Species disperse, colonize, and suffer stochastic local extinctions. In addition to these natural processes, some species are introduced intentionally or accidentally by humans, and sometimes their establishment allows other species to invade as well. 2. All variation is accounted for in the key. As stated above, identification keys use specific, primary, diagnostic characters. Problems in identification are compounded by taxa that: (a) have different character states at different times; (b) only have diagnostic characters at certain life stages or in certain genders; and/or (c) have severely truncated morphology (often due to lack of sexual selection) and lack morphological characters to separate the species. Furthermore, new variation within taxa is continually developing, and thus, one cannot assume that species are immutable or develop tools predicting those changes. 3. The key is a sufficient identification tool in and of itself. A key is just a tool. The fact that one has a bolt that needs removing and a wrench of the correct size does not mean that the bolt can be loosened. Similarly, identification keys are tools to aid in taxon identification. They are primarily tools to eliminate incorrect taxa from the range of possible choices, narrowing the field to the names that may be applicable. Keys are the process of elimination. The possibility that the specimen to be identified is new, a hybrid, anomalous, or a recent invasive colonist is always a possible answer. This is fundamental to using any identification key. Once one arrives at a name or group of possible names for a specimen in hand, the specimen should then be compared against descriptions, distribution maps, and figures of that and other taxa in that group. The descriptions, figures, and maps are other tools to be used in identification. Direct comparison of the specimen at hand with identified museum material or using molecular comparisons is also sometimes necessary for a correct identification. Species are not immutable, fixed in location and form. They change constantly and will continue to do so, confounding keys and any other identification method, such as trait tables, character matrices, or even genetic analyses. This is why biology is far behind physics in the development of unified theories: biology is far more complex than physics, as it involves more interacting parts and processes.

KEY TO KINGDOMS AND PHYLA IN THIS VOLUME A major change in the identification keys for our fourth edition has been to include multiple keys per chapter that generally start with a class level key and proceed to finer and finer divisions. These allow users to work at their

3

levels of interest, need, and skill without having to wade through extraneous taxa not in the direct line to the taxon of interest. The following key was derived in part from Chapter 1 in Volume I of the fourth edition. It is meant to allow you to move to the next level of keys, which will be in individual chapters.

Freshwater Invertebrate Kingdoms and Phyla 1 Multicellular, heterotrophic organisms as individuals or colonies (sometimes with symbiotic autotrophs)........................... kingdom Animalia .................................................................................................................................................................................................................................. 2 1’ Unicellular (or acellular) organisms present as individuals or colonies with nuclei irregularly arranged; heterotrophic and/or autotrophic; multiple phyla within the autotrophic protozoa phyla ....................................................................................... kingdom Protista [Chapter 2] 2(1) Radially symmetric or radially asymmetric organisms living individually or in colonies ............................................................................. 3 2’ Individuals bilaterally symmetric ................................................................................................................................................................... 4 3(2) Surface not porus; oral tentacles always present around a closeable mouth; colonial or single, mostly single polyp forms (primarily hydra) or rarely medusoid form (freshwater jellyfish); adults with a single central body cavity opening to the exterior and surrounded by cellular endoderm, acellular mesoglea, and cellular ectoderm ....................................................................................... phylum Cnidaria [Chapter 4] 3’ Surface porus; colonial; tentacles absent; no closable orifices; without discrete organs; cellular-level (or incipient tissue-level) construction; variable, non-distinct colony shapes, including encrusting, rounded, or digitiform growth forms; skeleton of individual siliceous spicules and a collagen matrix; internal water canal system; may contain symbiotic algae; the sponges ....................... phylum Porifera [Chapter 3] 4(2) Oral region with numerous tentacles or cilia distributed around the mouth; organism never with eversible jaws and never vermiform as adult ................................................................................................................................................................................................................ 5 4 Oral region with two or no tentacles, or tentacles behind the mouth ............................................................................................................. 7 5(4) Oral region with tentacles, organisms in gelatinoids or branching colonies .................................................................................................. 6 5’ Oral region ringed with cilia, muscular pharynx (mastax) with complex set of jaws; single free swimming, or semi-sessile living singly or in small colonies; wheel animals, or rotifers ...................................................................................................... phylum Rotifera [Chapter 8] 6(5) Oral tentacles (the lophophore) in a “U” or “horseshoe” shape around mouth; anus opens outside of lophophore; colonial animals, often in massive colonies attached to hard surfaces; true bryozoans ....................................................... phylum Ectoprocta (Bryozoa) [Chapter 13] 6’ Both mouth and anus open within lophophore; individual (non-colonial) animals with a calyx containing a single whorl of 8–16 ciliated tentacles ........................................................................................................................................................ phylum Entoprocta [Chapter 14] 7(4) Not with the combination of characteristics described below ........................................................................................................................ 8 7’ Small (50–800  μm), spindle- or tenpin-shaped, ventrally flattened with a more or less distinct head bearing sensory cilia; cuticle usually ornamented with spines or scales of various shapes; posterior of body often formed into a furca with distal adhesive tubes; gastrotrichs (pseudocoelomates) ...................................................................................................................................... phylum Gastrotricha [Chapter 7] 8(7) Anterior mouth and posterior anus present ..................................................................................................................................................... 9 8’ Flattened or cylindrical, acoelomate worms with only one, ventral digestive tract opening; sometimes with evident head; turbellarian flatworms (commonly called planaria, a non-specific, and usually incorrect name) .................................. phylum Platyhelminthes [Chapter 5] 9(8) Vermiform or not, eversible oral proboscis not present, although eversible jaws or other mouthparts may occur ...................................... 10 9’ Long, flattened, unsegmented worms with an eversible proboscis; ribbon worms ......................................... phylum Nemertea [Chapter 6] 10(9) Body not enclosed in a single, spiraled shell or in a hinged, bivalved shell; or if a bivalved shell is present, then animal has jointed legs .......... 11 10’ Soft-bodied coelomates whose viscera is covered (in freshwater species) by a single or dual (hinged), hard calcareous shell; with a ventral muscular foot; fleshy mantle covers internal organs; snails, clams, and mussels ........................................... phylum Mollusca [Chapter 11] 11(10) Segmented legs absent in all life stages; if jaws are present, then body with at least 20 segments .............................................................. 12 11’ Adults and most larval stages with legs; if larvae without legs or prolegs (some insects), then cephalic region with paired mandibles, or eversible head, always with less than 15 body segments .............................................................................................................................. 14 12(11) Organism vermiform, not segmented ............................................................................................................................................................ 13 12’ Organism vermiform or not, body segmented ................................................................................................. phylum Annelida [Chapter 12] 13(12) Body cylindrical, usually tapering at both ends; cuticle without cilia, often with striations, punctuations, minute bristles, etc.; 1 cm long (except family Mermithidae, 50%). The highest number of shared species with other biogeographic regions is with the Palaearctic (13 species) followed by Neotropical (10), Oriental (6), and Afrotropical and Australian (5). No species are shared with the Pacific Oceanic Island Region. Despite the extreme rarity of some species of Spongillida, no species are currently protected by laws or international conventions in the Nearctic. No reliable data exist, due to the scarcity of studies, on the conservation status

of freshwater sponges despite their key functional role in inland water ecosystems as both benthic active filter-feeders and as natural mesocosms for other taxa.

LIMITATIONS The Nearctic freshwater sponges are well known and readily identified to species level. Fundamental studies and monographs (Potts, 1887; Penney, 1960; Penney & Racek, 1968; Smith, 2001; Reiswig et al., 2010) provide exhaustive data on the inland Porifera fauna of the region, although further investigations in inadequately explored habitat/areas should yield new taxa. Spongillida families presently occurring in the Nearctic Region are the Metaniidae with two species and the Spongillidae with 28 species. However, middle Eocene fossil remains of the family Potamolepidae were recently reported from the Giraffe kimberlite maar in northern Canada (Pisera et al., 2013), and recent surveys in the poorly known Tennessee and southern Appalachians streams and rivers yelded the discovery of a rich and diversified sponge fauna,

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385028-7.00003-2 Copyright © 2016 Elsevier Inc. All rights reserved.

39

Phylum Porifera

Eunapius51 Pottsiela53 Stratospongilla53 Anheteromeyenia55 Radiospongilla56 Spongilla59 Trochospongilla63 Corvospongilla65 Heteromeyenia68 Racekiela74 Ephydatia76 Dosilia80

Introduction39 Limitations39 Terminology and Morphology 42 Material Preparation and Preservation 42 Acknowledgments43 Keys to Spongillida of the Nearctic Region 43 References46 Appendix 3.1 - Taxonomic Accounts of  Nearctic Porifera 48 Family Metaniidae48 Corvomeyenia48 Family Spongillidae49 Duosclera49

40

Thorp and Covich’s Freshwater Invertebrates

Phylum Porifera FIGURE 3.1  All Nearctic species of sponges. Synthetic representation of the spicular complements of Nearctic freshwater sponge species. Drawings were made from the most authoritative sources available. Dimension scales are most precise as possible. Modified from Reiswig et al., 2010.

Chapter | 3  Phylum Porifera

41

TABLE 3.1  Checklist and Geographic Range of Nearctic Freshwater Sponge Species SPONGILLIDA Manconi & Pronzato, 2002 Metaniidae Volkmer-Ribeiro, 1986 Corvomeyenia Weltner, 1913

NA-NT

 *C. carolinensis Harrison 1971

NA

 *C. everetti (Mills, 1884)

NA

Anheteromeyenia Schröder, 1927

NA-NT

 *A. argyrosperma (Potts, 1880)

NA

Corvospongilla Annandale, 1911

PA-NA-NT-AT-OL

 *C. becki Poirrier, 1978

NA

 *C. novaeterrae (Potts, 1886)

NA

Dosilia Gray, 1867

OL-AT-NA-NT

  D. palmeri (Potts, 1885)

NA-NT

 *D. radiospiculata (Mills, 1888)

NA

Duosclera Reiswig & Ricciardi, 1993

NA

 *D. mackayi (Carter, 1885)

NA

Ephydatia Lamouroux, 1816

PA-NA-AT-OL-AU-NT-PAC

  E. fluviatilis (Linnaeus, 1759)

PA-NA-AT-OL-AU

 *E. millsi (Potts, 1887)

NA

  E. muelleri (Lieberkuhn, 1855)

PA-NA

 *E. subtilis Weltner, 1895

NA

Eunapius Gray, 1867

PA-AT-OL-AU-NA-NT

  E. fragilis (Leidy, 1851)

PA-NA-AT-NT-OL-AU

Heteromeyenia Potts, 1881

NA-PA-NT-AU-PAC

  H. baileyi (Bowerbank, 1863)

NA-PA-NT-PAC

 *H. latitenta (Potts, 1881)

NA

 *H. longistylis Mills, 1884

NA

 *H. tentasperma (Potts, 1880)

NA

 *H. tubisperma (Potts, 1881)

NA

Pottsiela Volkmer-Ribeiro et al., 2010

NA-PA

  P. aspinosa Potts, 1880

NA-PA

Racekiela Bass & Volkmer-Ribeiro, 1998

PA-NA-NT

 *R. biceps (Lindenschmidt, 1950)

NA

  R. ryderi (Potts, 1882)

PA-NA-NT

Radiospongilla (Penney & Racek, 1968)

AU-PAC-NT-PA-AT-OL-NA

  R. cerebellata (Bowerbank, 1863)

AU-AT-OL-PA-NA

  R. crateriformis (Potts, 1882)

NA-NT-PA-AU-OL

Spongilla Lamarck, 1816

PA-NA-OL-AT-AU-NT

Phylum Porifera

Spongillidae Gray, 1867

Continued

42

Thorp and Covich’s Freshwater Invertebrates

TABLE 3.1  Checklist and Geographic Range of Nearctic Freshwater Sponge Species—cont’d   S. alba Carter, 1849

OL-AT-AU-NT-PA-NA

  S. cenota Penney & Racek, 1968

NT-NA

  S. lacustris (Linnaeus, 1759)

PA-NA

 *S. wagneri Potts, 1889

NA

Stratospongilla Annandale, 1909

OL-AT-PA-AU-NA

 *S. penneyi (Harrison, 1979)

NA

Trochospongilla Vejdowsky, 1883

PA-NA-NT-OL-AU-AT

  T. horrida (Weltner, 1893)

PA-NA

  T. leidyi (Bowerbank, 1863)

NA-NT

 *T. pennsylvanica (Potts, 1882)

NA

*Nearctic endemics. OL = Oriental Region; NA = Nearctic Region; PA = Palaearctic Region; AU = Australian Region; AT = Afrotropical Region; NT = Neotropical Region; PAC = Pacific Islands Region.

Phylum Porifera

including a new genus and a new species of Potamolepidae (Copeland et al., 2015). A few Nearctic Spongillida species (e.g., Radiospongilla cerebellata, R. crateriformis, Spongilla alba, S. lacustris, and Ephydatia fluviatilis) are widespread, but the majority of taxa show a restricted geographic range. Smith (2001) reported that several species have become locally extint, mainly due to the pollution. The following keys focus on the distribution of sponges within the Nearctic bioregion but include some overlap with the Palaearctic (Beringia, Greenland) and the Neotropical (Baja Peninsula at Tropic of Cancer, Mexican Lowlands, Southern Florida Peninsula). We also refer to species distributions reported in Bânârescu (1992) and Manconi & Pronzato (2007, 2008, 2009).

TERMINOLOGY AND MORPHOLOGY Nearctic freshwater sponges range in size from 1 mm in length ........................................................................................................................................................ Hydridae [p. 88] 2’ Medusae, or polyps 11 μ in length; embryotheca either not tiled or not roughly spherical ......................................... 2 1’ Polyps green; stenotele nematocysts 9–11 μ in length; surface of embryotheca roughly spherical and cobbled with tiles (Fig. 4.3 A) ............................................................................................................................................................................... viridissima group 2(1) Holotrichous isorhiza nematocysts narrowly oval or slipper-shaped, less than half as wide as long (Fig. 4.4 B); embryotheca not flattened ......................................................................................................................................................................................................................... 3 2’ Holotrichous isorhiza nematocysts broadly lemon-shaped, at least half as wide as long (Fig. 4.5 B); embryotheca flattened against a substratum (Fig. 4.3 C)...................................................................................................................................................................... braueri group 3(2) Stenotele nematocysts appear all the same length (within 1 μm); embryotheca smooth and thin (Fig. 4.3 B) ........................ oligactis group 3’ Stenotele nematocysts varied in length, typically about 13–18 μ; embryotheca thick with numerous radial spines (Fig. 4.3 D) (one or several confused species. See section on limitations above) ................................................................................................................. vulgaris group

Hydra Species Groups: Viridissima: Hydra: Species Probably all green hydras in our area represent H. viridissima. H. hadleyi is based on the character of an extra chamber in the embryotheca, but this is now known to be present in H. viridissima. 1 Embryotheca with one large empty chamber proximal to the embryo ............................................................. Hydra hadleyi (Forrest, 1959) 1’ Embryotheca with only a narrow chamber proximal to embryo .................................................................... Hydra viridissima Pallas, 1766

Hydra Species Group: Oligactis: Hydra: Species 1. Internal tubule of the holotrichous isorhiza nematocysts with about 3 prominent transverse coils (Fig. 4.4 B) ........................................ ................................................................................................................................................................... Hydra canadensis Rowan, 1930 1’ Internal tubule of the holotrichous isorhiza nematocysts plied longitudinally throughout (Fig. 4.5 D) ............. Hydra oligactis Pallas, 1766

Hydra Species Group: Braueri: Hydra: Species

Phylum Cnidaria

1 Desmoneme nematocyst length 7 μm, approximately as long as the holotrichous isorhiza nematocyst (Fig. 4.5 B) ..................................... ........................................................................................................................................................... Hydra hymanae Hadley & Forrest, 1949 [Canada; northern USA] 2(1) Budding polyps 3 mm in length .................................................................................................................... Hydra utahensis Hyman, 1931 [Canada; northern USA]

Chapter | 4  Phylum Cnidaria

89

3(2) Embryotheca sometimes spherical ................................................................................................................ Hydra lirosoma Campbell, 1987 [USA: Georgia] 3’ Embryotheca always flattened ............................................................................................................................ Hydra minima Forrest, 1963 [USA: New Jersey]

Cnidaria: Hydrozoa: Olindiidae: Species 1 Medusae, or polyps without tentacles ............................................................................................. Craspedacusta sowerbii Lankester, 1880 1’ Polyps with tentacles consisting of a single cell each ................................................................... Calpasoma dactyloptera Fuhrmann, 1939

Campbell, R.D. 1987. A new species of Hydra (Cnidaria: Hydrozoa) from North America with comments on species clusters within the genus. Zoological Journal of the Linnean Society of London 91: 253–263. Cartwright, P., N.M. Evans, C.W. Dunn, A.C. Marques, M.P. Miglietta, P. Schuchert & A.G. Collins. 2008. Phylogenetics of Hydroidolina (Hydrozoa: Cnidaria). Journal of the Marine Biological Association of the United Kingdom 88: 1663–1672. Collins, A.G., B. Bentlage, A. Lindner, D. Lindsay, S.H.D. Haddock, G. Jarms, J.L.Norenburg, T. Jankowski & P. Cartwright. 2008.

Phylogenetics of Trachylina (Cnidaria: Hydrozoa) with new insights on the evolution of some problematical taxa. Journal of the Marine Biological Association of the United Kingdom 88: 1673–1685. Jankowski, T., A.G. Collins & R. Campbell. 2008. Global diversity of inland water cnidarians. Hydrobiologia 595: 35–40. Martinez, D.E., A.R. Iñiguez, K.M. Percell, J.B. Willner, J. Signorovitch & R.D. Campbell. 2010. Phylogeny and biogeography of Hydra (Cnidaria: Hydridae) using mitochondrial and nuclear DNA sequences. Molecular Phylogenetics and Evolution 57: 403–410. Schulze, P. 1917. Neue Beiträge zu einer Monographie der Gattung Hydra. Archiv für Biontologie 4: 31–119.

Phylum Cnidaria

REFERENCES

Phylum Platyhelminthes

Chapter 5

Phylum Platyhelminthes Carolina Noreña Departamento Biodiversidad y Biología Evolutiva, Museo Nacional de Ciencias Naturales (CSIC), Madrid, Spain

Cristina Damborenea, Francisco Brusa División Zoología Invertebrados, Museo de La Plata, CONICET, La Plata, Argentina

Chapter Outline Introduction91 Limitations91 Terminology and Morphology 92 Material Preparation and Preservation 92

INTRODUCTION

Keys to Platyhelminthes 96 Acknowledgments109 References109

“Turbellaria” is a class constituted by a very heterogeneous group of orders. Besides the body plan, all orders share a free-living life style, similar features of the body wall (simple epithelium, muscle network, absence of cuticle) and regeneration based on stem cell-like neoblasts (Tyler & Hooge, 2004). These characters justified the grouping of three main clades: Acoelomorpha, Catenulida, and Rhabditophora (Tyler & Hooge, 2004). Recently, the Acoelomorpha (Acoela and Nemertodermatida) were placed outside the Platyhelminthes, with the status of a new phylum, because the singular molecular composition of the mitochondrial and nuclear genes and morphological characters such as duet-spiral cleavage, bi-spiral segmentation and an epithelium with an own root system network (Ruiz-Trillo et al., 2004; Baguñà & Riutort, 2004a,b).

The flatworms of the phylum Platyhelminthes comprise free-living (“Turbellaria”) and obligate parasitic organisms (Monogenea, Digenea, Aspidogastrea, and Cestoda, today grouped in Neodermata). “Turbellaria” includes an amazing variety of forms, but built in a similar way. All have the following characteristics: bilateral symmetry, organs embedded in a solid cellular matrix (the parenchyma), a sac-like gut without an anus, a nervous system with an anterior “brain” and lateral nerve chords, and internal fluids that are regulated by protonephridia. Most are cross-­ fertilizing hermaphrodites, with morphologically diverse structures like sclerotic parts of the male copulatory organ that forms stylets of different shape and complexity, and with endolecithal or ectolecithal eggs. Variation also can be found in the location and shape of the pharynx along the LIMITATIONS main body-axis. Most “Turbellaria” are small with a body size of around The keys were built with users in mind who are not trained 1 mm (microturbellaria), but others, such as Tricladida and in the characteristics of species of freshwater “Turbellaria.” Polycladida, have a body-size of 5–10 cm (macroturbel- They use characters that can be easily seen, essentially in laria). They show a worldwide distribution and inhabit most whole mounts of living animals by inexperienced observkinds of freshwater, brackish, and marine habitats. ers and applied at higher systematic levels. In lower levels, Based on their simple body structure, the “Turbel- however, identification requires more detailed observalaria” has been considered one of the most basal bilateria tions. Once the identification is reached, it must be checked (Littlewood & Bray, 2001). This hypothesis is supported by against the original description of the taxon or later the results obtained from molecular phylogenetic analysis descriptions in monographs such as Ferguson (1954) and (Ruiz-Trillo et al., 1999, 2004; Baguñà & Riutort, 2004a). the website created by Shärer (2014) for Macrostomida. Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385028-7.00005-6 Copyright © 2016 Elsevier Inc. All rights reserved.

