Freshwater Algae of North America: Ecology and Classification (Aquatic Ecology)

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Freshwater Algae of North America Ecology and Classification

Freshwater Algae of North America Ecology and Classification Edited by

JOHN D. WEHR Louis Calder Center—Biological Station and Department of Biological Sciences Fordham University Armonk, New York

and

ROBERT G. SHEATH Office of Provost and Vice President for Academic Affairs California State University, San Marcos San Marcos, California

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

Front cover image: Kandis Elliot Institute of Applied Science.

This book is printed on acid-free paper.

Copyright © 2003, Elsevier Science (USA) All Rights Reserved. No part of this publication may be reproduced or trasmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2002 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. $35.00 Explicit permission from Academic Press is not required to reproduce a maximum of two figures or tables from an Academic Press chapter in another scientific or research publication provided that the material has not been credited to another source and that full credit to the Academic Press chapter is given.

Academic Press An imprint of Elsevier Science. 525 B Street, Suite 1900, San Diego, California 92101-4495, USA http://www.academicpress.com

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We acknowledge the numerous phycological pioneers who blazed many trails for those of us who followed in the study of North American freshwater algae, especially Gilbert M. Smith, Gerald W. Prescott, George J. Schumacher, Larry A. Whitford, Francis Wolle and Hoaratio C. Wood. John D. Wehr Robert G. Sheath

My heartfelt thanks and love go to Deborah Donaldson for her faith and support. John D. Wehr

My love and appreciation to Mary Koske who supported me through those many collecting trips to far-flung places and the myriad of career and location changes. Robert G. Sheath


Contents

Contributors Preface xv

xiii

III. IV. V. VI.

1

Lotic Environments 28 Wetlands 38 Thermal and Acidic Environments Unusual Environments 42 Literature Cited 45

40

INTRODUCTION TO FRESHWATER ALGAE Robert G. Sheath and John D. Wehr I. Introduction 1 II. Classification 5 III. Taxonomic Chapters in This Book Literature Cited 9

3 8

2 FRESHWATER HABITATS OF ALGAE John D. Wehr and Robert G. Sheath I. What is Fresh Water? 11 II. Lentic Environments 12

COCCOID AND COLONIAL CYANOBACTERIA Jirˇí Komárek I. II. III. IV. V. VI.

Introduction 59 Morphology and Diversity 60 Ecology and Distribution 63 Collection, Preparation, and Culture 67 Key and Descriptions of Genera 68 Guide to Literature for Species Identification 110 Literature Cited 110 vii

viii

Contents

4

III. Ecology and Distribution 257 IV. Collection and Preparation for Identification 258 V. Key and Descriptions of Genera VI. Guide to Literature for Species Identification 307 Literature Cited 307

FILAMENTOUS CYANOBACTERIA Jirˇí Komárek, Hedy Kling, and Jaroslava Komárková I. II. III. IV. V.

Introduction 117 Morphology 118 Ecology 120 Methods 121 Key and Descriptions of Genera Note Added in Proof 189 VI. Guide to Literature for Species Identification 191 Literature Cited 191

259

121

8 FILAMENTOUS AND PLANTLIKE GREEN ALGAE David M. John

5 RED ALGAE Robert G. Sheath I. II. III. IV.

Introduction 197 Diversity and Morphology 197 Ecology and Distribution 202 Collection and Preparation for Identification 206 V. Key and Descriptions of Genera 207 VI. Guide to Literature for Species Identification 221 Literature Cited 221

6

I. II. III. IV. V. VI. VII.

Introduction 311 Diversity and Morphology 311 Classification of Green Algae 312 Ecology and Distribution 313 Collection and Preparation of Samples Key and Descriptions of Genera 316 Guide to Literature for Species Identification 347 Literature Cited 349

9 CONJUGATING GREEN ALGAE AND DESMIDS Joseph F. Gerrath

FLAGELLATED GREEN ALGAE Hisayoshi Nozaki I. II. III. IV.


I. II. III. IV.


10 PHOTOSYNTHETIC EUGLENOIDS

7 NONMOTILE COCCOID AND COLONIAL GREEN ALGAE L. Elliot Shubert I. Introduction 253 II. Diversity and Morphology

254

James R. Rosowski I. II. III. IV.

Introduction 383 Diversity and Morphology 387 Ecology and Distribution 405 Collection, Culturing, and Preparation for Identification 408

315

Contents

V. Key and Descriptions of North American Genera 410 VI. Guide to Literature for Species Identification 415 Literature Cited 416

VI. Guide to Literature for Species Identification 519 Literature Cited 519

14 SYNUROPHYTE ALGAE

11 EUSTIGMATOPHYTE, RAPHIDOPHYTE, AND TRIBOPHYTE ALGAE Donald W. Ott and Carla K. Oldham-Ott I. II. III. IV. V.

General Introduction 423 Eustigmatophytes 424 Raphidophytes 427 Tribophytes 429 Collection and Preparation for Identification 463 VI. Guide to Literature for Species Identification 465 Literature Cited 466

Peter A. Siver I. II. III. IV.

Introduction 523 Diversity and Morphology 524 Ecology and Distribution 534 Collection and Preparation for Identification 539 V. Keys to Genera and Common Species from North America 541 VI. Guide to Literature for Species Identification 551 Literature Cited 552

15 CENTRIC DIATOMS

12 CHRYSOPHYCEAN ALGAE Kenneth H. Nicholls and Daniel E. Wujek I. II. III. IV.

Introduction 471 Diversity and Morphology 473 Ecology 485 Collection and Preparation for Identification 490 V. Key and Descriptions of Genera 491 VI. Guide to Literature for Species Identification 503 Literature Cited 503

Eugene F. Stoermer and Matthew L. Julius J. P. Kociolek and S. A. Spaulding (Introduction) I. II. III. IV. V. VI. VII. VIII.

General Introduction to the Diatoms 559 Introduction to Centric Diatoms 562 Classification 563 Morphology and Physiology 565 Ecology and Evolution 568 Collection and Study Methods 570 Key and Descriptions of Genera 571 Guide to Literature for Species Identification 587 Literature Cited 588

16 ARAPHID AND MONORAPHID DIATOMS

13 HAPTOPHYTE ALGAE Kenneth H. Nicholls I. II. III. IV.

Introduction 511 Diversity and Morphology 512 Ecology and Distribution 513 Collection and Preparation for Identification 514 V. Key and Descriptions of Genera 515

John C. Kingston I. II. III. IV.


ix

x

Contents

17 SYMMETRICAL NAVICULOID DIATOMS J. P. Kociolek and S. A. Spaulding I. II. III. IV.

Introduction 637 Ecology and Distribution 638 Key and Descriptions of Genera Guide to Literature for Species Identification 651 Literature Cited 651

639

II. Morphology and Diversity 687 III. Ecology and Distribution 699 IV. Collection and Preparation for Identification 702 V. Key and Descriptions of Genera 703 VI. Guide to Literature for Species Identification 709 Literature Cited 710

21 CRYPTOMONADS

18 EUNOTIOID AND ASYMMETRICAL NAVICULOID DIATOMS J. P. Kociolek and S. A. Spaulding I. II. III. IV.

Introduction 655 Diversity and Morphology 656 Ecology and Distribution 661 Key and Descriptions of North American Genera 662 V. Guide to Literature for Species Identification 666 Literature Cited 666

Paul Kugrens and Brec L. Clay I. II. III. IV. V.

Introduction 715 Unique Features of Cryptomonads 716 Origin of Cryptomonads 736 Ecology 738 Collection, Preparation for Isolation, and Culturing 740 VI. Classification, Key, and Descriptions 740 VII. Availability of Cryptomonads 749 VIII. Family Katablepharidaceae 749 Literature Cited 751

22

19 KEELED AND CANALLED RAPHID DIATOMS Rex L. Lowe I. II. III. IV.

Introduction 669 Diversity and Morphology 670 Ecology and Distribution 671 Collection and Preparation for Identification 674 V. Keys and Descriptions of Genera 675 VI. Guide to Literature for Species Identification 682 Literature Cited 682

BROWN ALGAE John D. Wehr I. II. III. IV.

Introduction 757 Diversity and Morphology 758 Ecology and Distribution 763 Methods for Collection and Identification 766 V. Key and Descriptions of Genera 767 VI. Guide to Literature for Species Identification 771 Literature Cited 772

23

20 DINOFLAGELLATES

USE OF ALGAE IN ENVIRONMENTAL ASSESSMENTS

Susan Carty

R. Jan Stevenson and John P. Smol

I. Introduction

685

I. Introduction

775

Contents

II. Goals of Environmental Assessment with Algae 776 III. Sampling and Assessing Algal Assemblages for Environmental Assessment 778 IV. Developing Metrics for Hazard Assessment 786 V. Exposure Assessment: What Are Environmental Conditions? 790 VI. Stressor–Response Relations 794 VII. Risk Characterization and Management Decisions 795 VIII. Conclusions 796 Literature Cited 797

24 CONTROL OF NUISANCE ALGAE Carole A. Lembi I. Introduction 805 II. Problems Associated with Algae 805 III. Control Methods for Nuisance Algae 812 Literature Cited 826 Glossary

835

Author Index

849

Subject Index

885

Taxonomic Index

897

xi


Contributors

Number in parentheses indicate the pages on which the authors’ contributions begin.

Susan Carty (685) Department of Biology, Heidelberg College, Tiffin, Ohio 44883. Brec L. Clay (715) CH Diagnostic and Consulting Service, Loveland, Colorado 80538. Joseph F. Gerrath (353) Department of Botany, University of Guelph, Guelph, Ontario, Canada N1G 2W1. David M. John (311) Department of Botany, The Natural History Museum, London SW7 5BD, United Kingdom. Matthew L. Julius (559) Department of Biological Sciences, St. Cloud State University, St. Cloud, Minnesota 56301. John C. Kingston (595) Center for Water and the Environment, Natural Resources Research Institute,

University of Minnesota Duluth, Ely, Minnesota 55731. Hedy Kling (117) Freshwater Institute, Winnipeg, Manitoba, Canada, R3T 2N6. J. P. Kociolek (559, 637, 655) Diatom Collection, California Academy of Sciences, Golden Gate Park, San Francisco, California 94118. Jaroslava Komárková (117) Hydrobiological Institute, Academy of Sciences of the Czech Republic, Faculty of Biological Sciences, University of South Bohemia, CZ-37005 Cˇeské Budeˇ jovice, Czech Republic. Jirˇí Komárek (59, 117) Institute of Botany, Academy of Sciences of the Czech Republic, Faculty of Biological Sciences, University of South Bohemia, CZ-37982 Trˇebonˇ, Czech Republic. Paul Kugrens (715) Department of Biology, Colorado State University, Fort Collins, Colorado 80523. xiii

xiv

Contributors

Carole A. Lembi (805) Department of Botany and Plant Pathology, Purdue University, West Lafayette, Indiana 47907. Rex L. Lowe (669) Biological Sciences, Bowling Green State University, Bowling Green, Ohio 43403 and University of Michigan Biological Station, Pellston, Michigan 49769. Kenneth H. Nicholls (471, 511) S-15 Concession 1, RR #1 Sunderland, Ontario, Canada L0C 1H0. Hisayoshi Nozaki (225) Department of Biological Sciences, Graduate School of Science, University of Tokyo, Hongo, Bunkyo-ku, Tokyo 113-0033, Japan. Carla K. Oldham-Ott (423) Department of Biology, University of Akron, Akron, Ohio 44325. Donald W. Ott (423) Department of Biology, University of Akron, Akron, Ohio 44325. James R. Rosowski (383) School of Biological Sciences, College of Arts and Sciences, University of Nebraska–Lincoln, Lincoln, Nebraska 68588. Robert G. Sheath (1, 11, 197) Office of Provost and Vice President for Academic Affairs, California State University, San Marcos, San Marcos, California 92096.

L. Elliot Shubert (253) Department of Botany, The Natural History Museum, London SW7 5BD, United Kingdom. Peter A. Siver (523) Botany Department, Connecticut College, New London, Connecticut, 06320. John P. Smol (775) Department of Biology, Paleoecological Environmental Assessment and Research Laboratory (PEARL), Queen’s University, Kingston, Ontario, Canada K7L 3N6. S. A. Spaulding (559, 637, 655) Diatom Collection, California Academy of Sciences, Golden Gate Park, San Francisco, California 94118. R. Jan Stevenson (775) Department of Zoology, Michigan State University, East Lansing, Michigan, 48824. Eugene F. Stoermer (559) Michigan Herbarium University of Michigan, Ann Arbor, Michigan 48109. John D. Wehr (1, 11, 757) Louis Calder Center— Biological Station and Department of Biological Sciences, Fordham University, Armonk, New York 10504. Daniel E. Wujek (471) Department of Biology, Central Michigan University, Mt. Pleasant, Michigan 48859.

Preface

The study of freshwater algae in North America has a long and rich history, with some of the early monographic works dating back to the late 1800’s. In recent years, there has been an enormous and remarkable level of research on this very diverse and heterogeneous collection of organisms, making any definitive taxonomic or ecological treatise always out of date. Nonetheless, it is our goal with this book to synthesize and update much of this vast knowledge, and to provide a practical and comprehensive guide to all of the genera of freshwater algae known from throughout the continent, in one volume. Chapters also provide guides to other publications and specialized works for the identification and ecological information at the species level. Our intent is to combine the necessary ecological and taxonomic information in a practical book that can be used by all scientists working in aquatic environments, whether their specialty is in environmental monitoring, ecology, evolution, system-

atics, biodiversity, or molecular biology. This is the first book of its sort covering the entire continent. We also hope that this book will serve to encourage new generations of aquatic biologists to explore freshwater algae carefully, rather than regarding phytoplankton or benthic algae as simply quantities of chlorophyll or carbon. The enormous variety of algae in lakes, rivers and other aquatic habitats is part of the ecological content of aquatic communities, and their ecosystem function varies with the species that occur there. Many of the previous monographs dealing with a broad geographic region are still useful, such as Smith’s (1950) Freshwater Algae of the United States, but most are decades old and do not contain recent taxonomic changes. Our approach is to include chapters authored by experts who have specialized in the study of specific groups of freshwater algae. Given the great quantity of research that has been produced on all of the major algal taxa, it is no longer possible for one or two xv

xvi

Preface

authors to produce an authoritative book of this kind, and one which will span the entire range of taxonomic and ecological detail that is now known about all the organisms termed algae. This volume is modeled closely after the book by Thorp and Covich on freshwater invertebrates (Ecology and Classification of North American Freshwater Invertebrates), also published by Academic Press. The organization of this book includes an introduction to the freshwater algae (with a guide to the taxonomic chapters that follow), an overview of freshwater habitats, 20 taxonomic chapters, and finally chapters on the use of algae in environmental assessments and control of nuisance algae. More than 770 genera are described and illustrated in this book, and each taxonomic chapter includes an introduction to the key terms and characteristics of the group, ecological distribution, and a guide to the taxonomic literature to distinguish species within each genus. While we have undoubtedly omitted some less common or yet unrecorded freshwater genera, this compilation represents an increase in the taxonomic scope and geographic coverage of the freshwater algae of North America. This compares with roughly 490 genera recorded from the United States by Smith (1950), about 335 in Prescott’s (1962) coverage of the Western Great Lakes region, and nearly 380 genera from the southeastern U.S. reported by Whitford and Schumacher (1984). Since not all algal groups are equally well studied, coverage in the present volume varies among taxa and chapters. We hope that students, scientists working in water management agencies, and experienced phycologists will use this book thoroughly and provide us with feedback, such as missing taxa or incomplete geographic information. We will endeavor to incorporate this information into a future edition. We are extremely grateful to the contributors of this volume who took much time and effort to research and prepare the chapters and follow through with the reviewer’s and editorial suggestions. We extend our sincere thanks to the reviewers whose helpful comments enhanced the quality of the final presentation. The reviewers were as follows: Robert Andersen, J. Craig Bailey, Barry Biggs, Alan Brook, Alain Couté, Eileen Cox, David Czarnecki, Gary Dillard, Gary

Floyd, Paul Hamilton, Kyle Hoagland, Ronald Hoham, Jeffrey Johansen, John Kingston, Dag Klaveness, Hedy Kling, Lothar Krienitz, Jorgen Kristiansen, Elsadore Kusel-Fetzmann, Carole Lembi, Rex Lowe, David Mann, Richard McCourt, Øjvind Moestrup, Orlando Necchi Jr., Kenneth Nichols, Gianfranco Novarino, Hans Paerl, Russell Rhodes, Frank Round, Robert Sheath, Alan Steinman, Eugene Stoermer, Francis Trainor, Richard Treimer, Herb Vandermeulen, Morgan Vis, James Wee, John Wehr, Robert Wetzel, Ruth Willey, and David Williams. We wish to thank our editor at Academic Press, Dr. Charles R. Crumly, and his assistant, Ms. Christine Vogelei, for their continued support throughout the creation and completion of this book. We also thank Ms. Geri Mattson at Mattson Publishing Services for providing professional assistance in the production of the final copy of the book. We also wish to acknowledge the many colleagues who generously permitted reproduction of published and unpublished material; these are acknowledged in the individual chapters. Kandis Elliot designed the beautifully illustrated cover. The production of this book was partially supported by the Routh Endowment of the Louis Calder Center for JDW and NSERC grant number RGP 0105629 to RGS, as well as general support from Fordham University, University of Guelph and California State University, San Marcos. Help in manuscript production from Pam Anderson, Petra DelValle (Fordham), Marcy Boyle (California State University, San Marcos), and Toni Pellizzari (Guelph) is appreciated. Lastly, we would like to thank our spouses, Deb Donaldson (Wehr) and Mary Koske (Sheath) for their understanding and support through this long process. John D. Wehr and Robert G. Sheath

LITERATURE CITED Prescott, G.W. 1962. Algae of the Western Great Lakes Area, 2nd Edn. W.C. Brown, Dubuque, Iowa. Smith, G. M. 1950. The Freshwater Algae of the United States, 2nd Edn. McGraw-Hill, New York. Whitford, L.A. & Schumacher, G.J. 1984. A Manual of Fresh-Water Algae, Revised Edn., Sparks Press, Raleigh, NC.

1

INTRODUCTION TO FRESHWATER ALGAE Robert G. Sheath

John D. Wehr

Office of Provost and Vice President for Academic Affairs California State University, San Marcos San Marcos, California 92096

Louis Calder Center—Biological Station and Department of Biological Sciences Fordham University, Armonk, New York 10504

I. Introduction II. Classification A. Cyanobacteria B. Red Algae C. Green Algae D. Euglenoids E. Eustigmathophyte, Raphidiophyte, and Tribophyte Algae F. Chrysophycean Algae

G. Haptophyte Algae H. Synurophyte Algae I. Diatoms J. Dinoflagellates K. Cryptomonads L. Brown Algae III. Taxonomic Chapters in This Book A. Key Literature Cited

I. INTRODUCTION Algae are treated in this book in the same sense as they are in many introductory phycology texts (e.g., Van den Hoek et al., 1995; Sze 1998; Graham and Wilcox, 2000); that is, they are considered to be a loose group of organisms that have all or most of the following characteristics: aquatic, photosynthetic, simple vegetative structures without a vascular system, and reproductive bodies that lack a sterile layer of protecting cells. As such, algae are no longer regarded as a phylogenetic concept, but still represent an ecologically meaningful and important collection of organisms. Both prokaryotic (cells that have no membrane-bound organelles) and eukaryotic taxa (cells with organelles) are included. In addition, there is a wide range of vegetative morphologies, including the following: 1. Unicells: species that occur as solitary cells that may be nonmotile or motile. Motile cells may have one or more flagella or they may glide. A wide variety of forms exists among unicells, including those contained within a gelatinous sheath (Fig. 1A), with intricate cell walls Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

(Fig. 1B), having flexible cell shapes (Fig 1C), with two flagella of unequal length (Fig. 1D) or two equal flagella (Fig. 1E), with cells drawn out into hornlike extensions (Fig. 1F), and having cells contained in a hardened case or lorica (Fig. 1G). 2. Colonies: an aggregation of cells that are held together either in a loose (Fig. 1H and I) or tight, well organized fashion (Fig. 2B, D, and E). Depending on the algal taxon, colonies may contain a variable number of cells or they may be constant throughout their development (Fig. 2B). Colonies may contain flagellated or nonflagellated cells. The basis for cellular connection varies among colonies, including a surrounding gelatinous matrix (Fig. 1H and I), gelatinous stalks (Fig. 2A), common parental wall (Fig. 2B), and direct attachment at the cellular edges (Fig. 2C) or at the middle portion of each cell (Fig. 2C). Alternately, cells may be connected via their loricae (Fig. 2E). 3. Pseudofilaments: an aggregation of cells in an end-to-end fashion. The cells are not directly connected to each other; rather, they are spaced 1

2

Robert G. Sheath and John D. Wehr

FIGURE 1 Unicellular and colonial forms of freshwater algae. A. Gloeocapsa (cyanobacterium), a unicell to small grouping of cells contained within concentrically layered gelatinous sheaths (arrows). B. Micrasterias (green alga, desmid), a unicell with many regular cell wall incisions (arrows) that form a series of lobes and lobules. C. Euglena (euglenoid), a unicell that does not produce walls and can readily change shape. D. Ochromonas (chrysophycean alga), a unicell with one long and one short apically inserted flagellum (arrows). E. Pyrenomonas (cryptomonad), a unicell with two equal subapically inserted flagella. F. Ceratium (dinoflagellate), a unicell with a theca composed of cellulose plates and cellular extensions or horns (arrows). G. Strombomonas (euglenoid), a flagellated unicell within a rigid lorica (arrow). H. Coelosphaerium (cyanobacterium), a colony with spherical cells loosely arranged at the periphery of a gelatinous matrix. I. Dermatochrysis (chrysophycean alga), a colony with spherical cells in a single layer scattered in a gelatinous matrix that has distinct perforations (arrows). Scale bars = 10 µm.

apart and contained in a common gelatinous matrix (Fig. 2F). 4. Filaments: a chain or series of cells in which the cells are arranged in an end-to-end manner, where adjacent cells share a common cross wall

(Figs. 2H–J, and 3B and C). Linear colonies can be distinguished from true filaments by the fact that abutting colonial cells each possess their own entire walls (Fig. 2D). Filaments may be arranged in a single series (uniseriate or

1. Introduction to Freshwater Algae

3

FIGURE 2 Colonial, pseudofilamentous, and filamentous forms of freshwater algae. A. Porphyridium (red alga), a colony with spherical cells attached together by gelatinous strands (arrows). B. Crucigenia (green alga), a colony with consistent groups of four cells produced inside the walls of the parent cells. C. Tabellaria (diatom), a colony with cells attached at their edges in a zig-zag fashion. D. Asterionella (diatom), a linear colony with cells attached only at the central region. E. Dinobryon (chrysophycean alga), a colony with cells attached by their loricae (arrows). F. Chroodactylon (red alga), a pseudofilament with cells arranged in an endto-end pattern in a common gelatinous matrix (arrows), but not directly connected to each other. G. Zygnema (green alga), an unbranched filament without a gelatinous matrix. H. Lyngbya (cyanobacterium), an unbranched filament that is contained in a gelatinous sheath that is evident at the filament tip (arrow). I. Scytonema (cyanobacterium), a filament that produces double false branches (arrows) that result from breakage and further growth of each fragment. J. Bangia (red alga), a multiseriate filament in part with at least two cells across (arrows). Scale bars = 10 µm.

uniaxial) (Fig. 2G–I) or they may be in more than one series of cells (multiseriate or multiaxial) (Fig. 2J). Filaments may be unbranched (Figs. 2G and H) or they can produce branches in a new plane that are morphologically similar to the main axis (Fig. 3B) or that are quite distinct (Fig. 3C). Branching may be dichotomous or forked (Fig. 3B), alternate (Fig. 3C), opposite, or whorled (Fig. 3D). False branches are formed in

some cyanobacteria, such as Scytonema (Fig. 2I), by fragmentation and continued growth of one or both fragments. Other types of filaments include those that are heterotrichous, that is, they have a distinct prostrate system with attached erect branches. Differentiated filaments have specialized cells within the chain. The main axis may have a surrounding layer of small cells termed cortication (Fig. 3A).

4


FIGURE 3 Filamentous, saclike, crustose, pseudoparenchymatous, and siphonous forms of freshwater algae. A. Compsopogon (red alga), a filamentous form with small cortical cells (arrows) covering the main filament. B. Cladophora (green alga), a filament that has dichotomous (forked) branches (arrows). C. Draparnaldia (green alga), a filament that has tuftlike lateral branches with cells that are considerably smaller than those of the main axis. D. Batrachospermum (red alga), a filament with whorllike lateral branches (arrows). E. Boldia (red alga), a saclike thallus that consists of a single layer of cells. F. Heribaudiella (brown alga), a crust that is tightly adherent to the rock substratum. G. Hildenbrandia (red alga), a cross section of a crust that shows vertical files of cells (arrows). H. Caloglossa (red alga), a pseudoparenchymatous thallus composed of a main filamentous axis (arrow) with tightly compacted lateral branches. I. Vaucheria (yellow–green alga), a siphonous thallus without cross walls separating the nuclei. Scale bars = 10 µm, except B = 250 µm, E = 1 cm, and F = 2 cm. Figure A courtesy of Tara Rintoul; Figure B from Vis et al. (1994) reprinted with permission of University of Hawaii Press; Figure E from Sheath (1984) with permission; Figure G courtesy of Alison Sherwood.

5. Pseudoparenchymatous structures: tissue-like thalli that consist of closely appressed branches of a uniseriate or multiseriate filament (Fig. 3H). Crustose forms may be composed of short, compacted filaments, such as the brown alga Heribaudiella (Fig. 3F) and the rhodophyte Hildenbrandia (Fig. 3G). 6. Parenchymatous forms: true tissues composed of a solid mass of cells that is three dimensional, variously shaped, and not filamentous in construction. The cells may be differentiated

into an outer photosynthetic layer (the cortex) and an inner non-photosynthetic region (the medulla). Most tissue-like forms in freshwater habitats are simple, such as the saccate red alga Boldia, which consists of a single layer of cells (Fig. 3E). 7. Coenocytic or siphonous forms: large multinucleate forms of various shapes without cross walls to separate the nuclei or other organelles. An example is the yellow–green alga Vaucheria (Fig. 3I).


Freshwater algae exhibit all of these morphologies, but the macroscopic pseudoparenchymatous and parenchymatous forms tend to be smaller than those found in marine systems (Sheath and Hambrook, 1990). In addition, planktonic (floating) forms are typically small and microscopic, and mostly consist of the simpler forms. In contrast, benthic (attached) algae include the entire range of morphologies, although flagellated taxa are less common than in plankton.

II. CLASSIFICATION Algae do not represent a formal taxonomic group of organisms, but rather constitute a loose collection of divisions or phyla with representatives that have the characteristics noted previously. The divisions are

5

distinguished from each other based on a combination of characteristics, including photosynthetic pigments, starchlike reserve products, cell covering, and other aspects of cellular organization (e.g., Van den Hoek et al., 1995; Sze, 1998; Graham and Wilcox, 2000). There is little consensus among phycologists as to the exact number of algal divisions; 8–11 have been recognized in recent texts (Van den Hoek et al., 1995; Sze, 1998; Graham and Wilcox, 2000). The 12 major algal groups (divisions and classes) recognized in this book are distinguished from each other in Table I. Each of the major groupings is briefly presented in the following sections, but the reader should refer to the relevant chapter(s) for more details. The number of freshwater genera now reported (>800) from North America, as discussed in Chapters 3–22, has greatly increased from earlier treatises (e.g., Smith,

TABLE I Major Distinguishing Features of the Major Algal Groups Presented Herein Algal group (chapter number)

Photosynthetic pigmentsa

Chloroplast outer Thylakoid membranes associations

Starch-like reserveb

External coveringc

Cyanobacteria (3 & 4)

chla, PE, PC, APC

0

0

Cyanophycean

Pepitoglycan matrices or walls

0

Red algae (5)

chla, PE, PC, APC

2

0

Floridean

Walls with a galactose polymer matrix

0

Green algae (6–9)

chla, b

2

2–6

True

Cellulosic walls, scales

0 – many

Euglenoid Algae (10)

chla, b

3

3

Paramylon

Pellicle

1–2 emergent

Yellow–green and related algae (11)

chla, c

4

3

Chrysolaminarin

Mostly cellulosic walls

2 unequal if present

Chrysophyte algae (12)

chla, c fucoxanthin

4

3

Chrysolaminarin

None, scales, lorica

2 unequal

Haptophyte algae (13)

chla, c fucoxanthin

4

3

Chrysolaminarin

Nonsiliceous scales

2 equal + haptonema

Synurophyte algae (14)

chla, c fucoxanthin

4

3

Chrysolaminarin

Siliceous scales

2 unequal

Diatoms (15–19)

chla, c fucoxanthin

4

4

Chrysolaminarin

Siliceous frustule

1, reproductive cells only

Dinoflagellates (20)

chla, c peridinin

3

3

True

Theca

2 unequal

Cryptomonads (21)

chla, c PC or PE

4

2

True

Periplast

2 equal

Brown algae (22)

chla, c fucoxanthin

4

3

Laminarin

Walls with alginate matrices

2 unequal

Flagella

Source: Various phycology textbooks (e.g., Sze, 1998; and Graham and Wilcox, 2000). a chl = chlorophyll (green); PE = phycoerythrin (red); PC = phycocyanin (blue); APC = allophycocyanin (blue); fucoxanthin and peridinin (golden to brown). b All of the reserves are polymers of glucose. They differ by their linkages: cyanophycean and floridean α1, 4 and α1, 6 branches; true starch with amylose α1, 4 and amylopectin α1, 4 and α1, 6 branches; paramylon β1, 3; chrysolamin and laminarin β1, 3 and β1, 6 branches. Only true starch stains positively with iodine (purple to black). c Pellicle and periplast within plasma membrane; the rest are external to it.

6


1950; Prescott, 1962), but is still highly tentative and likely to be an underestimate of the region’s biodiversity.

A. Cyanobacteria Cyanobacteria or blue–green algae are prokaryotes, that is, cells that have no membrane-bound organelles, including chloroplasts (Table I; Chap. 3). Other characteristics of this division include unstacked thylakoids, phycobiliprotein pigments, cyanophycean starch, and peptidoglycan matrices or walls. There are 124 genera reported from inland habitats in North America, of which 53 are unicellular or colonial (Chap. 3) and 71 are filamentous (Chap. 4). However, the taxonomy of this division is currently in a state of flux, as noted in Chapter 3, and the number of genera should be considered to be tentative. Cyanobacteria inhabit the widest variety of freshwater habitats on Earth and can become important in surface blooms in nutrient-rich standing waters (Chaps. 3 and 4). Some of these blooms can be toxic to zooplankton and fish, as well as livestock that drink water containing these organisms. Inland cyanobacteria also occur in extreme environments, such as hot springs, saline lakes, and endolithic desert soils and rocks.

B. Red Algae Rhodophyta or red algae represent a division that is characterized by chloroplasts that have no external endoplasmic reticulum and unstacked thylakoids, phycobiliprotein pigments, floridean starch, and lack of flagella (Table I; Chap. 5). They are predominantly marine in distribution; only approximately 3% of over 5000 species occur in truly freshwater habitats. In North America, 25 genera are recognized in inland habitats (Chap. 5). Freshwater red algae are largely restricted to streams and rivers, but also can occur in other inland habitats, such as lakes, hot springs, soils, caves, and even sloth hair (Chap. 5).

C. Green Algae Chlorophyta or green algae constitute a division that has the following set of attributes: chloroplasts with no external endoplasmic reticulum, thylakoids typically in stacks of two to six, chlorophyll-a and -b as photosynthetic pigments, true starch, and cellulosic walls or scales (Table I). This is a diverse group in inland habitats of North America that includes 44 flagellated genera (Chap. 6), at least 129 coccoid and

nonmotile colonies (Chap. 7), 81 filamentous and plantlike genera (Chap. 8), and 48 conjugating genera and desmids (Chap. 9). Some members of the green algae (Charophyeae) are part of a lineage that is thought to be ancestral to higher plants. Green algae are widespread in inland habitats, but certain groups may have specific ecological requirements. For example, flagellated chlorophytes tend to be more abundant in standing waters that are nutrient rich (Chap. 6). Coccoid unicells and colonies are common in the plankton of standing waters and slowly moving rivers when nutrients, light and temperature are reasonably high (Chap. 7). The majority of filamentous and plantlike Chlorophyta are attached to hard surfaces in standing or flowing waters, but some can exist in the floating state or on soils or other subaerial habitats (Chap. 8). Filamentous conjugating green algae are most frequent in stagnant waters of roadside ditches and ponds, and in the littoral zones of lakes, where they can form free-floating mats or intermingle with other algae in attached or floating masses (Chap. 9). Desmids are more common in ponds and streams that have low conductance and moderate nutrient levels, and often intermingle with macrophytes.

D. Euglenoids Photosynthetic Euglenophyta or euglenoids have chloroplasts surrounded by three membranes, thylakoids in stacks of three, chlorophyll-a and -b as photosynthetic pigments, paramylon, and a pellicle (Table I; Chap. 10). Ten genera are reported from North American freshwater habitats (Chap. 10). Euglenoids are particularly abundant in the plankton of standing waters rich in nutrients and organic matter, and they can be associated with sediments, fringing higher plants, and leaf litter, although some may dominate in highly acidic environments (Chap. 10).

E. Eustigmatophyte, Raphidiophyte, and Tribophyte Algae Eustigmatophyte, raphidiophyte, and tribophyte algae comprise a loose group of algae that share the following characteristics: chloroplasts with four surrounding membranes, thylakoids in stacks of three, chlorophyll-a and -c as the typical photosynthetic pigments, and chrysolaminarin as the photosynthetic reserve product (where known) (Table I; Chap. 11). The yellow–green algae are quite diverse in freshwater habitats of North America: at least 90 genera have been reported, whereas the eustigmatophytes and raphdiophytes are relatively small groups that comprise


eight and three genera, respectively (Chap. 11). Many of these genera seldom have been reported. Members of this group of algae have been collected from a wide variety of habitats, but many are collected primarily in northern habitats (Chap. 11). They are both planktonic and associated with a variety of substrata.

F. Chrysophycean Algae Chrysophyceae or chrysomonads are distinguished by chloroplasts that have four surrounding membranes, thylakoids in stacks of three, fucoxanthin that typically masks chlorophyll-a and -c, and chrysolaminarin as the photosynthetic reserve. At least 72 genera are reported from inland habitats of North America (Chap. 12). Chrysophycean algae are typically associated with standing bodies of water that have low or moderate nutrients, alkalinity, and conductances, and a pH that is slightly acidic to neutral (Chap. 12). In addition, the majority of genera tend to be planktonic; attached forms occur to a lesser extent.

G. Haptophyte Algae Haptophyceae are characterized by chloroplasts that have four surrounding membranes, thylakoids in stacks of three, fucoxanthin that masks chlorophyll-a and -c, chrysolaminarin as the photosynthetic reserve, and a unique appendage associated with the flagellar apparatus, the haptonema (Table I; Chap. 13). Only three freshwater genera are found in North America (Chap. 13). The two common genera are planktonic in lakes and ponds, and occasionally form predominant blooms, particularly in areas with low conductance (Chap. 13). Chrysochromulina breviturrita has been used as an indicator of moderately acidic water.

H. Synurophyte Algae Synurophyceae is characterized by chloroplasts that have four surrounding membranes, thylakoids in stacks of three, fucoxonthin that masks chlorophyll-a and -c, chrysolaminarin as the photosynthetic reserve product, and siliceous scales (Table I; Chap. 14). Three genera are found in North American freshwater habitats (a fourth is known only from Australia), but the genera are species-rich, such as Mallomonus and Synura (Chap. 14). Synurophytes are exclusively freshwater phytoplankters in lakes, ponds, and slowly flowing rivers (Chap. 14). Habitats that support the largest flora are slightly acidic, low in conductance, alkalinity,

7

and nutrients, and have moderate amounts of humic substances.

I. Diatoms Bacillariophyceae or diatoms are distinguished by chloroplasts that have four surrounding membranes, thylakoids in stacks of three, fucoxanthin that masks chlorophyll-a and -c, chrysolaminarin as the photosynthetic reserve product, and a siliceous frustule that makes up the external covering (Table I; Chap. 15). The diatoms are a complex and diverse group in terms of frustule morphology. The North American freshwater genera consist of 25 centrics (Chap. 15), 28 araphid and monoraphid diatoms (Chap. 16), 37 symmetrical naviculoid taxa (Chap. 17) 14 eunotioid and asymmetrical naviculoid diatoms (Chap. 18), and 14 keeled and canalled forms (Chap. 19). Diatoms are found in all freshwater habitats, including standing and flowing waters, and planktonic and benthic habitats, and they can often dominate the microscopic flora. Because diatoms inhabit a broad array of habitats but many have specific habitat requirements, they have been used in freshwater environment assessment and to monitor long-term changes in ecological characteristics (Chap. 23).

J. Dinoflagellates Pyrrhophyta or dinoflagellates are characterized by chloroplasts that have three surrounding membranes, thylakoids in stacks of three, peridinin that masks chlorophyll-a and -c, true starch, a nucleus that has condensed chromosomes in cell cycle phases, a theca covering, and frequently a transverse and posterior flagellum. (Table I; Chap. 20). There are 37 genera in North American freshwater habitats (Chap. 20). The dinoflagellates are typically minor components of the phytoplankton of lakes and ponds, but sometimes form dense blooms, particularly in the presence of high levels of nitrates and phosphates (Chap. 20).

K. Cryptomonads Cryptophyta, cryptomonads or cryptophyte algae, have chloroplasts that have four surrounding membranes in which a nucleomorph occurs between the outer and inner two membranes, thylakoids in loose pairs, phycocyanin or phycoerythrin that masks chlorophyll-a and -c, true starch as the photosynthetic reserve, a periplast, and two subapical flagella (Table I; Chap. 21). There are 12 genera reported from the inland waters of North America (Chap. 21).

8


Cryptomonads are typically planktonic in lakes and ponds, and are particularly diverse in temperate regions (Chap. 21).

L. Brown Algae Phaeophyceae or brown algae are distinguished by chloroplasts that have four surrounding membranes, thylakoids in stacks of three, fucoxanthin that masks chlorophyll-a and -c, laminarin as the photosynthetic reserve, and alginates commonly as the wall matrix component. There are six genera of freshwater brown algae, four of which have been collected in North America (Chap. 22).

Brown algae are predominantly marine in distribution; less than 1% of the species are from fresh water. The inland species are benthic, either in lakes or streams, and distribution is quite scattered (Chap. 22).

III. TAXONOMIC CHAPTERS IN THIS BOOK The approach of this book is to break the major algal groups into manageable taxonomic units, resulting in multiple chapters for those divisions that have many freshwater representatives. The following key gives major characteristics to allow the reader to immediately proceed to the appropriate chapter to determine an unknown algal sample.

A. Key 1a.

Cells with no chloroplasts; typically blue–green colored throughout (occasionally black, purple, brown, or reddish).......................2

1b.

Cells with variously colored pigments localized in one or more chloroplasts..................................................................................3

2a.

Organisms unicellular or colonial (coccoid cyanobacteria)...................................................................................................Chapter 3

2b.

Organisms filamentous (filamentous cyanobacteria)............................................................................................................Chapter 4

3a.

Cells stain positively (purple to black) with iodine for true starch.................................................................................................4

3b.

Cells do not stain positively (orange to reddish brown) with iodine for starch..............................................................................9

4a.

Green-colored chloroplasts with chlorophyll-a and -b as predominant photosynthetic pigments....................................................5

4b.

Chloroplasts with other colors and predominant photosynthetic pigments........................................................................................8

5a.

Organisms flagellated in the vegetative state (flagellated green algae)...................................................................................Chapter 6

5b.

Organisms nonflagellated in the vegetative stage................................................................................................................................6

6a.

Organisms coccoid (nonmotile unicells of various shapes) or colonial in forms without conjugation (coccoid and colonial nonmotile green algae).......................................................................................................................Chapter 7

6b.

Organisms filamentous, plantlike, or with sexual reproduction by conjugation.................................................................................7

7a.

Organisms with filamentous, bladelike, or plantlike morphologies without conjugation (filamentous and plantlike green algae).................................................................................................................................Chapter 8

7b.

Organisms with coccoid or filamentous morphologies with conjugation (conjugating green algae filaments and desmids) .............................................................................................................................................................................................Chapter 9

8a.

Cells with golden-colored chloroplasts with peridinin as the predominant photosynthetic pigment; two separate flagella typically with transverse and posterior insertions (dinoflagellates)....................................................................................................Chapter 20

8b.