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Readers should also consult the following texts: Gilbert (1938) and Houben et al. (2014) for Phaenocora, Kenk (1974, 1989) for Tricladida, Luther (1955) for Dalyelliidae, Luther (1960) for Catenulida, Macrostomida, Lecithoepitheliata, Prolecithophora, and Proseriata, Luther (1963) for Typhloplanida, Nuttycombe & Waters (1938) for Stenostomidae (Catenulida), and Nuttycombe (1956) for Catenulidae. More general information on turbellarians is available in Cannon (1986), Tyler et al. (2006–2015), and Kolasa & Tyler (2010). Table 5.1 lists the freshwater turbellarian species known in the Nearctic Region.

TERMINOLOGY AND MORPHOLOGY The following terminology is intended to assist in understanding the morphology and the use of turbellarian keys included in the chapter. It is based primarily on Cannon (1986), Rieger et al. (1991), and Richter et al. (2010). Adenodactyl: auxiliary glandular bulb in male reproductive system of some groups, mainly Tricladida. Adhesive glands: cell glands found in different parts of the body, which secrete adhesive substances. Ascus: glandular pocket near the gonopores. Ciliated pits: sensory structures (e.g., anterior end of Stenostomidae). Copulatory organ: the terminal part of the male reproductive system to deliver the sperm into the body of a partner at copulation. It is often bulbous (copulatory bulb), containing the sperm duct and elements of the prostate glands. It is a penis (penis papilla) if it is capable of protrusion but not armed with hard structures. It may be enclosed in an inner chamber (penis pocket) the walls of which can form a conical prominence that may serve to guide the penis (a penis sheath). The penis is said to be armed when a “cuticular” or “sclerotic” structure is present. A stylet is a single sclerotic tube or channel of variable shape; a cirrus is an eversible male duct; often spiny. Excretophore: large vacuolated cells of the intestine epithelium. Handles: structures in the basal part of the stylet. Light refracting bodies: unpigmented refracting bodies found in species of Stenostomidae. These bodies are formed by a variable number of spherical granules; some species have fewer than five and others more than 15. The light refracting bodies are frequently associated with the posterior cerebral lobes, or sometimes with the anterior lobes. Generally only one pair is found, although in some cases, more pairs are present. Neoblast: pluripotent cells responsible for the normal replacement of cells, cell substitution during regeneration, and stem cells for the gonads. Pharynx: the glandulo-muscular structure between mouth and intestine inwards. It is used to capture prey and may be of considerable taxonomic value. The main terms for practical use are: simplex (simple)—a ciliated mouth tube; plicatus (folded/plicate)—an annular fold which is variable in length, lacks a delimiting septum, and has a large protrusion capacity; bulbosus (bulbous)—a

Thorp and Covich’s Freshwater Invertebrates

glandulo-muscular bulb delimited by a more or less muscular septum, only the end of the pharynx may be protruded; it could be rosulatus (rosulate)—bulbous and usually globular and dorso-ventrally oriented; or doliiformis (doliiform)—bulbous and usually barrel-shaped and horizontally (fronto-caudally) oriented; or variabilis (variable)—bulbous, but with variable shape and a weakly differentiated septum (see Volume I, Chapter 10). Proboscis: anterior protrusible organ, which is a notable feature of the rhabdocoel group Kalyptorhynchia. Prostatic vesicle: part of the male reproductive system. Structure containing gland cells and its secretion is combined with the sperm during their release. Prostomium: body region anterior to mouth; usually containing the brain. Protonephridium: organ containing one or more flame cells and connecting ducts, which presumably functions in an osmoregulatory fashion. Rhabdite: rod-shaped bodies found in the epidermal cells or below the ectoderm. They are characteristic for Platyhelminthes. Statocyst: an orientation organ of the nervous system; it contains a space enclosing a granule (the statolith) and is surrounded by sensory tissue. Vitellaria: gland cells that secrete nutrient material which is included into egg capsules. Zooid: individuals formed by paratomy.

MATERIAL PREPARATION AND PRESERVATION In vivo observations are very important for the study of turbellarians because certain structures are only visible in living specimens. Once collected, the specimens will need to undergo different group-specific treatments for subsequent identification and preservation. For large planarians, external characteristics are easily observable under stereomicroscope. Specimens should then be fixed (in Steinman’s, 10% formaldehyde or Bouin fixative) for later histological analysis. For microturbellarians, the specimens must first be observed live and their size, shape, and color noted. After placing the individual on a slide with a drop of water and either Vaseline or plasticine smeared on the coverslip edges, slowly squash it with the coverslip. The water excess can be removed with filter paper. Diagnostic structures of the internal organs of the transparent microturbellarians can be observed with this method. Some specimens must be fixed and preserved in polyvinyl lactophenol, which makes them more transparent, thus allowing the observation of sclerotized diagnostic structures (e.g., spines, stylets, and the cirrus). Microturbellarians can be fixed in a variety of conventional fixation fluids (e.g., Bouin, 5% formaldehyde) prior to being processed for histological analysis of the internal structures of diagnostic importance. With few exceptions, the characters used in the following keys are viewable by transparency (squash methodology).

93

TABLE 5.1  List of Neartic Species of Turbellarians Catenulida

  Stenostomum ventronephrium Nuttycombe, 1932

  Family Catenulidae

  Stenostomum virginianum Nuttycombe, 1931

  Catenula confusa Nuttycombe, 1956

Macrostomida

  Catenula lemnae Duges, 1832

  Family Macrostomidae

  Catenula leptocephala Nuttycombe, 1956

  Macrostomum acutum Ax, 2008

  Catenula sekerai Beauchamp, 1919

  Macrostomum appendiculatum Fabricius, 1826

  Catenula turgida (Zacharias, 1902)

  Macrostomum bellebaruchae Ax, 2008

  Catenula virginia Kepner & Carter, 1930

  Macrostomum bicurvistyla Armonies & Hellwig, 1987

  Family Chordariidae

  Macrostomum carolinense Ferguson, 1940

  Chordarium europaeum Schwank, 1980

  Macrostomu collistylum Ferguson, 1939

  Family Stenostomidae

  Macrostomum curvistylum Ferguson, 1939

  Myostenostomum gigerium (Kepner & Carter, 1931)

  Macrostomum curvituba Luther, 1947

  Rhynchoscolex platypus Marcus, 1945

  Macrostomum frigorophilum Ferguson, 1940

  Rhynchoscolex simplex Leidy, 1851

  Macrostomum gilberti Ferguson, 1939

  Stenostomum anatirostrum Marcus, 1945

  Macrostomum glochistylum Ferguson, 1939

  Stenostomum anops Nuttycombe & Waters, 1938

  Macrostomum granulophorum Ferguson, 1940

  Stenostomum arevaloi Gieysztor, 1931

  Macrostomum hystricinum Beklemischev, 1951

  Stenostomum beauchampi Papi, 1967

  Macrostomum lewisi Ferguson, 1939

  Stenostomum bicaudatum Kennel, 1888

 *Macrostomum norfolkensis Jones & Ferguson, 1940

  Stenostomum brevipharyngium Kepner & Carter, 1931

  Macrostomum ontarioense Ferguson, 1943

  Stenostomum ciliatum Kepner & Carter, 1931

  Macrostomum orthostylum Braun, 1885

  Stenostomum cryptops Nuttycombe & Waters, 1935

  Macrostomum phillipsi Ferguson & Stirewalt, 1938

  Stenostomum glandulosum Kepner & Carter, 1931

  Macrostomum recurvostylum Ferguson, 1940

  Stenostomum grande Child, 1902

  Macrostomum reynoldsi Ferguson, 1939

  Stenostomum kepneri Nuttycombe & Waters, 1938

  Macrostomum riedeli Ferguson, 1939

  Stenostomum leucops (Duges, 1828)

  Macrostomum ruebushi Ferguson, 1940

  Stenostomum mandibulatum Kepner & Carter, 1931

  Macrostomum tenuicauda Luther, 1947

  Stenostomum membranosum Kepner & Carter, 1931

 *Macrostomum thermophilum Riedel, 1932

  Stenostomum occultum Kolasa, 1971

  Macrostomum schmitti Hayes & Ferguson, 1940

  Stenostomum pegephilum Nuttycombe & Waters, 1938

  Macrostomum sensitivum Silliman, 1884

  Stenostomum predatorium Kepner & Carter, 1931

  Macrostomum shenandoahense Ferguson, 1940

  Stenostomum pseudoacetabulum Nuttycombe & Waters, 1935

  Macrostomum tennesseensis Ferguson, 1939

  Stenostomum saliens Kepner & Carter, 1931

  Macrostomum truncatum Ferguson, 1940

  Stenostomum simplex Kepner & Carter, 1931

  Macrostomum tuba Graff, 1882

  Stenostomum sphagnetorum Papi in Luther, 1960

  Macrostomum vejdovskyi Ferguson, 1940

  Stenostomum temporaneum Kolasa, 1981

  Macrostomum virginianum Ferguson, 1937

  Stenostomum tuberculosum Nuttycombe & Waters, 1938

  Family Microstomidae

  Stenostomum uronephrium Nuttycombe, 1931

  Microstomum bispiralis Stirewalt, 1937 Continued

Phylum Platyhelminthes

Chapter | 5  Phylum Platyhelminthes

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TABLE 5.1  List of Neartic Species of Turbellarians—cont’d  *Microstomum caudatum Leidy, 1851

  Fulinskiella bardeaui (Steinböck, 1926)

  Microstomum lineare (Müller, 1773)

 *Gieysztoria blodgetti (Silliman, 1884)

 *Microstomum philadelphicum Leidy, 1852

  Gieysztoria choctaw Van Steenkiste, Gobert & Artois, 2011

Lecithoepitheliata

  Gieysztoria cuspidata (Schmidt, 1861)

  Family Prorhynchidae

  Gieysztoria dodgei (Graff, 1911)

  Geocentrophora applanata (Kennel, 1888)

 *Gieysztoria eastmani (Graff, 1911)

  Geocentrophora baltica (Kennel, 1883)

  Gieysztoria minima (Riedel, 1932)

  Geocentrophora cavernicola Carpenter, 1970

  Gieysztoria ornata (Hofsten, 1907)

  Geocentrophora marcusi Darlington, 1959

  Gieysztoria pavimentata (Beklemischew, 1926)

  Geocentrophora sphyrocephala de Man, 1876

 *Gieysztoria pseudoboldgetti Luther, 1955

  Prorhynchus stagnalis Schultze, 1851

  Gieysztoria triangulata (Robeson, 1931)

Prolecithophora

  Microdalyellia abursalis (Ruebush, 1937)

  Family Plagiostomidae

 *Microdalyellia armigera (Schmidt, 1861)

  Hydrolimax bruneus Girard, 1891

  Microdalyellia circulobursalis (Ruebush, 1937)

  Hydrolimax grisea Haldeman, 1843

 *Microdalyellia deses (Riedel, 1932)

 *Plagiostomum planum Silliman, 1885

  Microdalyellia fairchildi (Graff, 1911)

Bothrioplanida

  Microdalyellia gilesi Jones & Hayes, 1941

  Family Bothrioplanidae   Bothrioplana semperi Braun, 1881 Proseriata   Family Coelogynoporidae   Coelogynopora falcaria Ax & Sopott-Ehlers, 1979   Family Monocelididae   Monocelopsis carolinensis Ax, 2008

 *Microdalyellia groenlandica (Riedel, 1932)   Microdalyellia mohicana (Graff, 1911)  *Microdalyellia rheesi (Graff, 1911)  *Microdalyellia rochesteriana (Graff, 1911)   Microdalyellia rossi (Graff, 1911)   Microdalyellia ruebushi Luther, 1955

  Family Otomesostomidae

  Microdalyellia schockaerti Willems, Artois, Jocque, & Brendonck, 2007

  Otomesostoma auditivum (Du Plessis, 1874)

  Microdalyellia sillimani (Graff, 1911)

Rhabdocoela

  Microdalyellia tennesseensis (Ruebush & Hayes, 1939)

Dalyellioida

  Microdalyellia virginiana (Ruebush, 1937)

  Family Provorticidae

  Pseudodalyellia alabamensis Van Steenkiste, Gobert & Artois, 2011

  Provortex virginiensis Ruebush & Hayes, 1939   Vejdovskya pellucida (Schultze, 1851)

Typhloplanoida

  Family Dalyelliidae

  Family Typhloplanidae

 *Castrella cylindrica Riedel, 1932

  Acrochordonoposthia conica Reisinger, 1924

 *Castrella graffi Hayes, 1945

  Adenoplea nanus Sayre & Wergen, 1994

  Castrella groenlandica Riedel, 1932

  Amphibolella segnis Findenegg, 1924

  Castrella pinguis (Silliman, 1884)

  Ascophora elegantissima Findenegg, 1924

  Castrella truncata (Abidgaard, 1789)

  Bryoplana xerophila Van Steenkiste, Davison & Artois, 2010

 *Dalyellia alba Higley, 1918

  Bothromesostoma personatum (Schmidt, 1848)

  Dalyellia idahoensis Knapp, 1954

  Castrada affinis Hofsten, 1907

  Dalyellia viridis (Shaw, 1791)

  Castrada borealis Steinböck, 1931

95

TABLE 5.1  List of Neartic Species of Turbellarians—cont’d   Castrada hofmanni Braun, 1885

  Tetracelis marmorosa (Müller, 1773)

  Castrada inermis Hofsten, 1911

  Typhloplana minima (Fuhrmann, 1894)

  Castrada libidinosa Hofsten, 1916

  Typhloplana viridata (Abildgaard, 1789)

  Castrada luteola Hofsten, 1907

  Typhloplanella halleziana (Vejdovsky, 1880)

  Castrada lutheri Kepner, Stirewalt & Ferguson, 1939

Kalyptorhynchia

  Krumbachia minuta Ruebush, 1938

  Family Polycistididae

  Krumbachia virginiana (Kepner & Carter, 1931)

  Gyratrix hermaphroditus Graff, 1831

  Limnoruanis romanae Kolasa, 1977

  Opisthocystis goettei (Bresslau, 1906)

  Mesocastrada fuhrmanni Volz, 1898

Tricladida

  Mesostoma angulare Higley, 1918

  Familia Dendrocoelidae

  Mesostoma arctica Hyman, 1938

  Procotyla fluviatilis Leidy, 1857

  Mesostoma californicum Hyman, 1957

  Procotyla typhlops Kenk, 1935

  Mesostoma columbianum Hyman, 1939

  Dendrocoelopsis alaskensis Kenk, 1953

  Mesostoma craci Schmidt, 1858

  Dendrocoelopsis americana (Hyman, 1939)

  Mesostoma curvipenis Hyman, 1955

  Dendrocoelopsis hymanae Kawakatsu, 1968

  Mesostoma ehrenbergii (Focke, 1836)

  Dendrocoelopsis piriformis Kenk, 1953

  Mesostoma georgianum Darlington, 1959

  Dendrocoelopsis vaginata Hyman, 1935

  Mesostoma macropenis Hyman, 1939

  Familia Dugesiidae

  Mesostoma macroprostatum Hyman, 1939

  Cura foremanii (Girard, 1852)

  Mesostoma platygastricum Hofsten, 1924

  Girardia arizonensis Kenk, 1975

  Mesostoma vernale Hyman, 1955

  Girardia dorotocephala (Woodworth, 1897)

  Microcalyptorhynchus virginianus Kepner & Ruebush, 1935

  Girardia jenkinsae (Benazzi & Gourbault, 1977)

  Olisthanella coeca (Silliman, 1885)

  Girardia tigrina (Girard, 1850)

  Opistomum pallidum Schnidt, 1848

  Schmidtea polychroa (Schmidt, 1861)

 *Phaenocora agassizi Graff, 1911

  Familia Kenkiidae

  Phaenocora aglobulata Houben, van Steenkiste & Artois, 2014

  Kenkia glandulosa (Hyman, 1956)

  Phaenocora falciodenticulata Gilbert, 1938

  Kenkia lewisi (Kenk, 1975)

  Phaenocora gilberti Houben, van Steenkiste & Artois, 2014

  Kenkia rhynchida Hyman, 1937

  Phaenocora highlandense Gilbert, 1935

  Sphalloplana (Speophila) buchanani (Hyman, 1937)

  Phaenocora kepneri Gilbert, 1935

  Sphalloplana (Speophila) chandleri Kenk, 1977

  Phaenocora lutheri Gilbert, 1937

  Sphalloplana (Speophila) hoffmasteri (Hyman, 1954)

 *Phaenocora megacephala (Higley, 1918)

  Sphalloplana (Speophila) hubrichti (Hyman, 1945)

  Phaenocora sulfophila (Gilbert, 1938)

  Sphalloplana (Speophila) hypogeal Kenk, 1984

  Phaenocora virginiana Gilbert, 1935

  Sphalloplana (Speophila) pricei (Hyman, 1937)

  Prorhynchella minuta Ruebush, 1939

  Sphalloplana (Speophila) weingartneri Kenk, 1970

  Protoascus wisconsinensis Hayes, 1941

  Sphalloplana (Speophila) californica Kenk, 1977

  Rhynchomesostoma rostratum (Müller, 1773)

  Sphalloplana (Speophila) consimilis Kenk, 1977

  Strongylostoma elongatum Hofsten, 1907

  Sphalloplana (Speophila) culveri Kenk, 1977

  Strongylostoma gonocephalum (Silliman, 1884)

  Sphalloplana (Speophila) evaginata Kenk, 1977

  Styloplanella strongylostomoides Findenegg, 1924

  Sphalloplana (Speophila) georgiana Hyman, 1954 Continued

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TABLE 5.1  List of Neartic Species of Turbellarians—cont’d   Sphalloplana (Speophila) percoeca (Packard, 1879)

  Phagocata nordeni Kenk, 1977

  Sphalloplana (Speophila) subtilis Kenk, 1977

  Phagocata oregonensis Hyman, 1963

  Sphalloplana (Polypharyngea) mohri Hyman, 1938

  Phagocata notorchis Kenk, 1987

  Familia Planariidae

  Phagocata procera Kenk, 1984

  Hymanella retenuova Castle, 1941

  Phagocata pygmaea Kenk, 1987

  Paraplanaria dactyligera (Kenk, 1935)

  Phagocata spuria Kenk, 1987

  Paraplanaria occulta (Kenk, 1969)

  Phagocata tahoena Kawakatsu, 1968

  Phagocata angusta Kenk, 1977

  Phagocata velata (Stringer, 1909)

  Phagocata bulbosa Kenk, 1970

  Phagocata vernalis (Kenk, 1944)

  Phagocata bursaperforata Darlington, 1959

  Phagocata virilis Kenk, 1977

  Phagocata carolinensis Kenk, 1979

  Phagocata woodworthi Hyman, 1937

  Phagocata fawcetti Ball & Gourbault, 1975

 *Polycelis coronata coronata (Girard, 1891)

  Phagocata gracilis (Haldeman, 1840)

  Polycelis coronata monticola Kenk & Hampton, 1982

  Phagocata hamptonae Kenk, 1982

  Polycelis coronata brevipenis Kenk, 1972

  Phagocata holleri Kenk, 1979

  Seidlia remota (Smith, 1988)

  Phagocata morgani morgani (Stevens & Boring, 1906)

  Seidlia sierrensis (Kenk, 1973)

  Phagocata morgani polycelis (Kenk, 1935)

The arrangement of orders and families follows the taxonomic listing of Tyler et al. (2006–2015). *Indicates a doubtful species.

  Phagocata nivea Kenk, 1953

KEYS TO PLATYHELMINTHES Platyhelminthes: Orders 1 Pharynx simple ............................................................................................................................................................................................... 2 1’ Pharynx plicatus or bulbous ............................................................................................................................................................................ 4 2(1) Intestine ill defined; statocyst with one statolith (Acoela) or more (Nemertodermatida) ........................................................ Acoela [p. 97] 2’ Intestine well defined; freshwater; planktonic and benthonic ........................................................................................................................ 3 3(2) Protonephridium unpaired; excretory duct central and excretory pore caudal .................................................................. Catenulida [p. 97] 3’ Protonephridium paired ............................................................................................................................................... Macrostomida [p. 97] 4(1) Pharynx plicatus .............................................................................................................................................................................................. 5 4’ Pharynx bulbous .............................................................................................................................................................................................. 6 5(4) Pharynx plicatus directed ventrally; intestine tubular; with statocyst or prominent paired ciliated pits; testes and ovaries dispersed ............. ............................................................................................................................................................................................. Proseriata [p. 97] 5’ Pharynx plicatus directed backwards; intestine triradiate; without statocyst or prominent paired ciliated groves; with paired central eyes or numerous marginal eyes ...................................................................................................................................................... Tricladida [p. 97] 6(4) Oral pore and genital pore separated............................................................................................................................................................... 7 6’ Oral pore and male pore joined; frontal. Male copulatory organ with stylet; female gonads (ovaries and yolk glands) joined; diffuse .......... .................................................................................................................................. Lecithoepitheliata, one family Prorhynchidae [p. 102] 7(6) Pharynx bulbous variabilis (with variable shape); oral pore frontal; male pore caudal; without female copulatory organs; female gonads opens into the masculine atrium ................................................................................. Prolecithophora, one family Plagiostomidae [p. 103] 7’ Pharynx bulbous rosulatus (dorsoventral orientated) or doliiformis (frontocaudally); with male and female copulatory organs; male and female pore ventral or caudal; female gonads (ovaries and yolk glands) separated .................................................... Rhabdocoela [p. 105]

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Platyhelminthes: Acoela: Families 1 Body shape elongated, thin; statocyst with one statolith; colorless or slighty pigmented; without pharynx, small rounded eyes well separated; oral pore at the middle of the ventral side ......................................................................... Mecynostomidae, one genus Limnoposthia   [Palaearctic] 1’ Body shape pedunculate with a small tail and enfolded sides; statocyst with one statolith; gray or whitish pigmented; pharynx simple; orange or yellowish small crescent-shaped eyes behind the pharynx; oral pore at the anterior end ................................................................. ............................................................................................................................................................ Convolutidae, one genus Oligochoerus   [Palaearctic]

Platyhelminthes: Catenulida: Families 1  Brain compact; oval; with or without statocyst; ciliated furrow separate the anterior end (prostomium) from the posterior region............................................................................................................................................................................................................... 2 1’ Brain lobed; with anterior and posterior lobes; without statocyst; without ciliated furrow ........................................ Stenostomidae [p. 97] 2(1) Oral pore near the prostomium; often forming chains of numerous and compact zooids (Figs. 5.1 A, B and 5.8 C)........................................ ..................................................................................................................................................................... Catenulidae, one genus: Catenula 2’ Oral pore away from the prostomium; often forming chains of not more than two or four zooids (Figs. 5.1 C and 5.8 A, B)......................... ............................................................................................................... Chordariidae, one species: Chordarium europaeum Schwank, 1980

Platyhelminthes: Catenulida: Stenostomidae: Genera 1 With ciliated pits; prostomium of variable length; generally with paired light refracting bodies associated to the posterior brain lobes; forming zooids ................................................................................................................................................................................................ 2 1’ Without ciliated pits; with a long prostomium; without light refracting bodies; without zooids (Figs. 5.1 D and 5.8 D) ....... Rhynchoscolex 2(1) Without muscular belt between pharynx and intestine (Fig. 5.1 E) ............................................................................................ Stenostomum 2’ With muscular belt between pharynx and intestine (Fig. 5.8 E, F).................................Myostenostomum gigerium (Kepner & Carter, 1931)

Platyhelminthes: Macrostomida 1 Body shape oval; with two eyes; without ciliated pits; adhesive posterior end; paired testes and ovaries; stylet tube or funnel-like; sexual reproduction. (Fig. 5.1 F).............................................................................................................. Macrostomidae, one genus: Macrostomum 1’ Body shape elongated (often forming zooids); anterior and posterior end pointed; without eyes; with ciliated pits; ovaries unpaired; stylet turned; corkscrew-like; with sexual and asexual (preferred) reproduction (Fig. 5.1 G) ................ Microstomidae, one genus: Microstomum

Platyhelminthes: Proseriata 1 With two pairs of ciliated pits; ovaries behind the pharynx ............................................................................................................................ 2 1’ Without ciliated pits; ovaries before the pharynx ........................................................................................................................................... 3 2(1) Without statocyst; pharynx tubular directed backwards; intestinal branches at pharynx level; common gonopore; posterior adhesive glands (Fig. 5.2 C) ................................................................................................ Bothrioplanidae, one species: Bothrioplana semperi Braun, 1881 2’ With statocyst; pharynx directed ventral; without intestinal branches; female and male gonopore; without posterior adhesive glands (Fig. 5.2 E) .............................................................................. Otomesostomidae, one species: Otomesostoma auditivum (Du Plessis, 1874) 3(1) Pharynx tubular directed backwards; female and male gonopores separate; statocyst anterior to the brain; without sensory pits (Fig. 5.2 F) ............................................................................................................ Monocelididae sp., one species: Monocelopsis carolinensis Ax, 2008 3’  Pharynx globular (dorsoventrad orientated); common gonopore; statocyst before the brain; with sensory pits (Fig. 5.2 D) ...................................................................................... Coelogynoporidae, one species: Coelogynopora falcaria Ax & Sopott-Ehlers, 1979

Platyhelminthes: Tricladida: Families 1 Usually unpigmented; with anterior glandulomuscular organ ........................................................................................................................ 2 1’ Usually pigmented; without anterior glandulomuscular organ ....................................................................................................................... 3 2(1) Testes scattered; with two eyes; with adenodactyls.................................................................................................... Dendrocoelidae [p. 98]

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FIGURE 5.1  Schematic representation of some genera of Catenulida and Macrostomida from the freshwater environments. (A) Catenula sp. (Catenulida, Catenulidae) length: 1–5 mm; (B) Catenula turgida (Catenulida, Catenulidae) length 0.2–0.36 mm; (C) Chordarium europaeum (Catenulida, Chordariidae) length: 0.6–1 mm; (D) Rhynchoscolex sp. (Catenulida, Stenostomidae) length: 4–6 mm; (E) Stenostomum sp. (Catenulida, Stenostomidae) length: 0.5–4 mm; (F) Macrostomum sp. (Macrostomida, Macrostomidae) length: 1–3 mm; (G) Microstomum sp. (Macrostomida, Microstomidae) length: 0.4–4 mm. Abbreviations: b, brain; cp, ciliated pits; ex, excretophore; ey, eyes; fg, female gonads; i, intestine; lrb, light refracting bodies; m, mouth; o, ovary; ph, pharynx; pr, protonephridial duct; sc, statocyst; st, stylet; sv, seminal vesicle; t, testes. 2’ Testes before the pharynx; without eyes and without adenodactyls ................................................................................... Kenkiidae [p. 99] 3(1) Head triangular; one pairs of eyes; white auricle and eyefields ......................................................................................... Dugesiidae [p. 99] 3’ Head rounded, no triangular; two or numerous marginal eyes, sometimes absent; without white auricles or eyefields .................................. ......................................................................................................................................................................................... Planariidae [p. 100]

Platyhelminthes: Tricladida: Dendrocoelidae: Genera 1 Penis bulb elongated (Fig. 5.10 A) ..................................................................................................................................................... Procotyla 1’ Penis bulb rounded (Fig. 5.10 B) .......................................................................................................................................... Dendrocoelopsis

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FIGURE 5.2  Schematic representation of freshwater genera and some freshwater species of Lecithoepitheliata and Proseriata. (A) Geocentrophora sp. (Lecithoepitheliata, Prorhynchidae) length: 1–4 mm; (B) Prorhynchus stagnalis (Lecithoepitheliata, Prorhynchidae) length: 2–7 mm; (C) Bothrioplana semperi (Proseriata, Bothrioplanidae) length: 1–3 mm; (D) Coelogynopora sp. (Proseriata, Coelogynoporidae) length: 5–10 mm; (E) Otomesostoma auditivum (Proseriata, Otomesostomidae) length: 3–5 mm; (F) Monocelopsis sp. (Proseriata, Monocelididae) length 1–1.5 mm. Abbreviations: b, brain; co, copulatory organ; cov, common oviduct; cp, ciliated pits; ey, eyes; fg, female gonads; i, intestine; o, ovary; ph, pharynx; sc, statocyst; sp, spines; st, stylet; sv, seminal vesicle; t, testes; v, vitellaria.

Platyhelminthes: Tricladida: Kenkiidae: Genera 1 Body elongated; flat; with a well-developed postpharyngeal section (Fig. 5.11 A) ................................................................... Sphalloplana 1’ Body oval elongated-shaped; with reduced postpharyngeal section ....................................................................................................... Kenkia

Platyhelminthes: Tricladida: Dugesiidae: Genera 1 Head triangular; pharynx pigmented; testes pre and post pharyngeal............................................................................................................. 2 1’ Head bluntly triangular; pharynx unpigmented; testes pre pharyngeal; male copulatory organ small; penis papilla digitiform; bursa sac not developed (Fig. 5.10 C) .................................................................................................................................... Cura foremanii (Girard, 1852)

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FIGURE 5.3  Schematic representation of characteristic genera of Dalyellioida (Rhabdocoela). (A) Castrella sp. (Dalyelliidae) length: 0.5–1 mm; (B) Dalyellia sp. (Dalyelliidae) length: 1.5–5 mm; (C) Fulinskiella sp. (Dalyellidae) length: 0.7–1 mm; (D) Gieysztoria sp. (Dalyelliidae) length: 0.7–1.35 mm; (E) Microdalyellia sp. (Dalyelliidae) length: 0.5–1.5 mm; (F) Pseudodalyellia sp. (Dalyelliidae) length: 0.6–1 mm; (G) Provortex sp. (Provorticidae) length: 0.5–1 mm; (H) Vejdovskya sp. (Provorticidae) length: 0.3–1 mm. Abbreviations: bc, bursa copulatrix; eg, eggs; ey, eyes; i, intestine; o, ovary; ph, pharynx; rs, receptaculum seminis; st, stylet; sv, seminal vesicle; t, testes; v, vitellaria.

2(1) Head low triangular; testes forming dorsal clusters (Fig. 5.10 E) .......................................................... Schmidtea polychroa (Schmidt, 1861) 2’ Head high triangular; testes numerous; usually ventral and scattered (Fig. 5.10 D) ......................................................................... Girardia

Platyhelminthes: Tricladida: Planariidae: Genera 1 Numerous eyes ................................................................................................................................................................................................ 2 1’ One pair of eyes or absent ............................................................................................................................................................................... 3

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FIGURE 5.4  Schematic representation of characteristic genera of some subfamilies of Typhloplanidae (Typhloplanoida, Rhabdocoela). (A) Acrochordonoposthia sp. (Protoplanellinae) length: 0.9–1.5 mm; (B) Amphibolella sp. (Protoplanellinae) length: 1.4–1.8 mm; (C) Bryoplana sp. (Protoplanellinae) length: 0.4–0.5 mm; (D) Krumbachia sp. (Protoplanellinae) length: 1.7–2.8 mm; (E) Microcalyptorhynchus sp. (Protoplanellinae) length: 0.8–1 mm; (F) Prorhynchella sp. (Protoplanellinae) length: 0.6–1 mm; (G) Olisthanella sp. (Olisthanellinae) length: 1–1.5 mm. Abbreviations: b, brain; bc, bursa copulatrix; co, copulatory organ; ey, eyes; o, ovary; ph, pharynx; rs, receptaculum seminis; rt, rhabdite tracks; sv, seminal vesicle; t, testes; v, vitellaria.

2(1) Eyes located along the anterior edge of the head (Fig. 5.11 D).......................................................................................................... Polycelis 2’ Eyes located in two groups on both sides of the head (Fig. 5.11 E)...................................................................................................... Seidlia 3(1) Penis papilla long .............................................................................................................................................................................................4 3’ Penis papilla remarkable short; large male atrium .................................................................................... Hymanella retenuova Castle, 1941

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FIGURE 5.5  Schematic representation of characteristic genera of some subfamilies of Typhloplanidae (Typhloplanoida, Rhabdocoela). (A) Castrada sp. (Rhynchomesostominae) length: 0.5–1.5 mm; (B) Mesocastrada sp. (Rhynchomesostominae) length: 1–2.5 mm; (C) Opistomum sp. (Opistominae) length: 2–4.5 mm; (D) Rhynchomesostoma sp. (Rhynchomesostominae) length: 1–3.5 mm; (E) Tetracelis sp. (Rhynchomesostominae) length: 1–1.5 mm; (F) Protoascus sp. (Ascophorinae) length: 0.4–1.5 mm; (G) Ascophora sp. (Ascophorinae) length: 1–3 mm. Abbreviations: ago, atrial glandular organ; bc, bursa copulatrix; co, copulatory organ; eg, eggs; ey, eyes; i, intestine; o, ovary; ph, pharynx; rs, receptaculum seminis; rt, rhabdite tracks; sv, seminal vesicle; t, testes; v, vitellaria. 4(3) Adenodacty and bursal canal lead in the vicinity of gonopore (Fig. 5.11 B) ............................................................................. Paraplanaria 4’ Without adenodactyl (Fig. 5.11 C) .................................................................................................................................................. Phagocata

Platyhelminthes: Lecithoepitheliata: Prorhynchidae: Genera 1 Anterior end elongated; sometimes light truncated; no fan expanded; without eyes; male stylet straight (Fig. 5.2 B) .................................... .............................................................................................................................................................. Prorhynchus stagnalis Schultze, 1851 1’ Anterior end fan-like; with eyes except in some cave forms; male stylet curved; claw-like (Fig. 5.2 A) ............................. Geocentrophora

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FIGURE 5.6  Schematic representation of characteristic genera of some subfamilies of Typhloplanidae (Typhloplanoida, Rhabdocoela). (A) Adenoplea sp. (Typhloplaninae) length: 0.42–0.6 mm; (B) Limnoruanis sp. (Typhloplaninae) length: 0.4–0.5 mm; (C) Strongylostoma sp. (Typhloplaninae) length: 0.75–1.5 mm; (D) Styloplanella sp. (Typhloplaninae) length: 0.8–1.3 mm; (E) Typhloplana sp. (Typhloplaninae) length: 0.7–1 mm; (F) Typhloplanella sp. (Typhloplaninae) length: 1.5–2.5 mm; (G) Bothromesostoma sp. (Mesostominae) length: 3–5 mm. Abbreviations: b, brain; bc, bursa copulatrix; co, copulatroy organ; ey, eyes; fgl, frontal glands; nt, nerve trunks; o, ovary; pgo, prepharyngeal glandular organ; ph, pharynx; rs, receptaculum seminis; rt, rhabdite tracks; st, stylet; sv, seminal vesicle; t, testes; u, uterus; v, vitellaria.

Platyhelminthes: Prolecithophora: Plagiostomidae: Genera 1 Female gonads forming two well defined characteristic “bodies” along the main body axis; ovaries follicular; prostatic vesicle very large; unarmed penis (Fig. 5.7 C) ............................................................................................................................................................ Hydrolimax 1’ Female gonads on both sides of the intestine; ovaries compact; prostatic vesicle small; penis sometimes armed or reinforced ..................... .............................................................................................................................................................. Plagiostomum planum Silliman, 1885

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FIGURE 5.7 Schematic representation of characteristic genera of some subfamilies of Typhloplanidae (Typhloplanoida, Rhabdocoela), and of freshwater genera of Kalyptorhynchia (Rhabdocoela) and Prolecithophora. (A) Phaenocora sp. (Phaenocorinae) length: 2–6  mm; (B) Phaenocora sulfophila (Phaenocorinae) length: 3.5– 4.8 mm; (C) Hydrolimax grisea (Plagiostomidae, Prolecithophora) with a partial sample of their pigmentation; length: 10–15 mm; (D) Gyratrix hermaphroditus (Polycistididae Kalyptorhynchia) length: 0.5–12  mm; (E) Opisthocystis goettei (Polycistididae, Kalyptorhynchia) length: 2.5–3 mm. Abbreviations: b, brain; bc, bursa copulatrix; co, copulatory organ; eg, eggs; ey, eyes; i, intestine; o, ovary; pb, proboscide; ph, pharynx; pv, prostatic vesicle; rs, receptaculum seminis; st, stylet; sv, seminal vesicle; t, testes; u, uterus; v, vitellaria.

(A)

(C)

(B)

(D)

(E)

(F)

FIGURE 5.8  Microphotographs of live microturbellarians. (A) General view of Chordarium sp.; (B) detail of the frontal region of Chordarium sp.; (C) anterior region of Catenula turgida; (D) anterior region of Rhynchoscolex sp.; (E) Myostenostomum sp.; (F) detail of the muscular intestinal zone of Myostenostomum sp.

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(A)

(C)

(B)

(D) (E)

FIGURE 5.9  Microphotographs of live microturbellarians. (A) mature Mesostoma ehrenbergii with several dormant eggs in the uterus; (B) anterior region of Mesostoma ehrenbergii with several subitaneous eggs in the uterus; (C) Mesostoma ehrenbergii while mating; (D) Gieysztoria sp. with the developed ovary and vitellaria; (E) Mesostoma ehrenbergii with several eggs in the uterus and with some Cladocera in the intestine.

Platyhelminthes: Rhabdocoela: Families 1 Pharynx bulbous rosulatus (dorsoventral orientated) ..................................................................................................................................... 2 1’ Pharynx bulbous doliiformis (fronto-caudally orientated) .............................................................................................................................. 3 2(1) Without true proboscis.............................................................................................................................................. Typhloplanidae [p. 105] 2’ With true proboscis...................................................................................................................................................... Polycistididae [p. 108] 3(1) Ovary single; male copulatory organ with a spiny stylet ................................................................................................. Dalyellidae [p. 108] 3’ Ovary paired; male copulatory organ with a straight tubular stylet (bucket-like) without spines .............................. Provorticidae [p. 108]

Platyhelminthes: Rhabdocoela: Typhloplanidae: Genera 1 Pharynx ventral directed ......................................................................................................................................................................... 2 1’ Pharynx with other orientation ..................................................................................................................................................................... 22 2(1) Testes dorsal to the vitellaria ........................................................................................................................................................................... 3 2’ Testes ventral to the vitellaria ......................................................................................................................................................................... 7 3(2) Testes sac-like, in front of the pharynx ........................................................................................................................................................... 4 3’ Testes mostly follicular, behind or on both sides of the pharynx, pharynx in the mid-body (second third) with two eyes...................... 6 4(3) Without eyes; with ascus ................................................................................................................................................................................. 5 4’ Without eyes (sometimes with pigment patches); without ascus; male and female organ posterior; at the last third of the body (Fig. 5.4 G) ................................................................................................................................................................. Olisthanella coeca (Silliman, 1885) 5(4) Pharynx at the anterior end (first third of the body); female and male copulatory organ in the second third (Fig. 5.5 F) ................................ ............................................................................................................................................................. Protoascus wisconsinensis Hayes, 1941 5’ Pharynx in the middle of the body (second third); female and male copulatory organs close behind the pharynx (Fig. 5.5 G) ...................... ....................................................................................................................................................... Ascophora elegantissima Findenegg, 1924 6(3) Dense dark brownish pigmentation on the dorsal side; with anterior pigment loss eyes fields; eyes masked; with a prepharyngeal glandular organ (Fig. 5.6 G)................................................................................................................... Bothromesostoma personatum (Schmidt, 1848) 6’ Different tonalities (transparent; beige; caramel) and pattern (with spots or with thin stripes) of the dorsal pigmentation, eyes visible, without pre-pharyngeal glandular organ (Fig. 5.9 A–C, E) .................................................................................................................. Mesostoma

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FIGURE 5.10  Schematic representation of some characteristic genera of Dendrocoelidae (Tricladida) and Dugesiidae (Tricladida). (A) Procotyla sp. (Dendrocoelidae) length: 10–12 mm; (B) Dendrocoelopsis sp. (Dendrocoelidae) length: 14–22 mm; (C) Cura sp. (Dugesiidae) length: 7–15 mm; (D) Girardia sp. (Dugesiidae) length: 6–30 mm; (E) Schmidtea sp. (Dugesiidae) length: ±20 mm. Abbreviations: au, auricles; bc, bursa copulatrix; co, copulatory organ; e, eyes; i, intestine; ib, intestinal branches; ph, pharynx; vd, vas deferens.