Cells with blue-, brown-, or red-colored chloroplasts with either phycocyanin or phycoerythrin as the predominant photosynthetic pigment; flagella subapical (cryptophyte algae) ..................................................................................................................Chapter 21

9a.

Cells with blue- or red-colored chloroplasts with phycocyanin or phycoerythrin as the predominant photosynthetic pigment (red algae).............................................................................................................................................................................Chapter 5

9b.

Cells green or golden colored with other predominant photosynthetic pigments..............................................................................10

10a.

Motile green-colored cells with a pellicle (layer below the plasma membrane that often appears as spiral strips on the cell) (euglenoids)........................................................................................................................................................................Chapter 10

10b.

Nonmotile or motile, yellow–green- or golden-colored cells; naked or walled cells without a pellicle ..............................................11

11a.

Yellow–green-colored chloroplasts with chlorophyll-a and -c as the predominant photosynthetic pigments (eustigmatophyte, raphidiophyte, and tribophyte algae).....................................................................................................Chapter 11

11b.

Golden-colored chloroplasts with fucoxanthin as the predominant photosynthetic pigment............................................................12

12a.

Cells with a silica frustule covering (diatoms)...................................................................................................................................13


9

12b.

Cells with no covering or with one that is not a siliceous frustule (may be siliceous scales) .........................................................17

13a.

Frustules in the valve view are radially symmetrical or symmetrical in more than two planes (centric diatoms)..................Chapter 15

13b.

Frustules symmetrical in the valve view in one or two planes ..........................................................................................................14

14a.

Frustules without a raphe or with one-on-one valve only (araphid and monoraphid diatoms)............................................Chapter 16

14b.

Frustules with two raphes................................................................................................................................................................15

15a.

Raphe in an elevated keel or a canal (keeled and canalled raphid diatoms).........................................................................Chapter 19

15b.

Raphe not in a keel or canal............................................................................................................................................................16

16a.

Frustules bilaterally symmetrical in valve view (biraphid symmetrical diatoms) .................................................................Chapter 17

16b.

Frustules not bilaterally symmetrical in valve view (eunotioid and asymmetrical biraphid diatoms)...................................Chapter 18

17a.

Cells with a specialized appendage, the haptonema, in vegetative and/or reproductive stages (haptophyte algae)...............Chapter 13

17b.

Cells without a haptonema in any stage...........................................................................................................................................18

18a.

Vegetative cells with siliceous scales (synurophyte algae) ...................................................................................................Chapter 14

18b.

Vegetative cells without siliceous scales ....................................................................................................................................19

19a.

Exclusively benthic; often macroscopic thalli with no unicellular or colonial representatives; cell walls with alginates (brown algae).....................................................................................................................................................................Chapter 22

19b.

Mostly planktonic with some attached representatives; numerous unicellular or colonial representatives; mostly microscopic thalli; cell walls without alginates (chrysophyte algae)..................................................................................................................Chapter 12

LITERATURE CITED Graham, L. E., Wilcox, L. W. 2000. Algae. Prentice–Hall, Upper Saddle River, NJ, 640 pp., glossary, literature cited, and index. Prescott, G. W. 1962. Algae of the Western Great Lakes area. W. B. Brown, Dubuque, IA, 977 p. Sheath, R. G. 1984. The biology of freshwater red algae. Progress in Phycological Research 3:89–157. Sheath, R. G., Hambrook, J. A. 1990. Freshwater ecology, in: Cole, K. M., Sheath, R. G., Eds. Biology of the red algae. Cambridge University Press, Cambridge, UK, pp. 423–453.

Smith, G. M. 1950. Freshwater algae of the United States, 2nd ed. McGraw–Hill, New York, 719 p. Sze, P. 1998. A biology of the algae 3rd ed. McGraw–Hill, Boston, 278 p. Van den Hoek, C., Mann, D. G., Jahns, H. M. 1995. Algae. An introduction to phycology. Cambridge University Press, Cambridge, UK, 623 p. Vis, M. L., Sheath, R. G., Hambrook, J. A., Cole, K. M. 1994. Stream macroalgae of the Hawaiian Islands: A preliminary study. Pacific Science 48:175–187.


2

FRESHWATER HABITATS OF ALGAE John D. Wehr

Robert G. Sheath

Louis Calder Center — Biological Station and Department of Biological Sciences Fordham University, Armonk, New York 10504

Office of Provost and Vice President for Academic Affairs California State University, San Marcos San Marcos, California 92096

I. What is Fresh Water? II. Lentic Environments A. Major Lakes of North America B. Lake Basins C. Lake Community Structure and Productivity D. Ponds, Temporary Pools, and Bogs E. Phytoplankton of Lakes and Ponds F. Benthic Algal Assemblages of Lakes III. Lotic Environments A. Major Rivers of North America B. Geomorphology of Rivers C. The River Continuum and Other Models D. Benthic Algal Communities of Rivers E. Phytoplankton Communities of Rivers

IV. Wetlands A. Functional Importance of Algae in Wetlands B. Algal Diversity in Freshwater Wetlands C. Algal Communities of Bogs V. Thermal and Acidic Environments A. Thermal Springs B. Acid Environments VI. Unusual Environments A. Saline Lakes and Streams B. Snow and Ice C. Other Unusual Habitats Literature Cited

I. WHAT IS FRESH WATER? The study of freshwater algae is really the study of organisms from many diverse habitats, some of which are not entirely “fresh.” Although the oceans are clearly saline (⬇ 35 g salts L–1) and most lakes are relatively dilute (world average < 0.1 g L–1; Wetzel, 1983a), there is enormous variation in the chemical composition of the nonmarine habitats that algae occupy. Conditions in lakes and rivers vary not only in salinity, but also in size, depth, transparency, nutrient conditions, pH, pollution, and many other important factors. Aquatic ecologists also use the term “inland” waters to encompass a greater range of aquatic ecosystems. Even this term may be unsatisfactory, because algae occupy many other habitats, such as snow, soils, cave walls, and symbiotic associations (Round, 1981). Organisms grouped together in this volume as Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

freshwater algae fall into a large, but ecologically meaningful collection of environments: all habitats that are at least slightly wet, other than oceans and estuaries. One reason for such a broad scope is that inland saline lakes, snow and ice, damp soils, and wetlands are studied by phycologists and ecologists who also examine more traditional freshwater environments. Some genera with terrestrial species, such as Vaucheria, Nostoc, Chlorella, and Prasiola, also have species found principally in streams or lakes (Smith, 1950; Whitton, 1975). In North America, the variety of freshwater habitats colonized by algae is very rich, and offers an enormous and fascinating range of environments for their study. The distinction between marine and freshwater habitats is revealed in the variety of algae that occur in these environments. There are no exclusively freshwater divisions of algae, but certain groups exhibit 11

12

John D. Wehr and Robert G. Sheath

greater abundance and diversity within fresh waters, especially Cyanobacteria, Chlorophyta, and Charophyta (Smith, 1950). Within the green algae, conjugating greens and desmids (Zygnematales, Chap. 9) comprise a very rich collection of species that almost exclusively occupy fresh water. Other groups, such as the diatoms and chrysophytes, are well represented in both spheres. Other groups, particularly the Phaeophyta, Pyrrophyta, and Rhodophyta, exhibit greater diversity in marine waters (Smith, 1950; Bourrelly, 1985). Most freshwater algae are best described as cosmopolitan, although there are reports of endemic chrysophytes, green algae, rhodophytes, and diatoms (Tyler, 1996; Kociolek et al. 1998), and at least some species of cyanobacteria (Hoffmann, 1996). Many algal taxa have particular environmental tolerances or requirements, and are ecologically restricted, but still geographically widespread. The euglenophyte Colacium is almost exclusively epizooic on aquatic invertebrates, but is widely distributed throughout North America (Smith, 1950; Chap. 10). The chrysophyte Hydrurus foetidus is an exclusive inhabitant of cold mountain streams, but is distributed worldwide (Smith, 1950; Whitton, 1975, Chap. 12). Even specialized taxa such as Basicladia chelonum (Chlorophyceae), which is restricted mainly to the shells of turtles, has been collected from many habitats throughout eastern North America (Smith, 1950; Prescott, 1962; Colt et al., 1995). The actual distribution of apparently disjunct freshwater species must therefore be viewed with some caution until detailed surveys have been conducted (see, for example, Linne von Berg and Kowallik, 1996; Müller et al., 1998). Inland waters represent only about 0.02% of all water in the biosphere, and nearly 90% of this total is contained within only about 250 of the world’s largest lakes (Wetzel, 1983a). Nonetheless, it is fresh water that is most important for human consumption and is most threatened by human activities. Algal ecologists play an important role in the understanding of aquatic ecosystems, their productivity, and water quality issues (Round, 1981; Brock, 1985a; Hoffmann, 1998; Dow and Swoboda, 2000; Oliver and Ganf, 2000, Chaps. 23 and 24). This chapter examines the habitats of freshwater algae and how differences in these systems affect algal communities.

II. LENTIC ENVIRONMENTS Lentic environments include standing waters from the smallest ponds (a few square meters) to enormous bodies of water (e.g., Laurentian Great Lakes: 245,000 km2). Their formation, geography, limnology, and conservation have been covered in several texts

(Hutchinson, 1957, 1967, 1975; Frey, 1963; Wetzel, 1983a; Cole, 1994; Abel et al., 2000). This section summarizes some features of lentic environments as they pertain to the ecology and distribution of freshwater algae.

A. Major Lakes of North America Worldwide, the single largest volume of freshwater — nearly 20% of the world’s total — is located in Lake Baikal, Siberia (23,000 km3), but the North American Great Lakes (Fig. 1A) collectively represent the largest total volume of nonsaline water on Earth, approximately 24,600 km3 (Wetzel, 1983a). North America is home to many spectacular large and deep freshwater systems, nearly half of all the world’s lakes greater than 500 km2 (Hutchinson, 1957). Two of the most impressive lakes are subarctic: Great Slave Lake (28,200 km2; 614 m deep; deepest in North America) and Great Bear Lake (30,200 km2; > 300 m deep) in the Northwest Territories (Hutchinson, 1957). Crater Lake in Oregon (Fig. 1B) is much smaller (64 km2 in area), but is the deepest lake in the United States (608 m) and seventh deepest in the world (Edmondson, 1963). The largest lakes on the continent are located in northern and temperate regions, although Great Salt Lake (Utah) is a massive remnant lake (> 6000 km2) that has a mean depth (⬇ 9 m) and very high salinity (130–280 g L–1) that fluctuate with available moisture, and occupies a portion of the Pleistocene Lake Bonneville, which had an area > 51,000 km2 and a depth of 320 m (Hutchinson, 1957).

B. Lake Basins Sizes and shapes of lake basins (their morphometry) have profound effects on the physics, chemistry, and biology of lake ecosystems, and influence the composition of algal communities and their productivity. Lake basins differ in morphometry as a result of the forces that created them, many of which were catastrophic events from the past, principally glacial, seismic, and volcanic activity. Hutchinson (1957) distinguished 76 different lake types based on their origins; these were classified into a simpler scheme by Wetzel (1983a) that is summarized in Table I. Glacial activity is the most important agent in North America. It created millions of small and large basins from the arctic south to the southern extent of the Wisconsin ice sheet. In this period (15,000–5000 years BP), many basins became closed by morainal deposits, including the Laurentian Great Lakes. Some morainal lakes occur at the ends of long valleys after glaciers have receded, including the Finger Lakes of

2. Freshwater Habitats of Algae

13

FIGURE 1 Examples of different types of lakes in North America: A, Laurentian Great Lakes; B, Crater Lake, Oregon, a deep caldera lake; C, New York Finger Lakes; D, kettle lakes in Becher’s Prairie, central British Columbia; E, pothole lakes in Qu’appelle Valley, Saskatchewan; F, Louise Lake, a cirque in Mt. Rainier National Park, Washington. Photos A and C courtesy of U.S. Geological Survey EROS Data Center, reproduced with permission; photo B by R. G. Sheath; photo D by R. J. Cannings; photo E by P. R. Leavitt, reproduced with permission; photo F by J. D. Wehr.

New York (Fig. 1C), which are elongate, radially arranged basins that range from small ponds to large lakes, such as Seneca (175 km2 area, 188 m depth; Hutchinson, 1957; Berg, 1963). However, most glacially formed lakes are small kettles scattered across the continent (Fig. 1D and E). Glacial scouring in mountainous terrain may form deep amphitheater-like cirques (Fig. 1F), which are common from Alaska through the western mountain ranges south to tropical locations in Costa Rica (Haberyan et al., 1995). Glacial

basins within narrow valleys may form deep fjord lakes (Fig. 2A), or a chain of smaller lakes known as paternosters. Several forces, including glacial scour and lava flow, combined to form the small (9.9 km2) but deep (259 m) fjordlike Garibaldi Lake (Northcote and Larkin, 1963; Fig. 2B). Ice-formed (thermokarst) lakes, which result from freezing and thawing action in ice and soil, are common in the Arctic. All are shallow but vary from large elliptical basins (up 70 km2 area) to small (10–50 m diameter), “polygon” ponds (Fig. 2C),

14


TABLE I Major Lake Basin Types, Grouped According to the Principal Forces That Shaped Them Type of forces

Basins

Principal force

North American examples

Catastrophic

1. Glacial

Glacial scouring Moraine deposits Kettles Cryogenic Graben Uplift Landslide Caldera Maar Meteor

Great Slave Lake, NWT Finger Lakes, NY; Moraine Lake, AK Linsley Pond, MA; Cedar Bog Lake, MN Many polygon lakes, AK Lake Tahoe, CA-NV Lake Okeechobee, FL Mountain Lake, VA Crater Lake, OR Zuni Salt Lake, Mexico New Quebec Lake, PQ

Doline Salt collapse Oxbows Plunge Shoreline Deflation Beaver Human

Deep Lake, FL; limestone areas of KY-IN-TN Montezuma Well, AZ; Bottomless Lakes, NM Lake Providence, LA Fayetteville Green Lake, NY Along Laurentian Great Lakes; Cape Cod area Moses Lake, WA; Sandhills region, NB Many locations in northern regions Lake Mead, AZ-NV; Cherokee Reservoir, TN

2. Tectonic

3. Volcanic 4. Meteor Noncatastrophic

5. Solution 6. Rivers 7. Coastal 8. Wind 9. Organic

FIGURE 2 Other glacial and ice-formed lakes: A, Okanagan Lake (BC), a fjord lake; B, Garibaldi Lake, formed by glacial scour and lava damming; C, arctic polygon ponds. Photos A courtesy of NASA, reproduced with permission; photos B and C by R. J. Cannings, reproduced with permission.

which are estimated at more than a million in number (Livingstone, 1963; Sheath, 1986). Tectonic basins are formed by movements of the Earth’s crust. Among these, grabens form when fault lines create often enormous depressions, such as Lake

Tahoe, a symmetrical, deep (505 m), and steep-sided lake (Fig. 3A). Tahoe (double fault lines) and Lakes Baikal and Tanganyika (single faults) include the deepest lakes in the world, although less spectacular examples also occur. Lago de Peten (Guatemala) is


15

FIGURE 3 Tectonic lakes: A, Lake Tahoe, a graben, viewed from the northeast; B, Spirit Lake, Washington, a landslide lake near Mt. St. Helens; C, Lake Okeechobee, Florida (larger lake on lower right), and surrounding lakes. Photos A and B by J. D. Wehr; photo C reproduced with permission of the South Florida Water Management District.

the largest (567 km2) and deepest (> 32 m) lake on the Yucatan Peninsula (Covich, 1976), and the largest lake in Mexico, Lago Chapala (1109 km2), is also a tectonic trench (Serruya and Pollingher, 1983). Landslide lakes form when water flows through an existing depression and is blocked by rock or other material, as in as Spirit Lake, near Mount St. Helens (WA; Fig. 3B) and Mountain Lake (VA; Parker et al., 1975). In the Pliocene, shallow marine areas were raised above sea level and existing depressions filled with freshwater, as with Lake Okeechobee (Fig. 3C), a large (1840 km2), shallow (4 m) subtropical lake in Florida. Volcanic lakes are among the deepest and most steep-sided lakes on the continent. The exceptionally clear water in Crater Lake (OR), a collapsed caldera (Fig. 1B), has Secchi depths between 20 and 30 m, with 1% surface light down to 100 m (Larson et al., 1996). Lake Atitlán in Guatemala is an alpine tropical caldera (8.2 km2; 1550 m elevation) that reaches a depth of 341 m, making its ratio of depth to surface area four times greater than better known Crater Lake. Lake Nicaragua (Fig. 4A; 7700 km2 area; depth ⬇ 60 m) was formed by volcanic lava damming an existing valley (Cole, 1963). Some volcanic lakes, such as Yellowstone

Lake (WY) and Surprise (AK) Lake, have hydrothermal vents that influence temperature, pH, and O2 conditions, and may contribute trace metals (Pierce, 1987; Larson, 1989; Cameron and Larson, 1993). Other volcanic lakes formed from violent explosions of cinder cones (maars) and are often nearly circular in outline, such as Big Soda Lake in Nevada, Lago Chamico in El Salvador (Cole, 1963), and Laguna Hule in Costa Rica (Umana-Villalobos, 1993). Volcanic lakes also occur in Mexico, Guatemala, Nicaragua, and El Salvador (but not all are deep), and some were formed as recently as 500 year ago (Hutchinson, 1957; Cole, 1963). In limestone regions, solution or sinkhole lakes form from the dissolution of bedrock by surface and underground waters charged with CO2 (Cole, 1994). Sinks may be circular, elliptical, or irregular in outline, and occur throughout Kentucky, Indiana, Tennessee, Florida, Mexico, and Guatemala (Fig. 4C). Florida is especially rich in sinkholes, with several hundred lakes and ponds ranging from less than 1 ha to several square kilometers (Fig. 3C). In north Florida, some lakes are relatively dilute and colored with organic matter from pine litter, whereas others are clear,

16


FIGURE 4 Volcanic and solution lakes: A, Lakes Managua (near) and Nicaragua, two large calderas; B, Volcan Maderas (Nicaragua), a small caldera located on an island in Lake Nicaragua; C, unnamed, deep limestone sink in the Yucatan Peninsula; D, Montezuma Well (AZ), a collapsed travertine limestone sink. Photo A courtesy of NASA, reproduced with permission; photo B by A. Merola; photo C by L. P. Burney; photo D by D. W. Blinn, reproduced with permission.

hardwater systems (Shannon and Brezonik, 1972). Spring-fed sinkholes may become isolated or thermally constant, such as Montezuma Well (AZ; Fig. 4D), creating an environment with high algal productivity (600 g C m–2 y–1; Boucher et al., 1984) and unusual communities. Montezuma is a collapsed travertine system with several endemic invertebrates, but no fish, rotifers, or cladocerans (Cole, 1994). Lakes may form through wind action, whereby deposited sand blocks existing valleys (e.g., Moses Lake, WA) or forms depressions in dunes, as in Nebraska and Texas (Cole, 1963; Edmondson, 1963). River-formed lakes, including oxbows, occur across North America where rivers traverse level terrain, enabling siltation of meandering valleys (Fig. 5A). Other small basins form in the plunge pools of waterfalls (Fig. 5B). Fayetteville Green Lake is a relatively deep (59 m; 0.3 km2 area) plunge-pool lake in central New York that apparently has never fully mixed; it was formed during the Pleistocene when melting glaciers formed a vast waterfall (Berg, 1963; Brunskill and Ludlam, 1969). For many centuries the principal biological agent responsible for creating lakes in North America was

the beaver (Castor canadensis), which dams smaller rivers to form lakes and ponds (Hutchinson, 1957; Fig. 5C). Today, reservoirs are a more important group of lentic ecosystems, the size and number of which are increasing worldwide (Fig. 5D). The physical and chemical properties of reservoirs differ from natural lakes with respect to dendritic or eccentric morphometry (deepest near the dam), shorter flushing period, irregular water level, greater dissolved and suspended solids, and less stable littoral zone (Wetzel, 1990). Because reservoirs serve hydroelectric, flood control, or drinking water uses, they occur in many biomes. A few lakes may have been formed by meteor impact, such as New Quebec Lake, a nearly perfectly circular basin (3.4 km diameter) in a region of irregular, glacially formed lakes in northern Quebec (Cole, 1994). Carolina Bays, which are not truly bays, are a series of roughly 150,000 small, shallow, elliptical basins, with a distinctive NW–SE orientation, and concentrated along the Atlantic coast from New Jersey south to Florida. It is their directional orientation that has caused some to speculate that their origin may be from meteor showers, whereas others have suggested wind action or artesian springs (Hutchinson, 1967;


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FIGURE 5 Lakes formed by river action and other agents: A, an oxbow lake in Texas; B, a plunge pool below Waimea Falls, Hawaii; C, a beaver-dam lake (southern British Columbia); D, reservoir (eastern Colorado). Photo A by J. Cotner; photo B by D. Burney, reproduced with permission; photos C and D by J. D. Wehr.

Cole, 1994). Many are now filled, but those with aquatic habitats are shallow and have extensive macrophyte beds and low algal production (Schalles and Shure, 1989).

C. Lake Community Structure and Productivity 1. Lake Zones and Thermal Patterns Regions within lakes exhibit physical and chemical differences that affect algal communities. The open water region of lakes is termed the pelagic (or limnetic) zone, whereas close to shore is the littoral zone, where the greatest exchange between nutrient-rich sediments and the water occurs (Fig. 6). The littoral zone is colonized by submersed (e.g., Ceratophyllum, Potamogeton, and Vallisneria) and emergent (e.g., Scirpus and Typha) flowering plants, although some macroalgae (e.g., Chara, Nitella, and Batrachospermum) and nonflower-

ing plants (mosses, liverworts) also occur (Hutchinson, 1975). Vertical zones also develop in temperate regions. In early spring, most temperate lakes are well mixed, with similar temperatures and chemical conditions from top to bottom. As temperatures increase, upper mixed waters become thermally isolated from deeper and colder waters, a process which is termed stratification. The upper epilimnion continues to become warmer, receives greater irradiance, and is well mixed and oxygenated; deeper waters remain cool (ca. 4°C in deep lakes) and dissolved gases are consumed by microbial activity. At an intermediate depth (the metalimnion), temperature declines, often sharply, with reduced heat penetration and reduced mixing (= thermocline if ≥ ∆1° m–1). Density gradients formed by this thermal barrier may be sufficient to support dense algal populations (Pick et al., 1984); here light is adequate for photosynthesis coupled with a greater supply of nutrients. Algal production may create metal-

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FIGURE 6 Diagram that represents the zones and algal habitats within typical oligotrophic and eutrophic lakes (EZ = euphotic zone).

imnetic oxygen maxima in clear oligotrophic lakes (Wetzel, 1983a; Parker et al., 1991). The hypolimnion is a deeper, cooler region with greater nutrient supply, but reduced (approaching zero) oxygen levels; light may be too low for photosynthetic algal growth except in very clear lakes. Patterns of thermal stratification and mixing differ with altitude and across biomes. A dimictic pattern, in which lakes stratify in the summer, mix in the autumn, stratify in the winter after ice cover, and mix in the spring after ice-out, is most common in temperate climates. Warm monomictic lakes (stratification and one mixing period; summer epilimnion > 4°C) occur in warmer climates or in large basins without ice cover (Wetzel, 1983a). Examples include the Great Lakes, larger Finger Lakes, Lake Tahoe, lakes in warm or coastal climates, and subtropical, high altitude lakes. Cold monomictic lakes, with a single turnover in

summer or late spring, occur mainly in alpine and arctic areas (temperatures ≤ 4°C). Oligomictic lakes have rare mixing periods (less than once per year), where temperature strata (summer epilimnion > 4°C) may remain for some years; this pattern is most common in deep tropical lakes (Wetzel, 1983a). Polymictic lakes are shallow systems with frequent or continuous mixing, and occur in tropical and equatorial areas such as Lake Managua (Xolotlán), Nicaragua (Erikson et al., 1997). Amictic lakes, uncommon in North America (some in Greenland; common in the Antarctic), are perennially ice covered and do not turn over. A special class of lakes in which upper waters (mixolimnion) mix, but deeper waters (monimolimnion) never circulate, are termed meromictic (Wetzel, 1983a). These lakes have a very stable chemical and temperature density gradient, known as a chemocline, that results in anoxic conditions, H2S, and purple sulfur bacteria in


the water column. A strong depth to surface area ratio is usually necessary to maintain meromixis; Fayetteville Green Lake in New York and Hot Lake in Washington are examples (Hutchinson, 1957).

2. Lake Productivity Limnologists distinguish lakes according to a gradient of primary production (14C uptake, algal growth) or biomass, from oligotrophic (annual average < 50– 300 mg C m–2 d–1; < 0.05–1 µg chlorophyll-a L–1) to eutrophic (> 1000 mg C m–2 d–1; 15–100 µg chlorophyll-a L–1), and these levels are influenced, at least in part, by the properties of lake basins (Wetzel, 1983a; Likens, 1985). Oligotrophic lakes are poor in nutrients, usually deep and steep-sided, and have high transparency, a narrow littoral zone, abundant dissolved oxygen with depth, and larger relative hypolimnion volume. Lake Tahoe is an ultra-oligotrophic lake that has exceptional water clarity, although average Secchi depths have declined from about 30 to 23 m and primary productivity levels have more than doubled (40–100 mg C m–2 d–1) since the late 1950s, following increased nutrient loading from regional development (Goldman, 1988). Eutrophic lakes are nutrient-rich, often shallower with a broad littoral zone, and have depleted summer hypolimnetic oxygen and reduced transparency. Eutrophication of lakes often result when nutrients are added as sewage, detergents, or fertilizers (Wetzel, 1983a). Lake Mendota, Wisconsin, is an example of a larger lake (39 km2) that has an average depth of only 12 m and receives substantial input from agricultural and urban sources. The lake has experienced algal blooms and high nutrient levels for more than a century, and water transparency has been consistently low for more than 80 years (Brock, 1985a).

D. Ponds, Temporary Pools, and Bogs Smaller lentic environments, often called ponds, may seem very similar to lakes except for their size, but they have distinct properties. Ponds are shallow enough either to support rooted aquatic vegetation across the entire basin or to fail to stratify during the summer. The term “pond,” however, is not a precise concept, despite its frequent usage. In New England, the word is often applied to fairly substantial lakes, such as Linsley Pond (9.4 ha, max depth 14.8 m; Brooks and Deevey, 1963) and Long Pond (40 ha, 22 m; Canavan and Siver, 1995), both of which stratify in summer. Small limnetic systems can be divided into those that contain water year round, often called permanent, and temporary waters that become dry each year and are termed vernal ponds (Wetzel, 1983a).

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Temporary ponds are located in low-lying areas that fill with snow melt or spring runoff, but dry up in the summer (Wetzel, 1983a); resident algal populations may be quite substantial during the growing season and rely on resistant resting stages (e.g., dinoflagellate cysts [Chap. 20], cyanobacterial akinetes [Chap. 4], and zygospores of Zygnematales [Chap. 9]) capable of surviving extended adverse conditions (Reynolds, 1984a, b). Arctic tundra ponds typically freeze solid for many months, resulting in a growing season of only 60–100 days (Wetzel, 1983a; Sheath, 1986). Algal communities often consist of small flagellates (particularly Cryptophyceae and Chrysophyceae; see Chaps. 21 and 12–14) that bloom during the brief summer of long daylight, although many species apparently do not form resting stages and survive freezing conditions in their vegetative condition (Sheath, 1986).

E. Phytoplankton of Lakes and Ponds Assemblages of planktonic algae vary greatly among lake basins and biogeographic regions. They include members of all algal divisions except the Rhodophyta and Phaeophyta. Their ecology has been the subject of many reviews, including Hutchinson (1967), Kalff and Knoechel (1978), Round (1981), Reynolds (1984a), Munawar and Talling (1986), Sandgren (1988), Munawar and Munawar (1996, 2000), Stoermer and Smol (1999), and Whitton and Potts (2000).

1. Phytoplankton Diversity, Composition, and Seasonal Succession Every collection of freshwater algae is characterized by a fascinating and perplexing diversity of species, many of which are potential competitors for common resources. Up to several hundred algal species may comprise the phytoplankton community of a typical north-temperate lake (Kalff and Knoechel, 1978). This paradox of many potentially competing phytoplankton species coexisting in a relatively uniform habitat was proposed by Hutchinson (1961) to be possible because of the numerous niches within a lake, as well as variation within the environment over time. Many factors contribute to phytoplankton diversity and production, including temporal and spatial variations in nutrient supply, grazing, temperature, and parasitism (Turpin and Harrison, 1979; Crumpton and Wetzel, 1982; Sommer, 1984; Bergquist and Carpenter, 1986). The biomass of phytoplankton is thought to be driven mainly by nutrient supply and herbivory in all lakes, but their temporal dynamics differ in eutrophic and oligotrophic systems (Walters et al., 1987; Carpenter et al., 1993).

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Ecosystem-level studies of lake food webs in some cases have ignored phytoplankton species composition, and instead treat this component as a black box. This approach is incomplete because the functional properties of algal assemblages vary strongly with species composition. Taxonomic information is important for ecological studies because many of the features used to classify algae, such as photosynthetic pigments, storage products, motility, reproduction, cell ultrastructure, and even DNA sequence information, have functional importance. For example, among freshwater phytoplankton, only cyanobacteria and some cryptomonads possess the red accessory pigment phycoerythrin, which has an absorption maximum (540–560 nm) that broadens the photosynthetic capacity of cells and may facilitate growth at greater depths (Goodwin, 1974, Chap. 21). Similarly, only certain species of cyanobacteria are able to fix N2, mainly those that possess heterocysts (e.g. Anabaena, Aphanizomenon, and Nostoc), although some non-heterocystous forms with thick sheaths or that form dense aggregations (Gloeocapsa, Oscillatoria, Microcoleus, and Plectonema; see Chap. 4) also have this ability (Bothe, 1982; Paerl et al., 1989). Algal flagellates are a polyphyletic collection of different protists, but they form an important ecological group because of their tendency toward mixotrophy (Porter, 1988). This fact is not coincidental, because the flagellar apparatus is directly involved in the capture of bacterial prey (Andersen and Wetherbee, 1992). Diversity in size is also an important property of phytoplankton communities. One scheme (Sieburth et al., 1978) categorizes sizes into groups that differ over orders of magnitude: picoplankton (> 0.2–2 µm), nanoplankton (> 2–20 µm), microplankton (> 20–200 µm), and mesoplankton (> 200–2000 µm) include most algal cells and colonies in freshwater. Expressed in terms of volume, the sizes of freshwater phytoplankton span at least 8 orders of magnitude (Reynolds, 1984b). The bacteria-sized picoplankton have attracted recent interest because they have been found to dominate (at least numerically) many phytoplankton communities in lakes and marine systems (Stockner et al., 2000). They occur in great numbers (105–106 mL–1), possess rapid growth rates, and are highly productive; most often reported are cyanobacteria (e.g., Cyanobium, Cyanothece, Synechococcus, and Synechocystis; see Chap. 3) and some green algae (e.g., Nannochloris). Their importance (percentage of biomass or primary production) seems to be greatest in oligotrophic and least in eutrophic lakes (Burns and Stockner, 1991; Hawley and Whitton, 1991; Wehr, 1991), although there are exceptions (Wehr, 1990; Weisse, 1993). Autotrophic picoplankton have a strong competitive ability in

P-limited conditions (Suttle et al., 1987, 1988; Wehr, 1989) and are grazed mainly by micro-zooplankton (ciliates, flagellates), rather than cladocerans or copepods (Pernthaler et al., 1996; Hadas et al., 1998), making them important links between microbial and classical food webs (Christoffersen et al., 1990; Sommaruga, 1995). Size affects sinking rate and thus the ability of cells to remain in the euphotic zone. Smaller cells tend to be spherical or ellipsoid and thus sink more slowly, whereas larger forms have more elongate or complex shapes to reduce sinking. The dinoflagellate Ceratium hirundinella is a large planktonic alga (up to 400 µm) common in mesotrophic and eutrophic lakes with stable stratification (Chap. 20); the alga regulates its position in the water column by active migration and perhaps by changes in the shape and size of hornlike projections (Heaney and Furnass, 1980; Pollingher, 1987; Heaney et al., 1988). The silica walls of diatoms result in heavier cellular densities, making them susceptible to sinking. Diatoms are estimated to have 3–16 times faster sinking rates than nonsiliceous algae of equivalent sizes (Sommer, 1988). Larger colonial diatoms, such as Asterionella formosa, Fragilaria crotonensis, and Tabellaria fenestrata (see Chap. 16), have more elongate or elaborate morphologies, which may reduce sinking rates or cause some cells to rotate (Smayda, 1970; Barber and Haworth, 1981). Planktonic cyanobacteria and desmids produce extracellular mucilage, which may aid in buoyancy (Round, 1981). Some cyanobacteria, such as Anabaena flos-aquae and Microcystis aeruginosa, also maintian their position in the water column using gas vacuoles (Chaps. 3 and 4). Observed changes in the population size may be the result of differences in vertical migration and sinking to lower strata, rather than actual changes in numbers. Regular seasonal changes (seasonal succession) in phytoplankton populations over many years have been observed widely and reported in long-term limnological records. Few species have been documented as thoroughly as Asterionella formosa, a diatom found in mesotrophic, temperate lakes (Round, 1981; Reynolds, 1984a, Chap. 16). In Lake Windermere, populations increase after turnover and peak in the spring when light levels and temperatures are increasing and the Si:P ratio is near maximum (Fig. 7). Populations decline over the summer and have a second (usually smaller) peak in the autumn (Lund, 1964; Reynolds, 1984a; Neale et al., 1991). Asterionella was the spring dominant in Lake Erie from at least 1931 until about 1950, when nutrient enrichment selected for diatom genera such as Stephanodiscus, Aulacoseira (Melosira; see Chap. 15), and Fragilaria, and filamentous cyanobacteria such as Anabaena and Aphanizomenon (Davis,


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FIGURE 7 Seasonal periodicity in the abundances (cells per liter) of Asterionella formosa (solid line), Fragilaria crotonensis (dashed line), and Tabellaria flocculosa (dotted line), and concentrations of dissolved silica (upper, black; milligrams per liter) in Windermere, English Lake District 1945–1960. Reproduced with permission from A. Horne and C. R. Goldman, Limnology, 2nd ed. Copyright © 1994, McGraw–Hill, New York.

1964). As eutrophication receded, there was a 70–98% reduction in numbers of Stephanodiscus spp. and Aphanizomenon flos-aquae, and the reappearance of Asterionella (Makarewicz, 1993). Further details are given in Munawar and Munawar (1996). Population dynamics generally follow predictable changes in temperature, sunlight, nutrients, and other factors. However, algal population dynamics exhibit more abrupt changes than these gradual trends predict, suggesting that other factors may drive seasonal succession. Round (1971) described these as “shock” peri-

ods: times in the lake cycle, such as turnover, that lead to sharp changes in chemical or physical conditions. Lakes with less predictable conditions or more shock periods exhibit frequent changes in species composition and shorter growth peaks (Fig. 8A), but when conditions are more stable, populations may persist over longer time periods (Fig. 8B). Furthermore, coexistence (temporal overlap) among species will result if a greater variety of habitats are available, perhaps through stratification, basin complexity, or multiple inflows (Fig. 8C). These temporal patterns repeat only if nutrient

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FIGURE 9 Predicted (lines) and observed (points) outcomes of competition between Asterionella formosa (stars = dominant) and Cyclotella (Stephanocyclus) meneghiniana (diamonds = dominant) under varying levels of Si and P, indicating that coexistence (solid circles) is possible in an intermediate range of Si:P ratios (From Tilman, D.; Resource Competition and Community Structure. Copyright © 1982 by Princeton University Press. Reprinted by permission of Princeton University Press.

FIGURE 8 Diagrammatic representation of seasonal growth curves of freshwater phytoplankton species in lakes with different ecological conditions and habitat complexity: A, variable ecological conditions or frequent shock periods, with short peaks and low overlap; B, more stable conditions or longer stratification periods, with greater temporal overlap; C, complex lake basins with more habitats and peaks of different duration, overlap, and frequency. Redrawn and adapted from Round (1972).

or other conditions are stable and other disturbances are kept to a minimum. For example, in Lake Michigan Tabellaria sp. and Asterionella formosa exhibited regular peaks in the spring phytoplankton community for several decades, but Asterionella numbers have declined in more recent years, while Stephanodiscus and filamentous cyanobacteria have increased following increases in P supply (Makarewicz and Baybutt, 1981). Laboratory experiments with Stephanocyclus (Cyclotella) meneghiniana and Asterionella formosa, which have different Si and P requirements and consumption rates, have indentified the levels of these nutrients at which the two species may coexist (Fig. 9, Tilman, 1977, 1982). Studies have quantified resource competition among other species and with other

resources, such as C:P and N:P (Kilham and Kilham, 1978; Rhee and Gotham, 1980; Sommer and Kilham, 1985; Olsen et al., 1989). In situ manipulations of phosphorus and light have demonstrated clear differences among species: some respond positively to P addition alone (e.g., Synedra radians), whereas others, such as chrysophytes (e.g., Dinobryon sertularia and Synura uvella), increase under greater light (de Noyelles et al., 1980; Wehr, 1993). Species composition and size structure each can be influenced by nutrient supply. With higher N:P supply ratios, assemblages in an oligotrophic lake were dominated by pico-cyanobacteria (Synechococcus sp.; see Chap. 3), but at lower supply ratios, larger diatoms (Nitzschia and Synedra) dominated (Suttle et al., 1987). Whole-lake N and P additions to oligotrophic Kennedy Lake (BC), used to enhance production of sockeye salmon, also affected competitive interactions among phytoplankton (Stockner and Shortreed, 1988). Loadings at N:P ratios between 10 and 25 increased algal biomass with a summer community dominated by N2-fixing Anabaena circinalis; increasing the N:P ratio to 35 retained the higher biomass, but shifted the community dominance to small-celled Synechococcus spp. Manipulations of fish densities in a small Wisconsin lake varied predation pressure on zooplankton, which in turn created different levels of grazing pressure on phytoplankton; nested nutrient-permeable chambers separated effects of recycled nutrients from relaxed herbivory (Vanni and Layne, 1997). Phyto-


plankton biomass and the abundance of many algal taxa (e.g., Peridinium inconspicuum, Chrysochromulina sp., and Staurastrum dejectum) increased with greater fish biomass, although a similar effect was seen in grazer-free diffusion chambers for some species, which suggested that the positive effect of fish may have been mediated through recycled nutrients, rather than lower grazing pressure. Some algae occupy both planktonic and benthic habitats at different periods of the year or even daily. This strategy may involve overwintering as inactive stages (e.g., algal cysts) or active recruitment of benthic forms into the pelagic zone. In many systems, flagellates such as Ceratium, Cryptomonas, and Euglena, exhibit diurnal vertical migrations toward the surface by day and into deeper strata at night (Palmer and Round, 1965; Heaney and Talling, 1980; Hansson et al., 1994), responding to patterns in light availability, temperature cues, mixing conditions, or nutrient supply. In spring and early summer, the cyanobacterium Gloeotrichia echinulata colonizes shallow sediments or submersed plants in eutrophic lakes, but in late summer, gas-vacuolate colonies migrate into the pelagic zone, representing as much as 40% of the planktonic assemblage (Barbiero and Welch, 1992). In subtropical Lake Apopka, Florida, benthic and settled planktonic diatoms (and their resting stages) are regularly resuspended into the water column by wind action, making a major contribution to the chlorophyll budget of the lake (Schelske et al., 1995). In these shallow lakes, buoyant cyanobacteria contribute to recycling phosphorus (internal loading) from sediments back into the water column (Salonen et al., 1984; Pettersson et al., 1993; Moss et al. 1997).