7(2) Pores of the protonephridium open near the oral pore or in the genital pore ................................................................................................. 8 7’ Pores of the protonephridium open on body surface .................................................................................................................................... 17 8(7) Pores of the protonephridium open in the buccal cavity or near the pharynx ................................................................................................ 9 8’ Pores of the protonephridium open in the genital atrium ............................................................................................................................. 14 9(8) Copulatory organ at the distal end of the body ............................................................................................................................................. 10 9’ Copulatory organ at the middle of the body, close to the pharynx ........................................................................................................ 11 10(9) Male organ with a stylet funnel-like and smooth; body oval; without eyes and rounded (light truncated) anterior part (Fig. 5.6 A) .............. .......................................................................................................................................................... Adenoplea nanus Sayre & Wergen, 1994 10’ Male organ with a stylet crown-shaped with four curved spines; body elongated; with two eyes and an elongated anterior part (Fig. 5.6 B) ................................................................................................................................................................. Limnoruanis romanae Kolasa, 1977

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FIGURE 5.11  Schematic representation of some characteristic genera of Kenkiidae (Tricladida) and Planariidae (Tricladida) length: 7–15 mm. (A) Sphalloplana sp. (Kenkiidae) length: 5–20 mm; (B) Paraplanaria sp. (Planariidae) length: 9–13 mm; (C) Phagocata sp. (Planariidae) length: 10–14 mm; (D) Polycelis sp. (Planariidae) length: 7–12 mm; (E) Seidlia sp. (Planariidae) length: 11–17 mm. Abbreviations: ad, adenodactyl; ado, adhesive organ; bc, bursa copulatrix; co, copulatory organ; e, eyes; gp, gonopore; i, intestine; ib, intestinal branches; ph, pharynx; vd, vas deferens. 11(9) Caudal testes ................................................................................................................................................................................................. 12 11’ Frontal testes, behind and before the pharynx .............................................................................................................................................. 13 12(11) Ejaculatory duct developed as spiny cirrus; testes stretching at the posterior end; with bursa copulatrix; with eyes (Fig. 5.6 C) ................... ................................................................................................................................................................................................. Strongylostoma 12’ Ejaculatory duct with cuticular lining, without spines; testes near the pharynx; vitellaria at the posterior end; without bursa copulatrix; without eyes (Fig. 5.6 E) ............................................................................................................................................................... Typhloplana 13(11) Without eyes, whitish, opaque, sometimes light pigmented; ejaculatory duct with cuticular lining; with bursa copulatrix (Fig. 5.6 F) ......... ................................................................................................................................................... Typhloplanella halleziana (Vejdovsky, 1880) 13’ With eyes; transparent; with tube-like stylet (Fig. 5.6 D) ............................................... Styloplanella strongylostomoides Findenegg, 1924

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14(8) With eyes; male organ with numerous spines in the bursa copulatrix, in the blind sacs or in the male atrium ............................................ 15 14’ Without eyes; copulatory organs in the middle, or between the first and the second third of the body. Male organ with numerous spines in blind sacs or in the bursa copulatrix (Fig. 5.5 A) ............................................................................................................................... Castrada 15(14) With two eyes ................................................................................................................................................................................................ 16 15’ With four eyes (Fig. 5.5 E) ................................................................................................................... Tetracelis marmorosa (Müller, 1773) 16(15) With a retractable anterior end (telescope-like) (Fig. 5.5 D) ................................................... Rhynchomesostoma rostratum (Müller, 1773) 16’ Without retractable anterior end (Fig. 5.5 B) ......................................................................................... Mesocastrada fuhrmanni Volz, 1898 17(7) Pharynx in the second third of the body ....................................................................................................................................................... 18 17’ Pharynx anterior; in the first third of the body ............................................................................................................................................. 19 18(17) Testes before the pharynx; copulatory organs posterior to the pharynx; on the last third of the body (Fig. 5.4 D) ..................... Krumbachia 18’ Small testes behind the pharynx; copulatory organs close behind the pharynx on the second third of the body (Fig. 5.4 B) .......................... .............................................................................................................................................................. Amphibolella segnis Findenegg, 1924 19(17) Male copulatory organ without spines or cirrus ........................................................................................................................................... 20 19’ Male copulatory organ with a spiny cirrus (Fig. 5.4 A) ........................................................... Acrochordonoposthia conica Reisinger, 1924 20(19) Male and female copulatory organs on the last third of the body ................................................................................................................. 21 20’ Male and female organs close behind the pharynx; on the first third of the body (Fig. 5.4 F) ............. Prorhynchella minuta Ruebush, 1939 21(20) Testes paired; anterior, voluminous and with uneven development; without eyes; anterior end with a small anterior projection like a proboscis (Fig. 5.4 E) ............................................................................................ Microcalyptorhynchus virginianus Kepner & Ruebush, 1935 21’ Paired small posterior testes with equal development (Fig. 5.4 C)................. Bryoplana xerophila Van Steenkiste, Davison, & Artois, 2010 22(1) Pharynx forward directed; feathery vitellaria; male copulatory organ sometimes with spines ............................ Phaenocora (Fig. 5.7 A, B) 22’ Pharynx backwards directed; vitellaria smooth; male copulatory organ cirrus-like; with spines (Fig. 5.5 C) .................................................. ................................................................................................................................................................. Opistomum pallidum Schmidt, 1848

Platyhelminthes: Rhabdocoela: Polycistididae: Genera 1 Copulatory organs in the second third of the body, behind and near the pharynx; paired ovaries; short crooked funnel-like stylet; with excretory cup (Fig. 5.7 E).............................................................................................................................. Opisthocystis goettei (Bresslau, 1906) 1’ Copulatory organs in the last third of the body, behind and clear separate from the pharynx; posteriorly with a long fork-like stylet; with eyes (Fig. 5.7 D) .................................................................................................................................... Gyratrix hermaphroditus Graff, 1831

Platyhelminthes: Rhabdocoela: Dalyellidae: Genera 1 Stylet with two handles and two spiny branches............................................................................................................................................. 2 1’ Stylet with one handle or without handles ...................................................................................................................................................... 5 2(1) Stylet with two handles of similar size ........................................................................................................................................................... 3 2’ Stylet with two handles of different size (Fig. 5.3 F)........................... Pseudodalyellia alabamensis Van Steenkiste, Gobert & Artois, 2011 3(2) Stylet compose by two joined handles, branches and spines .......................................................................................................................... 4 3’ Stylet compose by two independent handles, branches and spines (Fig. 5.3 C)................................ Fulinskiella bardeaui (Steinböck, 1926) 4(3) Stylet with narrow handles, usually longer than the branches, branches with different kind of spines, with a middle channel; with only one egg; without zoochlorellae (Fig. 5.3 E) .................................................................................................................................... Microdalyellia 4’ Stylet with width handles, usually shorter than the branches, branches with robust spines, without a middle channel, several eggs stored in the parenchyma, sometimes with zoochlorellae (Fig. 5.3 B) ............................................................................................................. Dalyellia 5(1) Stylet with one handle and two spiny branches (Fig. 5.3 A) ............................................................................................................. Castrella 5’ Stylet without handles, with a wide belt and different kind of spines (Figs. 5.3 D and 5.9 D) ..................................................... Gieysztoria

Platyhelminthes: Rhabdocoela: Provorticidae: Genera 1 Tubular stylet long; very fine and needle-like (Fig. 5.3 H) .................................................................. Vejdovskya pellucida (Schultze, 1851) 1’ Tubular stylet short, width, truncated cone (Fig. 5.3 G)........................................................ Provortex virginiensis Ruebush & Hayes, 1939

ACKNOWLEDGMENTS We would like to thank Dr Thorp for inviting us to participate in this chapter, Dr Ana Maria Leal-Zanchet for allowing us to use the photographs of the catenulids (Fig. 5.8) and Gieysztoria (Fig. 5.9), and Sebastian Castorino for providing assistance in the vectorization of the drawings. This study received support from the I+D Project grant CGL 2010-15786/BOS and CGL2011-29916, which are financed by Spanish Ministry of Economy; from PIP 2010-390 and PIP 2013-2015-635(CONICET), and N11/728 FCNyM (UNLP).

REFERENCES Baguñà, J. & M. Riutort. 2004a. Molecular phylogeny of the Platyhelminthes. Canadian Journal of Zoology 82: 168–193. Baguñà, J. & M. Riutort. 2004b. The dawn of bilaterian animals: the case of acoelomorph flatworms. BioEssays 26: 1046–1057. Cannon, L.R.G. 1986. Turbellaria of the world. A guide to families & ­genera. Queensland Museum, Brisbane. 186 pp. Ferguson, F.F. 1954. Monograph of the Macrostomine worms of Turbellaria. Transactions of the American Microscopical Society 73: 137–164. Gilbert, C.M. 1938. A remarkable North American species of the genus Phaenocora. Zeitschrift fuer Morphologie und Ökologie der Tiere 33: 53–71. Houben, A.M., N. Van Steenkiste & T.J. Artois. 2014. Revision of Phaenocora Ehrenberg, 1836 (Rhabditophora, Typhloplanidae, Phaenocorinae) with the description of two new species. Zootaxa 3889: 301–354. Kenk, R. 1974. Chapter 2: History of the study of Turbellaria in North America. Pages 17–22 in: N.W. Rise and M.P. Morse. (eds.), Biology of the Turbellaria. McGraw-Hill Book Company, New York, NY. Kenk, R. 1989. Revised list of the North American freshwater planarians (Platyhelminthes: Tricladida: Paludicola). Smithsonian Contributions to Zoology 476: 1–10. Kolasa, J. & S. Tyler. 2010. Chapter 6: Flatworms: Turbellarians and Nemertea. Pages 143–161 in: J.H. Thorp and A.P. Covich (eds.), Ecology and classification of North American freshwater invertebrates (Third Edition), Academic Press, San Diego, CA.

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Littlewood, D.T.J. & R.A. Bray (eds.) 2001. Interrelationships of the Platyhelminthes. Taylor & Francis, London, New York. Luther, A. 1955. Die Dalyelliiden (Turbellaria Neorhabdocoela) Eine Monographie. Acta Zoologica Fennica 87: 1–337. Luther, A. 1960. Die Turbellarien Ostfennoskandiens. I. Acoela, Catenulida, Macrostomida, Lecithoepitheliata, Prolecithophora, und Proseriata. Fauna Fennica 7: 1–155. Luther, A. 1963. Die Turbellarien Ostfennoskandiens. IV. Neorhabdocoela 2. Typhloplanoida: Typhloplanidae, Solenopharyngidae und Carcharodopharyngidae. Fauna Fennica 16: 1–161. Nuttycombe, J.W. 1956. The Catenulida of the Eastern United States. American Midland Naturalist 55: 419–433. Nuttycombe, J.W. & A.J. Waters 1938. The American species of the genus Stenostomum. Proceedings of the American Philosophical Society 79: 213–301. Richter, S., R. Loese, G. Purschke, A. Schmidt-Rhaesa, G. Scholtz, T. Stach, L. Vogt, A. Wanninger, G. Brenneis, C. Döring, S. Faller, M. Fritsch, P. Grobe, C.M. Heuer, S. Kaul, O.S. Møller, C.H.G. Müller, V. Rieger, B.H. Rothe, M.E.J. Stegner & S. Harzsch. 2010. Invertebrate neurophylogeny: suggested terms and definitions for a neuroanatomical glossary. Frontiers in Zoology 7:1–49. Rieger, R.M., S. Tyler, J.P.S. Smith III, & G. Rieger. 1991. Chapter 2: Turbellaria. Pages 7–140 in: F.W. Harrison and B.J. Bogitsh (eds.), Microscopic Anatomy of Invertebrates. Vol. 3: Platyhelminthes and Nemertinea. Wiley-Liss, New York, NY. Ruiz-Trillo, I., M. Riutort, D.T.J., Littlewood, E.A. Herniou & J. Baguñà. 1999. Acoel flatworms: Earliest extant bilaterian metazoans, not members of Platyhelminthes. Science 283: 1919–1923. Ruiz-Trillo, I., M. Riutort, H.M. Fourcade, J. Baguñà & J.L. Boore. 2004. Mitochondrial genome data support the basal position of Acoelomorpha and the polyphyly of the Platyhelminthes. Molecular Phylogenetics and Evolution 33: 321–332. Schärer, L. 2014. Macrostomorpha taxonomy and phylogeny. http:// macrostomorpha.info. Consulted 2014-12-04. Tyler, S. & M. Hooge. 2004. Comparative morphology of the body wall in flatworms (Platyhelminthes). Canadian Journal of Zoology 82: 194–210. Tyler, S., S. Schilling, M. Hooge, & L.F. Bush (comp.) 2006–2015. Turbellarian taxonomic database. Version 1.7 http://turbellaria.umaine.edu. Consulted 2014-12-04.

Phylum Platyhelminthes

Chapter | 5  Phylum Platyhelminthes

Chapter 6

Phylum Nemertea Malin Strand The Swedish Species Information Centre, Swedish University of Agricultural Sciences, Uppsala, Sweden

Department of Zoology, University of Gothenburg, Gothenburg, Sweden

Chapter Outline Introduction111 Limitations111 Terminology and Morphology 112

INTRODUCTION There are around 1275 species of nemerteans (phylum Nemertea) described (Kajihara et al., 2008), with the vast majority being marine, and only 22 species are currently known from freshwater habitats. Of these, only two species are recorded from the Nearctic region: one from Lake Huron in Canada, and the other from a stream in Chester County, Pennsylvania. One of the species (Prostoma canadiensis Gibson & Moore 1978) was later recorded from Holland (Moore & Gibson, 1985). The other species, Prostoma asensoriatum (Montgomery, 1896), is only recorded from the type locality, and thus endemic for the region. The species are both benthic, like all other known freshwater species. Being rare and often overlooked, the ecological impact of these species is unknown.

LIMITATIONS More freshwater nemerteans have been named and described from the region, besides the two listed above. Coe (1901), for example, combined a number of previous names into Tetrastemma rubrum (later transferred to Prostoma). However, the descriptions of these species are inadequate and too vague to be acceptable, and Gibson & Moore (1976) disregarded P. rubrum (and others) leaving the P. canadiensis and P. asensoriatum as the only valid names for nemerteans in this region. The original description of Prostoma asensoriatum closely resembles P. eilhardi, but

Material Preparation and Preservation 112 Keys to Nemertea 112 References113

offers a number of anatomical differences. Although some characters listed are too variable to be used for species diagnostics (Moore & Gibson, 1976), others appear to be a reliable species indicator. We thus conclude that there are currently two valid species known from the region, but at the same time emphasize that the real number of species is uncertain and probably higher. We base this conclusion on experience from other regions, and especially marine ­habitats, where new species are commonly found whenever a taxonomist familiar with the group samples in an area. The two Nearctic nemerteans are both small (up to 4 cm), slender, and orange-reddish, and both are difficult to distinguish using external characters. The number of eyes (up to 8 arranged in 2–4 pairs) varies intraspecifically and also by age, with a positive correlation between the number and age/size. Although it is stated that P. canadiensis is slightly shorter than P. asensoriatum, this is obviously a character that varies with age and degree of contraction. Nemerteans are very contractile, and this also affects the interpretation of anatomical characters like extension of organs, positions, and volumes. It is often stated in the literature that nemerteans can, and have to be, identified from internal characters. However, as shown by Moore & Gibson (1976), and also argued in other papers, many characters are for various reasons impossible to use in this context. Furthermore, internal characters can only (with few exceptions) be interpreted from histological sections. In most cases, these sections are too time-consuming/expensive to provide, and the

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technique also requires special skills and equipment, which are lacking in many situations. We therefore conclude that use of internal characters is rare in reality, despite what being stated in many publications as a necessary requirement for proper identification of nemertean species.

histological preparation, and in most situations, it still may not be possible to key the species below the level of genus. There are currently no published molecular markers for either species, but these may prove especially valuable in the future.

TERMINOLOGY AND MORPHOLOGY

MATERIAL PREPARATION AND PRESERVATION

Phylum Nemertea

A nemertean is recognized by its smooth gliding way of locomotion over the ground, using the cilia covering the body, and the fact that it contracts easily if disturbed. Nemerteans are often confused with flatworms, moving in the same way and also sometimes of similar body shape. The unique character identifying Nemertea as a monophyletic taxon is the eversible proboscis. It will be difficult to distinguish the two species from external characters, although it is stated in the description of P. asensoriatum that the color is bright orange, while P. canadiensis is described as “bright and … shade of pink”. The species can, however, be distinguished by the internal character presence/absence of frontal organ, which we judge as a valid species identifier. There may be cases when it is possible to observe this character in live specimens by the opening in the snout, but it will require considerable experience working with this and use of a dissecting microscope. A firm species identification would, however, require

Nemerteans should always be studied alive whenever possible. The characters to be observed are presence/absence of eyes, number and pattern if present, color, position of mouth, and (in this case), the presence of a frontal organ. For histological sections, it is important to properly anesthetize the animal before fixation to minimize contractions (which will distort anatomical characters). MS 222 or MgCl2 is recommended. A common fixative for nemerteans is Bouin’s fluid/paraformaldehyde. For histological studies, fixed animals are embedded in paraffin wax (56 °C). Sections are in general 6 microns thick and stained by the Mallory trichrome method. Ethanol is required for later DNA extractions, but is less appropriate for histology. Another good fixative for RNA and DNA studies is RNAlater, which is easy to carry around (not being toxic nor flammable) and is especially suited for fieldwork. Store the animals at −20 °C or lower if their DNA is to be analyzed.

KEYS TO NEMERTEA The currently two known species from the Nearctic region can be conclusively identified by the presence/absence of a frontal organ (Fig. 6.1), such as: 1 Frontal organ present ............................................................................................................... Prostoma canadiensis Gibson & Moore 1978 1’ Frontal organ absent .................................................................................................................. Prostoma asensoriatum (Montgomery 1896)

FIGURE 6.1  The placement of the frontal organ.

REFERENCES Coe, W. R. 1901. Papers from the Harriman Alaska expedition. XX. The nemertean. Proceedings of the Washington Academy of Sciences 3: 1–110. Kajihara, H., A. V. Chernyshev, S-C. Sun, P. Sundberg & F. B. Crandall. 2008. Checklist of Nemertean Genera and Species published between 1995 and 2007. Species Diversity 13: 245–274.

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Moore, J. & R. Gibson. 1985. The evolution and comparative physiology of terrestrial and freshwater nemerteans. Biological Reviews 60: 257–312. Sundberg, P. & R. Gibson. 2008. Global diversity of nemerteans (Nemertea) in freshwater. Hydrobiologia 595: 61–66.

Phylum Nemertea

Chapter | 6  Phylum Nemertea

Chapter 7

Phylum Gastrotricha Tobias Kånneby Department of Zoology, Swedish Museum of Natural History, Stockholm, Sweden

Chapter Outline

INTRODUCTION Gastrotricha are small vermiform or tenpin-shaped acoelomate animals. The group is common to most aquatic environments and constitutes an important part of the meiofauna. To date, roughly 850 species have been described from all over the world. Freshwater gastrotrichs are, with very few exceptions, classified within the order Chaetonotida. They are among the smallest metazoans and can have a total body length of only 60 μm. However, most freshwater species attain larger body sizes, and some can reach lengths of up to 800 μm. Gastrotrichs are widely distributed in a variety of freshwater habitats. They are most common in epibenthic or periphytic habitats but may also be encountered in the interstitial. Bogs with Sphagnum spp. or small still nutrient rich ponds with Lemna spp. are especially rich and diverse in gastrotrichs. The following works are helpful in species identification: Brunson (1950, 1959), who keyed and illustrated species then known from North American freshwaters; ­Robbins (1965, 1973), who gave additional information and drawings of North American freshwater species; d’Hondt (1971), who gave a key to the species of Lepidodermella; Kisielewski (1981), who made a critical evaluation of morphological characters that must be measured to identify a species; Kisielewski (1986), who gave a recent treatment of Aspidiophorus; Schwank (1990), who provided illustrated keys (in German) for all known freshwater species worldwide; Kisielewski (1991), who described many Neotropical species and addressed several important issues in gastrotrich systematics; Balsamo & Todaro (2002), who provided a key to the freshwater genera of the world; Kånneby et al. (2009) who gave an illustrated key to Ichthydium; and Kånneby

Material Preparation and Preservation 117 Keys to Gastrotricha 119 References130

et al. (2013), who provided the first molecular phylogeny of Chaetonotidae to evaluate the homology of cuticular characters for generic and subgeneric classification. The identification key below includes all known genera of freshwater chaetonotidans worldwide. For monotypic genera, the species is always presented, although the genus may not have been reported from the Nearctic region. The key covers all Nearctic freshwater species published, as of December 2013. For more recent publications on the Nearctic freshwater gastrotrich fauna, the reader should refer to: (1) Schwank & Kånneby (2014), which redescribes seven species, from Ontario, Canada, previously considered nomina nuda; and (2) Kånneby & Wicksten (2014), which present the first record of a freshwater Macrodasyida, Redudasys sp., from the Northern hemisphere, found in Texas, USA. The identification key to species may sometimes seem too detailed; however, in this way new records or species for the region can be discerned. Rough distributions are given for each species as reported in the literature, but most species probably have a much wider distribution because of the few studies focusing on Nearctic freshwater Gastrotricha.

LIMITATIONS It is relatively easy to identify most Nearctic freshwater gastrotrichs to genus and very difficult to identify them to species. Great patience is needed, since their minute, transparent, and soft bodies make them very hard to work with. Moreover, most of the freshwater gastrotrichs of the Nearctic are undoubtedly not described. For species identification, animals need to be studied alive. Material preserved as whole mounts deteriorate rapidly over time, and important diagnostic characters may be extremely

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Introduction115 Limitations115 Terminology and Morphology 116

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hard or impossible to discern. When studying the live animal, a good light microscope is essential (preferably with differential interference contrast (DIC) or phase contrast). I also recommend using a digital camera (with video capability to capture multiple focal planes) to record the animal. The difficulty to work with the group have made gastrotrichs somewhat neglected by invertebrate zoologists and the group is in need of taxonomic revisions. The extent of cryptic species is poorly studied within the group, and it is possible that the true number of species is underestimated (see Kånneby et al., 2012). Current classification is, to a great extent, based on morphology, and many groups have recently been rendered non-monophyletic based on molecular data (Kånneby et al., 2013).

TERMINOLOGY AND MORPHOLOGY In general the body of a freshwater gastrotrich is tenpin-shaped and can be divided into four parts (Fig. 7.1): (1) the anterior lobed head with cephalic plates and cephalic sensory ciliary tufts and/or tentacles; (2) the neck that separates the head from the trunk; (3) the trunk which constitutes the bulk of the body; and (4) the furca carrying 0–4, but usually 2 adhesive tubes. Exceptions to this generalization are the macrodasyidan

Phylum Gastrotricha

FIGURE 7.1 Schematic representation of a hypothesized Chaetonotid gastrotrich showing important diagnostic characters and the different body regions. Dorsal view to the left and ventral view to the right. Broken lines represent internal structures or structures on the opposite side of the body: (1) Hypopleura; (2) Epipleuria; (3) Cephalion; (4) Ocellar granule; (5) Pharynx with weak posterior swelling; (6) Anterior dorsal sensory bristle; (7) Intestine; (8) Dorsal column of 22 keeled scales; (9) Dorsal row of 13–14 alternating keeled scales; (10) Egg; (11) Spine girdle with bifurcated spines; (12) Posterior sensory bristle anchored by double-keeled scale; (13) Parafurcal spine; (14) Caudal incision; (15) Rounded double-keeled scale typical for certain species of Chaetonotus (Hystricochaetonotus); (16) Posterior sensory ciliary tuft; (17) Anterior sensory ciliary tuft; (18) Mouth; (19) Pharyngeal tooth; (20) Hypostomium; (21) Transverse ventral scale bars of the interciliary area typical of Lepidodermella squamata and Chaetonotus maximus; (22) Ventral ciliation; (23) Ventrolateral row of alternating keeled scales; (24) Ventral interciliary scales; (25) Ventral terminal keeled scales. Redrawn from various sources.

Thorp and Covich’s Freshwater Invertebrates

gastrotrichs, with a vermiform body and more than four adhesive tubes (Fig. 7.2), and the semi-planktonic groups (e.g., Dasydytidae and Neogosseidae), with a sack-shaped body that lacks the posterior furca (Fig. 7.3 L–R, T, U). The head can be rounded (one-lobed), three-lobed, or five-lobed (Fig. 7.4 A–C). The lobes are covered by cephalic plates, which in a species with a five-lobed head consist of a cephalion (Fig. 7.1.3), a pair of epipleura (Fig. 7.1.2), and a pair of hypopleura (Fig. 7.1.1). Cephalic sensory ciliary tufts usually originate from between the lateral or ventrolateral borders of the cephalic plates (Figs. 7.1.16, 7.4 A–E, and 7.6 F). The width of the neck region varies a lot between species, e.g., certain species may have a very constricted neck, whereas others almost completely lack a neck constriction. In the neck scale, columns usually converge to later diverge in the trunk region. In most species, a pair of anterior sensory bristles, inserted between the scales or on small papillae, is also present in the neck region (Fig. 7.1.6). The trunk constitutes most of the body and is usually also the widest part. In the posterior trunk region, a pair of posterior sensory bristles is often present; the bristles are usually inserted on specialized double-keeled scales (Fig. 7.1.12). The furca typically consists of two furcal branches, each with a coneshaped basal part that bears the distal adhesive tube (Fig. 7.1).

Chapter | 7  Phylum Gastrotricha

ventrolateral columns (5 on either side of the animal); hence the total number of columns is 23–24. The shape (outline) of individual scales is also an important character. A common type of scale is the three-lobed scale; it comes in different flavors ranging from rounded or weakly three-lobed (Fig. 7.5 L) via three-lobed (Fig. 7.5 A, B) to strongly threelobed (Fig. 7.5 C, I). There are also rounded to oval scales (Fig. 7.5 F, G, R), five-lobed scales (Fig. 7.5 K, M), rhomboidal scales (Fig. 7.5 S), and polygonal scales (Fig. 7.5 U). Scales may be smooth (e.g., Lepidodermella) (Fig. 7.5 U), keeled (e.g., Heterolepidoderma) (Fig. 7.5 V), pedunculate (e.g., Aspidiophorus) (Fig. 7.5 T), or they can bear spines (e.g. Chaetonotus). The spine can be simple (Fig. 7.5 A), dentate (Fig. 7.5 B), bidentate (Fig. 7.5 Y), or carry lateral denticles and a bifurcate tip (Fig. 7.5 H). Certain species of Ichthydium and Arenotus lack scales and spines altogether and are said to have a naked cuticle. The most striking feature of the ventral side is the ventral ciliation, which is used for locomotion. This ciliation often consists of two longitudinal bands going from the anterior to posterior (Figs. 7.1.22 and 7.6 E), but it can also consist of tufts. The area between the ciliary bands is called the interciliary area. It may be naked but is more often covered by scales, which can be smooth, keeled, spined, etc. (Figs. 7.1.24, 7.5 X, and 7.6 E). At the posterior end of the interciliary area, a pair of ventral terminal scales are often present (Fig. 7.1.25). The mouth is situated terminally or subterminally in the anterior end (Fig. 7.1.18). It is connected to the pharynx, which is a muscular tube with (Fig. 7.4 B, C) or without terminal swellings (bulbs) (Fig. 7.4 A). The intestine is usually straight and may sometimes have a differentiated anterior part (Fig. 7.1.7).

MATERIAL PREPARATION AND PRESERVATION

FIGURE 7.2  Freshwater macrodasyidan gastrotrichs: (A) Marinellina flagellata; (B) Redudasys fornerise. AT = Adhesive tube, EG = Egg, PP = Pharyngeal pore. Based on Ruttner-Kolisko (1955); Tirjaková (1998); Todaro et al. (2012).

Gastrotrichs can be collected from almost all aquatic environments, but are most common in still, nutrient rich waters. Material is collected with a plankton net with a mesh size of 25 μm or by collecting sediment, aquatic plants, and mosses by hand. In the laboratory, the samples can be kept in small aerated aquaria. They yield the highest diversity and densities of gastrotrichs just after collection. Subsamples are sucked up from the aquaria with a large pipette and put in petri dishes. The subsample can be treated with an isotonic solution of magnesium chloride (1% for freshwater); this anesthetizes the animals and prevents them from adhering to surfaces with their adhesive tubes. Scan the subsamples under a dissecting microscope, preferably in transmitted light. Once an animal is located, transfer it with a micropipette to a clean microscope slide, and then mount the animal alive in a small amount of water. To prevent squashing of the animal, add a small amount of modeling clay under the edges of the cover slip. In order

Phylum Gastrotricha

It is usually straight; but in certain species, it is forcipate (Fig. 7.4 I). In Polymerurus, the branches of the furca are annulate, appearing segmented (Fig. 7.4 G). The body is covered by a cuticle that is often developed into various types of cuticular structures like scales, spines, and plates. These cuticular structures vary between species and are often diagnostic when identifying and describing species. An important character when determining a species is the number of scales. Columns of scales are parallel to the longitudinal body axis and counted from one side of the animal to the other. Rows of scales are perpendicular to the longitudinal body axis and are counted from anterior to posterior. The number of rows is usually higher than the number of columns. In Fig. 7.1, one dorsal column with 22 scales is depicted, hence the number of rows is 22. Moreover, in Fig. 7.1, one dorsal row with 13–14 alternating scales is depicted; hence the number of columns is 13–14. The number of columns is always counted where the body is widest (unless otherwise stated), and the number of rows is counted down the median of the animal. The dorsal number of columns is the columns seen on the dorsal side of the animal, while the total number of columns also includes the ventrolateral columns. In Fig. 7.1, there are 10

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FIGURE 7.3  Freshwater chaetonotidan genera representing Chaetonotidae (A–K), Dasydytidae (L–R), Dichaeturidae (S), Neogosseidae (T, U), and Proichthydidae (V, W): (A) Arenotus; (B) Aspidiophorus; (C) Chaetonotus; (D) Fluxiderma; (E) Heterolepidoderma; (F) Ichthydium; (G) Lepidochaetus; (H) Lepidodermella; (I) Rhomballichthys; (J) Polymerurus; (K) Undula; (L) Anacanthoderma; (M) Chitonodytes; (N) Dasydytes; (O) Haltidytes; (P) Ornamentula; (Q) Setopus; (R) Stylochaeta; (S) Dichaetura; (T) Kijanebalola; (U) Neogossea; (V) Proichthydioides; (W) Proichthydium. Adapted and modified from Strayer et al. (2009).

to observe important characters, the animal should be oriented in a dorso-ventral fashion. Proper orientation can be achieved by adding small amounts of water to the sides of the cover slip or by removing water with a filter paper. This procedure creates currents under the cover slip, which should alter the animal slightly. If the animal gets stuck on either the surface of the slide or the cover slip, a gentle tapping with a pair of forceps on the cover slip can help. When properly mounted, examine the animal under a light microscope equipped with DIC or phase contrast.

Documentation should include pictures/recordings of the habitus and taxonomically important characters, such as distribution and shape of scales, spines, adhesive tubes, cephalic plates, pharynx, etc. After documentation, the animal can be recovered from the slide for further treatment. The recovering procedure is tricky, but by adding water, the cover slip can be made to float and subsequently gently lifted with a fine insect needle. The animal is now free and can be sucked up with a micropipette and transferred to a vial containing the

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FIGURE 7.4  Schematic representation of different head-shapes (A–E), pharynx-shapes (A–C), caudal ends/furcas (F–I) and spine distribution (J): (A) Five-lobed head with two pairs of sensory ciliary tufts, and pharynx of equal thickness along its length; (B) Three-lobed head with one pair of sensory ciliary tufts, and pharynx with posterior swelling; (C) One-lobed or rounded head with one pair of sensory ciliary tufts, and pharynx with anterior and posterior swellings; (D) Three-lobed head where the posterior lobes are modified into dorsal flaps; (E) Five-lobed head where the middle and posterior lobes are modified into dorsal flaps; (F) The peg-like protuberances typical of Stylochaeta; (G) The long ringed (appearing segmented) furca typical of Polymerurus; (H) Furca with reduced/absent adhesive tubes typical of Chaetonotus (Wolterecka); (I) Forcipate furca present in for example Ichthydium and Chaetonotus; (J) Distribution of the conspicuous rows of longer thicker spines and spineless scales of many Chaetonotus (Hystricochaetonotus) spp. Redrawn from Brunson (1950) and Schwank (1990).

medium of choice. For whole mounts, the animal can be fixed in 10% buffered formalin and then dehydrated through a series of ethanol and glycerol to pure glycerol. Before mounting, a few drops of formalin can be added to the slide to prevent bacterial deterioration (Kånneby et al., 2009).

Animals can also be fixed in 2.5% glutaraldehyde, exposed to osmium tetraoxide, dehydrated through an ethanol series, and mounted in epon. For molecular studies, animals are stored in 95% ethanol in −18 to 20 °C until further treatment.

KEYS TO GASTROTRICHA Gastrotricha: Orders 1 Animal with at least three pairs of adhesive tubes (one anterior and two posterior) and a pair of pharyngeal pores (absent in Lepidodasyidae) (Fig. 7.2) .................................................................................................................................................................................... Macrodasyida [very rare; an almost entirely marine group, represented by only two nominal species in freshwaters; the genus Redudasys was recently reported from Texas, USA] 1’ Animal lacking adhesive tubes (Fig. 7.3 L–R, T–U) or with one pair (very rarely two pairs) of adhesive tubes posteriorly (Fig. 7.3 A–J), and no pharyngeal pores .............................................................................................................................................. Chaetonotida [p. 119] [common and widespread in fresh-water]

Gastrotricha: Chaetonotida: Families 1 Posterior end of body usually with furca and adhesive tubes; body usually tenpin-shaped or strap-shaped (Fig. 7.3 A–K, S, V, W) ........... 2 1’ Posterior end of body without furca or adhesive tubes, although sometimes bearing spines or pegs; body usually sac- or tenpin-shaped (Fig. 7.3 L–R, T–U) ................................................................................................................................................................................................. 4

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Phylum Gastrotricha

FIGURE 7.5  Schematic representation of scales and spines in some freshwater Chaetonotida: (A) Dorsal three-lobed scale with simple gently curved spine of Chaetonotus (C.) maximus; (B) Dorsal three-lobed scale with dentate spine of C. (C.) similis; (C) Dorsal strongly three-lobed scale with simple spine of C. (C.) larus; (D) Dorsal elongated three-lobed scale with simple spine of C. (C.) hirsutus; (E) Dorsal longitudinally oval scale with simple spine of C. (Primochaetus) mutinensis; (F) Dorsal transversely oval to weakly five-lobed scale with dentate spine bent at almost right angle of C. (P.) heideri; (G) Dorsal longitudinally oval scale with dentate spine of C. (P.) chuni; (H) Dorsal three-lobed scale with spine with two diametrically opposite lateral denticles and bifurcated tip of C. (Schizochaetonotus) schultzei; (I) Dorsal strongly three-lobed scale with dentate spine of C. (Hystricochaetonotus) hystrix; (J) Dorsal three-lobed scale with long thick dentate spine of Hystricochaetonotus sp.; (K) Dorsal five-lobed spineless scale of C. (H.) octonarius; (L) Dorsal semi-circular to weakly three-lobed scale with dentate spine of C. (H.) octonarius; (M) Dorsal weakly five-lobed spineless scale of Hystricochaetonotus sp.; (N) Dorsal polygonal spineless scale of C. (Zonochaeta) succinctus; (O) Dorsal girdle scale with simple spine of C. (Z.) succinctus; (P) Dorsal pedunculate oval scale of Polymerurus rhomboides; (Q) Dorsal scale with constriction and transverse lines thickenings, with thin dentate spine of C. (Captochaetus) robustus; (R) Round scale of Fluxiderma; (S) Rhomboidal scale of Rhomballichthys; (T) Pedunculate scale of Aspidiophorus sp., dorsal view to the left and lateral view to the right; (U) Dorsal smooth scale of Lepidodermella squamata; (V) Dorsal oval keeled scale with short simple spine of Heterolepidoderma ocellatum; (W) Double keeled three-lobed scale anchoring posterior sensory bristles; (X) Three types of ventral interciliary scales: round smooth scale to the left, rounded to weakly oval keeled scale with short simple spine in the middle, and transverse scale plate with jagged anterior edge present in the pharynx region of C. (P.) macrolepidotus ophiogaster; (Y) Small rounded rudimentary scale with strong bidentate spine of Stylochaeta fusiformis. Redrawn from Schwank (1990) and references therein.

2(1) Furca doubly branched; scales and spines sparse or absent (Fig. 7.3 S) ............................................. Dichaeturidae, one genus: Dichaetura [Palaearctic. Rare, not yet reported from the Nearctic] 2’ Furca singly branched; scales and spines present or absent; common and widespread.................................................................................. 3 3(2) Furca branches heavy, sickle-shaped, curved, and tapered, not distinctly divided into a cone-shaped basal part and a distal duct; head with long cilia that are not arranged in tufts; head plates absent (Fig. 7.3 V, W) ........................................................... Proichthydiidae [p. 121] [Neotropical and Palaearctic. Rare, not yet reported from the Nearctic] 3’ Furca branches usually with a cone-shaped base and a distal adhesive duct; body often with numerous spines, scales or spined scales; head with cilia arranged in tufts; cephalic plates present (Fig. 7.3 A–K) .......................................................................... Chaetonotidae [p. 121] [common and widespread] 4(1) Head with clavate tentacles (Fig. 7.3 T, U) .................................................................................................................. Neogosseidae [p. 128] [rare; semiplanktonic or pelagic] 4’ Head without clavate tentacles (Fig. 7.3 L–R); ............................................................................................................. Dasydytidae [p. 129] [rare; semiplanktonic or pelagic]

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FIGURE 7.6  Photographs of live specimens: (A) Aspidiophorus ophiodermus, dorsal view showing pedunculate scales and keeled non-pedunculate scales in posteriormost trunk region; (B) Chaetonotus (Primochaetus) acanthodes, dorsal view showing distribution of scales and the girdle; (C) C. (Hystricochaetonotus) hystrix, dosal view showing distribution of strongly three-lobed scales with dentate spines; (D) C. (H.) macrochaetus, dorsal view showing drastic increase in spine length; (E) C. (Chaetonotus) microchaetus, ventral view of anterior portion of body showing transverse scale plates in the pharynx region; (F) Chaetonotus sp., dorsal view showing distribution of three-lobed scales with simple spines; (G) Ichthydium podura, habitus showing the smooth cuticle. Scale bars: A-B, 20 μm; C, 10 μm; D-G, 20 μm. Photos: T. Kånneby.