2. Factors That Regulate Phytoplankton Production in Lakes A large body of data clearly indicates that phytoplankton production and biomass in most lentic systems is controlled by P supply (Schindler, 1978; Wetzel, 1983a; Hecky and Kilham, 1988). Although other factors (e.g., grazing, light availability, temperature) are clearly involved in regulating phytoplankton production, only nutrients are amenable to regulation (Schindler, 1978). This concept was demonstrated in Lake Washington, which received secondary sewage (a source of P) from the city of Seattle. Blooms of planktonic cyanobacteria, especially Planktothrix (Oscillatoria) rubescens, were common in past decades (Edmondson, 1977). Today eutrophication in Lake Washington has been largely reversed through monitoring and sewage diversion, which reduced P inputs to near zero, summertime chlorophyll-a from about 45 to 5 µg L–1 and the percentage of phytoplankton as

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cyanobacteria from ⬇100 to 10% or less (Edmondson and Lehman, 1981). Eutrophic lakes with low N:P ratios favor blooms of N2-fixing cyanobacteria; hence, nutrient cycling within these lakes is closely coupled these organisms (Schindler, 1977, 1985). Algal species have different micronutrient requirements, such as Si, Mg, Ca, Fe, Mo, and Se. Diatom dominance in phytoplankton assemblages is dependent on recycled Si following spring turnover; lakes with lower Si levels may lack a springtime diatom pulse altogether. The haptophyte Chrysochromulina breviturrita develops large populations in dilute lakes undergoing the early stages of acidification (Nicholls et al., 1982, Chap. 13). Its requirement for Se and NH4+, and its inability to use NO3– favor its success in oligotrophic lakes within the pH range 5.5–6.5 (Wehr and Brown, 1985; Wehr et al., 1987). Nutrient requirements may have important interactions, for example, Mo, which is an essential micronutrient for some cyanobacteria as a co-factor for N2 fixation; its assimilation may be inhibited by elevated SO42–, which is common in saline lakes (Howarth and Cole, 1985; Marino et al., 1990). Light supply within the water column is a critical factor that also affects phytoplankton production and species composition in lakes. Although the abundance of many species is greatest in the epilimnion where irradiance is greatest, other species, including several algal flagellates are adapted to deeper waters (Lund and Reynolds, 1982). Under relatively warm and calm conditions in Blelham Tarn, more than 90% of Uroglena sp. colonies and Eudorina elegans aggregated in the upper 2 m, but in October, Trachelomonas occupied a narrow band near the thermocline (6–8 m depth). In eutrophic or highly turbid waters, some algae remain buoyant using gas vesicles (e.g., Anabaena, Aphanizomenon, Coelosphaerium, and Microcystis; see Chaps. 3 and 4) or flagella (e.g., Ceratium, Chlamydomonas, and Euglena). Species also exhibit different photosynthesis–irradiance and temperature optima. Some species, such as Asterionella formosa, have especially high photosynthetic efficiency at low light (Reynolds, 1984a), whereas others, including species of Oscillatoria, may vary their chlorophyll-a levels and employ accessory pigments that saturate at lower and spectrally altered light levels (Mur and Bejsdorf, 1978). Flagellates such as Dinobryon, Poterioochromonas, Cryptomonas, and Ceratium may ingest bacteria under low light conditions or under ice; these mixotrophic species switch between photosynthetic and bacterivorous metabolism (Bird and Kalff, 1987; Porter, 1988; Berninger et al., 1992; Caron et al., 1993). This strategy may also serve to supplement inorganic nutrients as well as organic-C needs.

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F. Benthic Algal Assemblages of Lakes Benthic algae—those attached to or closely associated with various substrata or bottom surfaces— occupy an enormous variety of microhabitats, including stones, macrophytes, sediments, sand grains, and logs, as well as a variety of artificial substrata. All divisions of algae have benthic representatives, although many more freshwater species in Chrysophyta (Chaps. 12–14), Xanthophyta (Chap. 11), Cryptophyta (Chap. 21), and Pyrrophyta (Chap. 20) are planktonic, whereas all species of freshwater Phaeophyta and Rhodophyta are benthic (but rare in lakes). Perhaps the most widely used term for benthic algae is periphyton, a word that has obscure etymology and debatable usage (Sládeckova, 1962; Round, 1981; Wetzel, 1983a; Aloi, 1990; Stevenson, 1996a). The term may have been coined in the 1920s by Russian limnologists (Sládecková, 1962) to refer to a collection of organisms (bacteria, fungi, protozoa) and detritus (Wetzel, 1983c). The word is analogous to the German aufwuchs, which means “to grow upon,” but some authors (e.g., Round, 1981) argue against using it because it is often used incorrectly to describe only the algal community and is imprecise with regard to habitat. Its use probably will not be abandoned, however. We recommend the use of the most precise and descriptive terminology for particular algal communities (e.g., epilithic diatom) that includes the nature of the substratum. Otherwise the term “benthic” is perhaps most suitable for general uses or when the substratum is not defined. Modes of algal attachment are diverse. Some are firmly attached or encrusting, such as Chamaesiphon, Coleochaete, and Cocconeis, making them resistant to wave scour or other disturbances, but susceptible to competition from canopy-forming morphologies, such as Stigeoclonium or Ulothrix (Hoagland and Peterson, 1990; Maltais and Vincent, 1997; Graham and Vinebrooke, 1998). Diatoms, such as Frustulia, Navicula, Nitzschia, Pinnularia, and Stauroneis, maintain contact with various surfaces (and glide among these microhabitats) by means of a slit in the wall that is termed a raphe (Chaps. 16–19). Other diatoms (e.g., Cymbella and Gomphonema) attach by means of gelatinous pads or stalks. Some filamentous cyanobacteria, such as Oscillatoria, Hapalosiphon, Lyngbya, and Microcoleus, exhibit motility by gliding, although the mechanisms are not clear (Castenholz, 1982). Filamentous green algae, such as Cladophora, Oedogonium (see Chap. 8), Spirogyra, and Zygnema (Chap. 9) produce holdfast-like rhizoids that enable them to remain attached in turbulent conditions. Other benthic forms are loosely associated with plants or sediments, and include filamentous species like Mougeotia, flagellates

such as Cryptomonas, Euglena, and Eudorina, and chains of cells such as Tabellaria (Hutchinson, 1975; Graham and Vinebrooke, 1998). The range of sizes in freshwater benthic algae exceeds that of planktonic forms. The smallest include unicells like Nannochloris or Synechococcus (0.8–2 µm diameter) to actual macrophytes, such as Chara, Cladophora, and Hydrodictyon, which range from a few centimeters to more than a meter in length. In terms of length, this range is more than 6 orders of magnitude, and in biovolume, perhaps as great as a factor of 1010. The tremendous variety of microhabitats, morphologies, sizes, and architectures found in benthic algal associations has led to the suggestion that these organisms may represent a more diverse community and greater trophic complexity than phytoplankton (Havens et al., 1996; Stevenson, 1996a). In shallow lakes, production of epiphytic algae often equals or exceeds that of phytoplankton per unit area (Wetzel, 1983a). Much of the literature on freshwater benthic algae has been reviewed in several important works, including Hutchinson (1975), Round (1981), Wetzel (1983b), and Stevenson et al. (1996).

1. Epiphytic Communities Epiphytic algae colonize submersed and emergent plants. These are the most widely studied group of benthic algae in lakes, perhaps because of their obvious accumulation in the littoral zone. Larger forms, such as Cladophora, Chara, Hydrodictyon, and Oedogonium, serve as additional substrata for microalgae. Epiphytic algae are important in macrophyte communities, because greater densities may cover and shade their hosts (Losee and Wetzel, 1983). Evidence includes negative relationships between epiphyte and macrophyte biomass (Sand-Jensen and Søndergaard, 1981; Cattaneo et al., 1998) and more rapid host senescence with greater epiphyte cover (Neely, 1994). There are often differences in species composition and biomass of epiphytic algae among different macrophyte host species. Prowse (1959) recognized that densities of three common epiphytes, Gomphonema gracile, Eunotia pectinalis, and Oedogonium sp., differed among three macrophyte species in one small pond. Many subsequent studies have reported differences in epiphyte biomass or species composition on different plant hosts (Gough and Woelkerling, 1976; Eminson and Moss, 1980; Lodge, 1986; Blindow, 1987; Douglas and Smol, 1995; Hawes and Schwarz, 1996), although not in all cases (Siver, 1977). In one shallow lake, epiphyte biomass on submersed macrophytes (Myriophyllum spicatum, Ceratophyllum demersum, and Najas marina) was 10–40 times greater than on floating-leaved plants (Trapa natans), but


species diversity was less (Cattaneo et al., 1998). In the Great Lakes, Cladophora glomerata is a host to many microalgal epiphytes, but the red alga Chroodactylon ornatum (as C. ramosum) is attached only to this species (Sheath and Morrison, 1982). The three-dimensional architecture of epiphyte assemblages also varies with the type of substratum. Colonization by epiphytic algae has been compared to terrestrial plant succession, which comprises temporal changes in vertical structure and diversity, an increase in the dominance of larger organisms, and possible facilitative effects of earlier colonizers (Hoagland et al., 1982). The reasons for differences in epiphytic communities among host plant species can be attributed to features of the macrophyte, such as leaf orientation, texture, or chemical properties. One survey revealed a correspondence between epiphytic communities and species of submersed macrophytes in less productive lakes, but little pattern was observed in eutrophic lakes where nutrient macrophyte interactions might be less (Eminson and Moss, 1980). However, plants may inhabit different zones within lakes that indirectly offer different ecological conditions for algal colonization. Nonetheless, direct evidence shows that living macrophytes translocate and release small quantities of P (about 3.5 µg P g–1 macrophyte shoot), which can be taken up by algal epiphytes, and that algal species differ in their ability to sequester released P (Moeller et al., 1988). A Synedra–Fragilaria complex obtained more than 50% of released P, but erect forms such as Mougeotia and Lyngbya, and stalked Gomphonema obtained most of their P from the surrounding water. Alkaline phosphatase activity of epiphytic algae on artificial (plastic) plants was shown to be greater than on natural plants under similar conditions (Burkholder and Wetzel, 1990). Epiphytic communities are important and complex components of lake food webs. In mesotrophic Lake Mann (WI), herbivorous snails consume and regulate benthic algal biomass, but pumpkinseed sunfish also exert predatory control on snails (Brönmark et al., 1992). Algal-feeding snails, benthic insects, and other invertebrates also have a qualitative impact on epiphytic communities, because many consumers graze more effectively on erect or filamentous forms, thereby shifting the community toward more compact or adherent forms like Cocconeis placentula and Coleochaete spp. (Kesler, 1981; Lodge, 1986; Marks and Lowe, 1993). In eutrophic lakes, snails similarly avoid larger colonies of epiphytic Gloeotrichia (Cattaneo, 1983; Brönmark et al., 1992). In contrast, the limpet Ferrissia fragilis grazes mainly understory species, such as Epithemia spp., Cocconeis placentula, and Achnanthidium minutissimum, and avoids upright forms such as

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Synedra ulna and Fragilaria vaucheriae (Blinn et al., 1989). Grazers of epiphytic algae may have indirect effects on host plants by reducing shade and enhancing plant growth (Lodge et al., 1994). Because of the difficulties of sampling epiphytic algae, artificial substrata, such as glass slides, plastic flagging, styrofoam floats, plexiglas plates, and plastic aquarium plants, are employed. Advantages include reduced variability, known surface area, standardized conditions, and no nutritional or chlorophyll artifacts from the host. An implicit assumption in their use is that the community sampled is representative of the “true” epiphyte community on aquatic plants, but studies suggest this is rarely true (Tippet, 1970; Robinson, 1983; Aloi, 1990; Cattaneo and Amireault, 1992). Glass microscope slides were among the first materials used (Sládecková, 1962), but differences in biomass, seasonal patterns, and community structure (different species proportions ) suggest this approach may provide unreliable estimates (Tippet, 1970). Evidence suggests that biomass of most epiphytic algae is overestimated when some types of artificial substrata are used, although green algae and cyanobacteria may be undersampled (Aloi, 1990; Cattaneo and Amireault, 1992). Synthetic materials are much simpler in surface texture and chemistry than natural substrata, and this is likely to affect the grazing, production, and community structure of epiphytes. Although artificial substrata should not be assumed to mimic natural habitats fully, they can be useful in comparative analyses or replicated studies on the effects of disturbances on benthic algal communities (Robinson, 1983; Aloi, 1990).

2. Epilithic Communities Epilithic algae colonize stones, boulders, and bedrock in lakes, and may dominate wave-swept littoral zones and oligotrophic lakes that have minimal macrophyte cover (Loeb et al., 1983). Species composition differs more strongly from phytoplankton than do epiphytic communities, but do exhibit pronounced seasonality in response to changes in nutrients, temperature, and other factors (Hutchinson, 1975; Lowe, 1996). Epilithic communities in turbulent littoral habitats are distinct from epiphytic communities within the same lake and comprise species known mainly from streams, such as Chamaesiphon spp., Gongrosira incrustans, Hildenbrandia rivularis, Tolypothrix distorta, or Heribaudiella fluviatilis (Kann, 1941, 1978; Auer et al., 1983). Vertical zonation is often observed. Many epilithic species are restricted to the upper littoral zone, whereas others occur in deeper waters where wave action is less severe (Hoagland and Peterson, 1990; Lowe, 1996). Bangia atropurpurea and Ulothrix zonata

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occur on rocks in the upper splash zone of the Laurentian Great Lakes, just above a mat of Phormidium sp., whereas a Cladophora glomerata zone is found in deeper water (Sheath and Cole, 1980). In Lake Traunsee (Austria), Kann (1959) documented that zonation patterns can differ among regions in a lake according to differences in slope. Epilithic algae form distinct communities in different regions of oligotrophic Lac à l’Eau Claire (subarctic Quebec), that are influenced by ice scour and wave action (Maltais and Vincent, 1997). A Gloeocapsa-dominated community colonized shallow areas, while a filamentous community dominated by Ulothrix zonata occurred in open, southfacing shores. In a study of 35 arctic ponds, freezing and other habitat factors were found to be of greater importance to benthic communities than were chemical variables (Douglas and Smol, 1995). Light or other factors may interact with the effects of wave action. An epilithic population of U. zonata exhibited a greater photosynthesis irradiance optimum (1200 µmol photons m–2 s–1) than C. glomerata (300 µmol photons m–2 s–1) isolated from the same region of Lake Huron (Auer et al., 1983). Temperature tolerances and nutrient requirements may further interact with irradiance optima, and affect local and regional distributions (Graham et al., 1985). Bangia, recent invader to the Great Lakes, has displaced Ulothrix zonata in many locations, perhaps because of its ability to produce more durable holdfasts (Lin and Blum, 1977) or an ability to resist epiphyte cover by sloughing its cell wall (Lowe et al., 1982). Interestingly, Kann (1959) similarly observed Bangia occupying rocks in the splash zones of the Traunsee, but co-occurring with another filamentous green alga, Mougeotia. In the calmer epilithic community of Montezuma Well (AZ), only 8 of the 83 benthic diatom taxa identified were restricted to this habitat (Czarnecki, 1979). The upper littoral zone can be a harsh habitat, where algal cells experience abrasive turbulence or desiccation during an annual cycle. Controls on epilithic production vary among different lake types. The epilithon of softwater, oligotrophic Lakes in the Experimental lakes Area (ELA; ON) was dominated by diatoms and filamentous green algae; production levels tended to be low but quite variable (Stockner and Armstrong, 1971; Schindler et al., 1973). Comparing nutrient-amended, pH-manipulated, and reference lakes in the ELA, Turner et al. (1994) concluded that epilithon production is unrelated to N or P supply (despite positive effects on phytoplankton), but is limited by dissolved inorganic carbon (DIC). Experiments using nutrient-diffusing substrata determined that DIC and P supply were the most important influences on biomass and species composition in another oligotrophic, softwater lake in

Pennsylvania (Fairchild et al., 1989). The epilithon of meso-oligotrophic Flathead Lake (Marks and Lowe, 1993) was limited principally by N and P, but individual species differed in their response to nutrient and shading manipulations. In sublittoral Lake Tahoe, epilithic populations of Calothrix, Tolypothrix, and Nostoc were strongly N-limited and exhibited N-fixation activity, in contrast to resident phytoplankton (Reuter et al., 1986). Recent increases in atmospheric N deposition may change the nutrient economy toward P limitation (Jassby et al., 1995). Grazing by benthic invertebrates is also important to epilithic algae. Snails (Planorbis contortus) and limpets (Ancylus fluviatilis) that inhabit the stony littoral zone of a small calcareous lake consumed substantial quantities of algae and detritus, and each preferentially grazed certain algal species (Calow, 1973a, b). Selectivity and more intense grazing activity by limpets exerted greater effects on algal community structure than did snails. Light availability and grazing pressure are factors that logically would be expected to be more important than nutrients for epilithic algae in eutrophic lakes, but grazing had minor impacts on biomass and seasonal patterns of epilithic algae in Crosmere, a eutrophic lake in the English Midlands, although caddisfly larvae may have contributed to spatial patchiness of Cladophora (Harrison and Hildrew, 1998). Studies on epilithic food webs that consisted of crayfish, invertebrates, and macrophytes in one Swedish lake found little top-down control by crayfish on epilithic algae (Nyström et al., 1996). The general importance of benthic algae in lake food webs is not well established, in part because of difficulties in their quantification. However, a study of arctic, temperate, and tropical lakes using stable isotopes suggests that previous efforts may have underestimated the importance of algae in benthic food webs (Hecky and Hesslein, 1995). Prior studies were based on net production of phytoplankton, benthic algae, and macrophytes, instead of ease of grazing, edibility, or nutritional quality; all of these qualities were predicted to be greatest in benthic algae. The importance of epilithic algae in some lakes may be increasing, due to the expansion of filamentous algae in many acidifying lakes (Stokes, 1986; Turner et al., 1995). In one neutral lake, tadpoles suppressed the growth of filamentous algae on tiles, and favored communities of adherent and encrusting species (Coleochaete scutata, Achnanidium minutissimum); grazers had no such effects on epilithic communities in acidified lakes (Graham and Vinebrooke, 1998). Transplants of epilithic algae (on natural quartz tiles) across lakes of varying acidity, coupled with grazer exclosures confirmed that pH was the key factor that regulated algal communities,


but grazer control was important in neutral lakes (Vinebrooke, 1996).

3. Epipelic and Epipsammic Communities Algal communities that colonize sediments (epipelic) and sand (epipsammic) are among the least studied benthic associations. This knowledge gap is surprising, considering the large area of many lakes not covered by stones or aquatic plants. It is difficult to separate epipelic from epipsammic habitats because these substrata are often mixed and, due to wave action, are especially unstable (Hickman, 1978; Kingston et al., 1983). Methods used to quantify living algal cells from lake sediments require considerable care and may still result in fairly high relative error (30% or greater), which may not be consistent among taxa (Eaton and Moss, 1966). Epipelic and epipsammic communities occur in all lake types, but their relative importance is greatest in small, shallow systems. Diatoms are the most common algal group in most systems (numbers and biomass), although cyanobacteria, cryptomonads, desmids, euglenoids, and colonial and filamentous green algae often are observed (Gruendling, 1971; Round, 1972; Hickman, 1978; Roberts and Boylen, 1988). Epipelic algae may live on or within the first few millimeters of sediment and so must be able to tolerate conditions of very low light or oxygen, making motility a distinct advantage. Nonmotile epipelic and epipsammic forms may be capable of heterotrophic growth, which allows them to tolerate dark conditions and utilize greater levels of dissolved nutrients. Despite these constraints, epipelic and epipsammic communities are often very diverse. In a study of arctic ponds, Moore (1974) identified 357 algal taxa from sediments, of which 226 were benthic diatoms. Very high standing crops were also measured, some exceeding 108 cells cm–2, which perhaps was influenced by long photoperiods during arctic summers. A diverse assemblage of 255 taxa was observed along a depth gradient in Lake Michigan, where benthic forms predominated in shallow and middepth communities (6–15 m) and settled, living planktonic forms were more common in deep (23–27 m), low light conditions (Stevenson and Stoermer, 1981). Patterns of seasonal succession seem to differ from those observed for lake phytoplankton. Over a threeyear period, Round (1972) observed that populations of epipelic algae in two shallow ponds had distinct and reproducible periods of abundance peaks, but their duration was longer (months rather than weeks) and varied markedly among species. Stauroneis anceps and Oscillatoria sp. persisted for several winter months, whereas other species like Navicula hungarica and N. cryptocephala maintained sizeable populations for

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more than nine months. In acidified Woods Lake (NY), several epipelic species were abundant from May through October, such as Navicula tenuicephala at 1 m and Hapalosiphon pumilis at 7 m (Roberts and Boylen, 1988). Following liming, several acidophilic species were replaced by other taxa, but diatom species still dominated the epipelon (Roberts and Boylen, 1989). Round (1972) suggested that epipelic habitats provide greater habitat diversity than the water column, enabling more species to coexist.

4. Benthic Macroalgae Several species of macroalgae, those that form macroscopic or plantlike morphologies, can be important in benthic lake communities. Some investigators (e.g., Hutchinson, 1975) restrict this category to taxa that are attached by means of rhizoids, which include only charophytes (Chara, Lamprothamnium, Nitella, Tolypella, and Nitellopsis). From an ecological perspective it makes sense to include other algae, such as Cladophora, Enteromorpha, and Nostoc, which also may function as structuring elements within the benthic zone and as hosts for epiphytic microalgae. Astounding colonies of Nostoc greater than 30 cm in diameter (“mare’s eggs”) have been observed (Dodds et al., 1995), and tubes of Enteromorpha may exceed 1 m in length (Wehr, unpublished). Chara may form large underwater meadows in the littoral zone of calcareous, nutrient-poor lakes and frequently is encrusted with marl (CaCO3). In clear, oligotrophic lakes, charophytes colonize lake bottoms down to depths of 30 m or more. In nonturbid systems, wave action is suggested to be the primary limiting factor, rather than light (Schwarz and Hawes, 1997). Various species of Chara and Nitella differ in their depth distributions, presumably as a function of individual light requirements (Wood, 1950). The lower depth boundary for Nitella meadows in more productive lakes is influenced by total irradiance and the supply of red light; in deep, clear lakes, their distribution is limited by the availability of blue light (Stross et al., 1995). Reduced charophyte abundance in lakes with eutrophication has been attributed to excessive or even toxic levels of P (Hutchinson, 1975; Phillips et al., 1978). However, the macrophyte community of eutrophic Lake Luknajno (Poland) is dominated (90% of total dry mass) by seven charophyte species (codominants: C. aculeolata and C. tomentosa), where the mean biomass is greater than 1 kg dry mass m–2 (Krolikowska, 1997). Nitella hookeri was found to grow best at very high P concentrations, about 20 mg L–1 (Starling et al., 1974). Reduced charophyte abundance in eutrophic lakes may be the result of light limitation, given that some species do grow at greater

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depths but only in clear lakes (Blindow, 1992; Steinman et al., 1997). Recovery of a previously eutrophic lake resulted in a 15-fold increase in the benthic cover of charophytes in just a two-year period (Meijer and Hosper, 1997). The epiphytic algae that colonize charophytes can be dense and may differ in composition among specific charophyte taxa. In one deep oligotrophic lake, epiphytes on charophytes differed strongly with depth; filamentous green algae and stalked diatoms predominated in shallow water (≤ 5 m), while adnate forms, such as Cocconeis placentula, predominated at greater depths (Hawes and Schwarz, 1996). Littoral epiphyte densities in one eutrophic Swedish lake were greater on C. tomentosa, C. globularis, and Nitellopsis obtusa than on Potamogeton pectinatus (Blindow, 1987). Nitellopsis harbored the highest total densities, but differences in epiphyte species composition were most pronounced between the two Chara species, probably a result of differences in marl encrustation. Benthic invertebrates and zebra mussel larvae also colonize species of Chara more readily than other macrophyte hosts (Lewandowski and Ozimek, 1997; Van Den Berg et al., 1997).

III. LOTIC ENVIRONMENTS Running water ecosystems, from headwater streams to the largest rivers, are termed lotic environments and differ from standing waters in several important respects. Lotic systems are turbulent and generally well mixed; hence stratification is uncommon except for brief periods in slowly flowing, lowland rivers (Hynes, 1970). Waters with greater current velocity tend to have abundant dissolved oxygen; even large rivers with substantial current speed, such as the Ohio, are generally well mixed (Thorp et al., 1994). In shallow rivers or littoral regions of large rivers, organisms experience less temperature fluctuation than organisms in lakes of comparable depths. Many organisms adopt a benthic habit and are attached to a variety of substrata, the sizes of which are a function of current velocity and discharge. Rapid rivers are typified by large stones and boulders, whereas the bottom material in more slowly flowing systems consists of sand and silt. Rivers are intimately connected with the surrounding watershed, which is responsible for regulating water and chemical balances within the aquatic environment (Likens et al., 1977). Rivers are open systems that transport materials and energy from one part of the watershed to downstream areas. Descriptions of the physical and geological attributes of river systems are

given in the treatises by Leopold et al., (1964) and Morisawa (1968), and in syntheses by Hynes (1970) and Beaumont (1975). Several important reviews discuss ecological feature of rivers, in particular, those by Hynes (1970), Whitton (1975, 1984), Lock and Williams (1981), Allan (1995), and Petts and Callow (1996).

A. Major Rivers of North America The Mississippi–Missouri River system is longest in the world (6970 km), has the third largest drainage area (3270 ⫻ 103 km2), and is the sixth largest in terms of discharge (18,390 m3 s–1; Hynes, 1970; Milliman and Meade, 1983). At least 10 other rivers in North America exceed 1000 km in length. Most of them have been altered substantially by navigation or hydroelectric dams, channelization, wetland removal, and pollutants (Sparks, 1995; Wehr and Descy, 1998). The Missouri River (3770 km) has six major impoundments over 1230 km (33%) of its length; another 1200 km (32%) have been channelized. Only 35% (1330 km) of all river sections remain free-flowing, although discharge still is influenced by reservoir conditions upriver (Hesse et al., 1989). No major impoundments are located on the Ohio or Mississippi Rivers, but navigation dams and channelization have been built throughout their lengths to facilitate ship traffic.

B. Geomorphology of Rivers Streams and rivers are part of a network of connected, increasing tributaries that have hydrological features that vary in predictable ways. Many features, such as discharge, substratum size, stream width, and depth, affect the species composition and productivity of lotic algae and their consumers. For example, sizes of substrata available for colonization vary with differences in current velocity (Table II). Physical features of rivers may be described in a system of stream orders, which assign increasing numbers to streams when two tributaries of equal order join. The most widely adopted system (Strahler, 1957) defines a headwater stream or spring with no (permanent) tributaries as first order and the junction of two such streams a second order (Fig. 10). A second-order stream increases only when it is joined by another second-order stream and so on. Larger order streams are wider and longer segments, drain larger areas, and have a more gradual slope than smaller streams. The network of these stream segments forms a treelike structure that is used in hydrological models to predict average discharge, behavior of flood events, and quantity of suspended matter (Beaumont, 1975). A simpler scheme divides rivers into three zones


TABLE II Relationship between Minimum Current Velocity and Mean Size of Stone Substrata That Can Be Moved along a Streambed (based on Hynes, 1970; Reid and Wood 1976) Current velocity (m s-1)

Particle size (cm)

Stream bed characteristics

Habitat

3.0 2.0 1.0 0.8 0.5 0.2 0.1

180 80 20 10 5 1 0.2

Bedrock Boulders Large stones Stones and gravel Gravel & coarse sand Sand Silt

Torrential Rapids Riffles Riffles Run Run Pool

(Schumm, 1977). Zone 1 (erosional) includes headwater and small order streams that function as the source of water and sediments for downstream reaches. In Zone 2 (conveyance), water and sediments are transported along the mainstem with no net gain or loss of materials. Zone 3 (deposition) includes lower reaches that receive sediments from upriver, including river deltas and estuaries. Some studies suggest that benthic algal communities differ similarly along these zones (Rott and Pfister, 1988).

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Although many aspects of drainage basins follow an ordered structure within an individual watershed (catchment), physical conditions and biological communities in streams of equal order at two locations may be quite different. Current velocity, stream width, and substrata in a second- or third-order mountain stream in New England differ from a similar order cool-desert stream in the western Great Basin (Minshall, 1978; Minshall et al., 1983; Benke et al., 1988). Landscape factors such as climate, terrestrial vegetation, and external nutrient supplies, may exert a substantial effect on the biological properties of rivers. For example, rivers that flow through a limestone region will possess greater concentrations of certain ions (Ca, Mg) than would be found in streams flowing through a region composed of granite or basalt. River basins also differ in the amount of interaction between river channels and their watershed, which is mainly influenced by geological features of the region and amount of interface between groundwater and surface waters (Dahm et al., 1998). Some rivers are geologically constricted, such as the Hudson River and large sections of the Ohio and Columbia. Floodplain rivers have substantial watershed interaction, such as the lower Mississippi. The floodplain includes portions of the watershed (tributaries, adjacent wetlands, flood-

FIGURE 10 Structure of tributaries in a watershed, indicating numbering of stream orders.

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plain lakes, riparian zones) that are seasonally inundated during periods of high flow. Meandering rivers and those with more islands have greater littoral and floodplain interaction, and more complex and varied current regimes than rivers with less sinuous courses (Fig. 11A and B). This increased habitat complexity may offer refuge to larger, more slowly growing algae. Small “islands” also form in lowland rivers from large stands of submersed angiosperms (Butcher, 1933; Holmes and Whitton, 1977). Within a given reach, there are alternating regions of erosion and deposition. Bottom materials are eroded from river margins and deposited downstream in point bars. Differences in current velocity and substratum also create regular, alternating patterns of riffles and pools (Fig. 11C and D). Riffles are shallow sections with larger substrata and greater current velocity, and are spaced at fairly regular intervals, about five to

seven stream-widths apart (Hynes, 1970). These regions have greater turbulence and concentrations of dissolved gases, which may provide a physiologically richer habitat for benthic algae. Deeper pools form downstream of riffles, where organisms experience reduced current velocity, possibly depleted dissolved gases, and perhaps light limitation. Bottom materials consist of smaller particles, primarily sand and silt, and current velocity is reduced during average flow periods. However, during flooding, pools lack stable substrata, which makes them susceptible to scouring and erosion.

C. The River Continuum and Other Models For many decades ecologists lacked broad conceptual models for describing and testing patterns in the structure and function of river communities. Early concepts, some of which were developed by algal biolo-

FIGURE 11 Examples of river features: A, B, meanders in lowland rivers; C, riffle (arrows) and pool sequences in a larger (Firehole River, WY) and smaller (Little Beaverkill, NY) streams. All photos by J. D. Wehr.


gists, borrowed ideas from lake systems, such as the concept of oligotrophic–eutrophic gradients and climax communities (Blum, 1956; Hynes, 1970). These ideas do not adapt easily to lotic ecosystems, and a recent synthesis of nutrient and chlorophyll data from more than 200 temperate streams suggest that nutrient–algal biomass relationships are weaker than in lakes, perhaps because of the effects of nonalgal turbidity in running waters (Dodds et al., 1998). Some ecologists recognize different zones along the length of a river that possess different physical conditions and habitats for riverine organisms (Hawkes, 1975). A river’s features, however, do not fall into discrete zones, but rather vary continuously along a river’s course. The combination of hydrological principles with changes in biological and chemical processes along river gradients led to the development of the river continuum concept (RCC; Fig. 12; Vannote et al., 1980. The RCC characterizes lotic ecosystems as a network of streams with a continuum of longitudinally

FIGURE 12 Diagrammatic representation of the river continuum concept, predicting changes in P/R ratios, consumer groups, and sources of organic matter along the length of a river. Reprinted from R. L. Vannote et al., Canadian Journal of Fisheries and Aquatic Science 55:668–681, 1980, with permission from NRC Research Press.

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linked environmental (e.g., width, depth, flow) and resource (nutrients, light) gradients. Biological communities respond to longitudinal changes in geomorphology, water chemistry, and energy sources in several ways. In addition, organisms, by their own activities, influence conditions and communities downstream (Fig. 12). This is evidenced by changes in the types of invertebrate consumers that are localized in rivers of different sizes. The model was based on data largely from smaller, temperate forest streams, and broadened into a theory for rivers as a whole. Many other ideas and studies concerning nutrient spiraling, benthic invertebrates, fish production, and the influence of dams have emerged from the general framework laid out in the RCC (Newbold et al., 1981; Welcomme et al., 1989; Minshall et al., 1983; Ward and Stanford, 1983; Thorp and Delong, 1994). The role of algae and other primary producers was also considered in the RCC. The metabolism of first- to third-order streams was viewed to be largely dependent on external or allochthonous sources of terrestrial carbon. Hence, consumers in smaller streams were mainly shredders and collectors of coarse particulate matter. Algae were viewed as a minor component of food webs in headwater communities because of light limitation due to heavy riparian shading and subsidies of terrestrial organic matter. The overall effect is a net heterotrophic community, in which system-level respiration (R) exceeds in situ primary production (P; P : R < 1). Long-term studies confirm that the mass of allochthonous litter (leaves, wood) forms the dominant organic carbon source for small streams (Benke et al., 1988; Findlay et al., 1997), although food web studies suggest that benthic algae still may be an important or even dominant food source for some benthic animals, perhaps because of greater assimilation efficiency with algae than detritus (Fuller et al., 1986; Mayer and Likens, 1987). Algal production is predicted to increase in mid-sized (fourth to sixth order) rivers, in response to greater sunlight and a reduction in subsidies of allochthonous organic matter, resulting in P : R ≥ 1. In such systems, consumers are likely to be dominated by grazers of (mainly benthic) algal material and collectors of finer organic matter transported downstream from upper reaches. The model suggests that algal (plus macrophyte) primary production still may not come to dominate river metabolism in mid-sized rivers if the reach receives substantial supplies of allochthonous organic matter from smaller tributaries, which would increase system-level respiration and reduce light penetration. In larger rivers (greater than sixth order), the RCC predicts that even though the river basin is open to full sun, higher levels of fine particulate matter from upstream, greater depth, and resuspended sediments

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limit algal production. Although phytoplankton usually dominate the algal community in large rivers, these systems are predicted to be too turbid and too deep to support high levels of algal production. This forecast appears to be the case in the Hudson River, where depth and tidally driven mixing create a lightlimited environment for phytoplankton (Cole et al., 1992), but studies on other large rivers have found that substantial phytoplankton populations can develop (e.g., Descy and Gosselain, 1994; Lair and ReyesMarchant, 1997; Wehr and Descy, 1998). Ideas from the RCC have stimulated a great increase in research on and discussion of lotic ecosystems, but several exceptions have been raised (Wetzel, 1975; Lock and Williams, 1981; Benke et al., 1988). Rivers in different biomes have different climate, lithology, and amounts of riparian vegetation, which may shift the relative importance of autochthonous versus allochthonous production for river metabolism (Minshall, 1978; Minshall et al., 1983; Wetzel and Ward, 1992). Not all rivers (small or mid-sized) in North America are shaded heavily (e.g., high altitudes, deserts, agricultural areas) nor do they receive large subsidies of terrestrial organic matter from their watershed. In more open rivers, algal production may be substantial. A metanalysis of studies on 30 streams from five biomes showed that gross primary production (GPP) levels varied by more than 4 orders of magnitude; the greatest rates were in desert areas (Lamberti and Steinman, 1997). Variations in GPP were related to variables such as watershed area, inorganic P, temperature, discharge, and canopy cover, but not to stream order or latitude, further suggesting that variation among rivers may be greater than predicted by the RCC. River systems are also very patchy, from differences in sunlight within a local reach to watershed-level differences in sources of carbon or inorganic nutrients (Pringle et al., 1988). Subsequent syntheses have emphasized that lotic ecosystems also can be regulated by biotic interactions, such as predator–prey dynamics, competition, and food-chain length, rather than solely controlled by physical (e.g., hydraulic) factors (Power et al., 1988). In turn, physical factors, such as the frequency of floods, have significant effects on food web interactions at several levels, including algal biomass (Wootton et al., 1996). Other syntheses have focused on large rivers (greather than sixth order). Sedell et al. (1989) and Junk et al. (1989) pointed out that many large rivers receive materials, energy, and organisms from the adjacent floodplain, often in greater quantity than may have been transported from upriver. The influence of algal production was regarded as minimal because large rivers were generally turbid and light-limited. In

their flood-pulse model, Junk et al. (1989) suggested that large river metabolism may be driven more by batch processes—the pulses of resources from flood events—than by the continuous processes emphasized in the RCC. This process is undoubtedly true for rivers such as the Amazon and the lower Mississippi. However, not all large rivers occur in extensive floodplains. River basins also flow through constricted channels that have minimal floodplain influence. Thorp and Delong (1994) proposed a river productivity model that suggests that local autochthonous production may have been underestimated in large, constricted-channel rivers, which have firm substrata and less turbid water. In such systems, for example, a 350 km stretch of the Ohio River, substantial phytoplankton biomass and production have been observed, and during some seasons may be the principal carbon source for planktonic consumers, such as small-bodied cladocerans and rotifers (Thorp et al., 1994; Wehr and Thorp, 1997). Also, many, if not most large rivers worldwide have been altered substantially due to industrialization, which has had profound influence on ecological conditions. Flow regulation and nutrient inputs favor greater phytoplankton productivity in large rivers, such that P : R ratios may exceed 1.0, at least during the spring and the summer (Admiraal et al., 1994; Descy and Gosselain, 1994; Wehr and Descy, 1998).

D. Benthic Algal Communities of Rivers Most studies of river algae concern benthic species. The necessity to remain in a stable position while water flows downstream is an important selective force for all benthic organisms in lotic environments. Earlier reviews pointed out the paucity of studies on lotic algae relative to those on lakes (Blum, 1956; Hynes, 1970; Whitton, 1975), yet the pace of work on benthic algae in streams and rivers has increased substantially. Much of what has been learned in the interim has been summarized in several reviews (Lock et al., 1984; Reynolds, 1996; Biggs, 1996; Steinman, 1996), which reveal an evolution from mainly descriptive studies to structural and functional analyses of benthic algae and their importance in lotic food webs.

1. Benthic Algal Diversity, Composition, and Biogeography In streams, diatoms often comprise the dominant algal group in terms of species number and biomass (Blum, 1956; Douglas, 1958; Round, 1981; Kawecka, 1981). Their diversity in species and growth form (upright frustules, short- and long-stalked, rosettes, tube-dwelling, filamentous, mucilaginous matrix, and


prostrate cells) enables them to colonize a variety of microhabitats. In more slowly flowing or less floodprone systems, filamentous species like Melosira varians and upright, stalked forms such as Gomphoneis herculeana may predominate, along with filamentous nondiatom species (Stevenson, 1996a; Biggs, 1996). In very rapid water, firmly attached diatoms, such as Cocconeis placentula, Achnanthidium minutissimum, and Hannaea arcus, may occur with encrusting nondiatoms, such as Hildenbrandia rivularis, Gongrosira spp., and Chamaesiphon spp., and corticated forms like Lemanea spp. (Fritsch, 1929; Whitton, 1975; Kawecka, 1980; Kann, 1978). One of the fascinations of studying benthic algae in rivers is that many species (although not the majority) are macroscopic and recognizable in the field (Holmes and Whitton, 1977; Kann, 1978; Entwisle, 1989; Sheath and Cole, 1992). These include cyanobacteria, and green and red algae, as well as chrysophytes, xanthophytes, and brown algae (Whitton, 1975; Sheath and Cole, 1992). Morphologies are diverse and include encrusting, turflike, filamentous, cartilaginous, mucilaginous, tubular, and bladelike thalli (Fig. 13). In some instances, diatoms, including species of Eunotia,

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Fragilaria, and Melosira, and stalked forms, such as Didymosphenia geminata and Cymbella spp., may be recognized by their gross appearance, but are identified only using microscopy (Holmes and Whitton, 1981; Steinman and Sheath, 1984; Sheath and Cole, 1992). Some benthic stream algae have limited distributions. Prasiola mexicana, a seaweed-like green alga (Fig. 13B), thus far has been recorded only in streams located in the arctic tundra and western coniferous biomes, whereas the encrusting brown alga Heribaudiella fluviatilis has been confirmed in North America from streams in western Canada and the United States only (Sheath and Cole, 1992; Wehr and Stein, 1985). Some taxa have been regarded to be limited in geography or habitat, only to be found later in other locations. Thorea violacea is a large red alga that was thought to be restricted to streams in warmer biomes or in temperate areas only during the summer (Smith, 1950; Sheath and Hambrook, 1990, Chap. 5), but it was discovered growing profusely in the upper Hudson River in cool (15°C), rapidly flowing water (Pueschel et al., 1995). Few long-term studies of benthic stream macroalgae exist that may help to explain the dispersal patterns of these organisms. In one of the few cases of

FIGURE 13 Examples of different forms of macroalgae from rivers: A, Zygnema sp., simple, flexible filaments (arrow = direction of flow); B, Prasiola mexicana, a flat, seaweed-like thallus; C, Lemanea fluviatilis, corticated tubes; D, Nostoc verrucosum, mucilaginous colonies. Photos A and D by J. D. Wehr; photos B and C by R. G. Sheath.