Gastrotricha: Chaetonotida: Proichthydiidae: Genera 1 Head bearing a row of cilia shorter than the head (Fig. 7.3 W); .................................................... Proichthydium coronatum Cordero, 1918 [Neotropical. Rare, not yet reported from Nearctic] 1’ Cephalic cilia row much longer than the head (Fig. 7.3 V); ........................................................... Proichthydioides remanei Sudzuki, 1971 [Palaearctic. Rare, not yet reported from Nearctic]

Gastrotricha: Chaetonotida: Chaetonotidae: Genera 1 Furca branches not annulate (not appearing segmented) and furca amounts to less than 20 percent of the total body length....................... 2 1’ Furca branches annulate (appearing segmented) (Figs. 7.3 J and 7.4 G), except for one uncommon European species, and amounts to at least 20–25 percent of total body length; body often large and without a distinct neck ........................................................ Polymerurus [p. 122] [common]

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2(1) Body dorsal surface without spines or scales, sometimes a few scales or spines at the furca bases and/or scales anchoring posterior sensory bristles (Fig. 7.3 A, F)...................................................................................................................................................................................... 3 2’ Body dorsal surface with numerous scales, spines, or spined scales .............................................................................................................. 4 3(2) Cuticle very thick and smooth, distinct from the epidermis, entirely without scales (Fig. 7.3 A); mouth with large mouth ring and strong pharyngeal teeth (Fig. 7.1.19) ................................................................................................................ Arenotus strixinoi Kisielewski, 1987 [Neotropical. Rare monotypic genus not yet reported from Nearctic waters] 3’ Cuticle not especially thick, occasionally with cuticular folds and sometimes with scales near the base of the furca or the bases of posterior sensory bristles (Figs. 7.5 W and 7.6 G), or with minute cuticular structures; mouth ring small and without pharyngeal teeth (Fig. 7.3 F) ������� �������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������������� Ichthydium [p. 123] [common and widespread] 4(2) Spines or spined scales present and often numerous (Fig. 7.3 C, G) .............................................................................................................. 5 4’ Spines absent (occasionally a few spines are present at the base of the furca) ............................................................................................... 6 5(4) Ventral interciliary scales different in shape from dorsal scales; spines of various types (Figs. 7.3 C and 7.6 B–D, F); ................................. ........................................................................................................................................................................................ Chaetonotus [p. 123] [common and widespread] 5’ Ventral interciliary scales similar in shape to dorsal scales; posterior part of body with several long spines that reach beyond the end of the furca (Fig. 7.3 G) ............................................................................................................................ Lepidochaetus zelinkai (Grünspan, 1908) [Canada: New Brunswick, Ontario] 6(5) Furca with adhesive tubes; body strap-shaped or tenpin-shaped; without groups of long cilia on head and posterior body.......................... 7 6’ Furca without adhesive tubes; body markedly tenpin-like, with groups of long cilia on the head and posterior part of the body (Fig. 7.3 K); ................................................................................................................................................................. Undula paraënsis Kisielewski, 1991 [Neotropical. Rare monotypic, semiplanktonic genus not yet reported from Nearctic] 7(6) Body with dorsal scales not pedunculate ........................................................................................................................................................ 8 7’ Body with dorsal surface covered with pedunculate scales (Figs. 7.3 B and 7.5 T), in some rare species scales of posterior dorsal surface non-pedunculate (Fig. 7.6 A)....................................................................................................................................... Aspidiophorus [p. 127] [common]

Phylum Gastrotricha

8(7) Scales not keeled ............................................................................................................................................................................................. 9 8’ Scales elongate to suboval, with longitudinal keels (Figs. 7.3 E and 7.5 V) .......................................................Heterolepidoderma [p. 128] [common and widespread] 9(8) Scales numerous, flat and polygonal............................................................................................................................................................. 10 9’ Few scales, circular; rare (Figs. 7.3 D and 7.5 R) ............................................................................. Fluxiderma concinnum Roszczak, 1935 [USA: New Hampshire, New Jersey, Maine?] 10(9) Scales clearly rhomboid (Figs. 7.3 I and 7.5 S) ................................................................................................................... Rhomaballichthys [Palaearctic. Rare, not yet reported from Nearctic] 10’ Scales smooth and rounded, some with squared edges (Figs. 7.3 H and 7.5 U) ...................................................... Lepidodermella [p. 128] [Common and widespread]

Gastrotricha: Chaetonotida: Chaetonotidae: Polymerurus: Species 1 Cuticle with pedunculate or spined scales....................................................................................................................................................... 2 1’ Cuticle without scales and spines but with numerous small pointed excresences; furca almost as long as half of the total body length......... ................................................................................................................................................................ Polymerurus callosus Brunson, 1950 [USA: Arizona, Illinois, Indiana, Michigan] 2(1) Cuticle with elongated oval, pedunculate scales (Fig. 7.5 P); annulate part of furca without hairs/bristles...................................................... .......................................................................................................................................................... Polymerurus rhomboides (Stokes, 1887) [Canada: Ontario. USA: Illinois, New Jersey, Virginia] 2’ Cuticle with spined scales; spines reach their greatest length at the posterior body end; annulate part of furca with or without hairs/bristles (Fig. 7.4 G) ......................................................................................................................................... Polymerurus nodicaudus (Voigt, 1901) [Canada: Ontario. USA: Indiana, New Jersey, North Dakota, Virginia]

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Gastrotricha: Chaetonotida: Chaetonotidae: Ichthydium: Species 1 Head lobes developed as dorsal flaps (Fig. 7.4 D, E)...................................................................................................................................... 2 1’ Head lobes not developed as dorsal flaps........................................................................................................................................................ 3 2(1) Head with one pair of dorsal flaps (Fig. 7.4 D) ....................................................................................... Ichthydium auritum Brunson, 1950 [USA: Indiana, Michigan] 2’ Head with two pairs of dorsal flaps (Fig. 7.4 E); adhesive tubes approximately 1/3 of the total body length and with pointed tips................ ................................................................................................................................................ Ichhydium macropharyngistum Brunson, 1949 [USA: Michigan] 3(1) Head wider than body, single lobed and rounded to oval or rounded rectangular (Fig. 7.4 C)....................................................................... 4 3’ Head of equal width or narrower than body, three- or five-lobed (Fig. 7.4 A, B) .......................................................................................... 5 4(3) Head rounded to oval; furca extremely short, only 1/25 of the total body length; pharynx with thick medial swelling ................................... ........................................................................................................................................................... Ichthydium brachykolon Brunson, 1949 [USA: Michigan] 4’ Head rounded rectangular; furca forcipate (Fig. 7.4 I) and of normal size; pharynx without swellings........................................................… ......................................................................................................................................................... Ichthydium cephalobares Brunson, 1949 [USA: Michigan] 5(3) Furca distinctly forcipate (Fig. 7.4 I) .............................................................................................................................................................. 6 5’ Furca straight or weakly forcipate with hollow caudal cutting ....................................................................................................................... 7 6(5) Head distinctly five-lobed; total body length less than 100 μm; pharynx pear-shaped .......................... Ichthydium minimum Brunson, 1950 [USA: Michigan] 6’ Head weakly five-lobed, but may appear rounded; total body length more than 100 μm; adhesive tubes fine; posterior sensory bristles anchored by papillae..................................................................................................................................Ichthydium forficula Remane, 1927 [Canada: Ontario. USA: Michigan] 7’ Head weakly three-lobed; furca straight; body plump; cuticle with minute cuticular ridges (Fig. 7.6 G)........................................................ .................................................................................................................................................................... Ichthydium podura (Müller, 1773) [Canada: Ontario? USA: New Jersey] 8(7) Lateral edges of cuticle accordion-like; body covering with 35–40 transverse cuticular ridges ............ Ichthydium sulcatum (Stokes, 1887) [Canada: Ontario. USA: Indiana, Michigan, New Jersey, Virginia?] 8’ Lateral edges of body smooth; base of furca branches slightly enlarged; body covering apparently without transverse cuticular ridges................................................................................................................................................ Ichthydium leptum Brunson, 1949 [USA: Michigan]

Gastrotricha: Chaetonotida: Chaetonotidae: Chaetonotus: Subgenera Members of the subgenera Chaetonotus (Wolterecka) and Chaetonotus (Schizochaetonotus) have been reported from Nearctic waters; however, the specimens were not determined to species or described (see Green, 1986 and Weiss, 2001). 1 Furca not reduced; adhesive tubes developed, never rudimentary .................................................................................................................. 2 1’ Furca more or less reduced, with rudimentary adhesive tubes (Fig. 7.4 H) and strong hooked spines dorsally; benthic; rare.......................... ................................................................................................................................................................................. Chaetonotus (Wolterecka) [rare] 2(1) Cuticle normally developed, uncolored and more or less transparent............................................................................................................. 3 2’ Cuticle thick, three-layered, divided into: outer irregular granular layer, middle layer of rhomboidal scales with simple spines, and inner layer of basal cuticle; cuticle of distinct orange-brown color ............................................................................................................................ .......................................................................................................... Chaetonotus (Tristratachaetus) rhombosquamatus Kolicka et al., 2013 [Palaearctic. Rare, not yet reported from the Nearctic] 3(2) Lateral denticles of spines present or absent, if present never inserted diametrically opposite ..................................................................... 4

Phylum Gastrotricha

7(5) Head distinctly five-lobed; furca weakly forcipate with hollow caudal cutting or base of furca branches slightly enlarged ........................ 8

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3’ Scales three-lobed; spines with two diametrically opposite lateral denticles (Fig. 7.5 H), sometimes a distal denticle also present; benthic and periphytic; one freshwater species ....................................................... Chaetonotus (Schizochaetonotus) schultzei Metschnikoff, 1865 [USA: Arizona] 4(3) Trunk dorsally and laterally almost always without a transverse row (girdle) of long simple-, dentate- or bifurcated spines; if present, girdle-like spines simple and anchoring scales similar to the others of dorsal surface (Fig. 7.6 B) ............................................................... 5 4’ Trunk dorsally and laterally with a transverse row of long simple- dentate- or bifurcated spines (Fig. 7.1.11); scales anchoring girdle spines different in size and shape from others of dorsal surface; at least one pair of long, usually thin parafurcal spines present (Fig. 7.1.13); ................................................................................................................................................................. Chaetonotus (Zonochaeta) [p. 124] [benthic and periphytic; relatively rare] 5(4) Scales rounded- to five-lobed, if three-lobed without longitudinal keel; lateral denticle/s of spines present or absent; scales and spines sometimes reduced in posteriormost dorsal trunk region ............................................................................................................................... 6 5’ Small species, 60–190  μm in total body length; scales three-lobed with distinct longitudinal keel, sometimes rounded three-lobed or more or less pentagonal (Figs. 7.5 K–M and 7.6 C, D); spines usually with one (sometimes two or seldom absent) lateral denticle; many taxa with reduced dorsal spines (and scales) and/or a conspicuous group of spines in trunk region (Fig. 7.4 J); benthic, periphytic and interstitial ���� ������������������������������������������������������������������������������������������������������������������������������������������������� Chaetonotus (Hystricochaetonotus) [p. 124] [common and widespread] 6(5) Total body length usually 100  μm in total body length; girdle with more than 5 spines ............................................................................................... 2 1’ Small species, 20 columns; >32 rows); keels not drawn out into a short simple spine.................................... 3 2’ Total number of dorsal keeled scales lower than above (15–20 columns; 20–25 rows); columns straight; scales with keels drawn out into a short simple spine (Fig. 7.5 V); ocellar granules present (Fig. 7.1.4) (sometimes absent in certain populations/individuals) ......................... ........................................................................................................................................ Heterolepidoderma ocellatum (Metschnikoff, 1865) [Canada: Ontario. USA: Illinois] 3(2) Body relatively long and slender; scales hexagonal or elongate oval in shape; pharynx without distinct swellings, but can widen posteriorly ......................................................................................................................................................................................................................... 4 3’ Body stout and short; scales numerous, round to suboval; ocellar granules absent; pharynx with anterior and posterior swelling ................. ............................................................................................................................................... Heterolepidoderma illinoisensis Robbins, 1965 [USA: Illinois] 4(3) Head five-lobed; 20–25 dorsal columns of delicate hexagonal scales; keels only developed in middle trunk region, sometimes absent altogether; interciliary field naked; ocellar granules present or absent .............................................. Heterolepidoderma gracile Remane, 1927 [Canada: Ontario. USA: Illinois] 4’ Head three-lobed; approximately 25 columns of elongate oval scales; interciliary field with a pair of ventral terminal scales and 6–11 ­columns of scales; ocellar granules absent ..................................................................................... Heterolepidoderma majus Remane, 1927 [Canada: Ontario]

Phylum Gastrotricha

Gastrotricha: Chaetonotida: Chaetonotidae: Lepidodermella: Species 1 Head rounded or five-lobed with two pairs of sensory ciliary tufts................................................................................................................. 2 1’ Head distinctly three-lobed with one pair of sensory ciliary tufts (Fig. 7.4 B) ............................... Lepidodermella triloba (Brunson, 1950) [USA: Indiana, Michigan] 2(1) Seven to nine dorsal columns of smooth scales (Fig. 7.5 U); interciliary area with smooth transverse cuticular scale plates in pharynx region (Figs. 7.1.21 and 7.6 E) ............................................................................................................... Lepidodermella squamata (Dujardin, 1841) [Canada: Ontario. USA: Illinois, Indiana, Michigan, New Hampshire, New Jersey, New York, Ohio, Virginia, Washington] 2’ Thirteen to seventeen dorsal columns of smooth pentagonal to hexagonal scales; interciliary area with >20 columns of fine keels .............. ...................................................................................................................................................... Lepidodermella zelinkai (Konsuloff, 1913) [Canada: Ontario]

Gastrotricha: Chaetonotida: Neogosseidae: Genera Members of the genus Kijanebalola have been reported from Nearctic waters; however, the specimens were not determined to species or described (see Krivanek & Krivanek, 1958). 1 Posterior end of body with two groups of long spines (Fig. 7.3 U); elements of mouth ring jointed .............................. Neogossea [p. 128] [rare] 1’ Posterior end of body with single medial group of spines (Fig. 7.3 T); elements of mouth ring unjointed ................................ Kijanebalola [USA: Louisiana]

Gastrotricha: Chaetonotida: Neogosseidae: Neogossea: Species 1 Several pairs of longer spines in mid-dorsal and mid-lateral trunk region; 6 pairs of caudal spines ................................................................ .............................................................................................................................................. Neogossea sexiseta Krivanek & Krivanek, 1958 [USA: Louisiana]

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1’ Pairs of longer spines absent from mid-dorsal and mid-lateral trunk region; 8–12 pairs of caudal spines ....................................................... …........................................................................................................................................................... Neogossea fasciculata (Daday, 1905) [USA: Louisiana]

Gastrotricha: Chaetonotida: Dasydytidae: Genera 1 Posterior end of body without peg-like protuberances.................................................................................................................................... 2 1’ Posterior end of body with pair of peg-like protuberances (Figs. 7.3 R, 7.4 F, and 7.5 Y).............................................. Stylochaeta [p. 129] [Rare] 2(1) Scales absent or small and inconspicuous; in one South American species scales are large and smooth ...................................................... 3 2’ Body enclosed in a “lorica” of large, thick, ornamented scales (Fig. 7.3 P) .................................Ornamentula paraënsis Kisielewski, 1991 [Neotropical. Rare, not yet reported from the Nearctic] 3(2) Head distinctly wider than neck; body with long lateral spines, some mobile; pharynx with one bulb or no bulb ...................................... 4 3’ Head much narrower than body and scarcely wider than neck; lateral spines absent or identical to dorsal spines, the latter may be reduced to a posterior pair only; pharynx with two bulbs (Fig. 7.3 L) ............................................................................................... Anacanthoderma [Palaearctic. Rare, not yet reported from the Nearctic] 4(3) Lateral spines without denticles or with denticles not directed inwards towards the body ........................................................................... 5 4’ Body with 2–3 pairs of lateral spine bundles; spines with 1–2 lateral denticles directed inward towards the body (Fig. 7.3 M) .................... ..................................................................................................................................................................................................... Chitonodytes [Palaearctic. Rare, not yet reported from the Nearctic] 5(4) Posterior end of body usually with caudal spines ........................................................................................................................................... 6 5’ Posterior end of body rounded, without caudal spines (Fig. 7.3 O) .................................................................................. Haltidytes [p. 129] [rare]

[rare] 6’ Lateral spines tapered, with at most 1 weak lateral denticle and never terminally bifurcated; pharynx without posterior bulb (Fig. 7.3 Q) ................................................................................................................................................................ Setopus bisetosus (Thompson, 1891) [USA: New Jersey]

Gastrotricha: Chaetonotida: Dasydytidae: Stylochaeta: Species 1 Peg-like protuberances (styli) with 3 pairs of equally long hairs (Fig. 7.4 F); three pairs of shorter ventrolateral bifurcated spines anchored by three-lobed scales, anterior pair anchored in anteriormost group of long ventral spines; middle pair anchored between anterior and middle group of long ventral spines and posterior pair anchored in middle group of long ventral spines ........................................................ ............................................................................................................................................................ Stylochaeta fusiformis (Spencer, 1890) [USA: New Jersey] 1’ Peg-like protuberances with 2 pairs of short (5 μm) and 1 pair of long (20–30 μm) hairs; ventrolateral bifurcated spines anchored by threelobed scales absent ................................................................................................................................ Stylochaeta scirtetica Brunson, 1950 [Canada: Ontario. USA: Arizona, Louisiana, Michigan, New Jersey]

Gastrotricha: Chaetonotida: Dasydytidae: Haltidytes: Species 1 One pair of ventral rigid spines ....................................................................................................................................................................... 2 1’ Three pairs of ventral rigid spines with thick bases; 5 pairs of dorsal movable spines ............................ Haltidytes crassus (Greuter, 1917) [Canada: Ontario] 2(1) One pair of rigid straight caudal spines; 4 pairs of ventrolateral movable spines that goes around and cross each other distally .................... ................................................................................................................................................................... Haltidytes saltitans (Stokes, 1887) [USA: New Jersey] 2’ Eight pairs of ventral spines of which 6 pairs are movable ....................................................................... Haltidytes ooeides Brunson, 1950 [USA: Arizona, Michigan]

Phylum Gastrotricha

6(5) Lateral spines with 1–3 lateral denticles and often terminally bifurcated; spines of uniform thickness from their base to the last lateral denticle; pharynx usually with a distinct posterior bulb (Fig. 7.3 N) ............................................................................... Dasydytes [p. 130]

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Gastrotricha: Chaetonotida: Dasydytidae: Dasydytes: Species 1 Two rows of dentate spines present in neck region; two pairs of shorter spines on head; dorsal spines bidentate, not bifurcated ................... ....................................................................................................................................................................... Dasydytes monile Horlick, 1975 [USA: Illinois] 1’ Spines in neck region do not form two rows, one pair of longer spines on head; dorsal spines dentate with bifurcated tips ........................... ...............................…................................................................................................................................. Dasydytes goniathrix Gosse, 1851 [USA: Indiana]

REFERENCES

Phylum Gastrotricha

Balsamo, M. 1983. Gastrotrichi (Gastrotricha). Guide per il Riconoscimento delle Specie Animali delle Acque Interne Italiane 20. Consiglio Nazionale delle Richerce. Balsamo, M. & M.A. Todaro. 2002. Gastrotricha. Pages 45–61 in: S.D. Rundle, A.L. Robertson & J.M Schmid-Araya (eds.). Freshwater meiofauna: biology and ecology. Backhuys Publishers, Leiden. Brunson, R.B. 1950. An introduction to the taxonomy of the Gastrotricha with a study of eighteen species from Michigan. Transactions of the American Microscopical Society 69:325–352. Brunson, R.B. 1959. Gastrotricha. Pages 406–419 in: W.T. Edmondson (ed.). Fresh-water biology. Second edition. John Wiley and Sons, New York. Green, J. 1986. Associations of zooplankton in six crater lakes in Arizona, Mexico and New Mexico. Journal of Zoology, London (A) 208: 135–159. Greuter, A. 1917. Beiträge zur Systematik der Gastrotrichen in der Schweiz. Revue Suisse de Zoologie 25:35–76. d’Hondt, J.L. 1971. Note sur les quelques Gastrotriches Chaetonotidae. Bulletin de la Société, Zoologique de France 96:215–235. Horlick, R. 1975. Dasydytes monile, a new species of gastrotrich from Illinois. Transactions of the Illinois State Academy of Sciences 68:61–64. Kånneby, T., M.A. Todaro & U. Jondelius. 2009. One new species and records of Ichthydium Ehrenberg, 1830 (Gastrotricha: Chaetonotida) from Sweden with a key to the genus. Zootaxa 2278:26–46. Kånneby, T., M.A. Todaro & U. Jondelius. 2012. A phylogenetic approach to species delimitation in freshwater Gastrotricha from Sweden. Hydrobiologia 683:185–202. Kånneby, T., M.A. Todaro & U. Jondelius. 2013. Phylogeny of Chaetonotidae and other Paucitubulatina (Gastrotricha: Chaetonotida) and the colonization of aquatic ecosystems. Zoologica Scripta 42:88–105. Kånneby, T. & M.K. Wicksten. 2014. First record of the enigmatic genus Redudasys Kisielewski, 1987 (Gastrotricha: Macrodasyida) from the Northern hemisphere. Zoosystema 36(4): 1–12. Kisielewski, J. 1981. Gastrotricha from raised and transitional peat bogs in Poland. Monografie Fauny Polski 11:1–143. Kisielewski, J. 1986. Taxonomic notes on freshwater gastrotrichs of the genus Aspidiophorus Voigt (Gastrotricha: Chaetonotidae), with descriptions of four new species. Fragmenta Faunistica 30:139–156.