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a long-term (more than 40 years) database, Holmes and Whitton (1977, 1981) were able to characterize species as previously overlooked, currently increasing, or recently extirpated from the River Tees (UK). No such studies are known from North America. Ecological endemism in some species of river algae may be caused by the spread of marine taxa into estuarine environments, followed by adaptation to lower salinity. Such may be the case with genera such as Audouinella, Hildenbrandia, Prasiola, and Enteromorpha: genera that each has marine and freshwater species and are found in rivers but only rarely in lakes (Flint, 1955; Whitton, 1975; Sheath and Cole, 1980; Sheath et al., 1985; Hamilton and Edlund, 1994). However, other species appear to be human-accelerated invaders from other freshwater systems. Populations of the red alga Bangia, which invaded North American freshwaters in the 1960s through the St. Lawrence River, are not closely related to any marine populations (based on rbc-L, RuBisCo spacer, and 18s rDNA sequences) and show strong affinities with freshwater European populations (Müller et al., 1998). These data suggest that some vector, such as a ship’s ballast water, enabled invasion, rather than gradual spread and adaptation from a marine environment. Biogeographic data suggest that green algae are the most common group of stream macroalgae across all biomes in North America. Based on 1000 stream segments studied, the lowest species diversity was found in streams in arctic tundra and the greatest in boreal forests (Sheath and Cole, 1992). There was no increase in macroalgal species diversity from the arctic to the tropics (contrary to marine species), which may be the result of periodic flooding common in streams across all regions. Arctic streams tend to have more species of macroalgal cyanobacteria, whereas tropical streams have more species of Rhodophyta (Sheath and Cole, 1992; Sheath et al., 1996; Sheath and Müller, 1997).

2. Factors That Regulate Benthic Algae in Rivers Several reviews (Blum, 1956; Hynes, 1970; Whitton, 1975; Biggs, 1996; Stevenson, 1997) discuss many ecological factors, including current velocity, substratum, geology, nutrient conditions, grazers, temperature, pollutants, and light availability, and their effects on benthic algae in rivers. Factors often interact to affect algal growth and survival, and multivariate analyses have been useful in determining the key environmental factors that affect species composition (e.g., Hufford and Collins, 1976; Wehr, 1981; Lowe and Pan, 1996). The following discussion provides a brief overview of current perspectives on the influence of habitat variables on benthic stream algae and, in particular, efforts to integrate structural aspects (species

composition, diversity, architecture) with functional studies (production, food web dynamics, nutrient cycling). In all river systems, there are proximate variables (those we measure) that affect organisms, such as light availability or nutrient supply, and larger scale factors such as climate, watershed features, and land use practices, that drive proximate variables (Stevenson, 1997). Biggs (1996) and Biggs et al. (1998) proposed a conceptual, disturbance–resource supply–grazer model that categorizes controls on benthic algal production in streams in terms of two processes: factors that affect (1) biomass accrual and (2) biomass loss. The model recognizes how ecosystem-level changes, such as floodplain modifications, can affect proximate controls, such as current velocity. An energetic balance sheet is constructed for each side of the ledger that can be used to predict to algal production and species composition (Fig. 14). In this scheme, biomass accrual increases as a function of resource supply, whereas biomass losses are a function of disturbance and grazing. In rivers with infrequent, low-intensity floods (= disturbance) and modest grazing intensity, the model predicts that biomass accrual dominates to a level dictated by resources. Under low resource supplies, growth continues at lower rates, favoring an adnate and turflike community dominated by filamentous cyanobacteria (e.g., Schizothrix, Phormidium, and Tolypothrix; Chap. 4), red algae (e.g., Audouinella; Chapt. 5), and many benthic diatoms (e.g., Epithemia and Navicula; Chaps. 17–19). Under similar hydraulic and grazing conditions but greater nutrient supply, a greater biomass of filamentous taxa, such as Cladophora, Ulothrix, or Melosira, is expected. For each combination of plus and minus factors, specific predictions can be made about the most important variables that drive algal production and community composition. Empirical evidence supports these predictions. In the Colorado River, where nutrients and sunlight are generally nonlimiting, release of water from Glen Canyon Dam (greater disturbance) decreased the biomass and relative importance of Cladophora glomerata, and reduced total benthic primary productivity (Blinn et al., 1998). The physiognomy of epiphytic algae changed from upright assemblage (Diatoma vulgare, Rhoicosphenia curvata) to more closely adherent forms (Achnanthes spp. and Cocconeis pediculus; Hardwick et al., 1992). Herbivorous invertebrates have been widely shown to reduce the density and biomass of benthic algae, but losses due to grazer activity come from scouring as well as consumption (Allan, 1995). Furthermore, ingestion and assimilation of benthic algae vary widely (30–70%), depending on the species of both the alga and the consumer (Lamberti et al.,


FIGURE 14 Conceptual model of ultimate (landscape) and proximate (stream) variables that control benthic algal communities, their consumers, and interactions in streams (solid lines = strong effects; dashed lines = weaker effects; double arrows indicate feedback interactions. Reproduced with permission from Biggs, B. J., in Stevenson, R. J. et al., Eds., Algal Ecology: Freshwater Benthic Ecosystems. Copyright © 1996 by Academic Press.

1989; Pandian and Marian, 1986). Resultant biomass and compositional changes, in turn, have effects on benthic food webs, because many invertebrates find an adherent community less easily grazed (Colletti et al., 1987). The Riviere de L’Achigan (Quebec) is an unshaded stony stream interspersed with a chain of small lakes that alter flow conditions immediately downstream (lower disturbance; Cattaneo, 1996). Biomass and species composition of benthic algae on gravel varied inversely with distance from lake outlets, yet this impoundment effect was unimportant for algae that colonize boulders. Only boulders supported communities of large filamentous and plumose forms such as Draparnaldia (Chlorophyceae), Stigonema (Cyanobacteria), and Batrachospermum (Rhodophyta). An experimental study in Big Sulphur Creek (CA) demonstrated the interactive effects of shading (resource supply) and invertebrate grazing on epilithic algal communities (Feminella et al., 1989). Algal biomass in low-grazer conditions declined by 75% with greater (15–95%) canopy cover, but was unaffected by light availability at normal grazer densities. The food web— resource supply interaction in Big Sulphur Creek is complicated by the fact that densities of trichopteran grazers (e.g., Gumaga nigricula) declined significantly in shaded conditions, while other species were unaffected. Abundances of algal grazers are often driven by food supply, but differences in irradiance also affect the

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chemical composition of algal assemblages, including changes in protein, total lipids, and fatty acid composition (Steinman et al., 1988). In general, cyanobacteria have different complements of fatty acids, including greater oleic, linoleic, and linolenic acids than diatoms (McIntire et al., 1969). Reductions in light availability to Kingsley Creek (NY) caused significant reductions in epilithic algal biomass and densities of a herbivorous mayfly (Baetis tricaudatus), but did not affect densities of a filter-feeding blackfly larva (Simulium spp.), which mainly consumed detritus (Fuller et al., 1986). However, most field studies on grazing are conducted under summer base-flow conditions when disturbance is low and resource factors are less important (Feminella and Hawkins, 1995). Current velocity is of great importance to benthic algae. Species that colonize areas of rapid current velocity are firmly attached to substrata using rhizoidal or holdfast-like structures (Israelson, 1949; Whitton, 1975). Greater current velocity provides a continuous replenishment of nutrients from upstream and a steeper diffusion gradient near the cell surface (Whitford, 1960; Horner et al., 1990). Species restricted to riffles with strong current velocity (e.g., Hydrurus foetidus and Lemanea fluviatilis) may have a higher metabolic demand for nutrients, which must be met by a greater physical supply. Studies suggest that there is no simple positive relationship between current velocity and algal metabolism or growth rate; greater current speed may also decrease algal biomass through scouring (Borchardt et al., 1994; Stevenson, 1996b). Few correlative studies are able to attribute differences in algal communities to current velocity effects because many variables change along a river’s course. A more clearcut study of parallel streams draining the same reservoir found that the regulated stream had greater diatom cover and large populations of Prasiola fluviatilis, while the stream that lacked flood control had populations of Hydrurus foetidus and Ulothrix zonata, and greater species number, but a lower biomass of diatoms (Kawecka, 1990). Current velocity affects the growth form of individual algae, such as in Cladophora, which develops compact tufts with narrow branching in greater current velocity and develops plumose forms with widely branched filaments in calmer flow (Whitton, 1975; Dodds and Gudder, 1992, Chap. 8). The architecture of algal communities affects their response to current speed. A mucilaginous, stalked diatom community (e.g., Gomphoneis herculeana and Navicula avenacea) exhibited increased production with greater near-bed current velocity, whereas a community consisting of long filamentous algae (Oedogonium sp. and Phormidium sp.) experienced decreaseed biomass accrual (Biggs

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et al., 1998). The morphology of Nostoc parmelioides (a filamentous colonial cyanobacterium) in stony streams is altered from spherical to ear-shaped colonies by the presence of an endosymbiotic midge larva (Ward et al., 1985), and only ear-shaped colonies exhibit greater photosynthesis and N2-fixation rates with greater current velocity (Dodds, 1989). Such studies that aim to link metabolic patterns with differences in community composition or form are needed to improve our understanding of the factors that regulate lotic communities. Several studies of benthic stream algae also have considered the influence of substratum. Several decades ago, Douglas (1958) recognized that different sizes of stones supported different densities and species of epilithic algae, which was likely the result of differences in their susceptibility to flood disturbance. A comparison of epilithic communities on different substratum sizes and similar in nutrient conditions found that most of the variation in algal biomass was explained by total-P and seston levels, but size of the substratum also exerted a significant effect (Cattaneo et al., 1997). Stones that are disturbed or scoured during floods may have algal crusts or propagules that still remain on their surfaces (Power and Stewart, 1987). The degree to which algae can recolonize disturbed substrata in a stream is a function of (1) their resilience, through immigration and greater growth rates, or (2) their resistance, as influenced by the species’ morphology and community physiognomy (Peterson, 1996). Benthic algal biomass and species composition are also influenced by substratum–current interactions. Diatom immigration onto bare substrata may increase with either reduced current speed or greater surface complexity (Stevenson, 1983). Substrata conditioned with a simulated mucilage (agar coating) were colonized twice as rapidly as clean surfaces, but responses were speciesspecific; some increased (Navicula gregaria and Synedra ulna), while others declined (Achnanthidium minutissima and Diatoma vulgare) or were unaffected (Diatoma tenue and Gomphonema olivaceum). A matrix of organic matter and bacteria probably facilitates colonization by most benthic algae in rivers (Karlström, 1978; Korte and Blinn, 1983; Sheldon and Wellnitz, 1998). A comparison of benthic diatom assemblages that colonize natural rocks, sterilized rocks, and clay tiles in Fleming Creek (MI) found greater total densities and species diversity on natural rocks (Tuchman and Stevenson, 1980). Otherwise identical natural substrata constructed from three rock types (basalt, sandstone, and limestone) that occur in Oak Creek (AZ) were compared for their effects on diatom colonization of riffles (Blinn et al., 1980). Densities on sandstone substrata were 60–80% greater than on basalt and limestone, but species composition and

diversity were similar. In Mack Creek (OR), greater biovolume (but not chlorophyll-a) and diversity of benthic algae were observed colonizing pieces of wood than clay tiles (Sabater et al., 1998). Certain taxa, such as Cymbella minuta, Hannaea arcus, and Zygnema sp., were more abundant on wood, while some closely adherent forms, such as A. minutissima and A. lanceolata, were more abundant on clay tiles. The physical structure of the stream bed also influences benthic communities. Experimental manipulations of rocks and bricks with different density and surface texture resulted in significant changes in the diversity and the abundance of benthic invertebrates and epilithic algae in the Steavenson River, Australia (Downes et al., 1998). Greater densities of Audouinella hermannii were observed on substrata without large crevices, but total biomass was greatest on surfaces that were roughened, independent of the presence of crevices.

E. Phytoplankton Communities of Rivers Although benthic algae typically dominate rocky streams and smaller rivers, phytoplankton become important in larger rivers and lowland streams (Rosemarin, 1975; Reynolds and Descy, 1996). A long history of studies on river phytoplankton dates back to at least the 1890s, when Zacharias (1898) coined the term “potamoplankton,” to refer to the suspended organisms in flowing waters. In North America, phytoplankton have been studied since the early years of limnological research in rivers including the Illinois (Kofoid, 1903, 1908), Mississippi (Reinhard, 1931), Ohio (Eddy, 1934), San Joaquin (Allen, 1921), and Sacramento (Greenberg, 1964). Much of the early research focused on whether a true phytoplankton community (populations that survive and reproduce within rivers) actually existed, as opposed to dislodged benthic forms or plankton washed in from lakes within the watershed. Indeed, plankton in most rivers consists of all three components in varying proportions (Reynolds, 1988), but in a single river sample, it is difficult to distinguish these sources, although certain algal taxa may be considered typical of each. Many benthic algae become suspended. These meroplanktonic forms can be washed out from sediments, plants, or other substrata. A metanalysis of 67 studies suggests that about 50% of suspended algal taxa in rivers are either benthic or meroplanktonic (Rojo et al., 1994). Among diatoms, most raphe-bearing species are likely nonplanktonic, but it is difficult to distinguish true potamoplankton from those that originated from lakes (Reynolds, 1988). One distinction may be found in species’ responses to flow regime and other physical factors, as has been attempted for phyto-


plankton species in the Ohio River (Peterson and Stevenson, 1989; Wehr and Thorp, 1997). The abundances of most species were negatively related to discharge, but tributary rivers were found to have little effect on species composition. However, during low flow (summer to early autumn), there were significantly greater amounts of colonial cyanobacteria (e.g., Aphanocapsa saxicola) and certain diatoms (e.g., Stephanocyclus [= Cyclotella] meneghiniana) in the river downstream of these tributaries, suggesting that some populations may have originated from outside the main river. The fact that chlorophyll-a concentrations in large rivers usually vary inversely with discharge (Schmidt, 1994) strongly suggests that potamoplankton is not derived primarily from benthic habitats. This pattern was observed for individual species in the River Thames, although smaller cells were less affected or even increased with greater discharge (Ruse and Love, 1997). In general, smaller forms appear to be more successful members of the potamoplankton, perhaps due to greater growth rates and surface area to volume ratios (Reynolds, 1988; Rae and Vincent, 1998). Diatoms are clearly the most diverse and abundant group, with Cyclotella and smaller species of Stephanodiscus especially common in larger rivers worldwide (Chap. 15). Algal flagellates rarely achieve large numbers in river plankton, except some cryptomonads, chrysophytes, and members of the Volvocales (see Chap. 6). In some instances, poor preservation techniques may cause underreporting (see Chaps. 12 and 21). Two major limitations to survival and growth of river phytoplankton are the continuous removal of organisms by downstream flow (so-called washout) and mixing within the water column, which places cells in variable and often aphotic light fields. Hence, most studies conclude that riverine phytoplankton production is controlled by discharge (Baker and Baker, 1979; Soballe and Kimmel, 1987; Cole et al., 1992; Reynolds and Descy, 1996). Assuming no other limiting resources, rivers must be sufficiently long and/or the flow rate sufficiently low for net positive algal growth rates. This principle was demonstrated clearly in early studies on the Sacramento River in which peaks of abundance in potamoplankton became progressively more pronounced further downstream, a region that provided reduced current velocity and more time for populations to develop (Greenberg, 1964). A similar increase was seen along the Rhine (Germany– Netherlands; de Ruyter van Steveninck et al., 1992). In the lowland River Spree (Germany; Köhler, 1993, 1995), phytoplankton biomass declined in mid-river in response to increased turbidity and Fe precipitation

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of P, but then increased further downstream as a result of impoundments and flow regulation. A rather different longitudinal pattern is seen in the St. Lawrence River, in which a gradient of increased P and reduced current velocity is counterbalanced by greater suspended matter, causing a net decrease in river plankton densities downstream (Hudon et al., 1996). The contrary and complex influences of discharge and algal growth rates have been discussed in detail by Reynolds (1988, 1995). Despite these limits on algal growth, large accumulations of phytoplankton frequently develop in summer and other low-flow periods. With greater nutrient supplies, surface blooms of cyanobacteria (Microcystis, Anabaena and Aphanizomenon) may occur, although their prevalence appears to be greater in warmer climates or in temperate zones under lowflow conditions (Baker and Baker, 1979; Krogmann et al., 1986; Paerl and Bowles, 1987). Unlike many lakes, nutrient limitation is uncommon in larger rivers (Reynolds, 1988; Reynolds and Descy, 1996; Wehr and Descy, 1998). Therefore, the principal factor that regulates phytoplankton production in rivers is discharge, which regulates dilution rates, turbidity, and mixing of cells within the water column (Reynolds and Descy, 1996). Algal productivity in larger and lowland rivers can be substantial despite frequent turbidity and continuous mixing of algal cells within the water column (Descy et al., 1987, 1994; Reynolds and Descy, 1996). A delicate balance exists between phytoplankton production and respiratory losses during periods of higher turbidity (Descy et al., 1994; Reynolds and Descy, 1996). In freshwater tidal sections of the Hudson River, turbidity is further complicated by tidally driven mixing, resulting in a net heterotrophic balance for most of the year (Cole et al., 1992). Following invasion of the Hudson by zebra mussels (Dreissena polymorpha), phytoplankton biomass declined from a summertime mean of about 30 to about 5 µg chlorophyll-a L–1 and species composition shifted from colonial cyanobacteria to diatoms (Caraco et al., 1997; Smith et al., 1998). In the River Spree, a positive autotrophic balance is established during the spring (mainly diatoms), but in the summer, cyanobacteria dominate (Köhler, 1995). How large river systems maintain large phytoplankton populations throughout the year is still something of a mystery, but the main channel may receive subsidies of algae and nutrients from tributaries, wetlands, or backwaters (Owens and Crumpton, 1995; Reynolds, 1996). Species composition may provide a clue: slower growth rates of larger colonial species (e.g., Aphanizomenon and Planktothrix) may have higher respiratory costs (P : R < 1) for maintaining populations than smaller centric diatoms (e.g., Cyclotella spp. and

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Stephanodiscus hantzschii; see Chap. 15), but may be stable if they are ineffectively grazed by small-bodied zooplankton (Gosselain et al., 1998). Only zebra mussels, which utilize a wider particle size range, may crop these larger forms, as in the Hudson River. In the Meuse, Rhine, Danube, and upper Mississippi Rivers, higher levels of nutrients coupled with less turbid conditions enable high levels of phytoplankton production to be sustained for several months of the year (Baker and Baker, 1979; Descy et al., 1987; Lange and Rada, 1993; Admiraal et al., 1994; Kiss, 1994). With greater primary production, potamoplankton influence the biogeochemical properties of large rivers, including dissolved O2 (Köhler, 1995; Reynolds and Descy, 1996), dissolved Si (Admiraal et al., 1993), and dissolved organic matter (Wehr et al., 1997). Phytoplankton are an important food source for zooplankton in rivers, even if grazers do not regulate algal biomass or production as effectively as in lakes. Grazing pressure is less important because the zooplankton community is usually dominated by smallbodied cladocerans and rotifers (Winner, 1975, Köhler, 1995). Rivers select for small-bodied zooplankton because of their ability to grow rapidly enough to compensate for downstream losses (Viroux, 1997). Biomass and density of zooplankton in larger rivers also may be less than in lakes (Pace et al., 1992; Thorp et al., 1994). Although discharge and turbidity typically drive phytoplankton production in rivers on an annual basis, zooplankton grazing may still be an important loss factor during summer low-flow periods (Gosselain et al., 1994). Grazing activity in the River Meuse (mainly Bosmina and rotifers) appears to exert strong control over summertime phytoplankton numbers and to cause a shift in phytoplankton size structure toward larger celled forms (Gosselain et al., 1998). Models designed to predict phytoplankton production in large rivers primarily have been devised for specific conditions, such as the influence of temperature and irradiance on phytoplankton production in one reach of the Great Whale River, Quebec (Rae and Vincent, 1998). Light and temperature explained between 74 and 98% of the variation in photosynthetic activity in this subarctic river. A larger model for the Rhine, which included the effects of irradiance, light attenuation, flow, nutrients, phytoplankton biomass, and grazing, was successful in predicting the fate of algal production, although some parameters (e.g., zooplankton grazing) were based on data from lakes (Admiraal et al., 1993). Efforts to develop nutrient– algal biomass models have been less effective than in lakes, owing to weaker relationships between N or P and chlorophyll-a, and the complicating effects of discharge and turbidity (Van Niewenhuyse and Jones,

1996; Dodds et al., 1998). Even more difficult to predict are changes in species composition of phytoplankton communities in rivers. This information is important for water management agencies, because certain algae affect taste and odor conditions of water that may ultimately be used for domestic consumption. Despite the inclusion of many physical and chemical variables, only about 20% of the total variation in phytoplankton species composition for the River Thames could be explained using a canonical correspondence model (Ruse and Hutchings, 1996). One model, which incorporates several physical factors and river order, found that hydrological conditions exert an overriding effect on potamoplankton development, but biological controls (mainly grazers) are important during low-flow conditions (Billen et al., 1994). Reynolds (1988, 1995) has suggested that more fundamental work is still needed to acquire the necessary data to build meaningful predictive models for plankton communities in rivers.

IV. WETLANDS Wetlands regulate nutrient fluxes between terrestrial and aquatic systems, serve as nurseries for many of the world’s fisheries, and are among the most productive and threatened ecosystems worldwide (Whittaker, 1975; Mitsch and Gosselink, 1993). Freshwater wetlands occur from arctic to tropical biomes across North America. Many are situated in the upper littoral zone of lakes and rivers, and also include marshes, peat bogs, fens, wet alpine meadows, cypress swamps, and forested lowlands. Unifying features of wetland include more or less continuously saturated soils, shallow water depth (≤ 2 m), fluctuating water levels, an accumulation of plant detritus and organic matter, and vegetation adapted to wet conditions (Mitsch and Gosselink, 1993; Goldsborough and Robinson, 1996).

A. Functional Importance of Algae in Wetlands Most wetland algae are benthic, loosely associated with emergent plants (metaphyton), attached to plants, or colonized in sediments; most suspended forms have been dislodged from various surfaces. A five-year study of epipelic, epiphytic, metaphytic, and planktonic primary production in Delta Marsh, Manitoba, determined that metaphyton contributed roughly 70% of the total algal productivity, compared with only 6% for phytoplankton (Robinson et al., 1997). Values for algal productivity (400–1100 g C m–2 y–1) over the year are comparable to or exceed that of the emergent


macrophytes present (aboveground: 100–1700 g C m–2 y–1). Among several constructed wetlands in Illinois, benthic algae are estimated to contribute between 1 and 65% of the total system primary production (Cronk and Mitsch, 1994). However, not all wetland algal communities are highly productive, presumably a result of nutrient limitation (Murkin et al., 1991; Goldsborough and Robinson, 1996). Algal production is important for many invertebrate consumers that preferentially consume algal material over either live or detrital macrophyte tissues (Campeau et al., 1994; Goldsborough and Robinson, 1996). Attached algal material may be especially important in winter when emergent macrophytes are dead or senescent (Meulemans and Hienis, 1983). In a wetland along western Lake Superior, δ13C data suggest that entrained algae are an important primary food source for the grazing food web in addition to macrophyte detritus (Keough et al., 1998). Given the greater proportion of structural tissues in emergent plants and the greater turnover rates among algal cells, the importance of algae in wetland food webs is often substantial. Nutrient additions designed to enhance algal biomass in wetland enclosures also resulted in greater densities of cladocerans and copepods in nearshore water, as well as benthic invertebrates such as snails and chironomids (Gabor et al., 1994). Using pigment tracers, Bianchi et al. (1993) estimated that benthic diatoms comprise a major food source for invertebrates in Hudson River wetlands and, combined with lower C:N ratios, may be a better resource for benthic consumers than detritus. Algal biomass may, however, be spatially variable. In wetlands of Lake Gooimeer (Netherlands), stable isotope data indicate that within Phragmites beds, macrophyte detritus is the major carbon source for benthic invertebrates, but algal material dominates littoral food webs outside the reed bed (Boschker et al., 1995). The littoral zone of lakes is an important region of nutrient exchange between nearshore and pelagic zones, and attached algal communities are important components of this exchange (Mickle and Wetzel, 1978; Aziz and Whitton, 1988; Moeller et al., 1988). During decomposition of wetland plants, attached algae may enhance breakdown (Neely, 1994), although other data suggest that dissolved organic carbon (DOC) released during this process may inhibit algal growth (Cooksey and Cooksey, 1978).

B. Algal Diversity in Freshwater Wetlands The communities of algae in freshwater wetlands are nearly as diverse as those found in lakes. As in other systems, nutrient conditions, climate, and geology influence species composition, but in wetlands,

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water level, macrophyte plant composition, and degree of mixing with other water bodies are also important (Goldsborough and Robinson, 1996). In the Everglades, benthic algae colonize many macrophyte and sediment surfaces; species composition and production vary with species of macrophyte, water level, nutrient inputs, and degree of CaCO3 incrustation (Browder et al., 1994). Filamentous cyanobacteria, including Scytonema, Schizothrix, Oscillatoria, and Microcoleus, are often abundant. Filamentous green algae (Spirogyra, Bulbochaete, and Oedogonium), desmids, and diatoms (Cymbella, Gomphonema, and Mastogloia) are common in less calcareous conditions. Algae represent between 30 and 50% of primary producer biomass in these systems, and their activity is apparent in the large diurnal changes in dissolved O2 and CO2. Because of their close connection with water chemistry, benthic algae help regulate water quality in wetlands, especially P loading from agricultural and urban runoff (McCormick and Stevenson, 1998). In nutrient-rich wetlands, the algal flora is typified by filamentous green algae such as Stigeoclonium, Oedogonium, and Cladophora, cyanobacteria such as Lyngbya, Oscillatoria, and Nostoc, and many diatoms, including species of Amphora, Epithemia, Navicula, Nitzschia, and Surirella (Chaps. 17–19). Filamentous cyanobacteria Rivularia, Calothrix, Microcoleus, and Gloeotrichia (Chap. 4), and meadows of Chara (Chap. 8) typically dominate systems that have greater Ca2+ levels. In more oligotrophic systems, filamentous members of the Zygnematales (Chap. 9), including Mougeotia, Zygnema, and Spirogyra, commonly occur, along with epiphytic diatoms such as Tabellaria, Eunotia, and Fragilaria (see Chaps. 17 and 18), sediment-dwelling species such as Frustulia and Pinnularia (Chap. 17), and many desmids (Hooper-Reid and Robinson, 1978; Livingstone and Whitton, 1984; Goldsborough and Robinson, 1996; Pan and Stevenson, 1996, Chap. 9). Similar to patterns in softwater lakes, wetlands too may be impacted by acidic precipitation, leading to increases in Mougeotia, Zygnema, and, in severely impacted systems, Zygogonium (Stokes, 1986; Turner et al., 1995). Invertebrate grazers mediate species composition. Experimental removal of cladocerans and copepods from marsh enclosures resulted in a shift from a simple community dominated by Stigeoclonium to a more diverse and structurally complex assemblage of diatoms, filamentous green algae, and cyanobacteria (Hann, 1991). An important reason for the success of certain algal species in wetland habitats is their ability to tolerate variations in water level and desiccation. One model predicted specific wetland algal communities that depend on varying water level: dry, sheltered, or

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lakelike (Goldsborough and Robinson, 1996). Water levels may fluctuate several times in a few months or persist for several years. Algae that occupy a variable moisture regime must have adaptations to tolerate extremes of conditions. Some epipelic desmids (Closterium and Micrasterias species) are capable of surviving extended periods of drying and darkness (Brook and Williamson, 1988). Other filamentous forms (e.g., Oscillatoria, Lyngbya, and Oedogonium) may form thick mats during the open (flooded) state that protect algal cells during a later dry phase.

C. Algal Communities of Bogs Bogs are a special class of algal habitats, and include wetlands, streams, and ponds where water retention and chemistry are usually influenced by Sphagnum. Most bogs have lower pH (4.0–5.5), low Ca2+, are poor in nutrients, and have high levels of dissolved organic matter, which casts a yellow or brown stain to the water (Gorham et al., 1985; Cole, 1994). These dystrophic systems, along with lakes high in humic materials, accumulate poorly decomposed organic matter as peat. Ombrotrophic bogs are hydrologically isolated and depend on precipitation as their water source. Bogs are scattered throughout North America, especially in cool or cold regions that have an excess of moisture most of the year (Wetzel, 1983a). They are common across the subarctic, New England– Maritime region, northern Great Lakes, Pacific Northwest, and scattered alpine areas (Brooks and Deevey, 1963; Northcote and Larkin, 1963; Yung et al., 1986). Bogs are not restricted to cool climates, however. Many of the Carolina Bays are dystrophic ponds or bogs and exhibit low algal production (Schalles and Shure, 1989). Bogs also occur in semidesert, desert, and tropical areas, such as in east-central Texas, Arizona, and Costa Rica (Cole, 1963). Some bogs are islands of acidic waters and soils surrounded by an alkaline “sea” (Glime et al., 1982). In bog lakes, mats of vegetation (bryophytes, angiosperms, algae) may float out over the littoral zone and grow toward the center for many years as the bottom of the lake fills in with peat (Whittaker, 1975). Bogs are a stage in the long-term succession of some lake basins that are in the gradual process of filling in. A pioneering limnological study of Cedar Bog Lake (MN; Lindeman, 1941a, b, 1942) documented the vegetational history and aquatic food webs, and was one of the first estimates of freshwater algal productivity. Lindeman’s study helped launch the trophic–dynamic concept in ecology and served as the basis for subsequent energetic studies of freshwater food webs and development of the ecosystem concept (Cole, 1994).

Algal communities of bogs are typically speciespoor, although diversity, especially that of the desmids, may be much greater in systems connected to other lakes or streams (Woelkerling, 1976; Hooper, 1981; Mataloni and Tell, 1996). A tangle of filamentous green algae (Zygnematales and Ulotrichales) and desmids (see Chap. 9) are common, but contrary to common wisdom, although desmids are numerous and diverse, they are rarely important in terms of algal biomass (Yung et al., 1986). Further details on their distribution and ecology are given in Chapter 9. At least three major algal habitats are found in bogs: (1) pools and open water, (2) habitats associated with Sphagnum, and (3) epiphytic habitats of Nuphar or other macrophytes. Data for 31 eastern bog systems demonstrated that algal species richness increases (especially desmids) with proximity to the Atlantic coast, and is less in systems with greater color and lower pH (Yung et al., 1986). The flagellate Gonyostomum (Raphidophyceae; Chap. 11, Sect. III) and the red alga Batrachospermum turfosum (as B. keratophytum; Chap. 5) are two unusual species that are characteristic of boggy systems (Prescott, 1962; Bourelly, 1985; Yung et al., 1986; Sheath et al., 1994). Diatoms are generally less diverse than desmids, but certain species are frequently observed, including Anomoeoneis brachysira, Frustulia rhomboides var. saxonica, Eunotia elegans, E. exigua, Navicula subtilissima, Pinnularia viridis, and several types of Stauroneis spp. (Kingston, 1982; Cochrane-Stafira and Andersen, 1984; Mataloni and Tell, 1996). Diatom species composition may change with successional stage of the surrounding vascular plant community, and apparently are responsive to many of the same variables, such as Ca, pH, and specific conductance (Cochrane-Stafira and Andersen, 1984). Although it has been argued that cyanobacteria may be unable to tolerate lower pH, especially values less than 4.0 (Brock, 1973), there are several species that are common in these waters, including species of Aphanocapsa, Chroococcus, Dacytlococcopsis, Hapalosiphon, Microchaete, Nostoc, and Stigonema. Because many of these species are heterocystous, their N-cycling role in these nutrient-poor environments deserves attention. Some species of dinoflagellates, chrysophytes and synurophytes also may be found, although their abundance is usually low.

V. THERMAL AND ACIDIC ENVIRONMENTS A. Thermal Springs Thermal springs and streams (hot springs) are extreme environments in geologically active regions where temperatures are influenced by geothermal


sources and can range from 35 to 110°C. In North America, thermal springs are common from Alaska south to Costa Rica, and in scattered locations in Arkansas, Florida, Georgia, Virginia, and north to Massachusetts. Among the best known for aquatic organisms is the spectacular thermal area of springs, geysers, and fumaroles in Yellowstone National Park. Some thermal springs are less obvious, exerting their chemical and thermal effects on larger lakes and rivers, as in Yellowstone Lake (and adjacent lakes), and several streams and lakes in eastern Costa Rica (Pringle et al., 1993; Theriot et al., 1997). The geology, chemistry, and organisms of hot springs have been reviewed by Castenholz and Wickstrom (1975), Brock (1986), and Ward and Castenholz (2000). Temperatures can be fairly constant near the source, but can range from about 110°C (with a high concentrations of salts) to just above ambient, depending on the temperature and volume of thermal water, distance from the source, and volumes of nonthermal surface water entering a system. Temperature is not the only extreme condition for thermal organisms; most springs have elevated concentrations (50–150 mg L–1) of inorganic ions (Ca2+, Mg2+, Na+, HCO3–, SO42–, Cl–, Si, and H2S) and elevated pH (8–10). These conditions select for highly adapted organisms, especially chemoautotrophic and heterotrophic bacteria in very hot (> 70°C to ⬇94°C) conditions (Brock, 1985b). Among photosynthetic organisms, cyanobacteria are most common, with an upper limit between 70 and 73°C; eukaryotic algae are restricted to a maximum of about 55°C. Along a thermal stream, distinct zones of bright colors and morphologies that correspond to different species along the thermal gradient can be observed. A diversity of cyanobacteria dominate thermal waters. They include masses of coccoid species of Aphanocapsa, Chroococcus, Cyanobacterium, and Synechococcus, and filamentous species of Mastigocladus, Oscillatoria, and Phormidium (Ward and Castenholz, 2000, Chaps. 3 and 4). In cooler waters further from the source (35–50°C), diatoms (Achnanthes and Pinnularia) and green algae (Spirogyra and Mougeotia) proliferate (Stockner, 1967; Castenholz and Wickstrom, 1975). Cyanobacterial assemblages may form mats several centimeters thick and have extremely high rates of primary production (> 10 g C m–2 d–1; Castenholz and Wickstrom, 1975). In one hot spring in Costa Rica (62°C, pH 7.0), a species of Oscillatoria dominated, while nearby streams with less extreme temperatures (35–36°C; pH 7.8–8.0) had a greater diversity of algal species, including cyanobacteria (Oscillatoria, Phormidium and Lyngbya) and diatoms (Pinnularia; Pringle et al., 1993). Unusual con-

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sumers are associated with hot springs, because few metazoa tolerate temperatures greater than about 50°C. Invertebrates include ostracods, water mites, and rotifers, but little is known of their dynamics or food webs. Adult beetles and flies are successful in some systems. Brine flies (Paracoenia and Epihydra) lay eggs in microbial mats found in springs in Yellowstone Park within the 30–40°C range. Both adult and larval stages consume algal and bacterial material, which may in turn enhance primary productivity (Brock, 1967; Brock et al., 1969). Some thermal springs are highly acidic, which further limits their species diversity. A notable eukaryote, the red alga Cyanidium caldarium, is often the sole photosynthetic organism in very acid (pH 2–4) hot springs up to about 55°C. This somewhat enigmatic organism was variously classified as a cyanobacterium, green alga, cryptomonad, and an evolutionary link between red and green algae (Seckbach, 1991); today it is placed in the division Rhodophyta based on pigments, chloroplast structure, and molecular features (Steinmüller et al., 1983; Pueschel, 1990). In acid hot springs, other enigmatic rhodophytes including Cyanidioschyzon merolae and Galdiera sulphuraria (DeLuca and Moretti, 1983), also have been reported, also have. It seems to be unlikely that either high temperature or low pH is solely responsible for this peculiar flora, because alkaline hot springs and nonthermal acid springs have very different algal communities.

B. Acid Environments Most highly acid springs and streams are nonthermal and support a characteristic algal flora that is unlike those in other aquatic environments. Although bogs may exhibit relatively low pH (4.0–5.0), highly acidic environments typically are regarded as systems with H+ concentrations at least an order of magnitude greater, that is, pH values ≤ 3.0, and they usually receive acidic inputs from either geological or anthropogenic sources (Hargreaves et al., 1975). Nearly all have elevated concentrations of metals, including Al, Fe, Mn, Pb, Co, Cu, and Zn, which may be near saturation levels even for very low pH, resulting in the formation of metal salt precipitates along stream margins or on algal colonies. Acid springs with very high Fe concentrations also may have very low dissolved O2, due to Fe(OH)2 and FeO(OH)2 precipitates (Van Everdingen, 1970). Laguna de Alegría, a crater lake in El Salvador, is influenced by sulfur-rich fumaroles and exhibits pH values as low as 2.0 (Cole, 1963). The earliest detailed studies of algae in highly acidic systems in North America were conducted in acid mine drainages in Indiana, Kentucky, Ohio,

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Pennsylvania, and West Virginia (Lackey, 1938; Bennett, 1969; Warner, 1971). These studies, as well as those in the United Kingdom (Hargreaves et al., 1975), all reveal low species diversity and a remarkable similarity in composition. Euglena mutabilis is the most widespread and often most abundant species, occurring in systems as acidic as pH 1.5. E. mutabilis is also common in naturally acidic streams, such as the Rio Agrio (pH 2.3) in Costa Rica (Pringle et al., 1993), and in acidic ponds in the Smoking Hills region of the Northwest Territories (Sheath et al., 1982). The latter site also supports populations of Chlamydomonas acidophila. E. mutabilis is not apparent in natural acid springs in Kootenay Paint Pots (BC) although a few diatoms and green algae are present (Wehr and Whitton, 1983). Other common elements in many highly acidic environments include Klebsormidium (previously Hormidium) rivulare, Eunotia tenella, Pinnularia microstauron, P. braunii, and Gloeochrysis turfosa. Acid sites in West Virginia contain many of the same species found in the United Kingdom. (Bennett, 1969), although an apparent absence of G. turfosa from North American sites may be the result of the alga being overlooked. No studies report cyanobacteria in these highly acidic environments, in agreement with Brock’s (1973) recommendation for a lower pH limit of less than 4.0. Isolates of E. mutabilis, Chlamydomonas acidophila, Klebsormidium rivulare, Gloeochrysis turfosa, and Stichococcus bacillaris from one acid stream (pH 2.6–3.1) were able to tolerate and grow at pH levels less than the lowest measured in their collecting site (Hargreaves and Whitton, 1976a). In addition, an acid strain of Klebsormidium rivulare tolerated greater Zn and Cu concentrations in the pH range 3.0–4.0, than at pH ≥ 6.0, suggesting a H+–metal interaction (Hargreaves and Whitton, 1976b).