Kisielewski, J. 1987. Two new interesting genera of Gastrotricha (Macrodasyida and Chaetonotida) from the Brazilian freshwater psammon. Hydrobiologia 153:23–30. Kisielewski, J. 1991. Inland-water Gastrotricha from Brazil. Annales Zoologici 43, Supplement 2:1–168. Krivanek, R.C. & J.O. Krivanek. 1958. Taxonomic studies on the Gastrotricha of Louisiana. ASB Bulletin 5: 12. Remane, A. 1935–1936. Gastrotricha und Kinorhyncha. Klassen und Ordnungen des Tierreichs, Band 4, Abteilung 2, Buch 1, Teil 2, Lieferungen 1-2: 1-242, 373–385. Robbins, C.E. 1965. Two new species of Gastrotricha (Aschelminthes) from Illinois. Transactions of the American Microscopical Society 84:260–263. Robbins, C.E. 1973. Gastrotricha from Illinois. Transcations of the Illinois State Academy of Science 66: 124–126. Ruttner-Kolisko, A. 1955. Rheomorpha neiswestnovae und Marinellina flagellata, zwei phylogenetisch intressante Wurmtypen aus dem Süsswasserpsammon. Österreichische Zoologische Zeitschrift 6:55–69. Schwank, P. 1990. Gastrotricha. Pages 1–252 in: J. Schwoerbel & P. Zwick (eds.). Süsswasserfauna von Mitteleuropa, Band 3. Gustav Fischer Verlag, Stuttgart. Schwank, P. & T. Kånneby. 2014. Contribution to the freshwater gastrotrich fauna of wetland areas of southwestern Ontario (Canada) with redescriptions of seven species and a check-list for North America. Zootaxa 3811(4): 463–490. Strayer, D., Hummon, W.D., Hochberg, R. 2009. Gastrotricha. In: Thorp, J.H., Covich, A.P. (Eds.), Ecology and Classification of North American Freshwater Invertebrates, third ed. Elsevier Inc., New York, pp. 163–172. Sudzuki, M. 1971. Die das Kapillarwasser des Lückensystems Bewohnenden Gastrotrichen Japans. I. Zoological Magazine (Tokyo) 80: 256–257. Tirjaková, E. 1998. Zaujímavý nález Brušnobrvca (Gastrotricha) zčel’ade Dichaeuturidae. Folia Faun. Slovaca 3, 19–21. Todaro, M.A., Dal Zotto, M., Jondelius, U., Hochberg, R., Hummon, W.D., Kånneby, T., Rocha, C.E.F. 2012. Gastrotricha: a marine sister for a freshwater puzzle. PLoS ONE 7 (2), e31740. Weiss, M.J. 2001. Widespread hermaphroditism in freshwater gastrotrichs. Invertebrate Biology 120: 308–341.

Chapter 8

Phylum Rotifera Robert L. Wallace Department of Biology, Ripon College, Ripon, WI, USA

T.W. Snell School of Biology, Georgia Institute of Technology, Atlanta, GA, USA

E.J. Walsh Department of Biological Science, University of Texas at El Paso, El Paso, TX, USA

S.S.S. Sarma Universidad Naćional Autónoma de México Campus Iztacala, México

Hendrik Segers Royal Belgian Institute of Natural Sciences, Brussels, Belgium

Chapter Outline

INTRODUCTION Classification schemes differ slightly in how they regard the four groups of rotifers: Seisonidea, Bdelloidea, Monogononta, and Acanthocephala, the latter being an obligatorily parasitic taxon previously treated as a distinct phylum (Garey et al., 1996; Garey et al., 1998; Mark Welch & Meselson, 2000; Fontaneto & Jondelius, 2011). The peculiar problem is that outside of the phylogenetic literature, acanthocephalans and rotifers are still treated separately. Basically, each group has ignored much of the research done by the other. However, as evidenced in the works cited above, there is ample support to unite the two taxa within a single classification. First, they both share an important, unique, synapomorphic characteristic. Within the syncytial integument of both lies a curious layer of two proteins called the intracytoplasmic lamina. These proteins provide a degree of stiffness to body wall depending on its thickness. More importantly, molecular evidence supports

Material Preparation and Preservation 134 Key to Freshwater Rotifers (Class Eurotatoria) 135 Rotifera: Bdelloidea Families 136 Rotifera: Monogonata: Orders 139 Rotifera: Monogonata: Collothecacea: Families 140 Rotifera: Monogononta: Flosculariaceae: Families 142 Rotifera: Monogononta: Ploima: Families 148 References165

uniting these taxa; indeed, researchers are now arguing that acanthocephalans ought to be subsumed within Rotifera (Sørensen & Giribet, 2006; Segers, 2007; Wey-Fabrizius et al., 2014). Thus, a new classification is emerging that asserts that acanthocephalans and seisonids are sister groups within Pararotatoria (Min & Park, 2009; Fontaneto & Jondelius, 2011; Wey-Fabrizius et al., 2014). Pararotatoria and bdelloids then comprise Hemirotifera, with Hemirotifera and Monogononta comprising the Syndermata. Syndermata is then a sister taxon to the Gnathostomulida, thus creating a group of small, jawed animals, the Gnathifera (Sørensen, 2001; Leasi et al., 2012). The close affinity among these taxa has led some to propose that the Gnathifera be treated as a distinct phylum (Shiel et al., 2009). However, given the scope of this work, we ignore the problem of where to place the acanthocephalans and gnathostomulids, but look forward to additional study that will provide a better synthesis of these taxa.

Thorp and Covich’s Freshwater Invertebrates. http://dx.doi.org/10.1016/B978-0-12-385028-7.00008-1 Copyright © 2016 Elsevier Inc. All rights reserved.

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Introduction131 Systematics of Rotifer Classes 132 Class Pararotatoria, Family Seisonidea 132 Class Eurotatoria, Subclass Bdelloidea 132 Class Eurotatoria, Subclass Monogononta 132 Rotifers of the Nearctic 132 Limitations132 Terminology and Morphology 133

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At one time rotifers were divided into two classes: those with two gonads (seisonids and bdelloids) were considered to be orders within class Digononta, leaving the Monogononta as a separate class (Pennak, 1989). In this key, two classes, Pararotatoria and Eurotatoria, are recognized (Melone & Ricci, 1995; Smith, 2001; Segers, 2002; Wallace et al., 2006; Wallace & Smith, 2009; Wallace & Snell, 2010). Older classification schema that considers rotifers as a class within the phylum Aschelminthes should be abandoned.

Systematics of Rotifer Classes

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to as the limnoterrestrial (Devetter, 2008; Segers, 2008). Bdelloids also are present in communities that develop in tree holes (Devetter, 2008) and the pitfall traps of the carnivorous plant Sarracenia purpurea Linnaeus, 1753 (Bateman, 1987; Błędzki & Ellison, 1998). Many species are capable of becoming desiccated and then rehydrated (Ricci & Fontaneto, 2009; Wilson, 2011). Unfortunately, there has been no systematic review of Nearctic bdelloids, but there are several works that should be consulted for more information (Bartoš, 1951; Donner, 1965; Pourriot, 1979; Koste & Shiel, 1986; Melone et al., 1998; Ricci & Melone, 2000).

Class Pararotatoria, Family Seisonidea

Class Eurotatoria, Subclass Monogononta

This taxon comprises only two genera (Paraseison and Seison), each with two species. These are large (2–3 mm) dioecious, marine rotifers that live on the gills of crustaceans (Ricci, 1993; Sørensen et al., 2005; Leasi et al., 2011; Leasi et al., 2012). Sometimes described as aberrant, the corona of seisonids is very reduced and not used in locomotion. All species have paired gonads and a functional gut in both sexes. The jaws or trophi are described as being fulcrate. Females have ovaries without vitellaria. Sexes are of similar size and morphology.

The monogononts comprises the largest group of rotifers with >1500 species in more than 100 genera. While most are free-living, taking a benthic, free swimming, or sessile existence, a few genera are parasitic. Nearly all rotifers are found in inland waters, both fresh and alkaline, but some appear to be exclusive to the marine habitat. In many species, males have never been observed, but all monogononts are assumed to be dioecious. As the name implies, these rotifers possess a single gonad. Males are often structurally reduced with a vestigial gut that functions only in energy storage (Ricci & Melone, 1998). They have a shorter lifespan than females and are usually present in the plankton for only a few days or weeks each year. Monogononts are microphagous or raptorial, but a few are parasitic. The corona is more varied than in other rotifers; the corona may range from broad to narrow disks or possess ear-like lobes or be vase-shaped with reduced ciliation and long setae that are used in prey capture. This key recognizes two superorders: Pseudotrocha (order Ploima) and Gneisotrocha (orders Collothecaceae and Flosculariaceae).

Class Eurotatoria, Subclass Bdelloidea

Phylum Rotifera

This important subclass comprises 19 genera and, depending on the author, approximately 350–460 species (Ricci, 1987; Segers, 2002, 2007, 2008). A complicating factor in determining an accurate estimation of the number of species is that cryptic speciation (hidden biological diversity) appears to be widespread in bdelloids (Fontaneto et al., 2007a, 2009, 2011). Bdelloids possess a somewhat uniform body plan (Donner, 1965; Melone & Ricci, 1995) and are exclusively females, reproducing by parthenogenesis. Bdelloids are characterized by having paired ovaries with vitellaria, more than two pedal glands, and ramate trophi (Melone et al., 1998; Wallace & Ricci, 2002; Wallace & Snell, 2010). Nearly all bdelloids are microphagous with a corona of either two trochal discs or a modified ciliated field. Bdelloids often have a vermiform body with a pseudo-segmentation consisting of annuli that permits shortening and lengthening of the body by telescoping. Current taxonomy recognizes three orders: Adinetida (family Adinetidae), Philodinida (families Habrotrochidae and Philodinidae), and Philodinavida (family Philodinavidae) (Melone & Ricci, 1995). Bdelloids generally are not caught in plankton tows, although they may be found in waters with dense vegetation. On the other hand, they often occur in sediments, among plant debris, or crawling on the surfaces of aquatic plants. Some forms inhabit the capillary water films of soils (Pourriot, 1979; Devetter, 2010) or covering mosses (Burger, 1948; Peters et al., 1993; Ricci & Caprioli, 2005; Bielańska-Grajner et al., 2011); this habitat has been referred

Rotifers of the Nearctic Here we recognize Phylum Rotifera to comprise 35 families with >130 genera. Of these, >100 genera and >900 species have been reported from inland waters of the Nearctic (NEA). Because of the relatively little taxonomic work done in the Nearctic, we feel that this assessment is a gross underestimation of the true species, richness.

LIMITATIONS Except for the incomplete Guide Series (Segers, 1995a,b; De Smet, 1996; De Smet & Pourriot, 1997; Nogrady & Segers, 2002; Wallace et al., 2006), there has been no attempt to construct a comprehensive key to Rotifer. There are a variety of keys to rotifers, including those by Edmondson (1959) and Smith (2001), which cover some of the North American fauna to the level of genus, and Stemberger (1979) for the Laurentian Great Lakes, which provide a species-level key. Although specialized and covering other biogeographical

Chapter | 8  Phylum Rotifera

The obligatory marine taxon, class Pararotatoria (family Seisonidae) is not included herein. The following key is based on several important works (Koste, 1978; Segers, 2004; Wallace et al., 2006). The species records annotated herein for the Nearctic are noted in brackets; monospecific genera are designated as such. All data were derived from (Segers, 2007).

TERMINOLOGY AND MORPHOLOGY Most rotifer trophi are composed of seven hardened elements or sclerites (Fig. 8.1; also Fig. 13.19 in Volume I). Three of these, the manubrium (manubria), ramus (rami), and uncus (unci) are paired, while the fulcrum is unpaired (see also Fig. 13.10 in Volume I). The central most elements are the rami that may contain one or more chambers. The fulcrum, which is absent in bdelloids, is located caudally between the rami and articulates with them. Together, the fulcrum and paired rami comprise the incus. Located left and right of the rami are the manubrium and uncus; together they comprise the malleus (plural, mallei). These elements are all held together and are moved by muscular action. Besides the discussion in Volume I of this series, additional information on rotifer trophi may be found in the following works: (Donner, 1956; Edmondson, 1959; Koste, 1978; Sørensen, 2006). The nine different types of trophi were established based on the size, shape, and positioning of the seven elements described above, as well as the presence of other accessory parts and variations. The “Rotifer trophi web page” (Anon, 2014) also provides some excellent scanning electron photomicrographs. Of the nine types, the fulcrate trophi of seisonids is not considered in this key. The other eight types, which are important to this key (cardate, forcipate, incudate, malleate, malleoramate, ramate, uncinate, and virgate), are briefly discussed below. They are also described in detail in Chapter 13 of Volume I.

FIGURE 8.1  Schematic view of the basic elements of rotifer trophi. (See also Figs. 13.10 and 11 in Volume I for a complete overview of the trophi.)

Phylum Rotifera

realms, valuable keys can be found for Australian waters (Koste & Shiel, 1986, 1987, 1989a,b, 1990a,b, 1991; Shiel & Koste,1992, 1993; Shiel, 1995), the British Isles (Pontin, 1978), Italy (Braioni & Gelmini, 1983), and planktonic rotifers in general (Ruttner-Kolisko, 1974). Unfortunately, no key is comprehensive enough to cover all of the variations in size and morphology that are sometimes found within a species. Of all the efforts, Koste’s (1978) revision of Voigt for the rotifers of central Europe comes the closest; although over 35 years old, it is still important. We urge caution in using keys that are available in electronic form (e.g., Internet or on CD-ROMs). In most instances, these are far too simplistic, often reflecting a suite of species from a rather narrow region; in no way should they be considered thorough. Nevertheless, Jersabek et al. (2014) and Jersabek & Leitner (2013) are two very valuable online resources. A further limitation stems from the fact that too few species have photographic documentation. Thus, as with other keys to the Rotifera, we caution that the illustrations used in this one are not necessarily from specimens collected from the Nearctic, and we have had to rely on illustrations from the older literature. When the figures used to demonstrate the general form of a taxon are from the Nearctic, they are designed with the abbreviation NEA. For this work, we estimated the number of species in each genus using the assessment of Segers (2007) and Jersabek & Leitner (2013), including subspecific variants (subspecies), but not those species noted as species inquirenda—species requiring additional investigation before being accepted as a valid taxon. We also did not count species that appear to be strict halophiles (only inhabiting marine and brackish waters), recognizing that they could still be present in alkaline, inland waters. Finally, it should be noted that emerging molecular data supports the concept of cryptic speciation in rotifers. Thus, there may be 3–14 times as many species within any currently recognized morphospecies (Gómez, 2005; Fontaneto et al., 2007a,b; Schröder and Walsh, 2007; Garcia-Morales and Elias-Gutierrez, 2013). One of the fundamental differences among the higher taxonomic levels in the phylum Rotifera is the structure of the trophi that reside in a muscular pharynx called the mastax. Nine different types of trophi are recognized, along with various transitional forms (Koste, 1978; Koste & Shiel, 1987). Although the following key is designed with the non-specialist in mind, commensurate with the central importance of the trophi, we use both the structure of the trophi and other obvious characters of anatomy and morphology as principal points of separation of the taxa. Therefore, it is important to note that you may have to destroy one or more specimens to get a good look at the trophi. Consequently, because other morphological features also are critical to identification, it is important that you reserve several specimens to study whole, preferably alive. Taking photomicrographs of the live, intact animal before preservation or hydrolysis is always a prudent practice.

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Other important terms are defined below. Alula—Wing-shaped projections present at the base of each ramus. l  Basal chamber—Found in some trophi, a chamber or depression in the ramus with a window-like opening (basifenestra) that opens on the dorsal side. l Cardate trophi—Trophi with a wide fulcrum and rami in the shape of a lyre (U-shaped). l  Cirri—Intermediate length (cilia-like) structures that are present in the coronal region. l Forcipate trophi—Trophi in the form of forceps that can be extruded from the mouth (found in some creeping and parasitic species; no planktonic and semiplanktonic); compare to Incudate. l  Cryptic species—More than one population from the same or different habitats that with currently recognized morphological metrics are difficult or impossible to diagnose as species, but that may be differentiated by their genetic signatures. Together these form a species complex (e.g., Brachionus plicatilis species complex). l  Gelatinous tube—A thick secretion from glands in the body that form a gelatinous mass that may remain transparent or become occluded with bacteria, algae, and/or detritus. l  Hemiforcipate trophi—The specialized jaws of family Asciaporrectidae as described by De Smet (2006). l Hypopharyngeal muscle—Powerful muscle in the mastax that helps in creating a strong pumping action in some species. l  Incudate trophi—The trophi of family Asplanchnidae (planktonic and semiplanktonic) that possess large, pincer-like rami, but reduced manubria and unci. l Illoricate—A term used for rotifers with a thin body wall: one that is not thickened by the proteins of the intracytoplasmic lamina (compare loricate). l  Infundibulum—A cup-shaped corona found in Order Collothecacea. l  Intramalleus—An accessory part found in the trophi of some species; a connecting piece between the uncus and the manubrium. l Loricate—A term used for rotifers with a thick body wall or lorica: one that is thickened by the proteins of the intracytoplasmic lamina; compare to illoricate. l  Malleate trophi—Trophi with a short fulcrum, but the manubrium and rami are strong; unci with 4–7 teeth; compare to Malleoramate. l  Malleoramate trophi—Rami with strong teeth and unci with many slender teeth; compare to Malleate. l Modulus—A specialized organ located at the anterioventral end of some Floscularia that makes small, round, or bullet-shaped pellets (sometimes called pseudofecal pellets) that are composed of bacteria and tiny bits of detritus. The animal uses the pellets to construct a tall, turret-like l

structure as found in many medieval castles. These pellets should not be confused with the fecal masses that a few species embed within a more extensive gelatinous tube. l  Morphospecies—A species designated and clearly separated from other species based on recognized unequivocal morphological features. l  Oral plates—An accessory part found in some trophi; plates with tooth-like structures associated with the rami. l  Oviferon—A structure in Sinantherina and Pentatrocha that holds their parthenogenetic eggs. l Preunci—An accessory part found in some trophi; a separate set of teeth present under the teeth in the uncus. l Pseudunci—An accessory part of large hooked structures found in the trophi of family Birgeidae. l Ramate—The trophi of bdelloid rotifers possessing wide, curved to half moon-shaped unci, but lacking the fulcrum. l Rostrum—A small to prominent, beak-like structure that projects from the anterior end of rotifers, especially in some bdelloids. l  Setae—Very long cilia that line the edge of the corona (infundibulum) in Collotheca and the elongate arms (lobes) of Stephanoceros. l  Spurs—Short to long protrusions that project dorsally at the terminus of the foot. Note that the foot itself may be much shorter than the toes (compare toes). l  Subbasal chamber—A chamber or depression in the ramus with a window-like opening (subbasifenestra) that opens on the ventral side. l Subunci—An accessory part found in some trophi; a narrow rim of small fused bumps or ridges that are attached to the uncus near the teeth. l  Sulcus—A hollow depression where the lorica is not overly thickened, usually between two regions or plates where the lorica is thickened (see loricate). l Toes—Short to long protrusions that project from the terminus of the foot in many rotifers (compare spurs). l  Trochal pedicels—Paired parts of the bdelloid corona elevated by a flexible stalk above the rest of the body. l  Uncinate—The trophi of collothecid rotifers, possessing well-developed rami, but weak fulcrum and manubria; 3–5 teeth in the unci. l Virgate—Trophi with an action that functions in pumping; often asymmetrical with most elements, having a tendency towards being slender; fulcrum and manubrium long.

Phylum Rotifera

MATERIAL PREPARATION AND PRESERVATION Whenever possible, specimens should be examined alive and then studied after preservation to appreciate the consequence of fixation on body shape. This is especially important for bdelloids and illoricate monogononts, which will shrivel into a mass of tissue that is usually impossible

Chapter | 8  Phylum Rotifera

to identify. While the trophi of these specimens may be extracted and used identification, the details of the body are often needed as well. Solutions of sugar (40% sucrose) and 4% formalin (Haney & Hall, 1973) or ethanol (Black & Dodson, 2003), which are employed in the preservation of cladocerans, do not prevent contraction in rotifers. However, in loricate taxa, such as Lecane, formalin fixation is important. In this case a so-called, hard fixation, with higher concentrations of formalin (>4%) will cause the tissues to contract revealing the details of the surface of the lorica, which is critical to the identification to the level of species (Segers, 1995a). Visualization of preserved rotifers may be enhanced by the use of the stain Rose Bengal (Rublee & Partusch-Talley, 1995; Rublee, 1998). While molecular techniques are being used to identify cryptic species in rotifers, its use has not yet been applied to the routine identification of species. As of yet, there is no single anesthetic that has proved to be useful for a wide range of species. For example, carbonated water (club soda), 1% solution of MgCl2, and various other narcotizing agents have been tried with mixed success. For a review of these techniques, consult the following publications: Edmondson (1959), Nogrady & Rowe (1993), Wallace et al. (2006), as well as Chapter 13 in Volume I. Two older techniques that kill the specimen in a more-orless natural condition are the hot-water fixation technique of Edmondson (1959) and the formalin drop technique. However, both require much practice to master. In the later procedure, minute drops of formalin (400 individuals (>4 mm in diameter) have been reported from deep lakes (Wallace et al., 2015).]

Chapter | 8  Phylum Rotifera

143

FIGURE 8.18  Cupelopagis vorax. Side view: animal attached to the undersurface of a hydrophyte; insert: from above, animal attached in a Petri dish. Body, without corona ∼250 μm. (NEA)

FIGURE 8.20  Atrochus tentaculus. Ventral view. Bar = 100 μm. After Wierzejski, 1893.

FIGURE 8.21  Hexarthra. Bar = 100 μm. (NEA)

Phylum Rotifera

FIGURE 8.19  Acyclus inquietus. Animal within a colony of S. socialis (left); two adult Acyclus attached to a bit of substrate along with one S. socialis (right). Abbreviations: sse = S. socialis embryos; ssw = S. socialis warts; aie = A. inquietus embryos. Bars ∼100 μm. (NEA) Photomicrographs courtesy of Adele Hochberg, University Massachusetts at Lowell.

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FIGURE 8.22  Conochilus. Small colony of C. unicornis (left). Dashed lines partially outline the margin of the gelatinous matrix. Dark ovoid objects are euglenoid algae colonizing the gelatinous matrix. Individual C. unicornis (right). Arrow = fused antennae within the corona. Bar = 100 μm. (NEA)

FIGURE 8.23  Conochiloides: an adult with one juvenile. Dashed lines partially outline the margin of the gelatinous matrix. Dark ovoid objects are euglenoid algae colonizing the gelatinous matrix. Bar = 250 μm. (NEA)

2’ Corona perpendicular to body axis; antennae located dorsally outside of the coronal field; diapausing eggs are ornamented with spirals; colonies not spherical, typically an adult with 0–3 juveniles; colony’s gelatinous matrix usually colonized by euglenoid algae (Fig. 8.23) [4 NEA of 4 species] ……………………………………………………………………………. Conochilus (Subgenus Conochiloides) spp. [Genus Conochiloides has been subsumed within Conochilus (Ruttner-Kolisko, 1974; Koste, 1978; Segers & Wallace, 2001).]

Rotifera: Monogononta: Flosculariaceae: Flosculariidae: Genera [9 genera; 8 reported in NEA] 1 Dorsal antennae (paired), not conspicuously long but not visible when animal contracts ………………………………………………… 2 1’ Single, centrally placed, dorsal antenna, conspicuously long (longer than body width), visible when animal is contracted, ≤500 μm. (Fig. 8.24) [monospecific] ……………………… ……………………………………………………………... Beauchampia crucigera (Dutrochet, 1812) 2(1) Corona with one or more pairs of distinct ear-like lateral lobes …………………………………………………………………………… 3 2’ Corona without distinct ear-like lobes (corona circular, subcircular, or at most, lobes indistinct to cordiform) …………………………….... 6 3 Corona with more than one pair of distinct lateral lobes on each side of the animal; tube a gelatinous matrix with or without pellets or debris ……………………………………………………………………………………………………………………………... 4 3’ Corona with one pair of distinct lateral lobes on each side of the animal; animals usually solitary (occasionally small colonies, usually ≤12) in separate straight or slightly curved tubes, each formed as a distinct pipe of hardened secretions (Fig. 8.25); ≤1000 μm. [2 species with ringed tubes; 3 species with tubes as a granular matrix-(stucco-like)]; 3 NEA of 5 species] …………………………………... Limnias spp. 4(3) Corona very large with more than 4 lobes ……………………………………………….…………………………………………………. 5 4’ Corona large with two pairs of distinct lateral lobes (= 4 lobes), one pair above the other; animals often colonial (Fig. 8.26); ≤1500 μm. [7 NEA of 9 species] ………………………………………………………………………………………………………… Floscularia spp. [Some species make pellets employing a specialized organ (modulus) located near the mouth on the ventral side; these are used to construct a pellet tube in which they reside (e.g., F. conifera, F. ringens) (Edmondson, 1945; Fontaneto et al., 2003). One species (F. janus) embeds fecal pellets in a gelatinous tube (cf. Ptygura); in F. melicerta the modulus produces a gelatinous tube in layers without pellets, but debris may accumulate within the jelly. Consult Segers (1997) for additional information.]

Phylum Rotifera

5(4) Corona with four pairs of distinct lateral lobes (= 8 lobes), one pair smaller; solitary in a gelatinous tube (Fig. 8.27); 200–2000 μm. [monospecific] ……………………………………………………………………………………………... Octotrocha speciosa Thorpe, 1893 [While the report from the Nearctic by Edmondson (1959) was called into question by Segers et al. (2010), specimens of this species have been observed in samples from Wisconsin (E.J. Walsh, personal observation; Fig. 8.27) and by Sarma & Elías-Gutiérrez (1998) from Mexico. For comments and additional illustrations, consult Segers & Shiel (2008), Segers et al. (2010), Meksuwan et al. (2011).] 5’ Corona with 5 promanent and 2 small lobes; solitary or colonial in a gelatinous tube; ≤1300 μm. Consult (Meksuwan et al., 2011) for figures and additional details. [monospecific] …………………………………………………………... Lacinularoides coloniensis (Colledge, 1918) 6(2) Corona cordiform (reniform), subquadrangular, or occasionally round; animals usually forming spherical to ellipsoid colonies with many individuals, occasionally solitary; tube, when present, a continuous gelatinous matrix …………………………………………………… 7

Chapter | 8  Phylum Rotifera

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FIGURE 8.25  Limnias: L. melicerta (left) and L. ceratophylli (right). Bars = 100 μm. (NEA) FIGURE 8.24  Beauchampia. Bar = 100 μm. (NEA)

FIGURE 8.27  Octotrocha speciosa. Solitary individual in a gelatinous tube. Bar = 250 μm. (NEA) Photomicrograph courtesy of E.J. Walsh, University of Texas at El Paso.

Phylum Rotifera

FIGURE 8.26  Floscularia: F. conifera, small colony (left); F. ringens, anterior (center); F. janus (right). Bars = 100 μm. (NEA) Center photomicrograph courtesy of E.J. Walsh, University of Texas at El Paso.

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FIGURE 8.28  Ptygura: P. beauchampi (sessile), P. libera (planktonic, with two embryos), and P. pilula (sessile, with fecal pellets embedded in a gelatinous tube.) Bars = 50 μm. (NEA)

(A)

(B)

(C)

FIGURE 8.29  Sinantherina. Planktonic colony (left) ≥1 mm diameter and S. socialis (right). Bar = 200 μm. (NEA)

6’ Corona round to slightly elliptical, or weakly-lobed; animals usually solitary in separate, gelatinous tubes that may be covered by or infiltrated with debris (Fig. 8.28); 150–1000 μm. [19 NEA of 31 species] ………………………………………………………… Ptygura spp. [One common species (P. pilula) embeds ovoid, fecal pellets in its gelatinous tube (cf. Floscularia janus); another, P. (Floscularia) noodti, (not been reported from the Nearctic), embeds elongate (vermiform) fecal pellets within its gelatinous tube. Some species of Ptygura may form intra- or interspecific colonies with other sessile Flosculariidae (Wallace, 1987).] 7(6) Possessing a specialized egg bearing structure (oviferon, ovifer) below anus to which eggs attach; solitary or colonial; sessile or planktonic; gelatinous matrix is absent (minimal?); individuals: 400–2000 μm, colonies: ≤4 mm. (Fig. 8.29) [4 NEA of 5 species] ………… …………… ……………………………………………………………………………………………………………….. Sinantherina spp. [Pentatrocha gigantea Segers & Shiel, 2008 also possesses an oviferon, but it has not been identified from the Nearctic. However, Segers & Shiel (2008) suggest potential confusion of this species with Octotrocha speciosa (see above). Both genera deserve additional study.] 7’ Lacking an oviferon; usually colonial; sessile or planktonic; possessing gelatinous matrix; individuals: ≤2000 μm, colonies: ≤5 mm. (Fig. 8.30) [2 NEA of 6 species] …………………………….……………………………………………………………..... Lacinularia spp.

Phylum Rotifera

Rotifera: Monogononta: Flosculariaceae: Trochosphaeridae: Genera Consult Nogrady & Segers (2002) for details. [3 genera; all reported in NEA] 1 Body spherical-shaped (a round ball) or slightly elongate (oviform) ………………………………………………………………………. 2 1’ Rotifers not as above; possessing two movable anterior setae (bristles or spines) of varying lengths below the corona, often much longer than the body and one (rarely two) rigid caudal spines; foot absent; (Fig. 8.31); excluding setae ∼40–325 μm. [8 NEA of 15 species] …… ……………………………………………………………………………………………………………………………………. Filinia spp. [Filinia (formerly Family Filiniidae) has been subsumed within Family Trochosphaeridae (Nogrady and Segers, 2002).]

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FIGURE 8.30  Lacinularia. Bar = 250 μm. (NEA)

FIGURE 8.31  Filinia. Bar = 100 μm. Photomicrograph courtesy of M.V. Sørensen, University of Copenhagen.

(A)

(B)

FIGURE 8.32  Family Trochosphaeridae. (A) Trochosphaera. (B) Horaella. Bars = 100 μm. (NEA) Photomicrographs courtesy of S.S.S. Sarma, Universidad Naćional Autónoma de México Campus Iztacala, México.

2(1) Body spherically-shaped, corona as a circular band of cilia around the equator or towards one end; 320–1100 μm; viviparous; (Fig. 8.32 A). [2 NEA of 2 species] ……………………………………………………………………………………………………. Trochosphaera spp.

Rotifera: Monogononta: Flosculariaceae: Testudinellidae: Genera [3 genera, 2 in NEA] 1 Lorica greatly flattened dorsoventrally with dorsal and ventral plates fused along the lateral margin (Fig. 8.33 A); foot annulated and retractile; ≤250 μm. [18 NEA of 38 species] .................................................................................................................................. Testudinella spp. 1’ Lorica not greatly flattened, rather possess 4 nearly equal lobes in cross section; 1 pair frontal eyespots; foot absent; (Fig. 8.33 B); ≤120 μm. [3 NEA of 3 species] ............................................................................................................................................................... Pompholyx spp.

Phylum Rotifera

2’  Body not spherically-shaped, slightly elongate (oviform); corona a slight extension of the body; ∼125 × 160 μm; oviparous; (Fig. 8.32 B). [1 NEA of 2 species] ……………………………………………………………………... Horaella thomassoni Koste, 1973

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Rotifera: Monogononta: Ploima: Families This key does not include two familes: family Clariaidae (Claria), which comprises a single species of endoparasitic rotifers of terrestrial Oligochaeta (Segers, 2008), and the monospecific Cotylegaleatidae (Cotylegaleata) (De Smet, 2007). Neither has been reported from the Nearctic. Also,

the separation of two curious congenerics (Pourriotia, formerly Proales, Proalidae, currently Notommatidae), may be made based on the fact that both are parasitic in galls of the algal genus Vaucheria.

1 Rotifers with forcipate (Fig. 13.11 j in Volume I) or modified (protrusible) virgate trophi ………………………………………………… 2 1’ Rotifers with trophi other than above ………………………………………………………………………………………………………. 4 2(1) Not inhabiting shells of live testate amoeba ………………………………………………………………………………………………… 3 2’ Inhabiting shells of live testate amoeba (genus Diffugia); hemiforcipate trophi described by De Smet (2006). [1 genus; 2 NEA of 3 species] …………………………………………...………………………………………………... Asciaporrectidae, one genus: Asciaporrecta spp. 3(2) Stomach filled with zoochlorellae; gastric glands absent, but with cecae directed towards anterior end; trophi not protrusible. Consult (De Smet & Pourriot, 1997) for details. [1 genus; 4 NEA of 6 species] …………………………………………. Ituridae, one genus: Itura spp. 3’ Not as above ……………………………………………………………………………………………………… Dicranophoridae [p. 150] 4(1) Mostly littoral rotifers possessing cardate trophi (Fig. 13.11 u in Volume I) having a sucking action or trophi highly modified and stomach with zoochlorellae …………………………………………………………………………………………………………………………... 5 4’ Rotifers with trophi other than cardate ……………………………………………………….…………………………………………….. 6 5(4) Manubria of the trophi with hooks that may be determined in lateral view of the animal (Fig. 13.11 u in Volume I), body fusiform, 300–500 μm. (Fig. 8.34). Consult Nogrady and Segers (2002) for details. [1 genus; 11 NEA of 13 species] ………………………..……… …………………………………………………………………………………………………………….. Lindiidae, one genus: Lindia spp. 5’ Highly modified trophi, possessing pseudunci, stomach with zoochlorellae, gastric glands absent, a rare littoral, ∼275 μm. (Fig. 8.35) [monospecific] …………………………………………………………….. Birgeidae, one species: Birgea enantia Harring & Myers, 1922 6(4) Rotifers not possessing incudate or modified incudate trophi ………………………….…………………………………………………... 7 6’ Illoricate, saccate rotifers, with incudate or modified incudate trophi, (Fig. 13.11 g, r in volume I) .……………… Asplanchnidae [p. 152] 7(6) Rotifers possessing malleate trophi ………………………………………………………………………………………………………… 8 [In malleate trophi (Fig. 13.11 a–c, e, f, p in volume I), each uncus has only 4–7 teeth (unlike the condition found in malleoramate trophi). The fulcrum may be short (malleate) or long (submalleate)] 7’ Rotifers possessing virgate trophi …………………………………………………………………………………………………………. 15 [Virgate trophi are often asymmetrical (Fig. 13.11 d, k–o, q, s in Volume I), with a long fulcrum and manubria and generally small rami.] 8(7) Loricate rotifers: body wall thickened and firm …………………………………………………………………………………………….. 9 8’ Illoricate rotifers: body wall not thickened or firm ………………………………………………………………………………………... 13 [Interpreting whether the body wall is thickened (loricate) or not (illoricate) can be difficult, and it does take some experience to judge this characteristic in forms that do not easily fall one way or the other. To get an indication as to how firm the lorica is, follow this procedure. When working with fresh material, apply gentle pressure from the point of a pencil or probe onto the cover glass while observing the specimen. If the body puffs out under pressure and returns to its original shape when the pressure is released, the specimen is illoricate. Loricate forms will exhibit much less flexibility of the body wall: i.e., they do not puff out. NB: If too much pressure is applied to the cover glass, the animal may be crushed, becoming irreversibly damaged, perhaps rendering them unidentifiable. In preserved materials, illoricate forms tend to shrivel up, while the body wall of loricate rotifers will retain its shape even if the inner organs separate from the body wall, collapsing into a central mass of tissue. Some members of the Brachionidae that are weakly loricate will key to the Epiphanidae.] 9(8) Lorica possessing furrows, grooves, or sulci; or with a dorsal, semicircular head shield covering the corona; or with a very strongly developed lorica and a dorsal transverse ridge ………………………………………………………………………………………………….. 10 9’ Lorica lacking furrows, grooves, sulci, or dorsal head shield; without a strongly developed lorica and dorsal transverse ridge ………… 14 10(9) Lorica lacking medial, ventral furrow or notch, and lacking a head shield ……………………………………………………………….. 11

Phylum Rotifera

10’ Lorica with medial, ventral furrow (but no lateral furrow or grooves) extending the full length of the animal or with a ventral notch in which the foot lies, or with a dorsal, semicircular head shield covering the corona (includes former Colurellidae) .……………… Lepadellidae [p. 152] 11(10) Lorica not as below ………………………………………………………………………………………………………………………… 12 11’ Lorica with dorsal, medial sulcus (double keel) or a single dorsal keel ………………………………………………… Mytilinidae [p. 154] 12(11) Foot usually projecting from between dorsal and ventral plates at the posterior end of the lorica; dorsal and ventral plates separated by a deep furrow or groove …………………………………………………………………………………………………. Euchlanidae [p. 154] 12’ Foot projecting through a foramen (hole) in ventral plate at the posterior end of the lorica; dorsal and ventral plates connected by a weak furrow (groove); ≤200 μm (excluding toes) (Fig. 8.36) [1 genus, 105 NEA of ∼200 species] …………… Lecanidae, one genus: Lecane spp.

Chapter | 8  Phylum Rotifera

(B)

FIGURE 8.33  Family Testudinellidae: (A) Testudinella and (B) Pompholyx. Bars = 50 μm. (NEA) Photomicrograph of Pompholyx courtesy of C. Jersabek, University of Salzburg, Austria.

FIGURE 8.35  Family Birgeidae: Birgea. Whole animal (left); trophy (right). Bar = 100 μm. (NEA) After Harring & Myers, 1922.

FIGURE 8.36  Family Lecanidae: Lecane. Bar = 100 μm. (NEA) FIGURE 8.34  Family Lindiidae: Lindia. Top panel: left, whole animal; right, trophy. (NEA) (After Harring & Myers, 1922.) Lower panel: left, animals in a colony of the bluegreen bacteria, Gloeotrichia; right, close up. Bar ∼100 μm

Phylum Rotifera

(A)

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FIGURE 8.37  Family Scaridiidae: Scaridium. Bar = 250 μm. Photomicrograph courtesy of M.V. Sørensen, University of Copenhagen. [The genera Hemimonostyla (toes partially fused) and Monostyla (toes completely fused) has been subsumed within Lecane (with two toes that may be partially fused at the base) (Koste, 1978; Segers, 1995a). Use of these older names should be abandoned.] 13(8,) Mouth set in a funnel-shaped buccal field ……………………………………………………………………………. Epiphanidae [p. 156] 13’ Mouth set in an oblique, ciliated field on ventral side; body swollen, vermiform, or fusiform …………………………... Proalidae [p. 156] 14(9,) Lorica extending beyond body to head, foot, and toes ………………………………………………………………. Trichotriidae [p. 158] 14’ Lorica not extending beyond body …………………………………………………………………………………....Brachionidae [p. 158] 15(7,) Body not twisted as a partial helix and/or trophi not asymmetrical ………………………………………………………………………... 16 15’ Body twisted as a partial helix (asymmetrical) and/or trophi asymmetrical OR small, saccate animals, present within the colonial alga Volvox where they feed from the inside on the algal cells. ………………………………………………………….. Trichocercidae [p. 160] 16(15) Not as below ………………………………………………………………………………………………………………………………. 17 16’ Unci tips pointed outwards, capable of protruding through mouth; body elongate; head with two pairs of lateral lobes; corona with stiff, with preoral setae; foot with prominent muscles, foot and toes longer then rest of body; ≤450 μm. (Fig. 8.37). Consult Nogrady et al. (1995) for details. [1 genus, 3 NEA of 7 species] ………………………………………………………….. Scaridiidae, one genus: Scaridium spp. 17(16) Not as below ………………………………………………………………………………………………………………………………. 18 17’ Body spindle-shaped; long, knobbed, lateral antennae located near the base of the foot and with long setae; stomach separated from intestine by a circle of bulbous glands; ≤1000 μm (acid waters with a pH 1/5 total body length, toes end in rounded tips [3 NEA of 8 species] ……..…………………………………. Wierzejskiella spp. 13’ Short foot,  ½ total animal length; terminal foot segment longer than toes; ∼100–160 μm. (Fig. 8.46 [2 NEA of 2 species] ……………… …………………………………………………………………………. Paracolurella spp. 3’ Lorica lacking dorsal sulcus (continuous); foot + toes