VI. UNUSUAL ENVIRONMENTS A. Saline Lakes and Streams Saline lakes and streams make up a large and heterogeneous collection of water bodies that have elevated total dissolved salts (> 500 mg L–1; Williams, 1996). Many are closed basins or desert playas that gain salinity over the year as they lose water (Hammer, 1986; Evans and Prepas, 1996). The term “saline” does not mean simply greater concentrations of NaCl. The ion content of inland saline lakes is influenced by Na+, K+, Ca2+, and Mg2+, and the major anions are typically Cl–, SO42–, HCO3–, and CO32– (Wetzel, 1983a; Hammer, 1986; Cole, 1994). Systems are usually well buffered (high in HCO3– and/or CO32–) and neutral to alkaline in pH (7.5–10.0). Among 47 saline lakes in

the western United States and Canada, anion and cation chemistries vary significantly with latitude: lakes north of 47° latitude are dominated by SO42– and either Na+ or Mg2+, whereas lakes in more southern locations are dominated by CO32– or Cl– in conjunction with Na+ ions, reflecting climatic as well as geological characteristics of each region (Blinn, 1993). Saline lakes of athalassic (nonmarine) origin differ from oceanic systems in several important ways. Total dissolved salts vary considerably more (0.5–600 g L–1) than in oceans (35–40 g L–1), both among systems and over time. Most athalassic lakes are shallow, which makes seasonal and longer term (climatic, anthropogenic) changes in salinity important for algal survival, and results in low biotic diversity (Cole, 1994). A few are large and relatively deep, such as Pyramid Lake (532 km2; 102 m deep) and Big Soda Lake (1.5 km2; 64 m deep) in Nevada (Hutchinson, 1957), Mono Lake (150 km2; 40–50 m deep) in California (National Academy of Sciences 1987), and Soap Lake (3.6 km2; 27 m deep) in Washington (Castenholz, 1960). Because of differences in concentrations of salts with depth, many of the deeper saline lakes, such as Soap and Mono Lakes, are meromictic. Water level in Mono Lake varies substantially as a function of water diversion for human usage. In Redberry Lake, Saskatchewan, mean depth has decreased by about 37%, while salinity has increased by roughly 41% since the 1940s, due to changes in land use (Evans et al., 1996). Saline lakes are concentrated in arid environments, especially in the U.S. Southwest, Mexico, and interior regions of California, Oregon, Washington, northern prairies, and British Columbia. Some of the most extensive surveys of the chemistry and biology of saline lakes were conducted in Saskatchewan (Rawson and Moore, 1944; Hammer et al., 1983). Few inland saline lakes are found in eastern North America, although Onondaga Lake (12 km2, 20.5 m depth) is a saline lake in the New York Finger Lakes region. Levels of [Na+ + K+] and Cl–1 exceeded 500 and 1400 mg L–1, respectively, in part from salt springs, plus pollution from an adjacent soda ash facility (Berg, 1963; Sze and Kingsbury, 1972). Controls on salt waste have resulted in reduced total salinity (450 mg Cl– L–1), although levels are still greater than pre-industrial times (ca. 230 mg Cl– L–1; Effler and Owens, 1996; Rowell, 1996). Algal communities of saline lakes differ among systems and their diversity varies inversely with salinity (Blinn, 1993; Cole, 1994). Because many saline lakes are shallow and subject to wind-driven mixing, it is often difficult to distinguish between benthic and planktonic forms. Several studies have evaluated the influences of salinity and ion composition on algal communities (Castenholz, 1960; Hammer et al., 1983;


Blinn, 1993; Fritz et al., 1993; Evans and Prepas, 1996; Wilson et al., 1996). In mildly saline (total salts 500–2000 mg L–1) lakes, the algal flora is fairly rich and composed of a variety of diatoms (e.g., species of Amphora, Campylodiscus, Cyclotella, Epithemia, Fragilaria, Navicula, Nitzschia, and Rhopalodia), green algae (Crucigenia, Pediastrum, Oocystis, and Sphaerocystis), and cyanobacteria (e.g., species of Anabaena, Aphanizomenon, Chroococcus, Lyngbya, Merismopedia, Microcystis, and Oscillatoria), especially if N and P are high. In more strongly saline conditions (2–20 g L–1), many species are eliminated, but the community still includes taxa found in nonsaline waters, like Cladophora glomerata, Botryococcus braunii, Cocconeis placentula, Mastogloia spp., Nitzschia palea, Plagioselmis (as Rhodomonas) minuta, and several cyanobacteria (Aphanizomenon flos-aquae, Anabaena spp., Microcystis aeruginosa, and Oscillatoria spp.). Species typical of higher salinities also co-occur within this range, probably because concentrations can vary by an order of magnitude or more over a year in many lakes. Under hypersaline conditions (20–600 g L–1), diversity is very low, and includes some species that are restricted to higher salt levels, such as the diatoms Amphora coffeiformis, Anomoneis sphaerophora, Navicula subinflatoides, Nitzschia communis, and N. frustulum, cyanobacteria Nodularia spumigena and Aphanothece halophytica, and the filamentous green alga Ctenocladus circinnatus (Blinn, 1971; Herbst and Bradley, 1989; Wurtsbaugh and Berry, 1990; Kociolek and Herbst, 1992; Reuter et al., 1993). A few taxa that have marine distributions, such as Enteromorpha intestinalis, diatoms Dunaliella salina, D. viridus, Chaetoceros muelleri, and Thalassiosira pseudonana, and the coccolithophorid Pleurochrysis carterae, have been observed (Sze and Kingsbury, 1972; Stephens and Gillespie, 1976; National Academy of Sciences, 1987; Johansen et al., 1988). Experiments in which salinity was varied within Mono Lake mesocosms determined that diatom dominance, algal diversity, chlorophyll-a, and photosynthesis in benthic communities all declined at higher salinities, while Ctenocladus circinnatus and an Oscillatoria spp. dominated, although the diatom Nitzschia monoensis increased (Herbst and Blinn, 1998). Primary production by algae is often high (300– 1000 g C m–2 y–1), along with very high chlorophyll-a levels (50 to > 500 mg L–1), because many saline lakes are located in sunny locations and have surplus P levels (total P: 50 to > 1000 mg L–1; Hammer, 1981; National Academy of Sciences, 1987; Robarts et al., 1992). Much of this production comes from epipelic and epilithic assemblages (e.g., Ctenocladus) rather than phytoplankton (Wetzel, 1964). Surface blooms of cyanobacteria may at times represent a significant pro-

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portion of total production (Sze and Kingsbury, 1972; Robarts et al., 1992). Many lakes in the Canadian prairies have lower algal biomass and primary productivity than would be predicted using standard nutrient models (Campbell and Prepas, 1986), apparently because of greater densities of macrozooplankton that flourish in saline waters and perhaps due to fewer zooplanktivorous fish (Evans et al., 1996). In Great Salt Lake, weather-induced decreases in salinity (250–50 g L–1) resulted in an increase in grazers from one species (Artemia) to four, where rotifers and copepods were dominant (Wurtsbaugh and Berry, 1990). Paleoecological studies have considered whether these systems were saline in the past (Fritz et al., 1993; Rowell, 1996). A study of the diatoms from 219 lakes in British Columbia indicated that some species are limited by salinities as dilute as 0.02 g L–1 (e.g., Achnanthes pusilla), whereas others tolerate more than 500 g L–1 (Amphora coffeiformis and Nitzschia frustulum; Wilson et al., 1996). Models from these data have been used to infer temporal and spatial differences in precipitation, which in turn influences salinity in these closed basins. Long-term changes in fossil algal pigments suggest that as prairie lakes became more saline, phytoplankton communities have shifted from a diatom–chrysophyte–dinoflagellate community to one dominated by greens, cyanobacteria, and diatoms (Vinebrooke et al., 1998). Due to possible effects of global warming, semiarid regions of North America may experience an increase in the number of saline waters in the next few decades (Evans and Prepas, 1996).

B. Snow and Ice People who have hiked in alpine regions are familiar with red snow, especially where snowfields accumulate for most of the year. Aristotle also apparently observed red snow many centuries ago (Kol, 1968). This phenomenon is most often caused by the green alga Chloromonas (previously recorded as Chlamydomonas) nivalis (see Chap. 6). The red color is the result of an accumulation of secondary carotenoids, mainly astaxanthin, in resting cells (Round, 1981; Bidigare et al., 1993). Not all snow communities are colored red: orange, brown, and green patches are also seen, depending on the species present, dominant pigments, exposure to sunlight, and perhaps pH (Stein and Amundsen, 1967; Kol, 1968; Hoham and Blinn, 1979). Cells aggregate near the surface, and as successive snowfalls accumulate, layers or bands of pigmented algal communities can be seen in vertical cuts through a snow bank (Hoham and Mullet, 1977). Although motile stages can be found in snow, most cells occur either as thick-walled resting zygotes or as asexual

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hypnospores (Stein and Amundsen, 1967). Cryophilic flora include other algal flagellates, such as Chloromonas nivalis, C. brevispina, Carteria nivale (= planozygotes of Chloromonas sp.?), Scotiella cryophila, and Chromulina chionophila, and nonflagellated green algae, such as Raphidonema nivale and Stichococcus spp. (Stein and Amundsen, 1967; Kol, 1968; Hoham, 1975; Hoham and Blinn, 1979). The life cycles of many snow algae are incompletely known; thus, some cells previously identified as new taxa have been shown to be zygotes of other known species (Hoham and Blinn, 1979; R.W. Hoham, personal communication). Snow algae exhibit measurable but low photosynthesis rates, many of which reach a maximum near 10°C, although some peak near freezing and decline at higher temperatures (Mosser et al., 1977; Round, 1981). A strain of Chloromonas pichinchae from snowfields in Washington state grew best at 1°C and pH 6.0, whereas the optimum for an isolate of Raphidonema nivale from the same location was 4–5°C, with a pH optimum ≥ 7.0, despite lower environmental pH (ca. 5.0) and temperature (near 0°C; Hoham, 1975). Strains of Chloromonas isolated from snowfields in the Adirondack Mountains (exposed to decades of acid deposition) have significantly greater (1.5- to 2.2-fold greater) growth yields at pH 4.0 than Chloromonas isolated from the (less acidic) White Mountains of Arizona (Hoham and Mohn, 1985). Algal cells that colonize snow surfaces at high altitude experience extreme levels of solar (including UV) radiation. Carotenoid pigments provide photoprotection by reducing total radiation and filtering certain shorter wavelengths from photosynthetic pigments (Bidigare et al., 1993). Experiments with snow algae from Tioga Pass in the Sierra Nevada found that UV radiation inhibited photosynthesis in green snow by 85%, but only 25% in red snow (Thomas and Duval, 1995). Snow algal communities also are influenced by the amount of wind-blown soil (plus factor: nutrient source), exposure (plus or minus factor: direction and quantity of sunlight), snow albedo (usually minus factor: reflective property), and water content (plus factor) of the snow (Stein and Amundsen, 1967; Hoham and Blinn, 1979; Thomas and Duval, 1995). These communities have cryophilic food webs with protozoan consumers, and bacterial and fungal decomposers that are active at very low temperatures (Felip et al., 1995; Thomas and Duval, 1995).

C. Other Unusual Habitats Algae form thick mats on tank walls, outflow weirs, pipes, and other substrata in sewage treatment plants, where concentrations of nutrients greatly exceed

requirements (N: 1–20 mg L–1, P: 0.1–2 mg L–1). The most common taxon appears to be Stigeoclonium tenue; lesser abundances of Chlorella spp., Nitzschia palea, Oedogonium sp., Oscillatoria spp., Pleurocapsa minor, Pseudanabaena catenata, Scenedesmus quadricauda, and Tribonema spp. also are observed (Palmer, 1962; Sládecková et al., 1983; Davis et al., 1990a). Communities exhibit seasonal changes in composition: Tribonema dominates in the spring, and Oscillatoria, Scenedesmus, and Stigeoclonium reach their maxima in warmer months. These shifts are largely driven by changes in light and temperature, not nutrient supply (Davis et al., 1990a). Substantial biomass and rapid growth rates have made algae candidates for nutrient removal plans (Sládecková et al., 1983; Davis et al., 1990b). Treatment ponds have been devised in which > 95% of P may be removed by algal assemblages alone (Hoffmann, 1998). Algae colonize caves in many limestone or dolomite regions where fractures or underground streams form underground pockets or large caverns. Some cave-dwelling algae, including species of Gloeocapsa (Cyanobacteria), survive very low light levels by having densely packed thylakoids in their cells and very slow growth rates (Pentecost and Whitton, 2000). By some observers caves may be considered to be dark, unproductive environments, but substantial algal floras develop where surface are open to the sunlight (fissures, mouths) or in show caverns where artificial lighting has been added. Cyanobacteria (e.g., Aphanocapsa, Calothrix, Chroococcus, Gloeocapsa, Hapalosiphon, and Schizothrix) may be especially abundant near light sources (Claus, 1962; Round, 1981). Species diversity is typically low: Timpanogos Cave (UT) and Seneca Cavern (OH) each support an algal flora of fewer than 30 species (St. Clair and Rushforth, 1976; Dayner and Johansen,1991). Species include diatoms (Navicula tantula and N. contenta), unicellular green algae (Chlorella miniata), and the xanthophyte Pleurochloris commutata. CaCO3 from cave walls may become incorporated into algal colonies and thalli, which is why artificial lighting and the resultant algal colonization has been cited as a prime cause of cave wall destruction in show caves (Gurnee, 1994). The calcareous cave flora are similar to wet limestone seepages above ground, and exhibit an abundance of calcified cyanobacteria and diatoms (Pentecost, 1982; Pentecost and Whitton, 2000). Algae from pools adjacent to caves may be an important carbon source for food webs inside caves (Pohlman et al., 1997). Noncalcareous, temporary rock pools that form in weathered bedrock along lake and river margins also become algal habitats. These small systems (a few liters or less) experience great variations in water level and


nutrients, and extremes in solar radiation. The most common inhabitant is the algal flagellate Haematococcus lacustris, which accumulates red secondary carotenoids much like snow algae. This adaptation serves a similar photoprotective role and is not influenced by nutrient levels (Yong and Lee, 1991; Lee and Soh, 1991). Humans have created a widespread habitat for H. lacustris, namely birdbaths (Canter-Lund and Lund, 1995); hence, many people report red water in their birdbaths after a year or so of use. Endophytic algae live in cavities of higher plants or colonize plant tissues intracellularly. Algae colonize microhabitats created by bromeliad cups, and colonized by algae, and differ in pH and dissolved O2 levels among different plants (Laessle, 1961). Perhaps the most ecologically important endophytic algae in freshwaters are species of cyanobacteria that colonize various aquatic plants and bryophytes. The best known is Anabaena azollae, a symbiont within the water fern Azolla, which is used as a nitrogen source or “green fertilizer” for rice crops worldwide (Bothe, 1982). Roots of some cycads (Cyas, Encephalartos, and Macrozamia) are colonized by several cyanobacteria, especially Anabaena, Calothrix, and Nostoc (Huang and Grobbelaar, 1989); when sectioned, cyanobacteria appear as a healthy blue–green color. Green algae (e.g., Chlorella, Chlamydomonas) and some xanthophytes (see Frost et al., 1997, Chaps. 6 and 7) are symbionts in a variety of aquatic organisms, including freshwater sponges (e.g., Spongilla lacustris and Corvomeyenia everetti), cnidarians (e.g., Hydra spp.), ciliates (e.g., Euplotes and Ophrydium), and other organisms (Slobodkin, 1964; Frost and Williamson, 1980; Berninger et al., 1986; Sand-Jensen et al., 1997). In the spring, green, baseballsized (up to 20 cm) green masses can be observed in ponds and lake outflows that are actually mucilaginous masses of amphibian eggs (Amblystoma and Rana) colonized by the green alga Chlamydomonas (syn. Oöphila) amblystomatis. This association appears to provide N for the alga and added O2 for developing amphibian eggs (Goff and Stein, 1978; Bachmann et al., 1986; Pinder and Friet, 1994). Much more intimate symbioses are found among algae and cyanobacteria with fungi, as lichens. The treatise by Ahmadjian (1993) can be consulted for further information. Many algae colonize terrestrial habitats such as soils, trees, and other surfaces, and serve important ecological functions in soil and moisture retention, seed germination, nutrient dynamics, and succession of terrestrial vegetation (Carson and Brown, 1978; Bell, 1993; Vazquez et al., 1998). Many of the more common terrestrial species, appear as a green “felt” on stone walls, tree bark, and wooden fences such as Apatococcus and Trentepohlia (Chlorophyta), that may

45

be mistaken for moss (Canter-Lund and Lund, 1995). Epiphytic green algae have been used as biological indicators of air quality (Hanninen et al., 1993). A number of genera that colonize plant leaves, walls, stones, or soils are also common in fresh waters, including many cyanobacteria (Nostoc, Oscillatoria, Lyngbya, and Plectonema), green algae (Chlamydomonas, Chlorella, Chlorococcum, and Klebsormidium), and diatoms (Achnanthes, Hantzschia, and Navicula; Segal, 1969; Cox and Hightower, 1972; King and Ward, 1977; Hunt et al., 1979). Diatoms from these habitats are often desiccation-resistant and many are regarded as obligate aerial taxa (Johansen, 1999). Other genera, such as Prasiola and Zygogonium, have very distinct ecological requirements. Prasiola colonizes N-rich (e.g., guano) rocks and walls in aerial (often shaded) and even urban environments (Jackson, 1997; Rindi et al., 1999), whereas aquatic species colonize cool, nutrientpoor streams (Sheath and Cole, 1992; Kawecka, 1990; Hamilton and Edlund, 1994). A few terrestrial species colonize the interstitial spaces within crystalline rocks in arid or semiarid regions, where water supply may be limited (Bell, 1993; Johansen, 1993). Further details on the algal flora of soils and terrestrial habitats can be found in reviews by Fritsch (1922), Starks et al. (1981), and Johansen (1993). Some algae are parasitic within plant tissues (leaves and twigs), such as Cephaleuros, which is the cause of red rust disease in higher plants (Thompson and Wujek, 1997). Perhaps one of the most unusual of all algal habits is that of the red alga Rufusia, which lives within the hairs of twoand three-toed sloths, but apparently does not colonize nearby vegetation (Chap. 5).

ACKNOWLEDGMENTS Thanks are due Dr. R. Jan Stevenson (Michigan State University) and Dr. Robert G. Wetzel (University of Alabama) for helpful reviews and advice on relevant literature. Dr. Ronald W. Hoham (Colgate University) provided advice on species of snow algae. We also thank D. W. Blinn, D. Burney, R. J. Cannnings, S. J. Cannings, J. Cotner, P. R. Leavitt, A. Merola, NASA, and the South Florida Water District for permission to reproduce photos.

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COCCOID AND COLONIAL CYANOBACTERIA Jirˇí Komárek Institute of Botany Academy of Sciences of the Czech Republic Faculty of Biological Sciences University of South Bohemia CZ-37982 Trˇebonˇ, Czech Republic I. Introduction II. Morphology and Diversity A. Cellular and Colonial Morphology B. Cytology C. Cell Division and Reproduction D. Diversity and Classification III. Ecology and Distribution A. Soils B. Subaerial Environments

I. INTRODUCTION Cyanobacteria (cyanoprokaryotes, cyanophytes, and blue–green algae) represent an ancient, but diverse and abundant group of microorganisms that possess prokaryotic (bacterial) cell structure and predominantly CO2-dependent oxygen-evolving photosynthesis. Unlike all other algae, cyanobacteria are classified within the Eubacteria due to their simple, prokaryotic cells and gram-negative (peptidoglycan) cell walls (Stanier and Cohen-Bazire, 1977). Cells contain chlorophyll-a and several phycobilin–protein complexes that produce a variety of pigmentations, including the characteristic blue–green color. Many species possess nitrogenase and fix atmospheric N2; several are capable of precipitating calcium carbonate, and they form stromatolites and travertine deposits (Fogg et al., 1973; Carr and Whitton, 1973, 1982; Whitton and Carr, 1982). Therefore, they belong to very important photosynthetic microscopic organisms in natural environments. Cyanobacteria are thought to have evolved during the early Precambrian period, and during their long exisFreshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

C. Aquatic Environments D. Extreme Environments E. Geographic Distribution IV. Collection, Preparation, and Culture V. Key and Descriptions of Genera A. Key B. Descriptions of Genera VI. Guide to Literature for Species Identification Literature Cited

tence, they have colonized nearly all freshwater, marine, and terrestrial habitats on Earth. Many cyanobacteria occur in extreme habitats, such as hot springs (up to 70°C), hypersaline lakes, and hot and cold deserts. Some planktonic1 species form massive surface blooms, have the ability to produce toxins, and play an important role in water quality (Reynolds and Walsby, 1975). Cyanobacteria occur in a wide variety of morphologies and ecological forms. Hence, historically their classification has been based on simple morphological characteristics (e.g., Gomont, 1892; Geitler, 1932; Desikachary, 1959), whereas many bacteriologists have applied physiological and biochemical properties for those species that exist in culture (e.g., Castenholz and Waterbury, 1989; Castenholz, 2001). During the last four decades, ecological characteristics, ultrastruc-

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The adjective “planktonic,” derived from the Greek term “plankton,” is incorrect according to ancient and modern Greek grammars (it should be “planktic,” similarly to “benthic,” “metaphytic,” etc.), but it is the common term in English speaking countries.

59

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tural features, and molecular evidence have substantially influenced our knowledge of this group. As a result, species concepts and classification are undergoing radical changes (Anagnostidis and Komárek, 1985; Castenholz, 1992; Komárek, 1994; Castenholz, 2001). However, the cellular–morphological approach to classification (including ecospecies) cannot yet satisfactorily be replaced by any other criteria. The molecular and ultrastructural studies explain relationships well, but thus far encompass only a small portion of the cyanobacterial diversity and variability that occur in natural habitats. Many well-known taxa (e.g., from the genera Gloeocapsa, Gomphosphaeria, Woronichinia, Chroococcus, Hyella, and many others) have not yet been studied in culture, which is a first step to experimental revisions of the taxonomic status of species (Rippka et al., 1979; Komárek, 1994; Castenholz, 2001). There are also difficulties in identifying cyanobacteria from different geographic areas. Misinterpretations result with both field populations and isolated strains, due to the fact that so few collections and identifications have been made from certain habitats, especially tropical and subarctic regions, and from extreme environments (Frémy, 1930b; DiCastri and Younéz, 1994; Watanabe, 1999). Changes in the classification and identification of cyanobacteria can be expected in the future, but presently the traditional approach is still necessary, especially for field studies. The distribution and taxonomy of cyanobacteria in North America has been studied by a number of phycologists over many decades (e.g., Wood, 1872; Tilden, 1910; Smith, 1920, 1925, 1933, 1950; Gardner, 1927; Copeland, 1936; Drouet, 1936a, b, 1938, 1942; Daily, 1942, 1946; Drouet and Daily, 1948, 1952; Prescott, 1962; Golubic´, 1965, 1967a, b, 1980; Friedmann, 1971; Croasdale, 1973; Komárek and Anagnostidis, 1995). However, recent studies of natural populations are few and much needed to understand the level of cyanobacterial diversity in North America. Specialists are interested mainly in revisions and characterization of individual species and genera (Jaag, 1941; Komárek, 1970; Parker, 1982; Lukas and Golubic´, 1983; Wilmotte and Stam, 1984; Komárek and Hindák, 1989; GoldMorgan et al., 1994), or cyanobacterial diversity in particular habitats, some of which may employ molecular data (Komárek, 1999). Based on these historical and present-day studies, plus some preliminary studies by me and co-workers (Komárek and Montejano, 1994; Tavera and Komárek, 1996; Komárek and Komárková-Legnerová, 2002) on habitats in central America (e.g., volcanic regions, marine and brackish waters, subaerial habitats from wet and desert rocks, and soil communities), it is estimated that no more than 10% of all natural forms of cyanobacteria (species and

varieties) from North America have been described. Chapters 3 and 4 represent an effort to update the general knowledge of the North American cyanobacterial flora and to stimulate greater study of these organisms across the continent. Cyanobacteria have been traditionally classified into several orders. This chapter concerns coccoid unicelled and colonial cyanobacteria [order Chroococcales Wettstein 1924; see Geitler (1932, 1942), including Pleurocapsales Geitler 1925; for details see Geitler (1932), Waterbury and Stanier (1978), and Komárek and Anagnostidis (1986, 1998)]. This is a heterogeneous group, as concerns ultrastructural and molecular criteria, and will be most likely reclassified along several distinct evolutionary lines. However, the data available do not yet allow a full revision and therefore all coccoid cyanobacteria are considered and reviewed as one group in this chapter.

II. MORPHOLOGY AND DIVERSITY A. Cellular and Colonial Morphology The simplest cyanobacteria grow as solitary cells, which may be enveloped by a thin, diffuse or firm gelatinous layer (sheath), or may be clustered into colonies; very thin sheaths may not be apparent in the light microscope (Drews and Weckesser, 1982). Cell size varies from less than 1 µm in diameter (picoplankton; Platt and Li, 1986; Komárek, 1999; ˇSmajs and ˇSmarda, 1999) to cells greater than 20 µm in length. Solitary cells and/or specialized cells in a cluster or colony may reach a diameter up to 100 µm (Geitler, 1932; Komárek, 1976). Some colonial forms (e.g., species of Microcystis) produce massive accumulations in surface blooms that are recognizable with the naked eye. Cells vary from spherical, oval, fusiform, and rodlike, to irregular in shape; in some colonial genera, the shape may appear polygonal or elongate. An important evolutionary trend within chroococcal cyanobacteria is the polarization of cells and thalli. For example, in some forms the cells are heteropolar, morphologically differentiated in basal and apical parts, and possess specific patterns of cell division (Chamaesiphonaceae and Dermocarpellaceae; Figs. 16B–18). A heteropolar thallus occurs in more complicated forms, where a collection of heterogeneous cells forms a colony (Xenococcaceae and Hyellaceae; Figs. 20–23). Both amorphous and polarized colonies occur in this group; each develops from solitary cells and forms small clusters, which may grow into distinct colonies or macroscopic mats. Mucilaginous colonies of simple genera can be spherical, oval, or irregular. The structure of sheaths and envelopes around cells is diverse: the mucilage can form hyaline, amorphous mass, diffused

3. Coccoid and Colonial Cyanobacteria

and marginal envelopes, or structured and layered sheaths. Gelatinous envelopes (fine mucilage) and sheaths (firm or structured external layer) appear to have a protective function in habitats that are exposed to intense solar radiation; often they are colored by carotenoid pigments (scytonemin and gloeocapsin; Garcia-Pichel and Castenholz, 1991; Whitton, 1992).

B. Cytology The prokaryotic structure of cyanobacterial cells corresponds with other members of the Eubacteria. The distribution of DNA appears fairly uniform by electron microscopy (EM), but is actually concentrated in the central region, forming nucleoids that have shapes that are diagnostic among different genera (Cepák, 1993). The position and pattern of photosynthetic thylakoids seen with EM differ among various genera. In numerous species, three to six or more thylakoids are peripherally arranged in parallel and cause a dark peripheral region within cells that is recognizable with the light microscope (LM), and is sometimes termed chromatoplasm in older literature. The internal part without thylakoids is called the centroplasm (or nucleoplasm; Geitler, 1932, 1942). Genera with different thylakoid patterns probably belong to different evolutionary lines. Regularly widened thylakoids have been observed in several species (Komárek et al., 1975; Roussomoustakaki and Anagnostidis, 1991). Their occurrence and intensity depend on light intensities, but the function and metabolic processes connected with this variability are still unclear. Under the LM, cells appear to be pale or bright blue–green, olive green, greyish, pinkish, or violet, depending on the ratio of photosynthetic pigments, particularly chlorophyll-a and phycobilins, allophycocyanin, phycocyanins (blue), and phycoerythrin (red). Phycobilins are localized in phycobilisomes and their ratio can change in several species and within the same population (called photoacclimation or chromatic adaptation in older literature). In species or strains capable of photoacclimation, green light stimulates synthesis of phycoerythrin, which more efficiently captures these wavelengths than chlorophyll-a; conversely, increased red light stimulates phycocyanin synthesis (Cohen-Bazire and Bryant, 1982; Bryant, 1994). Different carotenoids have been found in several species from individual genera. Granules of various types occur throughout most cells. Phycobilisomes associated with thylakoids contain both phycobilins (Cohen-Bazire and Bryant, 1982). Assimilatory materials, polyphosphates, and carboxysomes (site of ribulose bisphosphate carboxylase/oxygenase RUBISCO) are the most common inclusions (Healy, 1982; Smith, 1982; Whitton, 1992). The character of

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most granular inclusions can be confirmed only with EM, although polyphosphate bodies can be recognized under the LM and their production can be induced under a variety of nutritional conditions (Sinclair and Whitton, 1977; Lawry and Simon, 1982). Physiological and evolutionary implications of various granules in cyanobacteria were reviewed by Jensen (1984, 1985). When observed with EM, ribosomes are also recognizable. Important intracellular inclusions, which are distinctive in many cyanobacteria, are gas vesicles, which are present in many planktonic species (e.g., Microcystis and several species of Woronichinia); their presence may be characteristic for certain genera and species. The gas vesicles (Walsby 1972, 1978) are gathered in clusters (hundreds or thousands per cell) and are visible in vegetative cells as dark brownish, irregular bodies of various sizes (aerotopes; the older term is gas vacuoles). When aerotopes are abundant, they may render the observed color of vegetative cells as brownish (due to refraction of light), which can be mistaken as the actual pigmentation. In such cases, slight or sometimes substantial pressure applied to the specimens or samples will collapse the vesicles to reveal the actual pigmentation (Walsby, 1972). This process is reversible and dependent on environmental conditions, including nutrients, light, temperature, and atmospheric pressure (Walsby, 1972, 1981; Fay, 1983). Cyanobacterial walls are mainly proteinous, with a peptidoglycan layer (electron opaque) that overlays the cytoplasmic membrane; an external sheath, as well as lipoproteins and lipopolysaccharides also may be present (Drews and Weckesser, 1982). This basic structure of cell walls is largely uniform in cyanoprokaryotes. A facultative crystalline S-layer is unique in a few unicellular and simple filamentous types; its structure differs ˇ marda, 1991; ˇSmarda et al., 1996, among genera (S 2002; Komárek, 1999). The cell surface is smooth and without sculpturing. Almost all cyanobacteria produce mucilaginous compounds that form variably structured slimy masses around cells. The form of characteristic colonies is dependent on these mucilaginous formations, the genetic stability of which is not known well. Gelatinous envelopes may be colored by various pigments (red, blue, yellow, or brown) and may be characteristic for species separation. However, the intensity and character depend on environmental conditions, particularly light and pH (Jaag, 1945). Details on the development and ultrastructure of cyanobacterial walls were reviewed by Drews and Weckesser (1982).

C. Cell Division and Reproduction Cells divide mainly by simple binary fission, in which the cell wall projects into the protoplast and the

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cell splits into two isomorphic or (less frequently) asymmetrical daughter cells. This process proceeds by one of two methods (Drews and Weckesser, 1982). The simplest (in gram-negative bacteria) is the constrictive type via simultaneous invagination (pinching) of all cell layers of the cell wall. In the other type, a septum forms from invagination of the cytoplasmic membrane and the peptidoglycan layer (septum type) with later invagination of the outer layer (cleavage). In general, cell division may occur in one, two, or three planes, which are more or less perpendicular to one another in successive generations. This process is regularly repeated and characteristic of different families. A special type of this cell division is a facultative asymmetrical binary fission in simple genera under suboptimal conditions, as seen in some members of the Synechococcaceae or simple Chamaesiphonaceae (Waterbury and Stanier, 1977; Rippka et al., 1979; Komárek and Anagnostidis, 1986; Figs. 4Ab and 17Ba), or regularly asymmetrical in the upper part of polarized cells (Stichosiphon; Montejano et al., 1997). Daughter cells usually grow to the original size (in several genera also to the original shape) before the next division. In more diversified genera, cell division is irregular, resulting in packet-like colonies (e.g., Gloeocapsopsis and Cyanosarcina), or division in one direction prevails in genera with polarized thalli (e.g., Chlorogloea, Entophysalis, and Hydrococcus) and pseudofilamentous forms (e.g., Pleurocapsa, Hyella). A more complicated type of cell division is an asymmetrical cell differentiation, in which the mother cell divides in rapid sequence in the upper part of heteropolar cells (Chamaesiphon and Chamaecalyx). Daughter cells of such distinctly and obligately polarized cells are called exocytes (exospores in early studies), which separate singly and/or successively in rows, and rarely in three-dimensional clusters from the apical cell end (Chamaesiphonaceae; Figs. 16B–18A). Exocytes have the capability for repeated divisions in a few genera (Stichococcus; Montejano et al., 1997). Another specialized type of cell division is multiple fission, in which the whole cell or a part of it (apical) divides in rapid sequence or almost simultaneously into numerous small daughter cells, which liberate from gelatinized or split envelopes and contribute to the reproduction. If the daughter cells develop from cells localized in fine, diffuse mucilage, they are called nanocytes (Komárek and Anagnostidis, 1998; Fig. 19B). Other daughter cells, termed baeocytes, originate from mother cells enclosed in firm sheaths (Waterbury and Stanier, 1978; Rippka et al., 1979; Figs. 18B, 19A, and 20). Both cell division patterns, binary and multiple fission, may be combined in several genera, where the cell division differs during growth of colonies, and

nanocytes/baeocytes predominantly contribute to the reproduction (Komárek and Anagnostidis, 1998, Table 4). In some genera, morphologically different developmental stages appear that seem to depend on environmental conditions (Microcystaceae, Chroococcaceae, and Entophysalidaceae; Jaag, 1945; Komárek, 1993). Several developmental (and dormant) stages of complicated forms are easily confused with different taxa (e.g., certain species of Asterocapsa, Gloeocapsa, and Gloeocapsopsis; Figs. 12A, and 13A and B). In morphologically complex families, especially in Hyellaceae, the thallus develops from initial cells into morphologically diverse cells in different parts of the thallus (Waterbury and Stanier, 1978; Lukas and Golubic´, 1981; Golubic´ et al., 1985; LeCampion-Alsumard and Golubic´, 1985). Reproduction in coccoid cyanobacteria proceeds by ordinary cell division in the unicellular types, by solitary cells or their small clusters liberated from colonies, by fragmentation of thalli, or by the production of exocytes, nanocytes, and baeocytes. Sexual processes have never been observed. Motility has been recognized in baeocytes and in solitary cells of several genera (Waterbury and Stanier, 1978; Waterbury and Rippka, 1989). This motility is probably facultative and its mechanism is not yet satisfactorily explained. No motility organelles have been found in any cyanobacteria.

D. Diversity and Classification The taxonomy and classification of cyanobacteria has been under study since about the middle of the 19th century using morphological and cellular criteria, similar to other microalgae (e.g., Kützing, 1849; Nägeli, 1849). However, more modern approaches in the last four decades have emphasized important structural and molecular characteristics of these bacterial organisms with plantlike metabolism (Stanier and Cohen-Bazire, 1977; Rippka et al., 1979). Diversification through ecological acclimation, adaptation and stabilization of diverse morpho- and ecotypes, as well as changes caused by mutation and possibly also genome transfers (Rudi et al., 1998), give rise to new cyanobacterial types in many habitats. Knowledge of the full diversity and evolutionary hierarchy of cyanobacterial taxonomic entities is still unclear. Thus far, DNA-based genotypes more or less correspond to traditional (morphologically based) genera, or to phenotypically distinct and definable clusters of species within known genera (Castenholz, 1992). However, the infrageneric diversity is heterogeneous, and stable traditional species that are


morphologically and ecologically well defined (e.g., Aphanothece, Chroococcus, Chamaesiphon, and Hyella) occur repeatedly in similar but distant habitats and over extended time intervals (Smith, 1950; Prescott, 1962; Whitford and Schumacher, 1969; Bourrelly, 1985). In contrast, variable taxa also exist and occur in numerous morphological forms, some of which are described as separate taxonomic species (e.g., Microcystis and Gloeocapsa). Nearly all populations of cyanobacteria from different locations differ to some degree from each other, and these deviations may stabilize in cultures (Kondrateva, 1968; Waterbury and Rippka, 1989; Kato and Watanabe, 1993). This process indicates that new forms continually develop and are stabilized under new constant conditions. Diversification within the cyanobacteria is a continuing process in which new types develop from continually modified cyanobacterial genotypes under various environmental situations. Recent data also reveal variation within cyanobacterial genomes, such as changes in the toxicity of certain strains (Neilan et al., 1997) and possible interchange of genetic material (e.g., Rudi et al., 1998). Other well-adapted forms, socalled traditional species, appear to persist over the long term and contain a wide spectrum of stable types for long periods (Komárek and Anagnostidis, 1998). Therefore, it is believed that a substantially greater diversity of cyanobacteria exists in nature than has been recognized up until now, making any current synopsis a low estimate, even with traditionally defined taxa. Arbitrary use of taxonomic names in strain designations, and in ecological and floristic studies also complicates this estimate. The present account of cyanobacteria (Chaps. 3 and 4) recognizes 53 coccoid genera (in 9 families) and 71 filamentous genera (in 16 families) from North America. Future studies will undoubtedly reveal more, especially as classification of known entities into genera, species, and other taxonomic or nontaxonomic units (e.g., strains) receives further research.

III. ECOLOGY AND DISTRIBUTION Cyanobacteria colonize nearly all habitats on Earth and have attracted the attention of ecologists for centuries. They can occur in great masses, and are important in many aquatic and terrestrial communities for their substantial biomass and primary production, N2 fixation, production of toxic compounds, creation of stromatolites, boring in limestone substrates, and importance in symbioses. Coccoid cyanobacteria contribute significantly to almost all the metabolic activities mentioned, as well as toward global carbon

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and oxygen budgets. The present review considers both coccoid and filamentous forms. For further information on the habitats and ecology of cyanobacteria, consult Fogg et al. (1973), Carr and Whitton (1973, 1982), Fay and VanBaalen (1987), Mann and Carr (1992), and Whitton and Potts (2000). Available data indicate at least some ecological specialization in all cyanobacterial taxa; no species is known to be ubiquitous across all or most freshwater environments.

A. Soils Species of cyanobacteria are widely distributed in nearly all types of soil. The species present may be specialized to particular habitats, but may still be quite broadly distributed geographically. Examples include species from the genera Synechococcus, Aphanothece, and the filamentous genera Leptolyngbya, Pseudophormidium, Phormidium, Mastigocoleus, Nostoc, and Schizothrix (Starks et al., 1981). Differences among plant communities, soil types, and local physical–chemical conditions affect the composition and abundance of algal taxa in soils. The most commonly reported factor that favors cyanobacteria in soils appears to be higher pH (Fairchild and Wilson, 1967; MacEntree et al., 1972; King and Ward, 1977). Some studies suggest that cyanobacteria are important in the early stages of primary succession processes in soils (Starks et al., 1981; Lukesˇová, 1993), especially because many species are N2 fixers, aid in stabilizing soils against erosion, or facilitate production of organic matter in new soils (Bailey et al., 1973; Jürgensen, 1973; Kubecˇkova et al., in press). Their importance appears to be greater in regions with sparse vegetation, such as deserts, semideserts, and polar regions (Cameron, 1963; Cameron et al., 1965; Forest and Weston, 1966), although more recent data suggest their abundance and productivity can also be substantial in agricultural and forest soils (Hunt et al., 1979; Shimmel and Darley, 1985). The majority of soil cyanobacteria are filamentous species, but several coccoid forms (particularly from the genus Synechococcus) are also common. Cyanobacterial assemblages are recognizable with the naked eye on wet soils, where they develop in sufficient quantity to form mats on the soil surface, or as crusts (microzones) under the surface, especially in sandy and periodically flooded soils (e.g., Eskew and Ting, 1978; GarciaPichel and Belnap, 1996), as well as on metal-contaminated soils (e.g., Maxwell, 1991).

B. Subaerial Environments Cyanobacteria are among the most important autotrophs that colonize periodically or continually

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moist soils, rocks, and walls. They are found together with lichens and mosses, which sometimes represent a later successional stages (Jaag, 1945; Golubic´, 1967a). Some cyanobacteria grow only under conditions of high humidity (aerophytic), in sites with periodic water supply (subaerial), or sites with a continual supply of water vapor (atmophytic). Their development depends on the periodicity of moisture, chemical composition of the substratum, intensity of illumination, temperature, and so forth. The composition of tropical algal communities of this kind is also distinctly different from those in temperate to polar zones (personal observation). Often, assemblages form dark stripes on rocky walls, in dripping water, and near waterfalls in mountainous areas. Tree bark may also serve as a habitat for cyanobacteria, especially in humid tropical forests (Frémy, 1930a, b; Desikachary, 1959). Our recent investigations indicate that specialized types of cyanobacteria occur in these habitats, but their taxonomy is still almost unknown. Various species from the genera Gloeocapsa, Gloeothece, Aphanocapsa, Aphanothece, Gloeocapsopsis, Asterocapsa, Chroococcus, and Chroococcidiopsis are common components of communities on wet rocks (Jaag, 1945; Komárek, 1993). The morphological variability and importance of various (possibly dormant) stages substantially complicate their taxonomic position and identification. Distinct differences in taxa from various geographically distant regions remain unclear, due to both lack of study and many taxonomic misinterpretations.

C. Aquatic Environments 1. Standing Waters A wide spectrum of cyanobacterial types occur in planktonic, benthic, metaphytic, and periphytic habitats, and the literature on phytoplankton, including cyanobacteria, based on these habitats is enormous (Bourrelly, 1966; Baker and Baker, 1979; Gibson and Smith, 1982; Whitton and Potts, 2000) and beyond the scope of this review (but see Chapters 2 and 24). Aquatic species are the best known examples of cyanobacteria, particularly as a result of their abundance and diversity in different lakes and reservoirs. Their importance typically increases with eutrophication (Pick and Lean, 1987), with accompanying surface blooms and massive planktonic biomass (Reynolds and Walsby, 1975; Paerl, 1996). Several species have worldwide distributions (e.g., from the genera Anabaena and Nodularia); others have distinctly limited distributions (several species of Aphanizomenon, Cylindrospermopsis, and others). A long list of specialized bloom-forming

nanoplanktonic and picoplanktonic species are known. Their abundance depends on a variety of ecological factors, particularly nutrient status, illumination, turbulence, and temperature. Distinct differences in environmental requirements exist between species; some occur in strictly saline habitats or oligotrophic freshwater habitats. Several species are characteristic of particular regions (e.g., temperate Gomphosphaeria aponina, tropical G. multiplex, and subpolar G. natans), but more species are known to be cosmopolitan. Some species (e.g., Aphanothece stagnina, or several picoplanktonic or water-bloom forming species) may have a specialized developmental cycle, where initial growth is in the benthos and they later become planktonic (Barbiero and Welch, 1992). Attached species may be restricted to one type of the substratum, such as limestone, submerged plants, mosses, or filamentous algae (e.g., different species of the genus Chamaesiphon). The following brief overview summarizes a few of the differences in cyanobacterial assemblages from the major habitats in which they are commonly found; other ecological details are discussed in Chapter 2.

2. Bloom-Forming Species Cyanobacteria that contain gas vesicles in their cells are well adapted to the planktonic habit and commonly occur in moderately productive to hypertrophic lakes throughout temperate and tropical regions. Their importance as biomass and toxin producers has been recognized for many years (Gorham et al., 1964; Paerl and Millie, 1996). Other cyanobacteria are responsible for taste and odor problems in surface waters (Jüttner, 1987). Specialized bloom-forming taxa also occur in coastal brackish and marine (also oceanic) waters (species of Nodularia and Trichodesmium). Of the coccoid forms, the genus Microcystis is one of the most widely reported (although taxonomically difficult), and typically forms blooms in the spring in temperate zones and may persist throughout the summer in eutrophic lakes (Reynolds et al., 1981; Doers and Park, 1988). Several species occur in tropical regions over the entire year. These species establish massive surface blooms (“hyperscums”) under eutrophic conditions, low turbulence (calm winds), and high irradiance (Zohary and Breen, 1989). Microcystis blooms can represent more than 50% of the total algal biomass in some lakes (Fallon and Brock, 1981; Zohary and Robarts, 1990), and some strains produce hepatotoxins (Carmichael, 1992, 1997; Codd, 1995; Chorus and Bartram, 1999) that can inhibit herbivorous zooplankton (Fulton and Paerl, 1987; Hietala et al., 1997). Other planktonic coccoid genera contribute to water blooms, such as species of Coelosphaerium, Snowella, and Woronichinia (Komárek and Komárková-Legnerová,


1992). There is a large literature on bloom-forming cyanobacteria (reviewed in Reynolds and Walsby, 1975; Paerl, 1996), but in many individual studies, the characteristics of species and populations are often incompletely documented. Further details concerning filamentous forms are discussed in Chapter 4.

3. Picoplanktonic and Nanoplanktonic Species Many other species of coccoid cyanobacteria that lack gas vesicles are well adapted to a planktonic mode of life. Several live as solitary cells and are very small (ca. 1–2 µm diameter); these members of the picoplankton are important or even dominant primary producers in oligotrophic waters (Fahnenstiel et al., 1986; Fogg, 1986; Stockner and Shortreed, 1991; Burns and Stockner, 1991; Corpe and Jensen, 1992) and occasionally in more productive systems (Platt and Li, 1986; Wehr, 1990; Weisse, 1993; Komárková, 2001). Their importance is typically greater in large oligotrophic or mesotrophic lakes and the open ocean, and at moderate or greater depths (Eguchi et al., 1996; Waterbury, 1979; Komárková and Sˇimek, 2003). Some of these tiny species are superior competitors for P (Suttle and Harrison, 1988; Wehr, 1989), and some populations may also be adapted to low light regimes or altered spectra (Hauschild et al., 1991; Wehr, 1992; Callieri et al., 1995). Picoplanktonic (unicellular) species have most commonly been assigned to the genus Synechococcus (some of them perhaps erroneously), but the genera Cyanobium and Synechocystis may be equally or more important (Stockner, 1988; Albertano and Capucci, 1997; ˇSmajs and ˇSmarda, 1999; Komárek, 1999). Other coccoid species form microscopic mucilaginous colonies that passively float in the water (nanoplankton). Nanoplanktonic species are common in mesotrophic and eutrophic waters, but rarely produce a substantial biomass. Common coccoid nanoplanktonic species include the genera Chroococcus, Cyanodictyon, Aphanothece, Rhabdoderma, Aphanocapsa, Merismopedia, Coelosphaerium, Coelomoron, Snowella, and Gomphosphaeria (Komárek and KomárkováLegnerová, 1992; Komárek and Anagnostidis, 1998). A few species are more common in waters with higher salinity (Pannus spumosus and several Aphanothece species), and appear to be halotolerant by means of osmoregulatory compounds (Reed et al., 1986).

4. Benthic and Epiphytic Species Benthic cyanobacteria occur widely in lentic and lotic ecosystems, in epipelic, epipsammic, and epilithic habits or are epiphytic on filamentous algae, mosses, and vascular plants. Benthic mats develop on sediments in standing waters and later may form floating mats (Aphanothece, Oscillatoria, and Phormidium). Many are specialized with respect to substratum and habitat

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(see Chap. 2). For example, some benthic cyanobacteria colonize specific types of rock (limestone, sandstone, granite). Characteristic communities with a dominant cyanobacterial flora develop in mountain streams, particularly in limestone areas, where mainly filamentous species form characteristic microzonation on stones (Geitler, 1932). Similar communities occur in splash zones of larger lakes, and include the coccoid genera Chamaesiphon subg. Godlewskia, Chlorogloea, and Hydrococcus, plus filamentous species of Homoeothrix, Schizothrix, Phormidium, Calothrix, and Rivularia (Golubic´, 1967a). Many epilithic species form distinctly colored macroscopic patches on stones, particularly species of Chamaesiphon (Geitler, 1932; Kann, 1973). Epiphytic communities typically include other species of cyanobacteria, such as members of the coccoid genera Stichosiphon, Chamaesiphon, Chamaecalyx, Cyanocystis, and Xenococcus, plus many filamentous species in the genera Leptolyngbya, Heteroleibleinia, Hapalosiphon, Cylindrospermum, Trichormus, and Oscillatoria, among others.

5. Transported Plankton in Rivers and Streams A number of cyanobacterial species occur in the potamoplankton of large streams and rivers, although their relative importance (and diversity) appears to be less than centric diatoms or coccoid green algae (Wehr and Descy, 1998). Among those commonly reported are species of Synechococcus, Chroococcus, Pseudanabaena, Planktothrix, and Oscillatoria (Reynolds and Descy, 1996). Current data suggest that, in general, their importance is greater in slower flowing and warm–temperate to tropical rivers (especially those with backwaters), where longer residence times permit their slower growth rates to attain greater population sizes (Reynolds and Descy, 1996). Well-studied examples include the Neuse River, North Carolina (Paerl and Bowles, 1987) and the Darling River, Australia (Hötzel and Croome, 1994), both of which receive large quantities of nutrients. Summertime cyanobacterial blooms (e.g., Microcystis, Anabaena, and Aphanizomenon) have also been reported periodically in the upper Mississippi and Potomac Rivers over at least five decades (Reinhard, 1931; Baker and Baker, 1981; Krogman et al., 1986). Some data suggest that picoplanktonic species may be quite numerous in large rivers, but apparently are not dominant in terms of biomass (Wehr and Thorp, 1997). The many studies over the last 100 years suggest that few, if any, species reported are strictly potamoplanktonic forms (Wehr and Descy, 1998); indeed many genera and their species (e.g., species of Anabaena, Chroococcus, and Oscillatoria) are also commonly observed in lakes. Species composition often depends on tributaries and especially

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on plankton from adjacent lakes or reservoirs (Reynolds and Descy, 1996), and is more dependent on the planktonic cyanobacterial flora typical for that region.

6. Wetlands Cyanobacteria are well represented in the metaphyton and periphyton of swamps, temporary pools, and bogs. Many such systems possess a rich cyanobacterial flora, especially alkaline systems, where species of Chroococcus, Aphanothece, Leptolyngbya, Phormidium, Lyngbya, Microcoleus, Scytonema, and Schizothrix are frequently reported, many of which become encrusted with CaCO3 (Browder et al., 1994; Goldsborough and Robinson, 1996). This flora has been described from alkaline wetlands in the Caribbean and Cuba (Gardner, 1927; Schiller, 1956), the Florida Everglades (McCormick and Stevenson, 1998), and coastal wetlands in Belize and Mexico (Rejmánková et al., 1996). Some of these communities contain cyanobacterial species that have been recognized as possibly endemic to the region. In contrast, a number of species from the genera Cyanothece, Aphanothece, Rhabdoderma, Merismopedia, Eucapsis, and Chroococcus are known mainly from acidic peat or salt swamps. Among the best studied wetland cyanobacteria may those that colonize rice fields, because many are important N2 fixers (Whitton, 1992). Species from the littoral regions of mesotrophic or eutrophic ponds and lakes are less specific, and often represent dislodged benthic forms or accumulations of entrained algae from pelagic regions.

Mastigocladus (M. laminosus), and Phormidium. These assemblages often develop an extremely large biomass, which often forms layered mats of various colors (Castenholz and Wickstrom, 1975). Many species experience not only extreme temperatures, but also high levels of sulfide and UV exposure (Castenholz, 1976; Miller et al., 1998).

2. Saline Inland and Coastal Habitats

Cyanobacteria are the main and often sole autotrophic organisms in many extreme environments, which include thermal, saline, arid, and endolithic habitats.

Cyanobacteria adapted to high salinity can often be the main autotrophs in hypersaline environments (Setchell and Gardner, 1905; Gardner, 1906; Taylor, 1928; Frémy, 1933; Feldman, 1958; Golubic´, 1980; Humm and Wicks, 1980; Komárek and Lukavsky´, 1988; Montoya and Golubic´, 1991; Anagnostidis and Pantazidou, 1991; Roussomoustakaki and Anagnostidis, 1991). Pools, lakes, and swamps with high salt concentrations that periodically dry and often contain unusual combinations of salts (sulfates, Mg salts, and sulfides; see also Chap. 2, Sect. VI.A), represent very specialized habitats for coccoid and filamentous cyanobacterial species. Lakes in desert regions, some volcanic lakes, and coastal pools (with periodic marine influences) belong to these environments. Some of the exclusively halophilic species are members of coccoid genera Aphanothece, Cyanothece, Merismopedia, Stanieria, and Synechocystis (Golubic´, 1980; Anagnostidis and Pantazidou, 1991; Roussomoustakaki and Anagnostidis, 1991; Palinska et al., 1996) or species of filamentous genera Arthrospira and Nodularia. Many of the species regarded as halophiles have been found to be salt-tolerant, and their occurrence in saltwater environments is often influenced by their acclimation to these conditions and less by competition with other phototrophic organisms (Komárek and Lukavsky´, 1988). Mechanisms of acclimation, and development of halophilic species of cyanobacteria has been reviewed by Golubic´ (1980).

1. Thermal Waters

3. Arid Environments

The average upper temperature limit for plant existence is slightly above 40°C, but species adapted to about 70°C exist among cyanobacteria (Castenholz, 1969a, b, 1977; see also Chap. 2, Sect. V). Although modern taxonomic studies are sorely needed for thermal cyanobacterial species, North American hot springs belong to the best known thermal sites in the world. Several coccoid forms in Yellowstone National Park were described more than five decades ago by Copeland (1936), who described many of the major representatives of high-temperature cyanobacteria, including species of Synechococcus (S. lividus and S. vulcanus) and Cyanobacterium (C. minervae). A number of important filamentous cyanobacteria in hot springs include species of Leptolyngbya (several species),

Several cyanobacteria live as endolithic species inside rocks (within the crystal lattice) in very arid, hot or cold deserts. Three types are recognized: (1) euendoliths, which bore into rocks (see the next section), (2) chasmoendoliths, which occur in fissures and cavities, and (3) cryptoendoliths, which colonize fractured or porous rocks, usually forming layers parallel to the rock surface (Whitton and Potts, 1982; Friedmann and Ocampo-Friedmann, 1984). The crypto- and chasmoendoliths are unique phototrophic organisms that occur mainly in very arid, hot or cold deserts. Their presence is dependent at least on traces of periodic humidity, but they represent some of the most interesting forms of autotrophic, oxygen-evolving life on Earth (Friedman, 1971, 1980; Friedman and Ocampo,

D. Extreme Environments


1985; Friedman and Ocampo-Friedman, 1984; Golubic´ et al., 1981; Büdel and Wessels, 1991; Büdel et al., 1994; Johansen and Rushford, 1985; Johansen, 1993; Johansen et al., 1993; Kubecˇková et al., 2003). Species of the genus Chroococcidiopsis are among the most commonly reported organisms in this unusual habitat.

4. Endolithic and Lithogenic Aquatic Habitats A specialized and unique ecological group of cyanobacteria occurs in both marine and freshwater regions that contain euendolithic and lithogenic species (Golubic´ et al., 1975). The boring types are known mainly from marine rocky littoral zones (Cyanosaccus sp. div., Hyella balani, H. tenuior, Scytonema endolithicum, and others), but a few species are known also from freshwaters. In contrast, under subaerophytic and submersed freshwater conditions and in submersed marine habitats, many species participate in travertine and stromatolite formation (Golubic´ et al., 1975, 1981). A few species of the coccoid genera (Chroococcidium, Bacularia, Chlorogloea, and Entophysalis) were described from such habitats (Geitler and Ruttner, 1935; Copeland, 1936; Komárek and Montejano, 1994).

E. Geographic Distribution In accordance with their considerable morphological diversity, many of the genera and species of cyanobacteria that have been described are considered to be distributed in many locations worldwide (Whitton, 1992). However, the geographic distribution of cyanbacterial species is nonetheless dependent on their ecological requirements. Some species, which are almost cosmopolitan in their distribution, occur in distinctly specialized habitats; hence, even very specialized taxa (e.g., species from hot springs) can still be cosmopolitan among suitable habitats. Other species are known to tolerate a wide range of special ecological factors (e.g., temperature and salinity), but are quite sensitive to other environmental factors (e.g., halophilic Nodularia harveyana and Arthronema africanum; Komárek and Lukavsky´, 1988; personal observation). A consistent pattern of tolerances to and requirements of ecological factors does not exist for cyanobacteria; therefore, the geographic distribution of various species and forms often varies. Data suggest there are many very specialized types, the occurrence of which is restricted to small defined habitats and localities; several of these distributions are described in Section V.B. Notable examples can be found in thermal springs, certain tropical habitats, volcanic lakes, and near marine coastal regions. For example Chlorogloea lithogenes is known from central Mexico, and Bacularia indurata

67

from Yellowstone National Park, Mantellum rubrum from volcanic lakes in Mexico, Aphanothece bacilloidea and Gomphosphaeria semen-vitis from alkaline swamps in Central America. Each appears to occupy restricted habitats (Gardner, 1927; Copeland, 1936; Komárek and Montejano, 1994; Tavera and Komárek, 1996). There are, however, forms with broader ecological requirements, but restricted geographic distribution. For example, a number of freshwater planktonic (or metaphytic) species have been confirmed only from certain locations in North America, such as Coelomoron regulare, Merismopedia smithii, Woronichinia klingae, Gomphosphaeria natans, Snowella rosea, Chroococcus multicoloratus, and Stanieria cyanosphaera (see the ecological data in Komárek and Anagnostidis, 1998). Some species occur more widely, but with morphologies that are phenotypically uniform in one area and variable in another (including more morphotypes). Examples of variable populations (in comparison with other regions) are Woronichinia naegeliana in Canadian lakes (Komárek and Komárková-Legnerová, 1992) and Microcystis wesenbergii in Northern Europe (Cronberg and Komárek, 1994). For these reasons, use of identification keys based on microflora from other regions (especially Europe) for North American (and especially tropical) cyanobacteria, should be used with utmost care.

IV. COLLECTION, PREPARATION, AND CULTURE Samples of natural cyanobacterial populations should be collected by methods widely used for other microalgae (e.g., Prescott, 1962; Whitford and Schumacher, 1969). For example, plankton nets (5, 10, 20, or rarely 45 µm) are well suited for most planktonic and metaphytic species. However, small nano- and picoplanktonic forms would be undersampled using nets and generally require collection of whole water samples, which can be concentrated by sedimentation or centrifugation. Epiphytic, epilithic, and endolithic species can be collected with parts of the substrata whereas parts of massive macroscopic colonies and mats can be collected directly into tubes of different sizes. The exact documentation and description of samples is an important requirement. The study of living material is preferred for species identification. Fixation with 2% formalin (final concentration) is preferred for most samples, because higher concentrations produce artifacts in the mucilaginous sheaths, and distort cell size and shape (Komárek, 1958). Lugol’s solution, commonly used in limnological studies for quantitative estimates (e.g., Utermöhl, 1958), is not recommended for taxonomic purposes, due to disintegration of colonies, changes in color, and distortions

68

Jirˇí Komárek

in sheaths or mucilage. Glutaraldehyde (to 2% concentration) is recommended for EM examination. In most cases, drying cyanobacterial samples is not recommended (especially for delicate forms), although some old herbarium specimens could be usable for comparisons based on general morphology and DNA-based methods. Periodically dried preparations (i.e., slides usable for microscopic treatment) are not commonly used, but may be used over longer periods if specimens are first preserved with formalin or glutaraldehyde. Isolation and culture is important for cyanobacterial taxonomy. Methods used to culture freshwater (and terrestrial) algae and cyanobacteria are described in several general treatises (Pringsheim, 1946; Venkataraman, 1969; Carr and Whitton, 1973). However, most species have yet to be isolated into pure culture. Obtaining cultures is particularly difficult for ecologically specialized forms, such as thermal, endolithic, and even some bloom-forming species (see the lists of strains in world collections). However, problems also exist for many common planktonic (e.g., Aphanothece, Lemmermanniella, Cyanodictyon, Woronichinia, and Microcystis) and benthic species (e.g., Chamaesiphon species from mountain streams) that have not yet been studied in culture. Another complication is that in culture, many species lose the characteristic colony shape or change cell size, making it difficult to link strains with known field populations (Van Baalen, 1965; Komárek, 1976). Much useful information could be gained in future studies by carefully recording morphologies and phenotypic variation of field-collected cyanobacteria (under natural conditions) prior to isolation. Currently cultures that contain similar, small, rodlike cells may belong to species or forms with different colony forms in nature; hence, standardization and simplification of media and growth conditions are not able to clarify different accounts of cyanobacterial morphology. There is a special problem with identification and designation of experimental strains that have unknown field populations. Strains should be identified before

isolation or during the course of the isolation process. Based on the collective experience in our laboratory, in more than 50% of the cyanobacterial strains from the world collections, the taxonomic name on the label was a different (nomenclatural or taxonomic) type (according to both botanical and bacteriological codes) than what we found inside the tube. Unfortunately, many experimental scientists (e.g., physiologists using a strain for particular enzyme assays or systematists measuring phylogenetic affinities) do not revise the phenotype identification. This can lead to numerous discrepancies in morphological and molecular data in modern cladograms, simply due to incorrectly identified or labeled strains. Specialized collections with well-defined strains (e.g., National Institute for Environmental Studies – NIES in Japan), are rare. The designation of strains may also lead to confusion. Several proposals (Komárek, 1969; Lhotsky´ and Komárek, 1981) aimed to unify methods for strain designations, but a single unified method was never accepted and each collection has its own code. Many strains thus occur in the literature under different numbers, for example, Nostoc sp. (strain NIVA-CYA 246) and Anabaena sp. (strain PCC 7120), which are derived from the same original strain (their identity was recently proved by molecular sequencing). It is recommended that researchers use at least the citation of the acronym of the collection (prescribed by the International Committee of Strain Collections; Garcia Reina, 1997; Sugawara and Miyazaki, 1999) and the full strain number of the corresponding collection. Despite these problems, the use of cultures is quite necessary for modern cyanobacterial taxonomy. Molecular analyses and almost all ultrastructural studies are impossible without defined strains, and several cyanobacterial types (picoplanktonic species) are not recognizable or identifiable without culturing. However, new isolates should always be added to collections with documentation and characterization of the original (natural) material, its phenotype features, and ecological parameters.

V. KEY AND DESCRIPTIONS OF GENERA A. Key2 1a.

Cells solitary (rarely aggregated in groups), heteropolar, with distinct basal and apical ends, usually attached by one (basal) end to the substratum (plants or stones); family Chamaesiphonaceae ……………………....………………………………….………………..31

1b.

Cells of different shape, solitary or in various colonies, but not distinctly diversified into basal (attached) and apical ends (cells may be polarized within colonies) ……………………………....…………………....…………………....………………….....…………...……2

2

Note: Cyanobacterial genera are characterized mainly by molecular sequencing and cell ultrastructure in modern taxonomy. It is sometimes difficult to find a single morphological or phenotypic feature that is diagnostic. Therefore, it is necessary to compare the entire set of characteristics from the key for identification.


69

2a.

Cells widely oval to cylindrical (rodlike), which divide exclusively perpendicular to one (usually longer) cell axis; family Synechococcaceae ...…………………………………………....…………………....…………………....…………………......….………....3

2b.

Cells spherical, hemispherical, ovoid, irregularly elongated, polygonal–rounded, or of irregular outline, dividing in two or more planes..……………………………………………………………………………………………………………………………………….....16

3a.

Cells solitary (or agglomerations of solitary cells)…………………………….………….…………………....…………………....……….4

3b.

Cells united regularly within mucilaginous colonies of various kinds...…………………………………………………………………....7

4a.

Cells cylindrical to long rodlike; cell content more or less homogeneous with only solitary granules; chromatoplasm sometimes barely visible (Fig. 4A)……………………………………………………………………………………………..........………Synechococcus

4b.

Cells widely oval to wide cylindrical….……………………………………………………………………………….………..……………..5

5a.

Cells with differentiated centro- and chromatoplasm, 0.4–4.5 (up to 10) µm long (Fig. 1A) ……………...........……….....Cyanobium

5b.

Cells usually with lengthwise striated or reticulate content, longer than (2)3.7 µm (up to 30–100 µm long) ………….......………….6

6a.

Cells with lengthwise striated content (special thylakoid arrangement), 2–30 µm long (Fig. 3B)………......................Cyanobacterium

6b.

Cells with reticulate content (radial arrangement of thylakoids), 6.2–40 (100) µm long (Fig. 5B)......................................Cyanothece

7a.

Cell length/width ratio = 1 to 2(3):1; colonies spherical or formless...................................................................................................8

7b.

Cells more than (2)3 times longer than wide; colonies irregular or distinctly elongated ...................................................................13

8a.

Colonies CaCO3 encrusted; component of travertine formations (not pictured)….……..…………........................…...……Lithomyxa

8b.

Colonies not CaCO3 encrusted; not in travertine formations..............................................................................................................9

9a.

Cells arranged only in microscopic, mucilaginous, spherical or irregular netlike colonies; at least partly in rows.............................10

9b.

Cells within or on the surface of irregularly spherical colonies or in amorphous gelatinous colonies; not in rows............................11

10a.

Colonies more or less spherical or slightly elongated, with cells clustered in the center and in short, radiating rows at the colonial periphery (Fig. 1C).............................................................................................................................................................Radiocystis

10b.

Colonies irregular–reticulate, composed of mucilaginous strands; cells more or less in not radiating rows (Fig. 1B).......Cyanodictyon

11a.

Oval cells situated on the surface of mucilaginous spheres (Fig. 2A)..........................................................................Epigloeosphaera

11b.

Cells inside of micro- to macroscopic gelatinous colonies, sometimes enveloped by their individual, sometimes lamellated envelopes.........................................................................................................................................................................................12

12a.

Cells entirely (or at least central cells) without individual envelopes (Fig. 2B)...............................................................Aphanothece

12b.

All cells with individual, occasionally lamellated envelopes (Fig. 3A).................................................................................Gloeothece

13a.

Cells cylindrical or fusiform, always longer than wide, randomly distributed within colonies (occasionally regularly oriented along longer axes in one direction); colony usually elongated with cells not in pseudofilamentous rows...................................................14

13b.

Cells shorter than long (cells divide perpendicular to shorter axis of cells), arranged in uniseriate pseudofilamentous rows, forming elongated mucilaginous colonies (Fig. 6A)................................................................................................................Johannesbaptistia

14a.

Colonial mucilage indistinct, colorless, diffuse; cells distant from one another.................................................................................15

14b.

Colonial mucilage distinctly limited, sometimes encrusted; cells arranged clearly and usually ± densely in one direction (Fig. 5A)................................................................................................................................................................................Bacularia

15a.

Cells cylindrical with rounded ends (Fig. 4B).................................................................................................................Rhabdoderma

15b.

Cells fusiform or with tapering cell ends (Fig. 4C)...........................................................................................................Rhabdogloea

16a.

Cells divide exclusively in two directions in successive generations; cells solitary or in flat to nearly spherical colonies; family Merismopediaceae............................................................................................................................................................................17

16b.

Cells divide in three or more directions (or irregularly) in successive generations, cells in colonies of various forms........................27

17a.

Solitary cells, always spherical (Fig. 6B)..........................................................................................................................Synechocystis

17b.

Cells in mucilaginous colonies; cells spherical or slightly elongated (oval, ovoid).............................................................................18

18a.

Colonies distinctly platelike (cells arranged more or less in one flat layer)........................................................................................19

18b.

Colonies three dimensional, irregular, or more or less spherical……................................................................................................22

19a.

Cells spherical, in perpendicular rows in colonies………..................................................................................................................20

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Jirˇí Komárek

19b.

Cells slightly but distinctly elongated, in short rows or irregularly arranged....................................................................................21

20a.

Cells in free-living, platelike colonies (Fig. 7A)...............................................................................................................Merismopedia

20b.

Flat colonies attached (monolayer), lying flat on substrata (Fig. 7B)...................................................................................Mantellum

21a.

Colonies always only few-celled, longer axis in the plane of colony; cells oval to ovoid (Fig. 7C).....................................Cyanotetras

21b.

Old colonies many-celled, longer axis perpendicular to plane of the colony; cells oval to almost rodlike (Fig. 8A)............Microcrocis

22a.

Colonies amorphous, irregular; cells irregularly (sometimes densely) arranged (Fig. 6C).................................................Aphanocapsa

22b.

Colonies more or less spherical; cells arranged peripherally and sometimes radially near the surface of mucilaginous spheres.........23

23a.

Colonial mucilage of older (larger) colonies more or less homogeneous; without visible gelatinous stalks (not revealed by any stain)................................................................................................................................................................................................24

23b.

Within colonies develops a gelatinous system of stalks; cells or cell groups joined to the ends of stalks (revealed sometimes only by stain)................................................................................................................................................................................................25

24a.

Cells spherical, arranged peripherally; colonies spherical (rarely irregular), mucilaginous (Fig. 8B)............................Coelosphaerium

24b.

Cells slightly elongated, arranged radially in the marginal parts in more or less spherical colonies, or several spherical colonies joined together (Fig. 8C)...................................................................................................................................................Coelomoron

25a.

Stalks thin, stringlike, pseudodichotomously divided, joined in the center to form a widened gelatinous cluster; cells more or less distant from one another (Fig. 9A).........................................................................................................................................Snowella

25b.

Stalks thick, not stringlike................................................................................................................................................................26

26a.

Stalk system composed of fine, gelatinous, parallel tubes radiating from center (phase contrast or staining); each stalk bears radially-arranged spherical or oval cells; cells may form dense peripheral layer in old colonies (Fig. 9B).................................Woronichinia

26b.

Stalks thick, divided and diffuse; widened at the ends and enveloping peripherally arranged cells; cells more or less obovoid and remain connected after division, often forming pairs of cordiform cells (Fig. 10A)...................................................Gomphosphaeria

27a.

Cells spherical, dividing regularly in three perpendicular planes in successive generations; family Microcystaceae...........................28

27b.

Cells hemispherical, polygonal, elongate, or irregular, dividing usually in three or more variously oriented planes..........................37

28a.

Cells arranged distinctly in three-dimensional, cubical colonies, in more or less regular, perpendicular rows (Fig. 10B).........Eucapsis

28b.

Cells not arranged in a cubical manner; colonies are irregular, sometimes forming amorphous masses............................................29

29a.

Cells without distinct individual sheaths, with aerotopes (groups of gas vesicles); planktonic colonies irregular in shape with irregularly (sometimes densely) scattered cells (Fig. 11)...............................................................................................................Microcystis

29b.

Cells (or small groups) with individual sheaths (may be colored), without aerotopes; subaerial, metaphytic, or benthic irregularly shaped colonies with irregularly arranged cells................................................................................................................................30

30a.

Cell groupings within colonies usually spherical, with more or less widened, spheroidal gelatinous envelopes; mainly subaerial species (Fig. 12A)...............................................................................................................................................................Gloeocapsa

30b.

Cell groupings within colonies in dense polyhedral clusters; mainly submersed species (Fig. 12B).................................Chondrocystis

31a.

Sessile cells reproduce by exocytes (daughter cells separate asymmetrically and individually, or remain in rows or clusters at the apex of the mother cell); family Chamaesiphonaceae.......................................................................................................................32

31b.

Sessile cells reproduce by baeocytes (formed from successive or almost spontaneous divisions of the whole mother cell; baeocytes disperse from gelatinized or ruptured sheaths); family Dermocarpellaceae.......................................................................................36

32a.

Sessile cells rodlike, without gelatinous sheaths; attached to the substratum by a small mucilaginous pad (revealed with staining or EM sections) (Fig. 16B)................................................................................................................................................Geitleribactron

32b.

Sessile cells always with firm gelatinous sheath (sometimes very thin and indistinct, or visible only during exocyte liberation)........33

33a.

Thin sheath (may be obscure) of vegetative cells narrowed at apex into conspicuous, short or long, hairlike projection (Fig. 16C)............................................................................................................................................................................Clastidium

33b.

Sheaths of vegetative cells without apical hairlike formations..........................................................................................................34

34a.

Cells (more or less cylindrical) produce long rows of exocytes that are retained within the sheath of the mother cell, forming pseudofilamentous formations (Fig. 17A).........................................................................................................................Stichosiphon

34b.

Cells (usually oval, ovoid to cylindrical) do not form long, pseudofilamentous rows of exocytes.....................................................35

35a.

Exocytes separate individually or arranged at the apex of the mother cell in a ± short single series (Fig. 17B) ...................................................................................................................................................Chamaesiphon subg. Chamaesiphon

35b.

Exocytes arranged apically in three-dimensional clusters (Fig. 18A)................................................................................Chamaecalyx


71

36a.

The first division plane is vertical during successive divisions of the mother cell into baeocytes (Fig. 18B)........................Cyanocystis

36b.

The first division plane into baeocytes is horizontal (Fig. 18C).....................................................................................Dermocarpella

37a.

Cells spherical to subspherical, gathered in small agglomerations without slime production; cell division solely into baeocytes: entire whole mother cell divides into numerous small daughter cells, which liberate from a ruptured sheath; family Dermocarpellaceae (Fig. 19A)...............................................................................................................................................................................Stanieria

37b.

Cells of variable shape, forming irregular colonies with slime production or a differentiated thallus; cell division irregular in various planes in exocytes or partially in one direction, sometimes combined with baeocyte production; several types of division are combined in some cases..........................................................................................................................................................................38

38a.

Cells within colonies divide only by binary fission, never by exocytes or baeocytes.........................................................................39

38b.

Cells within colonies divide by binary fission and via specialized cells, by exocytes or baeocytes.....................................................46

39a.

Colonies irregular, never polarized; all cells similar in character; family Chroococcaceae.................................................................40

39b.

Colonies often polarized and slightly differentiated, cells often with different shape in basal, central, and apical regions of colony..............................................................................................................................................................................................44

40a.

Cells with narrow, firm sheaths; gathered into dense packets with numerous cells...........................................................................41

40b.

Sheaths more widened, forming distinct gelatinous layers around cells; colonies usually few-celled (but sometimes clustered together)..........................................................................................................................................................................................42

41a.

Packets of cells with more or less cubical (or sarcinoid) arrangement; sheaths colorless (Fig. 14B).................................Cyanosarcina

41b.

Packets of clustered and ensheathed cells irregularly arranged; sheaths often colored (Fig. 13B)..................................Gloeocapsopsis

42a.

Sheaths (fine or firm) around cells often spheroidal and colored, and often with surface structure (granular or warty) (Fig. 13A)..........................................................................................................................................................................Asterocapsa

42b.

Sheaths usually fine, colorless, delimited or diffuse, in vegetative stages without surface structures.................................................43

43a.

Cells usually arranged in three-dimensional, spheroidal clusters (Fig. 14A)......................................................................Chroococcus

43b.

Cells arranged in very short, parallel rows, slightly distant from one another (Fig. 13C)..................................................Cyanokybus

44a.

Colonies slightly polarized, three-dimensional, forming large mucilaginous clusters on various substrata; family Entophysalidaceae ........................................................................................................................................................................................................45

44b.

Colonies more or less flat, forming discoidal, often one- or few-layered biofilms on substrata, growing radially, with morphologically different central and marginal (apical) cells; family Hydrococcaceae (Fig. 16A)...........................................................Hydrococcus

45a.

Central and apical cells usually spherical, in marginal or old colonial parts with individual gelatinous envelopes; mucilage mainly colorless, rarely colored (Fig. 15A)....................................................................................................................................Chlorogloea

45b.

All cells ± irregular, most (or small groups) with individual gelatinous sheaths, which are usually colored (Fig. 15B)......Entophysalis

46a.

Colonies multilayered, ± polarized, attached to substratum; most cells divide in apical areas as exocytes (remain attached to open sheaths of the mother cell), also by binary fission; family Chamaesiphonaceae (Fig. 17B) .........................................................................................................................................................Chamaesiphon subg. Godlewskia

46b.

Cells divide by binary fission and by baeocytes (rarely by nanocytes), never by exocytes.................................................................47

47a.

Thallus not distinctly polarized, usually three-dimensional, free-living or attached to substrata; baeocytes arise from specialized cells developing in parts of a colony; family Xenococcaceae…………………………………………………………………….......…..……...48

47b.

Thallus polarized (sometimes very short with few cells) with pseudofilamentous structure (epiphytic, epilithic, sometimes boring forms); baeocytes arise from specialized basal (old) or apical cells; family Hyellaceae......................................................................53

48a.

Cells of different size in one colony; cell groupings enveloped by fine, diffuse, gelatinous envelopes (Fig. 19B)..........Chroococcidium

48b.

Cells more or less uniform in size in one colony; cells enveloped by thin, but firm, distinct sheaths.................................................49

49a.

Colonies free-living, not directly attached to substrata but sometimes occurring in epiphytic or endolithic assemblages; three-dimensional but without any polarized cells...........................................................................................................................50

49b.

Colonies attached to substrata; at least portions of cells in colonies have distinct polarity...............................................................52

50a.

Packet-like or sarcinoid colonies, sometimes gathered together, with irregular cells (Fig. 20B).........................................Myxosarcina

50b.

Cells in irregular agglomerations, not forming packet-like colonies; cells subspherical.....................................................................51

51a.

Most cells of similar, more or less uniform spheroidal shape; only young and old cells differ in size; cells never in rows (Fig. 20A) ................................................................................................................................................................................Chroococcidiopsis

72

Jirˇí Komárek

51b.

Cells in colonies are different in size, sometimes arranged in short, irregular rows (Fig. 21B)......................................Chroococcopsis

52a.

Cells form a monolayer on substrata; a few may reproduce by baeocytes (Fig. 21A).........................................................Xenococcus

52b.

Cells in two or more layers on substrata, may form irregular hemispherical colonies; baeocytes produced from randomly arranged mother cells at periphery of colony (Fig. 21C)....................................................................................................................Xenotholos

53a.

Pseudofilaments erect, arising parallel from basal cells that creep along substrata; baeocytes occur in apical, enlarged cells (Fig. 22A)...............................................................................................................................................................................Radaisia

53b.

Pseudofilaments creeping along substrata, some are endophytic or endolithic; baeocytes do not arise from apical cells...................54

54a.

Pseudofilaments only creeping along substrata, never boring; baeocyte-forming cells arise in various parts of a thallus (Fig. 22B)….......................................................................................................................................................................Pleurocapsa

54b.

Pseudofilaments distinctly polarized, creeping on substrata and/or boring into limestone, or growing into the intercellular spaces of host plants (marine seaweeds); baeocytes arise from cells in oldest parts of thallus (Fig. 23)......................................................Hyella

B. Descriptions of Genera3 Synechococcaeae (Figs. 1–6A) Aphanothece Nägeli (Fig. 2B) Colonies are microscopic or macroscopic (up to several centimeters in diameter), multicellular, more or less spherical or mostly amorphous, greyish, greenish, blue–green, or brownish colored, with irregularly, sometimes densely arranged cells, which are embedded by colonial mucilage. Two subgenera are defined: (i) Anathece—cells in diffuse mucilage, without individual envelopes—and (ii) Aphanothece—mucilage usually distinctly delimited, and cells (especially marginal) enveloped facultatively within individual sheaths or envelopes. Cells are widely oval to cylindrical, straight or slightly arcuate, pale greyish blue–green, green to bright blue–green (rarely reddish), usually with apparent chromatoplasm in larger cells. Gas vesicles and aerotopes are present in some planktonic species. Reproduction by transversal binary fission of cells, which is perpendicular to the longer cell axis. A common genus with about 60 species (based on morphology) from diverse environments, but numerous morpho- and ecotypes are not identifiable. Ecological information is important for species identification, yet distribution data for various species is uncertain due to difficulties with identification and numerous misinterpretations in the literature. The genus is common in many North American environments (Prescott, 1962; Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979), with numerous species from wet rocks (A. castagnei, and A. saxicola), soils (A. pallida), benthos 3

Figures are largely based on North American material and literature. However, the documentation of several species had to be selected from non-North American papers. Localities are cited only in figures from this continent. Schemes of cell division, life cycles, and colony formations are selected according to various authors from Komárek and Anagnostidis (1998).

of ponds and pools (A. microscopica, A. nidulans, A. stagnina, and A. uliginosa), and thermal springs (A. bullosa, and A. thermalis), but many records need revision. Several species (possibly endemic and ecologically restricted) occur in alkaline swamps in subtropical and tropical areas, including Florida (Everglades), Cuba (Zapata peninsula, Ciénaga de Lanier), Belize, and Mexico (Yucatán). These species include A. bacilloidea, A. cylindracea, and A. opalescens (Gardner, 1927; Komárek, 1995). One tropical species with aerotopes, A. conglomerata, occasionally is present in surface blooms in reservoirs in Florida (original unpublished data). Some species colonize coastal and inland brackish environments, for example, A. utahensis and A. karukerae (Caribbean), and a few other species that require revision. Many species from the subgenus Anathece are common components of lake phytoplankton; A. minutissima, A. bachmannii, A. clathrata, and A. smithii are known from mesotrophic Canadian lakes (original unpublished data, H. Kling, personal communication). Bacularia Borzì (Fig. 5A) Cylindrical cells are arranged more or less in parallel (lengthwise, in one direction) in narrow, filamentous or tubular, elongated mucilaginous colonies, tapering and pointed or open at both ends. Cells are slightly distant, but not in rows or in pseudofilaments. The mucilage is fine, homogeneous, colorless, and usually distinctly delimited at the margin. Cells are always cylindrical, straight, with rounded ends, solitary or in pairs after division, pale blue–green, (2.2)3– 15(20) ⫻ 0.5–3 µm. Cells divide perpendicularly to the long cell axis in two, more or less isomorphic daughter cells. All five described species are from metaphytic and periphytic habitats in freshwater. Two species are known from thermal springs (Copeland, 1936; Frémy, 1949). Among them, B. indurata was described from thermal sites in Yellowstone National Park (62–70°C)


FIGURE 1 (A) Cyanobium; a. C. gracile, cross section through the cell with parietally localized thylakoids (t = thylakoids; bar = 0.3 µm; after Komárek, 1999); b. C. eximium and c. C. roseum (both described from thermal springs in Yellowstone National Park under Synechococcus; after Copeland, 1936). (B) Cyanodictyon; a, b. C. planctonicum (bars = 10 µm; photo after Hickel, 1981). (C) Radiocystis: a. R. geminata (after Skuja, 1948); b. R. elongata (bar = 10 µm; after Hindák, 1996, from lakes in central Canada).

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FIGURE 2 (A) Epigloeosphaera: a, b. E. glebulenta (bar = 10 µm; after Komárková-Legnerová, 1991, and Zalessky, 1926). (B) Aphanothece: a. A. (Anathece) smithii (bar = 10 µm; after Smith, 1920, from Wisconsin lakes, and Komárková-Legnerová and Cronberg, 1994); b. A. (Anathece) clathrata (after Smith, 1920, from Wisconsin lakes); c. A. (Aphanothece) stagnina (after Smith, 1920, from Wisconsin lakes); d. A. (Aphanothece) castagnei (after Komárek and Anagnostidis, 1998, and Smith, 1950, from North America; sub Anacystis rupestris).


FIGURE 3 (A) Gloeothece: a. G. heufleri (after Prichod’kova from Kondrateva et al., 1984); b. G. rupestris (after Bourrelly and Manguin, 1952, from Guadeloupe); c. G. interspersa (after Gardner, 1927, from Puerto Rico). (B) Cyanobacterium: a. C. cf. cedrorum, lengthwise section with characteristic position of thylakoids (t); b. C. minervae (after Copeland, 1936, from thermal springs of Yellowstone National Park); c. C. diachloros (after Skuja, 1939).

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FIGURE 4 (A) Synechococcus: a. Synechococcus sp., lengthwise section through a cell (t = thylakoids, bar = 1 µm); b. S. nidulans (after Komárek, 1989, from Cuba); c. S. lividus, diversity of cells in various populations (after Copeland, 1936, from Yellowstone National Park). (B) Rhabdoderma: a. R. lineare (after Smith, 1920; planktonic species from Wisconsin lakes); b. R. compositum (after Smith, 1920; planktonic species from Wisconsin lakes; sub Gloeothece linearis var. composita); c. R. zygnemicolum (after Copeland, 1936, epiphytic species from Yellowstone National Park). (C) Rhabdogloea: a. R. smithii (after Komárek, 1969), b. R. smithii (after Smith, 1920, from Wisconsin lakes; sub Dactylococcopsis raphidioides).


FIGURE 5 (A) Bacularia: a. B. gracilis (after Komárek, 1995, from Cuba); b. B. indurata (after Copeland, 1936, from hot springs in Yellowstone National Park under Bacillosiphon induratus). (B) Cyanothece: a. C. aeruginosa (after Geitler, 1925, under Synechococcus aeruginosus); b. C. maior (after Starmach, 1973, under Synechococcus maior); c. C. aeruginosa (t = thylakoids; bar = 2 µm; after Komárek and Cepák, 1998; cross section through the vegetative cell).

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FIGURE 6 (A) Johannesbaptistia: a. J. primaria (after Gardner, 1927, from Puerto Rico; sub Cyanothrix primaria); b. J. pellucida (after Gardner, 1927, from Puerto Rico; sub Cyanothrix willei); c. J. schizodichotoma (after Copeland, 1936, and Smith, 1950, from Yellowstone National Park; sub Heterohormogonium schizodichotomum). (B) Synechocystis: a. S. aquatilis (after Setchell and Gardner, 1919, from Pacific coast in California); b. S. willei (after Gardner, 1927, from Puerto Rico); c. S. thermalis (after Copeland, 1936, from Yellowstone National Park). (C) Aphanocapsa: a. A. incerta (after Smith, 1920, from Wisconsin lakes; sub Polycystis incerta); b. A. grevillei (after Smith, 1920, from Wisconsin lakes); c. A. farlowiana (after Drouet, 1942, from northern United States).


with lime-encrusted sheaths as Bacillosiphon. The metaphytic species B. gracilis was discovered in littoral zones of alkaline lakes in Cuba (Komárek, 1995), and occurs more in tropical America. Cyanobacterium Rippka et Cohen-Bazire (Fig. 3B) Cells are solitary or in groups and/or aggregates, do not form mucilaginous colonies, have no enveloping mucilage or only a very fine, indistinct, and narrow gelatinous layer around the cell surface. Cells are widely oval to cylindrical and rod-shaped, rarely slightly arcuate, after division in pairs. The cell content is homogeneous, sometimes slightly keritomized or lengthwise striated (parallel position of thylakoids, sometimes in entire cell volume), pale to bright blue– green or olive green, sometimes with distinct granulation, without obvious gas vesicles, (2)3.7–17(30) ⫻ (1)3–12(20) µm. Cell division only by perpendicular binary fission (mainly pinching); daughter cells grow into the original size and shape before the next division. This recently defined genus had a few species previously described under Synechococcus. Cyanobacterium now comprises eight well-known species, but further descriptions are expected. Species are not widely reported and most are from extreme environments. C. minervae (originally Synechococcus minervae) was described from hot springs in Yellowstone National Park, but is now known from thermal waters worldwide. C. cedrorum was recorded from soils in North Carolina (as Synechococcus cedrorum) by Whitford and Schumacher (1969), but this species occurs in various concepts in the literature. Other species occur in swamps and rice fields in subtropical zones (Florida and Cuba). Cyanobium Rippka et Cohen-Bazire (Fig. 1A) Cells are solitary or in pairs after division, without mucilage or with very fine, narrow, usually diffuse slimy envelopes around cells; never in colonies. Cells are oval to short rod-shaped, up to 4 µm ⫻ 1–2 µm, without gas vesicles, pale blue–green to olive green, greyish blue–green, or reddish, usually with distinctly visible chromatoplasm (= parietal thylakoids). Cell division by binary fission (usually by pinching), always transversely to the longer cell axis; daughter cells are more or less isodiametric, and grow to the original shape and size before the next division. This widely distributed but little-known genus occurs in plankton (may be a major component of freshwater and marine picoplankton), in subaerial and submersed benthic mats, and as an epiphyte (Drews et al., 1961; Waterbury et al., 1979; Komárek and Anagnostidis, 1998; Komárek et al., 1999). Only 12

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species currently have been described, but Cyanobium is likely present in many environments in North America. In older literature it is often identified as a “small Synechococcus species.” Earlier identifications from North America may require revision. A few types have been recorded from oligotrophic lakes in New York (Corpe and Jensen, 1992), but without species identification. Three thermophilic species, C. amethystinum, C. eximium, and C. roseum, are known from Yellowstone Park and other thermal springs in North America (Copeland, 1936; Komárek et al., 1999). Cyanodictyon Pascher (Fig. 1B) Cells occur in spherical to irregularly reticulate, slimy, microscopic flat or three-dimensional colonies, which are sometimes elongate, composed from irregularly branching and anastomosing mucilaginous strands, later forming an amorphous mass with holes. The mucilage is fine, colorless, and sometimes diffuse. Cells are arranged in strands in one or more rows, and later irregularly. Cells are more or less spherical, slightly elongated to rod-shaped, small, up to 4.5 µm long (or diameter), without obvious gas vesicles, and pale blue–green, greyish, or olive green. Cell division is by binary fission, always perpendicularly to one (longer) cell axis. Daughter cells are isomorphic and grow to the original shape and size before the next division. Cyanodictyon species are mainly planktonic in oligotrophic to mesotrophic (rarely dystrophic) water bodies; one species is endgloeic within the mucilage of Anabaena (Geitler, 1932). Probably widely distributed. Of eight described species, four were recorded from Canadian lakes: C. reticulatum, C. tubiforme, C. planctonicum, and C. filiforme (H. Kling, personal communication). Cyanothece Komárek (Fig. 5B) Cells are solitary or in pairs after division, rarely aggregated in groups, but never forming distinct colonies. They have no individual envelopes, but are surrounded by narrow, very fine, diffuse and structured colonial mucilage. Cells are widely oval to almost cylindrical with widely rounded ends, (6.2)7–45(100) ⫻ (6)7–30(76) µm, and a length/width ratio less than 3:1. Cells appear pale to bright blue–green, olive green, greyish, or pinkish. Higher magnifications show distinct netlike keritomy (= irregular to radial arrangement of thylakoidal fascicles, sometimes with intra–thylakoidas spaces). Involution cells are irregular. Cells divide by binary fission (cleavage), transversely to one (longer) cell axis, into two isomorphic hemispherical daughter cells, which grow to original size and shape before the next division. The ultrastructure of large cells is unlike

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most other morphologically similar genera (Komárek and Cepák, 1998). Six well-known and several uncertain species that occur mostly in metaphyton in various habitats have been identified. Two species are known from peaty, cold stenotherm swamps and lakes: C. aeruginosa (in previous literature Synechococcus aeruginosus) has cosmopolitan distribution and is well known from North America (Daily, 1942; Prescott, 1962; Whitford and Schumacher, 1969); C. major occurs in subarctic and alpine regions in Alaska and Canada (Smith, 1950; Croasdale, 1973). C. lineata was collected from small mesotrophic lakes in Mexico (Komárek and Komárková-Legnerová, 2002). Epigloeosphaera Komárková-Legnerová (Fig. 2A) Colonies are microscopic to (rarely, when old) macroscopic, composed of clusters of irregular mucilaginous spheres or elongated formations, with smooth surfaces, on which scarce and irregularly situated solitary cells are observed (from above). Mucilage is firm, homogeneous, colorless, and distinctly delimited. Cells are widely oval to cylindrical, up to 4.2 µm long, distant (after division in pairs), with no obvious gas vesicles, and pale blue–green. Cell division is only in one plane in successive generations, perpendicular to the longer cell axis. Populations typically develop in benthic habitats (epipelic, among plants) in oligotrophic and mesotrophic pools, ponds, and lakes; sometimes they float in metaphyton or in plankton. Of two species, E. glebulenta occurs rarely in clear, cold lakes in the northern temperate areas. In North America it was observed in small lakes in central Canada (our data). (The second species is known only from South Africa.) Gloeothece Nägeli (Fig. 3A) Colonies are composed of numerous groups of cells that have their own, usually lamellated and delimited mucilaginous sheaths or envelopes. They form slimy macroscopic layers on various substrata, sometimes surrounded by fine common mucilage. The mucilaginous envelopes are colorless or colored bluish, violet, reddish, or yellow–brown. Cells are widely oval to rod-shaped, pale or bright blue–green, olive green, or violet, with no obvious gas vesicles. Cell division is by binary fission, only transversely to the longer cell axis; reproduction is by disintegration of clusters of cells. Variability and taxonomic relationships of various populations are poorly known, and it has not yet in culture. Nanocytic cell division was described in several species (Geitler, 1942). The genus contains mainly subaerial (over 30 known) species growing on wet rocks; a few species are

found in aquatic habitats, and one species is nanoplanktonic (Skuja, 1964). Several species are recorded from North America (G. confluens, G. palea, G. linearis, G. fusco-lutea, G. rupestris, and others; Prescott, 1962; Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982), but because their taxonomy requires revision, other species are likely. Species cited in North American literature in various concepts (e.g., G. distans and G. membranacea) are taxonomically unclear. A few species are known from (possibly restricted to) subtropical regions (Puerto Rico, Cuba, and Mexico), including G. endochromatica, G. interspersa, G. opalothecata, and G. prototypa (Gardner, 1927). Taxonomic revision of the genus is needed. Johannesbaptistia De Toni (Fig. 6A) Short discoid cells are arranged in uniseriate pseudofilaments within wide, mucilaginous, tubelike strands. Strands may be simple, unbranched, or (rarely) pseudodichotomously divided and anastomozing, and straight or slightly wavy. Mucilaginous envelopes are homogeneous, colorless, usually distinctly delimited at the margin, rarely diffuse, and rounded at the ends. Cells are short discoid, but in a lateral view are narrow oval, slightly to distinctly distant, pale grey–blue with fine granular cell content and no gas vesicles, 1–5.5(6.5) ⫻ (1.2)6.6–8.3(10) µm. Apical cells are usually rounded. Numerous necridic cells are often in pseudofilaments. Cells divide transversely to their shorter axis, that is, perpendicularly to the pseudofilaments. Numerous species have been described (some under Cyanothrix; Gardner, 1927; Kiselev, 1947), but interspecific features are presently unclear. A common but variable species, J. pellucida, requires taxonomic attention. It occurs mainly in metaphyton of coastal swamps with elevated salinity (in Massachusetts; Drouet and Daily, 1956), in freshwaters in warmer regions (Florida, Louisiana, California, Caribbean islands, Mexico, and Belize), and in some freshwater ponds and small lakes in Connecticut, Iowa, and Indiana (Drouet and Daily, 1956). A second well-defined species, J. schizodichotoma, was described from slightly acidic mineral springs (as Heterohormogonium; Copeland, 1936) in Yellowstone National Park (28–42°C), and possibly is endemic. Lithomyxa Howe (Not pictured) Cells irregularly arranged in macroscopic, flat, originally slimy and later hard, crustose colonies, usually heavily encrusted by lime. Cells are enveloped by their own gelatinous envelopes. Cells are spherical to oval and short cylindrical, slightly distant from one another, rarely in groups, only slightly over 1 µm in


diameter, with no obvious gas vesicles, and pale blue–green. Cell division probably is in one plane in successive generations. Specimens of this poorly known monotypic genus (needing revision) were described (with the species L. calcigena) from the United States (Howe, 1932) as an important lithogenic and travertine-forming species. Not collected since the description, the herbarium type specimen does not contain similar cyanobacterial type. According to Drouet and Daily (1956), it belongs to the Eubacteria. Radiocystis Skuja (Fig. 1C) Colonies are microscopic, more or less spherical to slightly elongated, and have radially oriented rows of cells, which are later (particularly in the center of colonies) irregularly arranged. Colonial mucilage is fine, colorless, and diffuse (apparent in phase contrast or after any staining). Cells are slightly distant or in pairs, nearly spherical to oval, to 5 (in tropical species up to 8) µm in length (or diameter), sometimes with visible aerotopes, and pale greenish in color. Cells divide only in one plane in successive generations (perpendicularly to radiating rows); however, their radial position in colonies may be indistinct. Species are mainly nanoplanktonic and may be part of surface blooms in mesotrophic and eutrophic waters (Skuja, 1948; Hindák and Moustaka, 1988; Hindák, 1996). Of five described species, two are tropical (R. fernandoi occurs in tropical regions of central America; Komárek and Komárková-Legnerová, 1993), and two (R. geminata and R. elongata) are known from plankton of mesotrophic water bodies in north-temperate regions of North America, including Ontario (Duthie and Socha, 1976; Hindák, 1996; H. Kling, personal communication). Rhabdoderma Schmidle et Lauterborn (Fig. 4B) Cylindrical cells are arranged more or less in the same direction, but usually not chainlike, irregularly distant, in microscopic, irregularly oval or elongated mucilaginous colonies. The mucilage is fine, homogeneous, colorless, diffuse or delimited at the margin, and sometimes indistinct. Cells always are cylindrical, rod-shaped, or slightly curved, rounded at the ends, sometimes several times longer than wide, (2)4–12(33) ⫻ (0.5)1–3.5 µm, with pale blue–green, greyish, or olive-green cell content and no obvious gas vesicles. Cell division is perpendicular to the longer axis, and sometimes asymmetrical (particularly under suboptimal conditions). Daughter cells sometimes remain joined together in short pseudofilaments; long filamentous involution cells are known. This species never occurs in masses, but several species commonly occur in the plankton of large

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oligotrophic and mesotrophic lakes, particularly in northern regions of the temperate zone (Prescott, 1962; Stein and Borden, 1979; Komárek and Anagnostidis, 1998). R. lineare and R. compositum are common species in the plankton of various lakes and water bodies in the northern United States and Canada. R. curtum was described from saline coastal swamps and pools in California, and epiphytic R. zygnemicolum is known only from Yellowstone National Park in temperatures > 30°C (Copeland, 1936). Rhabdogloea Schröder (Fig. 4C) Fusiform cells are arranged irregularly, distant, rarely more or less in one direction, in microscopic mucilaginous colonies. Mucilage is fine, homogeneous, colorless, and usually diffuse or indistinct at the margin. Cells are fusiform or cylindrical with conical cell ends, sometimes pointed, often slightly curved, (1.8)3–12(22) ⫻ (0.5)1–3.5(6) µm, with pale blue–green or olivegreen content. Involution filamentous cells are up to more than 25 µm long (known only from culture). Cell division is by binary fission, transverse to the long axis into two more or less isomorphic daughter cells. Reproduction is by disintegration of colonies. Ten to fifteen species are known, mostly planktonic in lakes and reservoirs, but also endogloeic in Microcystis colonies (Komárek and Anagnostidis, 1998). One metaphytic species occurs in acidic peaty swamps. Planktonic R. smithii commonly is found in northern lakes in North America (our results) and R. hungarica was collected from snowfields in the Rocky Mountains (as Gloeothece transsylvanica; Garric, 1965). R. subtropica was described from the plankton of small reservoirs and swamps in Cuba (Hindák, 1984). Synechococcus Nägeli (Fig. 4A) Cells are solitary, or in irregular groups or agglomerations. They do not form distinct colonies, have no slimy envelopes or only a very fine, diffuse, and narrow gelatinous layer. Cells are cylindrical to long rodshaped, sometimes slightly arcuate, after division in pairs. Cell content is homogeneous, occasionally has a slightly recognizable centro- and chromatoplasm (= few to several parietal thylakoids), have pale blue– green, olive green, or reddish color, no gas vesicles, and sometimes prominent granules. Cells are (1.5)3–15(40) ⫻ 0.4–3(6) µm. Cell division is by perpendicular binary fission (usually cleavage), sometimes asymmetrically. Genus specific filamentous involution cells occur under stress conditions. This important, but possibly uncommon genus comprises several species. Cultures are used widely as experimental model organisms. Over 20 species are known after recent taxonomic revisions. S. nidulans is

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distributed in small water bodies, mainly in temperate zones, and the strain, isolated from the United States and known in the literature as Anacystis nidulans (Kratz and Allen’s strain; Allen, 1968) is an important experimental strain. A group of thermophilic types (S. bigranulatus, S. koidzumii, and particularly S. lividus and S. vulcanus) are adapted to temperatures of about 70°C (Copeland, 1936; Castenholz, 1969a, b, 1970) and have been recorded from hot springs in Yellowstone National Park. Morphologically similar species are known from other hot springs in North America, Japan, Africa, and Indonesia (Geitler and Ruttner, 1935; Hirose and Hirano, 1981). Several species are known from littoral zones of lakes, soils, and swamps, and could be expected in similar environments elsewhere on the continent, but their taxonomy and distribution are poorly known. S. sigmoideus is a characteristic species from nanoplankton of lakes, including those in North Dakota (Moore and Carter, 1923).

Merismopediaceae (Figs. 6B–10A) Aphanocapsa Nägeli (Fig. 6C) Aphanocapsa comprises microscopic to macroscopic, mucilaginous colonies with irregularly arranged cells of various densities that are spherical or irregular. Colonial mucilage is mainly homogeneous and colorless, and usually has a diffuse, rarely delimited margin. Cells have no individual envelopes or, rarely, have a fine surrounding slimy layer that is distinct from the common mucilage. Cells are spherical, but after division, they are hemispherical, pale or bright blue–green or olive green, rarely (in marine species) red or pinkish in color, with no gas vesicles, and 0.4–6(12) µm in diameter. Cells divide by binary fission in two perpendicular planes in successive generations (cleavage). Reproduction is by disintegration of colonies. Among many described species, almost 50 are well-defined. They colonize mainly subaerial, soil, and aquatic habitats (Komárek and Anagnostidis, 1998). A. delicatissima, A. incerta, A. holsatica, A. conferta, and A. planctonica, which are planktonic and grow in microscopic colonies in mainly mesotrophic lakes, are perhaps the most important species of this genus in North America (Duthie and Socha, 1976; Sheath and Steinman, 1982). A. farlowiana produces macroscopic colonies, is possibly endemic in the northern United States and Canada, and sometimes occurs in large masses in the littoral zone of ponds and smaller lakes (Drouet, 1942). A. thermalis, A. botryoides, A. protea, and A. tolliana are known from thermal springs (Copeland, 1936). Several species grow on wet rocks in mountains (A. muscicola) or are benthic in streams (sometimes identified as Polycystis montana f. minor); several of these need revision. A. arctica was described

from Canadian arctic regions (Whelden, 1947), and A. intertexta was obtained from Puerto Rico (Gardner, 1927). Other species occur in coastal marine, brackish, and saline swamps and pools (e.g., A. marina). This genus has a very simple morphology, leading to many misinterpretations in the literature. Coelomoron Buell (Fig. 8C) Cells are arranged more or less peripherally in one or a few layers (may be variable) near the surface of the microscopic, spherical, free-living colonies, which are sometimes composed of subcolonies. Colonial mucilage is fine, homogeneous, colorless, and usually diffuse at the margin, but usually more densely clustered in the center. Cells in young colonies are randomly and sparsely distributed, distant and variably arranged, and usually slightly shifted from one layer; later cells are arranged distinctly peripherally, forming one to three irregular layers. Cells are slightly elongated, widely oval to almost spherical, and radially situated in the colony (generic feature), pale blue–green or olive green, and 1–6.5 ⫻ 0.8–4 µm. One species (C. minimum) has distinct aerotopes (gas vacuoles). Cell division occurs in two planes in successive generations, perpendicular to one another and more or less to the colony surface. Reproduction is by disintegration of colonies. Seven described species (Komárek and Anagnostidis, 1998) exist, but further species may be discovered. The type species C. regulare was described from freshwater metaphyton in the United States (Buell, 1938; Geitler, 1942), but has not been seen again. Nanoplanktonic C. pusillum occurs in mesotrophic waters in temperate and subtropical zones (syn. = Coelosphaerium collinsii Drouet et Daily 1942, described from the United States). C. microcystoides and C. vestitum were described from tropical and subtropical regions (Komárek, 1989). The aerotopated, pantropical C. minimum was recorded from the plankton of tropical Lake Catemaco in Mexico (KomárkováLegnerová and Tavera, 1996). Coelosphaerium Nägeli (Fig. 8B) Coelosphaerium comprises free-living, spherical or oval mucilaginous colonies, in which the cells are arranged irregularly in one marginal layer (but sometimes slightly shifted to one another), or near the surface of the sphere. Colonies sometimes are composed of subcolonies. Mucilage is colorless and homogeneous. Cells are spherical, pale or bright blue–green, and 1–7 µm in diameter. One species has visible aerotopes. Cell division occurs in two planes in successive generations, perpendicular to one another and to the colony surface. Reproduction is by disintegration of a colony and by liberation of subcolonies.


FIGURE 7 (A) Merismopedia: a. M. punctata (after Smith, 1920, from Wisconsin lakes); b. M. angularis (after Thompson, 1938, from eastern Kansas); c. M. smithii (after Smith, 1920, from Wisconsin lakes; sub M. elegans var. maior); d. M. elegans (after Smith, 1920, from Wisconsin lakes). (B) Mantellum: a–c. M. rubrum (after Tavera and Komárek, 1996, from a volcanic lake in Mexico). (C) Cyanotetras: a–c. C. crucigenielloides (after Komárek, 1995; variation in form of colonies from swamps in Cuba).

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FIGURE 8 (A) Microcrocis: a. M. irregulare (after Lagerheim, from Smith, 1950); b. M. obvoluta (after Tiffany, 1934, from North America, Lake Erie); c. M. pulchella (after Buell, 1938, from North America). (B) Coelosphaerium: a. C. kuetzingianum (after Smith, 1950, from North America); b. C. subarcticum (after Komárek and Komárková-Legnerová, 1992, from Canadian lakes); c. C. aerugineum (bar = 10 µm; after Smith, 1920, from Wisconsin lakes, and after Komárek, 1958). (C) Coelomoron: a. C. tropicalis (after Senna et al., 1998, from Brazil); b. C. microcystoides (after Komárek, 1989, populations from Cuba).


FIGURE 9 (A) Snowella: a. S. fennica (after Komárek and Komárková-Legnerová, 1992, from Canadian lakes); b. S. litoralis (original photo by H. Kling, from lakes in Central Canada); c. S. litoralis (after Smith, 1950, from Wisconsin lakes; sub “Gomphosphaeria lacustris”). (B) Woronichinia: a. W. naegeliana (after Smith, 1950, from North America; sub Coelosphaerium naegelianum); b. W. naegeliana (bar = 10 µm; after Komárek and Komárková-Legnerová, 1992, from central Canadian lakes); c. W. klingae (bar = 10 µm; after Komárek and Komárková-Legnerová, 1992, from Manitoba, Canada).

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FIGURE 10 (A) Gomphosphaeria: a. G. aponina (after Smith, 1920, from Wisconsin lakes); b. G. natans (after Komárek and Komárková-Legnerová, 1992, from central Canadian lakes); c. G. semen-vitis (after Komárek, 1989, from alkaline swamps in Cuba and in Florida); d. G. virieuxii (after Komárek and Komárková-Legnerová, 1992, from central Canadian lakes). (B) Eucapsis: a. E. alpina (after Clements and Shantz, 1909, original drawing from Colorado); b. E. alpina (after Prescott and Croasdale, 1937, from Minnesota); c. E. alpina forma (after Thompson, 1938, from eastern Kansas); d. E. alpina (original photo from central Europe).


Ten species are known from plankton of lakes and reservoirs across North America and worldwide (Prescott, 1962; Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982; some = Woronichinia spp.); they rarely occur in metaphyton (Komárek and Anagnostidis, 1998). Nanoplanktonic C. minutissimum, C. aerugineum, and C. kuetzingianum are known from lakes in the temperate zone of North America. C. subarcticum occurs in clear lakes of central Canada, but is probably more widely distributed in similar environments. The occurrence of C. dubium and C. confertum in North America is unconfirmed. Cyanotetras Hindák (Fig. 7C) Cells occur in microscopic, free-floating colonies that are flat or platelike and composed from slightly elongated cells, the longer axis of which lies in the plane of the colony. Colonies typically are few-celled. The cells are arranged in short perpendicular rows, but have irregularities. The colony is surrounded by very fine, colorless, and diffuse mucilage. Cells are oval or ovoid, sometimes in twos or flat tetrads, have pale blue–green color and, homogeneous content. One species has gas vesicles and visible aerotopes; another species has iron precipitates within the mucilage. Cells are 1.5–5 µm long and 1–4.5 µm wide. Cells divide by binary fission in two planes in successive generations, perpendicularly to one another and to the plane of the colony. Three species have been described. Two are from the subtropical regions of North America (Komárek, 1995; Komárek and Komárková-Legnerová, 2002): C. crucigenielloides is common in the metaphyton of alkaline swamps, ponds, and lakes in the Caribbean region; C. aerotopa was discovered recently in plankton of mesotrophic and eutrophic waters in Mexico. Gomphosphaeria Kützing (Fig. 10A) Colonies are free-living, spherical or irregularly oval, and sometimes composed of subcolonies. They have a central system of thick mucilaginous stalks that are nearly pseudodichotomously divided and may be diffuse within the colony. Stalks widen at the ends and envelope individual cells with a thin mucilage layer. Cells are slightly elongate, obovate, or club-shaped, and radially oriented more or less at the colony periphery, which is sometimes enveloped by a fine, colorless, and diffuse mucilage. Cells have a homogeneous pale or bright blue–green, olive–green, or red content and are (4.2)6–12(15) ⫻ 2–8(13.2) µm. After division, the cells may remain joined together and form a characteristic cordiform shape. Cells in colonies are slightly distant and sometimes slightly radially displaced from

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one another. Cell division occurs in two planes in successive generations, perpendicular to one another and to the colony surface. Reproduction is by colony disintegration. Six species have been recorded from North America (Prescott, 1962; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). G. natans is planktonic in northern mesotrophic lakes (from Canada: Komárek and Hindák, 1988; Komárek and Komárková-Legnerová, 1992) and possibly endemic to the continent. G. virieuxii (planktonic in lakes), G. aponina (metaphytic in clear, slightly acidic swamps) and G. salina (metaphytic in saline pools and swamps), are known from across the temperate zones. G. multiplex and G. semen-vitis were recorded from alkaline swamps in subtropical North America (Caribbean; Komárek, 1989). G. cordiformis (sensu Smith, 1920) was recorded from Wisconsin lakes, but needs taxonomic revision and confirmation. Mantellum Dangeard (Fig. 7B) Cells occur in flat, microscopic formations attached to substrata (as epiphytes on other algae) in one layer and arranged more or less in perpendicular rows. Cells are spherical, pale blue–green or reddish in color, have homogeneous content or distinguishable centro- and chromatoplasm, and are 0.8–4 µm in diameter. Cells usually are enveloped by thin, very fine, colorless, and diffuse mucilage. Division is by binary fission, in two planes perpendicular to one another and to the substrate; from this process, flat formations arise on the substrate, but sometimes have slightly shifted cells. Only three epiphytic species, one marine and two freshwater, have been described to date. One freshwater species with reddish cells (M. rubrum) is known from deep volcanic lakes in central Mexico (Tavera and Komárek, 1996). Merismopedia Meyen (Fig. 7A) Merismopedia comprises flattened, free-living, platelike (rectangular), more or less rectangular colonies that have one layer of cells, arranged loosely or densely in perpendicular rows and enveloped by fine, colorless, usually indistinct, and marginally diffuse mucilage. Colonies are flat or slightly wavy, usually microscopic (except for a few species that are macroscopically visible), and sometimes composed of subcolonies. Cells are spherical or widely oval before the division, pale or bright blue–green, (rarely reddish), and sometimes have visible centro- and chromatoplasm (parietal thylakoids). Several planktonic species have gas vesicles (few or solitary in cell centers). Occasionally the cells have slimy envelopes. After division, the cells are hemispherical and (0.4)1.2–6.5(17) µm in diameter.

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When elongated, the longer axis is situated in the plane of the colony. Cell division (binary fission) occurs regularly in two planes perpendicular to the plane of colony; the daughter cells do not move from their position after division. Reproduction is by fragmentation of colonies. Over 30 species are known from freshwater and saline environments (Komárek and Anagnostidis, 1998). Several common species occur across temperate regions in the plankton and metaphyton of eutrophic and mesotrophic waters in North America, especially M. punctata and M. tenuissima; two common species, M. glauca and M. elegans occur in the metaphyton (rarely in plankton) of mesotrophic lakes, ponds, and swamps (Prescott, 1962; Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). M. smithii (syn. = M. major), is a species that has large cells and is typical of temperate swamps only in North America (not confirmed from other regions). M. angularis occurs in acidic swamps in cold temperate to subarctic regions. M. gardneri grows in pools on the American Pacific coast. M. convoluta is a conspicuous metaphytic species from warmer regions of the continent (North Carolina, Florida, and the Greater Antilles). Records of a species designated Agmenellum quadruplicatum in floras and ecological studies (according to Drouet and Daily, 1956) may be referred to either M. punctata or M. glauca. Strains designated by the same name in world collections and experimental papers (e.g., Van Baalen and O’Donnell, 1972) contain the species Synechococcus nidulans. Microcrocis Richter (Fig. 8A) Cells are arranged in free-living, flat, microscopic to macroscopic colonies, in one layer, and form perpendicular rows in young colonies; later they are irregularly aggregated. Colonies are enveloped by fine, colorless, homogeneous, and diffuse mucilage. Cells have no individual sheaths. A critical feature is the presence of elongated cells situated with the longer axes perpendicular to the colony plane. Cells are widely oval to rodlike with rounded ends, homogeneous content, and pale to bright blue–green color. The cell structure has been studied only in one species, in which the thylakoids were flexuous throughout (different from other related genera that have parietal thylakoids). Cell dimensions are (2)5–16(19) ⫻ 1.5–6(8) µm. Cells divide longitudinally in two perpendicular planes in successive generations. Reproduction is by fragmentation of colonies. This rare and poorly known genus has 10–15 species, of which several need revision (Komárek and Anagnostidis, 1998). Metaphytic, epipelic, and epipsammic species exist in freshwater and marine

environments. M. irregularis, M. obvoluta, M. gigas, and M. pulchella were described from North America (Komárek and Anagnostidis, 1998), but their taxonomy and ecology require confirmation. Snowella Elenkin (Fig. 9A) Snowella comprises free-living, spherical or oval microscopic colonies, sometimes composed of subcolonies, that have a central system of thin, nearly pseudodichotomously divided gelatinous stalks (may be confluent) that bear cells at their ends (periphery of colony). Colonies are enveloped by very fine, unstructured, and diffuse mucilage. Cells are spherical or slightly radially elongated, and pale blue–green, olive green, or yellowish in color; one species may be pinkish. Two species have one or a few central aerotopes. The cells are 0.6–4.2 µm in diameter (or long). Cell division is by binary fission in two planes, perpendicular to one another and to the colonial surface. Reproduction is by disintegration of colonies. Seven species are known from the plankton of temperate fresh and brackish waters, particularly from cold, northern, mesotrophic lakes (Komárek and Anagnostidis, 1998). S. septentrionalis and S. fennica were recorded from Canada (original unpublished data). The tychoplanktonic S. rosea has reddish cells and the nanoplanktonic S. litoralis and S. lacustris occur sporadically in the temperate zones of North America, although S. lacustris may also occur in subtropical regions (Komárek and Hindák, 1988). Synechocystis Sauvageau (Fig. 6B) Cells are solitary or in pairs short time after division, have no mucilage or a fine, narrow, colorless, indistinct, mucilage layer around the cells that is diffuse at the margins. Cells are spherical, rarely to widely oval before division, usually pale or rarely, bright blue–green or olive green, in few species, reddish violet or red. Cells are 0.7–15(30) µm in diameter. Cell content is more or less homogeneous, sometimes has solitary granules and a distinguishable centro- and chromatoplasm (parietal position of thylakoids). The species with irregularly distributed thylakoids and larger dimensions probably belong to another taxonomic type. Cell division is by binary fission (cleavage), regularly in two perpendicular planes in successive generations. Cells grow to the original size and shape before the next division. Over 20 species have been described. Most are planktonic and metaphytic, but some occur in thermal springs, mineral and saline environments, and from marine Ascidians (epizoic on the surface) and are symbiotic in dinoflagellates (Norris, 1967; Schulz-Baldes and Lewin, 1976; Komárek and Anagnostidis, 1998).


In North America, S. aquatilis and S. salina are planktonic in higher conductivity pools and lakes (Stein and Borden, 1979), S. sallensis occurs in northern peat bogs, and S. thermalis and S. minuscula are known from hot springs (Copeland, 1936). S. fuscopigmentosa, S. primigenia, and S. willei were recorded from swamps in the Caribbean region (Gardner, 1927, Kovácˇik, 1988). An interesting Synechocystis species was found in hollow hairs from polar bears in zoological gardens in California (Lewin and Robinson, 1979) and given the preliminary name Aphanocapsa montana, but its taxonomy is not resolved. Picoplanktonic forms that have spherical cells were recorded from clear lakes in New York (Corpe and Jensen, 1992) and probably belong to this genus. Woronichinia Elenkin (Fig. 9B) Woronichinia comprises spherical or irregularly oval, free-living colonies, commonly composed of subcolonies, that have a central system of simple, radially arranged, colorless, mucilaginous (sometimes tubelike) stalks, joined in the center of the colony. The colony is surrounded by a fine, diffuse, colorless mucilage. Cells are at (or within) the ends of mucilaginous stalks and radially arranged, forming a layer of cells at the colony periphery. Stalks are densely packed, causing radial lamellation, but sometimes are diffuse near the center. Cells may be nearly spherical, but are usually slightly elongate, widely oval, or obovoid; in old colonies the cells are densely arranged. The cells are pale blue–green, olive green, or slightly reddish in color, sometimes have gas vesicles and visible aerotopes, and are (1)2.5–7 ⫻ 1–4(5) µm. Cell division occurs in two planes, perpendicular in successive generations and to the colony surface. Reproduction is by disintegration of colonies and by solitary cells (“expulsion cells”), which sometimes liberate spontaneously from a colony. Of 15 described species, 3 contain gas vesicles and may form surface blooms in mesotrophic and eutrophic waters (Komárek and Anagnostidis, 1998). In temperate regions, the aerotopated (with gas vacuoles) and cosmopolitan W. naegeliana (syn. Coelosphaerium naegelianum) is widespread in the plankton of many lakes and may form numerous morphotypes, of which few are regarded as endemic in North America (Cronberg and Komárek 1994). A similar species named Gomphosphaeria wichurae has been recorded in North America, but probably falls within the range of variation in W. naegeliana. W. fremyi was described from the Caribbean region (Komárek, 1984). W. karelica and W. elorantae, originally described from the Baltic region in Europe, are nanoplanktonic species that occur in northern lakes in Canada (original unpublished data). W. klingae is an interesting plank-

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tonic species that has reddish cells and solitary gas vesicles, and possibly is endemic to stagnant, swampy waters in central Canada (Komárek and KomárkováLegnerová, 1992).

Microcystaceae (Figs. 10B–12) Chondrocystis Lemmermann (Fig. 12B) Colonies occur in microscopic or macroscopic masses, are free-living, irregular in outline, packet-like, gelatinous, in granular agglomerations, and composed of numerous subcolonies, sometimes with inner CaCO3 precipitates. The mucilage is firm, colorless to yellow–brown, and distinctly limited. Cells and their groups are surrounded by individual firm sheaths and envelopes, which are tightly and irregularly aggregated together. Cells are spherical, pale blue–green, 1.5–6 µm in diameter, have no gas vesicles, and divide by binary fission in three perpendicular planes in successive generations. Reproduction is by disintegration of colonies. Three halophilic (metaphytic) and one benthic species from limestone mountainous streams (C. dermochroa) are known (Komárek and Anagnostidis, 1998), but not yet confirmed from North America. C. schauinslandii from Hawaii and C. bracei from the Bahamas and Bermuda were described (Lemmermann, 1899; Howe, 1924). Eucapsis Clements et Shantz (Fig. 10B) Colonies are microscopic, mucilaginous, freeliving, more or less cubic in form, and sometimes composed of subcolonies with cells arranged three dimensionally (cubelike) in more or less regular perpendicular rows; rows may be disturbed in a few species. The mucilage is colorless, hyaline, and usually diffuse at the margin. Cells are spherical or slightly oval before division, pale or bright blue–green or olive green, 1–6(11) µm in diameter, and have no obvious gas vesicles. Cell division occurs regularly in three perpendicular planes in successive generations. Daughter cells more or less keep their position in a colony. Reproduction is by disintegration of colonies. Eight species and several varieties have been described. The often are metaphytic in swamps and bogs, but also are found in volcanic soils (Komárek and Hindák, 1989; Komárek and Anagnostidis, 1998). E. minor and E. alpina are found in peaty habitats in North America. The multicellular E. alpina var. maior (known only from Alaska; Prescott and Vinyard, 1965) probably represents a separate type of this genus (but the name is a later homonym and must be changed). Gloeocapsa Kützing (Fig. 12A) Colonies are microscopic, usually spherical, usually aggregated in a macroscopic mucilaginous, amorphous mass that colonizes wet stony substrates and, less

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FIGURE 11 (A) Microcystis: a. M. aeruginosa (after Teiling, 1941, Komárek, 1958, and Smith, 1950); b. M. pulchra (after Smith, 1920, from Wisconsin lakes); c. M. comperei (after Komárek, 1964, tropical species, from Central America); d. M. wesenbergii (after Teiling, 1941, and Wojciechowski 1971).


FIGURE 12 (A) Gloeocapsa: a. G. conglomerata (after Kützing from Geitler, 1925); b. G. alpina (after Geitler, 1932); c. G. sanguinea (from Anagnostidis and Komárek, 1998); d. G. gelatinosa (after Kützing from Tilden, 1910). (B) Chondrocystis; a. C. dermochroa (after Geitler, 1925, and Starmach, 1966; sub Gloeocapsa dermochroa); b. C. schauinslandii (after Lemmermann, 1905, from Hawaii Islands).

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frequently tree bark or aquatic habitats (metaphyton or plankton). Cells and their groups are surrounded by wide, spheroidal, concentrically layered gelatinous sheaths (with distinct or indistinct lamellation) and sharply delimited margins, which may be intensely colored (some species) yellow, yellow–brown, orange, red, blue, or violet. Sheath colors may change according to environmental pH (Jaag, 1945). Cells are spherical, but after division, are hemispherical; only in dormant stages are they irregular (not dividing). Cells have firm, gelatinous, usually rounded envelopes, with pale to bright blue–green or olive green protoplast, are 0.7–6(11) µm in diameter (without envelopes), and have no obvious gas vesicles. Cell division occurs in three perpendicular planes in successive generations. Daughter cells grow to the original size and shape before next division (Golubic´, 1965, 1967a, b). Reproduction is by disintegration of colonies. Of the more than 100 species described, perhaps 50 are clearly defined (Komárek and Anagnostidis, 1998). Most forms grow on wet rocks and walls, commonly in mountain areas. Several species are restricted to calcareous or acidic rocky substrates (Jaag, 1945; Golubic´, 1967). G. conglomerata, G. caldariorum, G. decorticans, G. gelatinosa, G. granosa, G. arenaria, and G. atrata (= G. alpicola) have colorless sheaths, G. nigrescens and G. alpina have blue or violet sheaths, G. sanguinea has red sheaths, and G. fusco-lutea, G. kuetzingiana, and G. rupestris have yellow or yellow– brown sheaths; all have been recorded in North America (Prescott, 1962; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). G. sparsa is known only from the eastern United States (Wood, 1869). G. thermophila was described from thermal springs in California, whereas G. acervata, G. calcicola, G. sphaerica, and several other spesies are known from limestone substrates in Puerto Rico (Gardner, 1927). Taxonomic revision of numerous species is needed; typical Gloeocapsa species are not yet in culture. Microcystis Kützing ex Lemmermann (Fig. 11) This genus has irregular micro- or macroscopic colonies that are free-floating, compact or clathrate, may be composed of clustered subcolonies, and has sparsely or densely, irregularly arranged cells. The mucilage is fine, colorless, and diffuse or distinctly delimited, sometimes forming a wide margin around the cells (rarely with indistinct structures), or delimited along cell agglomerations. Cells are spherical or hemispherical after division and pale blue–green, but they appear brownish due to aerotopes that mask the blue– green color of the protoplast. Cells are 0.8–6(9.4) µm in diameter and have no individual mucilaginous

envelopes. Cell division is by binary fission in three perpendicular planes in successive generations. The daughter cells grow to the original shape and size before the next division. Based on current revisions, about 25 planktonic species are known worldwide, and many form dense blooms in eutrophic waters (Reynolds and Walsby, 1975). Because many strains (species) are toxic, Microcystis is one of the most important cyanobacteria in limnological studies (Gorham and Carmichael, 1988; Chorus and Bartram, 1999). Several species have been recorded from North America, including toxic forms of M. viridis, M. aeruginosa, and M. ichthyoblabe (Smith, 1920; Prescott, 1950, 1962; Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979; Fallon and Brock, 1981; Doers and Park, 1988; our results). In a few North American species, including M. natans, M. smithii (as Aphanocapsa pulchra), M. flos-aquae, and M. wesenbergii, toxicity has not yet been recorded or definitively proved. Identification of species is difficult and further taxonomic investigation is needed. At least 50% of the species in the genus are restricted to tropical and subtropical regions (HuberPestalozzi, 1938; Desikachary, 1959; Komárek et al., 2002). Of these, M. comperei was described from Cuba and other species probably occur in southern locations. M. glauca, recorded from the United States by Smith (1950), and other species (denoted Polycystis firma, P. pulverea, and P. marginata) were recorded from North Carolina by Whitford and Schumacher (1969), but they are taxonomically uncertain.

Chroococcaceae (Figs. 13 and 14) Asterocapsa Chu (Fig. 13A) Cells are solitary or occur in spherical, microscopic colonies, sometimes agglomerated in granular, macroscopic, gelatinous masses. Cells and colonies are enveloped by distinctly delimited, usually spherical, firm mucilaginous sheaths, which are colorless or colored (blue, violet, orange, reddish), sometimes slightly concentrically lamellated and have a smooth or warty surface. Cells are subspherical, oval, irregular, or polygonal–rounded in outline, pale or bright blue– green or olive green, often have fine granulations in the protoplast, are 2–8 µm in diameter, and have no obvious gas vesicles. Cells divide by binary fission in various planes, irregularly. Reproduction is by cells that are liberated from split firm, gelatinous envelopes. Unicellular and colonial types are included in this genus, but may become classified as different genera. About 15 species have been described, most of which are subaerial on wet rocks (Komárek and Anagnostidis, 1998). Several interesting species are known from Mexico (Komárek, 1993, A. divina)


FIGURE 13 (A) Asterocapsa: a. A. divina (after Komárek, 1993, from San Luis Potosí, Mexico); b. A. sp. (after Gardner, 1927, from Puerto Rico; sub Anacystis nigroviolacea). (B) Gloeocapsopsis: a. G. magma (after Geitler, 1932); b. G. crepidinum (after Bornet and Thuret, and Hollerbach from Kosinskaja, 1948, sub Gloeocapsa crepidinum, marine species). (C) Cyanokybus: a. C. venezuelae (after Schiller, 1956, from Los Aves Islands, Venezuela); b. C. venezuelae (after Komárek, 1994, from Cuba).

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FIGURE 14 (A) Chroococcus: a. C. (Limnococcus) dispersus (after Smith, 1920, from Wisconsin lakes; sub C. limneticus var. subsalsus); b. C. (Limnococcus) sonorensis (after Drouet, 1942, from Baja California, Mexico); c. C. (Limnococcus) limneticus (after Smith, 1920, from Wisconsin lakes); d. C. (Chroococcus) mipitanensis (after Komárek and Novelo, 1994, from central America); e. C. (Chroococcus) polymorphus (after Komárek and Novelo, 1994, from central America); f. C. (Chroococcus) minutus var. thermalis (after Copeland, 1936, from Yellowstone National Park); g. C. (Chroococcus) yellowstonensis (after Copeland, 1936, from Yellowstone National Park); h. C. (Chroococcus) turgidus (after Smith, 1920, from Wisconsin lakes). (B) Cyanosarcina; a. C. sp. (after Hollenberg, 1939, from California; sub Microcystis splendens; marine species).


and Puerto Rico (Gardner, 1927, A. magnifica and A. pulchra). Chroococcus Nägeli (Fig. 14A) Cells or groups of cells (mainly two to four cells), are surrounded by mucilaginous envelopes, and usually occur in microscopic, spherical or composed colonies; rarely form agglomerations. Mucilaginous envelopes are colorless or yellowish, usually copying the cell form, sometimes lamellated, distinct or diffuse at the margin (subg. Chroococcus) or fine, homogeneous, and diffuse, in which the cells or cell groups are irregularly arranged (subg. Limnococcus). Cells at first are spherical or oval, and later are hemispherical or in the form of a segment of the sphere. The cells are 0.7–50 µm in diameter. The cell content is grey, blue–green, olive green, orange, or reddish violet and granular. Only in few planktonic species are there gas vesicles. Cell division is by binary fission in three or more planes, or irregular (in old colonies). Reproduction is by fragmentation of colonies. Many species have been described, and over 60 are well defined (Komárek and Anagnostidis, 1998); many occur in North America. Planktonic species from the subgenus Limnococcus are common in temperate and northern water bodies and include C. microscopicus, C. minimus, C. dispersus, C. distans, and C. limneticus. C. prescottii, C. refractus, and C. sonorensis were described from the United States, but especially C. prescottii is more common in cold, usually slightly acidic waters (our data). Most species from the subgenus Chroococcus are metaphytic, although the type species C. rufescens colonizes soils (Daily, 1942). From this group, C. varius, C. minutus, C. pallidus, C. schizodermaticus, C. multicoloratus, C. submarinus (halophilic), and C. turgidus are the most common. C. thermalis, C. tenacoides, C. yellowstonensis, and C. endophyticus are known from thermal springs (Copeland, 1936). Several species are typical of tropical and subtropical regions, including C. aeruginosus, C. cubicus, C. deltoides, C. heanogloios, C. mipitanensis, and C. polyedriformis (Komárek and Novelo, 1994). This genus has been widely reported across North America, including North Carolina (Whitford and Schumacher, 1969), the western Great Lakes region (Prescott, 1962), British Columbia (Stein and Borden, 1979), Ontario (Duthie and Socha, 1976), and the Northwest Territories (Sheath and Steinman, 1982). Cyanokybus Schiller (Fig. 13C) Colonies are few-celled, microscopic, and usually oval in outline. The cells are arranged in short perpendicular, irregular rows, more or less distant one from another, within a colorless, homogeneous, distinctly delimited mucilaginous envelope. The cells are hemi-

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spherical, rectangular, or polyhedral–rounded, have or do not have individual fine envelopes, are pale to bright blue–green, up to 15–20 µm in diameter, and have no obvious gas vesicles. Cell division is by binary fission in three planes, later repeatedly in one plane (cells in rows). Reproduction is by colony disintegration. This is a monotypic genus. The type species, C. venezuelae, was described from an island near the Venezuelan coast, but it is distributed sporadically in alkaline swamps throughout the Caribbean region (Schiller, 1956; Komárek, 1994). Cyanosarcina Kovácˇik (Fig. 14B) Colonies are microscopic, packet-like, sarcinoid or irregular, and have densely aggregated cells, enveloped by thin, colorless, firm envelopes that tightly enclose clusters of cells and later occasionally from macroscopic agglomerations. Cells are more or less spherical, subspherical, or irregularly rounded, enveloped by thin individual gelatinous layers, usually tightly packed, colored pale or bright blue–green, olive green, or reddish, finely granular, and 2–10 µm in diameter. Cells have no obvious gas vesicles. Cell division is by binary fission regularly in three, later in more planes; cells enlarge prior to divisions. Reproduction is by disintegration of colonies. Twelve species, usually metaphytic or periphytic, are described from various countries (Kovácˇik, 1988; Komárek and Anagnostidis, 1998). None is explicitly recorded from North America, but certain species have been described by Gardner (1927) from Puerto Rico (as Endospora rubra) and by Hollenberg (1939) from California (as Microcystis splendens), which probably belong to this genus. Further observations are needed. Gloeocapsopsis Geitler ex Komárek (Fig. 13B) Cells are usually densely aggregated in irregular, packet-like, microscopic colonies (rarely solitary), later forming a macroscopic flat or granular, gelatinous mass on substrata. Cells are subspherical or irregular– rounded in outline, sometimes slightly elongated, 2.5–14(20) µm in diameter, and usually have individual gelatinous envelopes. Colonies are surrounded by thin, firm, narrow, distinctly limited, and often colored (blue, red, or yellow–brown) sheaths that usually follow the irregular outline of cell aggregates by their shape. Cell content is pale or bright blue–green, finely granular, and has no obvious gas vesicles. Cell division is by irregular binary fission in various planes. Reproduction is by disintegration of colonies. Often confused with Gloeocapsa in older literature (because they resemble morphologically dormant stages of Gloeocapsa spp.), the geographic distribution of most of the 10 known species (Komárek and Anagnostidis, 1998) is still unclear. G. magma (which

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has reddish envelopes) may be widely distributed on wet rocks in high mountains (Tilden, 1910; Stein and Borden, 1979).

Entophysalidaceae (Fig. 15) Chlorogloea Wille (Fig. 15A) Colonies are multicellular, mucilaginous, spherical or irregular, and have a rough surface. Colonies usually are composed of subcolonies, that later form macroscopic gelatinous masses attached to various substrata; they are rarely epiphytic or metaphytic. Cells are arranged irregularly, but in the margin are more or less in radial rows (the result of terminal or heteropolar growth). Cells are spherical or slightly irregular–rounded in outline to polygonal–rounded, usually enveloped by individual envelopes, but later in homogeneous, colorless, and delimited mucilage that has a more or less homogeneous, pale grey–green, blue–green, or reddish content. The cells are 1–6 µm in diameter and have no gas vesicles. Cell division usually occurs in three perpendicular planes, less frequently irregularly, and sometimes with repeated division in one direction. Reproduction is by disintegration of colonies. This a poorly known genus comprises ecologically distinct types, most of which require revision. About 20 species have been described (Komárek and Anagnostidis, 1998), including three aquatic species (C. cuauhtemocii, C. epiphytica, and C. lithogenes) from alkaline environments in Mexico (Komárek and Montejano, 1994), and C. tuberculosa and C. regularis from saline environments in Pacific locations (Setchell and Gardner, 1924). Entophysalis Kützing (Fig. 15B) Colonies are multicellular, mucilaginous, microscopic to macroscopic, have polarized growth, are attached to the substratum, and are composed of groups of cells that are enveloped by their own distinct gelatinous and usually delimited envelopes. The gelatinous envelopes are firm, delimited, often layered, colorless or colored (red, dark violet, yellow–brown), and sometimes enveloped by distinct common mucilage. Cells (and their groups) are often arranged in irregular radial rows, spherical to irregular, often of variable size in the same colony, have pale blue–green, olive green, or yellowish content, are finely granular, (0.8)2–9 µm in diameter, and have no gas vesicles. Cells divide by binary fission, in various planes, but sometimes (in marginal parts) the perpendicular cell division repeats. Reproduction is by disintegration of colonies. More than 20 species are accepted in modern literature (Komárek and Anagnostidis, 1998), but few are well known in terms of variability, ecology, and distribution. E. cornuana was observed in U.S. mountain streams by Silva (Whitford and Schumacher, 1969). E. atrata and E. lithophila were described from stroma-

tolites in a saline volcanic lake in central Mexico (Tavera and Komárek, 1996), and E. willei was found in wet rocks in Puerto Rico (Gardner, 1927). Probably more forms belong to this genus, but taxonomic revision is needed. The very widely conceived E. lemaniae (recorded from North America by Drouet and Daily, 1956) contains a cluster of various types; the original E. lemaniae was described from the Baltic Sea and in this (taxonomically unclear) concept probably has not been collected from North America. Several marine species are known from Pacific coasts, North Carolina, and the Bahamas.

Hydrococcaceae (Fig. 16A) Several species from the genera Hormathonema Ercegovic´ and Placoma Schousboe ex Bornet and Thuret occur in North America, but are known only from marine environments. Hydrococcus Kützing (Fig. 16A) Colonies of cells are initially flat, in single layers, and pseudoparenchymatous (nematoparenchymatous at the margin) with more or less radially arranged cells and more or less circular outline. Aggregates of cells and erect pseudofilaments grow from a colonial center and form clusters of cells in older colonies. Pseudofilaments and groups of cells are embedded in thin, colorless, and confluent sheaths. Old colonies are flattened to hemispherical and are blackish green, brownish, or violet in color. In the center are groups of cells enveloped by individual mucilaginous sheaths, distant from one another, sometimes even forming sarcinoid packets. The cells are spherical, oval, irregular–rounded, or elongate at the ends of pseudofilaments, pale blue–green or olive green, and have more or less homogeneous content. They are 1.2–7.5 µm in diameter and have no obvious gas vesicles. Cell division occurs in various planes in successive cell cycles, but crosswise binary fission prevails in marginal parts of the colonies. Reproduction is by liberated solitary cells and clusters of cells. Several species have been described (partly under the synonymous genus Oncobyrsa Meneghini), of which H. cesatii and H. rivularis are known from North America (Smith, 1950); one morphotype was registered in Canadian Arctic (Whelden, 1947). Species have been reported on a variety of subtrata (e.g., rocks and wood) from streams in eastern and central Canada and the United States, Puerto Rico, and Hawaii (as Entophysalis rivularis; Drouet and Daily, 1956). However, the whole genus is little known.

Chamaesiphonaceae (Figs. 16A–18A) Chamaecalyx Komárek et Anagnostidis (Fig. 18A) Cells are heteropolar, solitary or in groups,


FIGURE 15 (A) Chlorogloea: a. C. epiphytica (after Komárek and Montejano, 1994, from San Luis Potosí, Mexico); b. C. lithogenes (after Komárek and Montejano, 1994, from San Luis Potosí, Mexico). (B) Entophysalis; a. E. willei (after Gardner, 1927, from Puerto Rico); b. E. lithophila (after Tavera and Komárek, 1996, from volcanic lakes in central Mexico).

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FIGURE 16 (A) Hydrococcus: a. H. cesatii (after Geitler, 1960); b. H. sp. (after Whelden, 1947, from Canadian Arctic). (B) Geitleribactron: a. G. periphyticum (after Komárek, 1975, and Hällfors and Munsterhjelm, 1982); b. G. crassum (bar = 10 µm; after Gold-Morgan et al., 1996, from central Mexico). (C) Clastidium: a. C. setigerum (after Starmach, 1966); b. C. cylindricum (after Whelden, 1947, from Canadian Arctic).


FIGURE 17 (A) Stichosiphon: a. S. exiguus (after Montejano et al., 1997, from San Luis Potosí, Mexico); b. S. willei (after Gardner, 1927, from Puerto Rico, sub Chamaesiphon willei); c. S. sansibaricus (after Whelden, 1941, from Florida, and, microphoto after Montejano et al., 1997, from central Mexico). (B) Chamaesiphon: a. C. (Chamaesiphon) incrustans (after Smith, 1950, from the United States); b. C. (Chamaesiphon) amethystinus (after Komárek, 1989, from Cuba); c. C. (Godlewskia) polonicus (from Geitler, 1932); d. C. (Godlewskia) subglobosus (after Waterbury and Stanier, 1977; sub Chamaesiphon sp., from culture).

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FIGURE 18 (A) Chamaecalyx: a. C. swirenkoi (after Geitler, 1932; microphoto from Gold-Morgan et al., 1996, from central Mexico); b. C. calyculatus (after Gold-Morgan et al., 1996, from San Luis Potosí, Mexico); c. C. suffultus (after Setchell and Gardner from Geitler, 1932, from California, marine species); d. C. clavatus (after Setchell and Gardner from Kosinskaja, 1948, from Baja California, Mexico; marine species). (B) Cyanocystis: a. C. valiae-allorgei (after Bourrelly, 1985, from thermal waters in Guadeloupe); b. C. mexicana (after Montejano et al., 1993, from central America); c. C. pacifica (after Setchell and Gardner from Smith, 1950; a marine species from North America; sub Dermocarpa pacifica). (C) Dermocarpella: a. D. hemisphaerica (after Lemmermann from Geitler, 1932; freshwater species from Chatham Islands, New Zealand).


obovoid to club-shaped, attached to substrata by narrowing basal ends (sometimes combined with a sheath pad), enveloped by a thin or slightly thickened, firm, colorless sheath (pseudovagina). The dimensions of older cells are 8–30(55) ⫻ 3.5–10.5(20) µm. The cell content usually is homogeneous (thylakoids are regularly distributed throughout the cells) and is pale blue–green, olive green, or reddish. Reproduction is by exocytes, which differentiate three dimensionally from the upper part of the cells after the first crosswise (horizontal) and the second vertical cell division, and which liberate from the sheaths of the mother cell by the apical opening. Differentiation of baeocytes is successive or simultaneous, while the basal part remains undivided; basal parts may grow into new mother cells, but rarely does the entire cell divide. Of the 12 known species, the freshwater C. swirenkoi and (possibly endemic) C. calyculatus were observed growing on filamentous algae in running waters in central Mexico (Gold-Morgan et al., 1996). Four species are known from marine coastal environments of North America (Setchell and Gardner, 1930; Komárek and Anagnostidis, 1986). Chamaesiphon A. Braun et Grunow in Rabenhorst (Fig. 17B) Cells are heteropolar, solitary or in groups (subg. Chamaesiphon), or form multilayered colonies with numerous cells (more or less parallely arranged) that form radial or shrublike, microscopic to macroscopic colonies (subg. Godlewskia) attached by their bases to plants, algae, or stony substrata. Cells are slightly or distinctly elongated, subspherical, oval, cylindrical up to slightly club-shaped, always enveloped by a sheath (pseudovagina) or (subg. Godlewskia) surrounded by sheaths and a common gelatinous, colorless or slightly yellowish to brownish matrix, (1)2.5–9(–70–200) ⫻ 1–8.5(13) µm in size, and have pale blue–green, olive green, greyish, pinkish, or reddish violet content, and no obvious gas vesicles, but sometimes a few prominent granules. Cells are sometimes slightly withdrawn from the narrowed basal sheaths. Cell division is asymmetric, crosswise near the apex; the exocytes separate solitarily or form short vertical rows before release from opened sheaths. In the subgenus Godlewskia, the exocytes attach to the margin of the opened sheaths or pseudovaginae (the origin of shrublike or multilayered colonies). This widespread and occasionally important genus in running waters has about 30 known species, mainly from alpine waters or from clear, cold streams worldwide (Geitler, 1925; Kann, 1972, 1973). C. britannicus (= C. regularis), C. incrustans, C. rostafinskii, C. polonicus, C. confervicolus, and possibly also C. geitleri are

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known from mountainous streams in North American temperate to subarctic zones (Smith, 1950; Stein and Borden, 1979; Sheath and Steinman, 1982); also a common epiphyte on Cladophora in hardwater lakes (e.g., Great Lakes). C. minutus occurs in mountains in Mexico (Gold-Morgan et al., 1996). C. amethystinus, C. fallax, and C. portoricensis occur in warmer regions (Cuba, Mexico, and Puerto Rico; Gardner, 1927; Komárek, 1989; Gold-Morgan et al., 1996). C. halophilus, a red-pigmented species, was described from a volcanic lake in Mexico and possibly is endemic (Tavera and Komárek, 1996). Clastidium Kirchner (Fig. 16C) Cells are heteropolar, solitary or in groups, elongated, and attached to the substrate by morphologically differentiated basal ends (usually narrowed and rounded with gelatinous pad). The apical part is conically tapered. The cells are enveloped by very thin, fine, colorless mucilaginous sheaths, which may be elongated into hairlike processes at the apex. The resulting cell form is oval, ovoid, ellipsoidal, cylindrical to slightly club-shaped, or pear-shaped, the cells are 2–15(38) ⫻ 1–4(6) µm, and the hairs are up to 52(75) µm long (different lengths among species). The cell content is pale blue–green, homogeneous, and has no obvious gas vesicles or prominent granules. Cell division is transverse, more or less simultaneously near the apex (rarely along the whole cell length) into a row of nearly spherical or oval and motile exocytes, which separate successively. The remnant of the mother cell may form the new vegetative cell. Five epilithic and epiphytic species are known from clear (Komárek and Anagnostidis, 1998), usually cold mountain streams and, rarely, lakes. Two species are known in this region: C. setigerum from the Northwest Territories, British Columbia, the northern United States, Alaska, Louisiana, and Mississippi (Drouet and Daily, 1956; Stein and Borden, 1979; Sheath and Steinman, 1982) and C. cylindricum from arctic Canada (Whelden, 1947). Geitleribactron Komárek (Fig. 16B) Cells are heteropolar, solitary or in groups, elongated, attached by one end to substrata (usually by indistinct, hyaline, gelatinous pad), and in groups or in stellate clusters. Cells are oval to cylindrical and rod-shaped, rounded at the apex, slightly and shortly attenuate or rounded at the base, have no sheaths, are pale blue–green or olive green, 4–25(50) ⫻ 1.3–6 µm in size, have finely granular content with no obvious gas vesicles, and sometimes have visible centro- and chromatoplasm (= parietal thylakoids). Cell division is by transverse, symmetrical or asymmetrical binary fision;

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rarely by two simultaneous divisions. Released daughter cells (exocytes) attach to substrata by either end. Three freshwater periphytic and epilithic species have been described (Komárek and Anagnostidis, 1998). G. crassum was described from running waters in central Mexico and is epiphytic on filamentous algae (Gold-Morgan et al., 1996). Stichosiphon Geitler (Fig. 17A) Cells are heteropolar, solitary or in groups, attached by the slightly narrowed base to substrata (usually also by means of a mucilaginous pad) and later form a long, uniseriate, straight or curved row of transversally differentiating exocytes at the apical end. The exocytes remain joined and form moniliform or trichome-like pseudofilaments, enveloped by thin, colorless sheaths that open at the apex. Exocytes may be irregularly clustered in the terminal part of a sheath. The pseudofilamentous rows of exocytes differ in the lengths among species, and measure (4)8–200(450) ⫻ (2.5)3–9 µm. Collectively, the complete cells with exocytes have the appearance of heteropolar, filamentous forms. Cells are shortly cylindrical or club-shaped when young, pale blue–green, olive green, or greyish in color, finely granular, have no gas vesicles, rarely have solitary granules, and usually have homogeneous or keritomized cell contents (thylakoids are located over the whole cell volume). Cell division is transverse with numerous exocytes in a close sequence; exocytes remain for extended periods in a filamentous formation within sheaths and can divide repeatedly. Exocytes are rarely rounded, usually slightly cylindrical, and after liberation attach to the substrate by either end. More than 10 species are known, mainly from warmer regions (Komárek and Anagnostidis, 1998). S. willei and S. gardneri have been described from swamps in the Greater Antilles; S. regularis and S. filamentosus were recorded in Mexico, where S. exiguus was also described from a limestone region in San Luis Potosí (Gardner, 1927; Komárek, 1989; Montejano et al., 1997). S. sansibaricus was described from wetlands in Florida (Whelden, 1941; Smith, 1950) and Mexico (Montejano et al., 1997).

Dermocarpellaceae (Figs. 18B and 19A) Cyanocystis Borzì (Fig. 18B) Cells are typically heteropolar, solitary or agglomerated in flat groups, usually slightly to distinctly elongated, widely oval, obovoid, club-shaped, or pearshaped, rarely nearly spherical, and rounded apically. The cells are attached to substrata by means of a sheath, which may be slightly widened at the base; sheaths (pseudovaginae) are thin, firm, and colorless.

Cells vary in size and have homogeneous, pale blue–green, olive green, or violet content, never have obvious gas vesicles, rarely have scattered prominent granules, and have very different sizes in various species, (5)6.2–30(90) ⫻ (0.7)3.6–21(30) µm. Cell division is by successive multiple fission into numerous spherical, nonmotile baeocytes, which liberate from the sheath by the rupture at the cell apex. The first division plane is always vertical from the apex to the base. About 17 species have been revised, mainly from marine, rarely freshwater environments (Komárek and Anagnostidis, 1998). Freshwater species C. pseudoxenococcoides (from Guadeloupe) and C. mexicana (from Mexico) were described from running waters of Central America (Bourrelly, 1985; Montejano et al., 1993). C. valiae-allorgei is known from thermal waters in Guadeloupe (Bourrelly and Manguin, 1952; Bourrelly, 1985). Several marine species have been recorded, particularly from the California coast (C. hemisphaerica, C. pacifica, and C. sphaeroidea); C. olivacea and C. violacea are probably distributed in marine coastal waters up to temperate zone (Hua et al., 1989). Many other similar species need revision. Dermocarpella Lemmermann (Fig. 18C) Cells are heteropolar, solitary or in groups, hemispherical, oval or club-shaped, rounded at the apex, and attached to substrata at the widened base or by narrowed ends. Sheaths are thin, firm, and colorless. Cells are more or less homogeneous or finely granular, pale or bright blue–green, olive green, or brownish, have no gas vesicles, and are (6)12–27(100) ⫻ (3)4– 27(40) µm in size. Cell division is only by complete, successive multiple fission into baeocytes, which liberate from the sheaths by the rupture at the apex. The first (and usually first several) division plane is always horizontal to the substrate. Only five species are confirmed, two of them from freshwaters (Komárek and Anagnostidis, 1998). Marine epiphytic species D. prasina and D. protea occur along both the Atlantic and Pacific coasts of North America. Stanieria Komárek et Anagnostidis (Fig. 19A) Cells are solitary or irregularly clustered in groups, more or less spherical, and attached to substrata by means of mucilaginous sheaths, but have no distinct cell polarity (attachment by any side). Sheaths are thin or slightly thickened, firm, and colorless. Cells are of variable size, pale blue–green, blue–green, yellowish, olive green, or pinkish-reddish, and have more or less homogeneous content and no prominent granules or


FIGURE 19 (A) Stanieria: a, b. S. cyanosphaera (after Komárek and Hindák, 1975, from Cuba; sub Chroococcidiopsis cyanosphaera). (B) Chroococcidium: a. C. gelatinosum (after Tavera and Komárek, 1996, from a volcanic lake in central Mexico).

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gas vesicles. Cell division (total) is by multiple fission, which proceeds in rapid sequence in various directions or almost simultaneously. The resulting numerous, motile spherical baeocytes liberate after splitting of the sheath. The difference in size of the small baeocytes and the mother cells is obvious. Baeocytes grow to the original size before the next division. Several marine species have been recognized. S. sphaerica was originally described from California and a few of them are not yet well defined taxonomically. S. cyanosphaera (one of three freshwater species) is known from alkaline swamps and littoral of lakes in Cuba (Komárek and Anagnostidis, 1998).

Xenococcaceae (Figs. 19B–21) Chroococcidiopsis Geitler (Fig. 20A) Cells are solitary or aggregated in irregular groups, enveloped by thin, firm, colorless sheaths, and usually attached to various, mostly stony substrates in a variety of subaerial and aquatic environments, sometimes under extreme conditions; endolithic species are also known. Cells are spherical, oval to irregular–rounded, of varying sizes, have more or less homogeneous pigmentation (thylakoids are distributed irregularly), pale or bright blue–green, rarely greyish or violet, and 1.5–20 (rarely more) µm diameter. Cells divide irregularly, successively, or in rapid sequence in cells of different size, or in numerous baeocytes, which liberate from ruptured sheaths; sheaths sometimes gelatinize. More than 20 morpho- and ecotypes (species?) are mentioned in the literature (Komárek and Anagnostidis, 1998), but sometimes are not clearly defined. C. thermalis has been recorded from hot springs and C. cubana has been noted from wetlands and littoral regions of standing waters in Florida, Cuba, and Mexico (Komárek and Hindák, 1975; Komárek and Anagnostidis, 1998). Taxonomically not well described forms occur in deserts of southwestern regions of North America, forming lichens (in Mexico; Büdel, 1985; Büdel and Henssen, 1983), or in various habitats in Puerto Rico (but assigned to Anacystis species; Gardner, 1927). Chroococcidium Geitler (Fig. 19B) Cells are clustered in irregular groups within thin, amorphous, colorless, and diffuse mucilage, usually attached to stony substrata; less frequently they are epiphytic. Cells are of different size, mainly spherical or irregularly rounded, sometimes with diffuse individual envelopes around cells and around cell groups, and have 3–18 µm diameter, with homogeneous cell content, yellow–green or blue–green color. Cell division is irregular in various planes, sometimes rapidly into

numerous nanocytes, which liberate by gelatinization of envelopes. One species, C. gelatinosum, originally described from Indonesia, occurs rarely in volcanic lakes in central Mexico (Geitler and Ruttner, 1935; Tavera and Komárek, 1996). Probably more species will be recognized in the future. Chroococcopsis Geitler (Fig. 21B) Cells are solitary, in few-celled clusters, or compact groups ensheathed by firm, thin, or slightly thickened envelopes; rarely they are organized in short, indistinct and irregular rows. Distinctly larger, terminal cells develop near the margins of older colonies. Sheaths are colorless and sometimes slightly layered. Cells are spherical, subspherical, oval, or irregular– rounded; marginal cells are club-shaped or pyriform (radially arranged). Cell size varies widely (2.5–36 µm in the whole genus). Cell content is homogeneous or finely granular and blue–green, olive green to slightly violet in color. Cell division is by irregular binary fission and, in older colonies (usually marginal), by multiple fission in numerous baeocytes, which liberate from the ruptured and/or gelatinized sheaths. Of five species, C. fluviatilis (probably a special morphotype) has been recorded from the United States (Smith, 1950; sub Pleurocapsa fluviatilis). An undescribed species was observed in Cuban streams on limestone substrata (Komárek, 1985). Myxosarcina Printz (Fig. 20B) Cells are densely agglomerated in packet-like, “sarcinoid” groups, irregular or polygonal–rounded, and sometimes in slightly flattened colonies. Subcolonies are aggregated sometimes in granular mats, attached to substrata or free-living among other algae or in detritus. Firm, mucilaginous envelopes are thin or distinct, and colorless, rarely yellowish brownish. Cells are homogeneous, dark or pale olive green or blue–green in color, rarely violet. Reproduction is by irregular binary fission in three or more planes in successive generations, sometimes obliquely, forming packet-like colonies; several cells in colonies divide rapidly into motile baeocytes, which usually liberate from the split sheaths. Colonies occasionally disintegrate. More than 10 species have been recognized (Komárek and Anagnostidis, 1998). At least two have been described from North America: M. amethystina from thermal springs in Yellowstone Park (Copeland, 1936) and M. gloeocapsoides from salt marshes in California (Gardner, 1918). M. rubra has been recorded from moist aerial environments (on rocks and wood near springs) from Puerto Rico (Gardner, 1927; Bourrelly, 1985), but its taxonomic identity needs


FIGURE 20 (A) Chroococcidiopsis: a. C. thermalis (after Geitler from Geitler and Ruttner, 1935); b. C. cubana (after Komárek and Hindák, 1975, from Cuba). (B) Myxosarcina: a. M. amethystina (after Copeland, 1936, from Yellowstone National Park); b. M. gloeocapsoides (after Setchell and Gardner in Gardner, 1918, from California; marine species; sub Pleurocapsa gloeocapsoides).

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FIGURE 21 (A) Xenococcus: a. X. willei (after Gardner, 1927, from Puerto Rico, and Montejano et al., 1993, from Mexico); b. X. bicudoi (after Montejano et al., 1993, from Mexico). (B) Chroococcopsis: a. C. fluviatilis (after Geitler, 1932, and Lagerheim from Geitler, 1942). (C) Xenotholos: a. X. kerneri (after Geitler, 1932); b. X. huastecanus (after Gold-Morgan et al., 1994, from San Luis Potosí, Mexico).

clarification. More species may be described from Caribbean habitats. Xenococcus Thuret in Bornet et Thuret (Fig. 21A) Cells are attached to substrata (usually filamentous algae) in clusters (sometimes densely arranged), enveloped by thin, firm, rarely gelatinizing, colorless or

yellowish–brownish sheaths. Cells in clusters are always in one layer, typically heteropolar, spherical to oval, pear-shaped, or club-shaped, and rounded at the apex. Cell content is homogeneous, pale or bright blue–green, yellow–green, or pinkish violet. Cells are 1.5–12(23) µm in diameter. Binary fission occurs in two or more planes, usually perpendicular to the sub-


stratum. Occasionally, cells divide by multiple fission in numerous small baeocytes, which liberate from split sheaths. Baeocytes are formed from whole cells or upper portions of mother cells. Twenty-six well described species, plus a similar number not yet revised (Komárek and Anagnostidis, 1998), are known from freshwater and saline environments worldwide. X. yellowstonensis was described from thermal waters (> 50°C) in Yellowstone (Copeland, 1936). X. bicudoi, X. lamellosus, and X. willei were described from freshwater habitats in Mexico and Puerto Rico, mainly as epiphytes on algae in streams (Gardner, 1927; Montejano et al., 1993; Gold-Morgan et al., 1994). X. candelariae, which has reddish cells, is a deep-water species from a volcanic lake in central Mexico that is epiphytic on Cladophora (Tavera and Komárek, 1996). Marine species X. pallidus, X. schousboei, X. pyriformis, X. angulatus, X. chaetomorphae, X. cladophorae, X. deformans, and X. gilkeyae have been recorded from various localities along the Atlantic and Pacific coasts (Gardner, 1918; Setchell and Gardner, 1924; Smith, 1950). Xenotholos Gold-Morgan, Montejano et Komárek (Fig. 21C) Cells are arranged in thalli (colonies) attached to the substrata. Colonies develop from solitary cells that are first disklike and later layered, usually forming a slightly globose (irregular–hemispherical) colony, in which the cells are organized more or less in radial rows, and finally in two or more layers. The colony is enveloped by a thin, firm, colorless sheath. Cells are hemispherical, polygonal–rounded to slightly elongated, sometimes with individual sheaths. Cells divide in various planes, occasionally (marginal parts) into baeocytes. Reproduction is by solitary cells and baeocytes. X. kerneri is not common, but is a widespread species in cold mountain streams, probably across North America (Smith, 1950, under Xenococcus kerneri). Three other species were described from mountains in central Mexico (Gold-Morgan et al., 1994).

Hyellaceae (Figs. 22 and 23) Hyella Bornet et Flahault (Fig. 23) Cells are organized in differentiated thalli composed of irregular clusters of cells and pseudofilaments, sometimes ramified. The thalli creep on calcareous substrata or are epiphytic creeping into intercellular spaces of host plants, or endolithic actively growing into substrata. Pseudofilaments are uni- to multiseriate, sometimes pseudodichotomously divided; on the surface (older thalli) they remain in nematoparenchymatous cell clusters. The developed thallus is usually composed

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from an epilithic (or epiphytic) component with irregularly arranged cells, and endolithic or endophytic component of pseudofilaments boring into substrata (or growing in intercellular spaces of seaweeds). Mucilaginous sheaths are firm, thick, and rarely gelatinizing. Cell morphology varies in each part of the thallus. Cells of very different sizes (2.5–55 µm in diameter) are more or less spherical, subspherical, polygonal–rounded, or elongate, pale or bright blue–green, olive green, or pinkish violet in color, and oval or club-shaped at the ends of pseudofilaments (= apical with respect to growth). Cells divide by binary fission in various planes, in pseudofilaments perpendicularly to the long axis, before branching longitudinally or obliquely. The oldest cells (basal, i.e., near the surface of stony substrata or of host plants) may divide into baeocytes—reproductive cells. About 30 species have been described (Golubic´ et al., 1975, 1981; Al-Thukair et al., 1994; Lukas and Golubic´, 1983; Lukas and Hoffman, 1984), usually from marine coastal environments (warm seas). H. kalligrammos has been collected from freshwater habitats in limestone areas of Mexico, but other forms occur in central and North America (e.g., Hyella cf. fontana in creeks in limestone areas in Cuba; Komárek, 1985). From marine locations, endolithic H. balani, H. gigas, H. pyxis, H. caespitosa, H. tenuior, H. linearis, H. littorinae, and H. vacans, and the epiphytic H. seriata are known, particularly from warmer seas (including the Bahamas and Bermuda; Gardner, 1918; Hollenberg, 1939; Lukas and Golubic´, 1983; Gektidis and Golubic´, 1996). Pleurocapsa Thuret in Hauck (Fig. 22B) Cells are arranged in irregular groups or rows and pseudofilaments, creep on substrata (mostly stones), and sometimes pseudodichotomously divide. Pseudofilaments are partly endolithic in several species or form crustose layers. Rows of cells are uni- to multiseriate and enveloped more or less by thin, firm, sometimes lamellate and yellow brownish, confluent sheaths. Cells are irregular, variable in size, sometimes slightly elongate, have homogeneous or slightly granular content, blue–green, pale blue–green, or pinkish color, and are (2.4)3–15(20) µm in diameter. Cells divide irregularly by binary fission in various planes; in pseudofilaments predominantly crosswise. Enlarged cells, which arise in different parts of a thallus, divide into baeocytes, which escape from the gelatinized and divided cells. Over 20 species have been described (Komárek and Anagnostidis, 1998), but require revision. The genus is poorly known and perhaps has two distinct groups (Waterbury and Stanier, 1978). P. minor occurs in

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FIGURE 22 (A) Radaisia: a. R. epiphytica (after Setchell and Gardner in Gardner, 1918, from California, marine species); b. R. gardneri (after Gardner, 1927, from Puerto Rico; sub Pleurocapsa epiphytica). (B) Pleurocapsa: a. P. minor (after Geitler, 1932); b. P. crepidinum (after Geitler, 1932, from coasts of North America; marine species); c. P. minuta (after Geitler, 1932; marine species).

calcareous streams in North America (Smith, 1950), but other freshwater species occur (e.g., taxonomically uncertain P. varia from numerous localities in the United States; Daily, 1942). Four marine species have been recorded from North American coasts (Setchell and Gardner, 1919; Setchell, 1924; Weber van Bosse, 1925).

Radaisia Sauvageau (Fig. 22A) Cells are arranged initially in flat, discoid, or irregular, nemato- or blastoparenchymatous layers on substrata, from which grow erect, more or less parallel pseudofilamentous rows of cells, which may be straight or slightly curved and sometimes divided. Old colonies form flat, gelatinous and crustose layers of pseudo-


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FIGURE 23 Hyella: a. H. fontana (after Smith, 1950, from North America); b. H. cf. fontana (after Komárek, 1985, from Cuba); c. H. seriata (after Hollenberg, 1939, from California; epiphytic marine species); d. H. caespitosa (after LeCampion-Alsumard and Golubic´, 1985; endolithic marine species); e. H. balani (after LeCampion-Alsumard and Golubic´, 1985; endolithic marine species; bar = 10 µm).

filaments in parallel arrangement, perpendicular to the substratum. Mucilaginous sheaths present around the pseudofilaments, are confluent later in a homogeneous mass. Cells are irregular, polyhedral–rounded, usually elongated toward the ends of the pseudofilaments, pale blue–green or reddish violet in color, and have 2.5–9(10) µm diameter. Cell division is irregular,

usually crosswise to the pseudofilament. Apical, usually larger cells divide into baeocytes, which arise by simultaneous or successive cell division and escape from the ruptured or gelatinized envelopes. About 10 species have been described (Komárek and Anagnostidis, 1998): three freshwater species are known from standing waters in Puerto Rico (R. conflu-

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ens, R. gardneri, and R. willei; Gardner, 1927; Komárek and Anagnostidis, 1998); and three marine species were described from coastal areas of California; one from Florida (Gardner, 1918, 1927; Weber van Bosse, 1926).

VI. GUIDE TO LITERATURE FOR SPECIES IDENTIFICATION This guide applies to cyanobacterial taxa covered in chapters 3 and 4. The taxonomy of cyanobacteria has changed drastically as a consequence of current molecular, biochemical, ecological, and ultrastructural data, but few of these changes are reflected even in newer identification keys. The reader must still rely on the older literature until these changes are incorporated into general keys. For North America, Tilden (1910) is still useful. Several cyanobacterial species in the Great Lakes region can be identified using Prescott (1962). Two other taxonomic works written in English are from India (Desikachary, 1959) and the British Isles (Whitton, 2002). Most other important works were not written in English, but should be consulted, especially Geitler (1932), as well as monographs covering continental Europe (Huber-Pestalozzi, 1938; Starmach, 1966), and Russia (Elenkin, 1936, 1938, 1949; Kosinskaja, 1948; Kondrateva, 1968). Some modern results are incorporated into in recent monographs (in English) on the Chroococcales (Komárek and Anagnostidis, 1986, 1998), and the filamentous orders Oscillatoriales, Stigonematales, and Nostocales (Anagnostidis and Komárek, 1985, 1988, 1990; Komárek and Anagnostidis, 1989). Castenholz (2001) has recently written (in English) an overview of cyanobacterial classification.

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Wehr, J. D. 1990. Predominance of picoplankton and nanoplankton in eutrophic Calder lake. Hydrobiologia 203:35–44. Wehr, J. D. 1992. Effects of experimental manipulations of light and phosphorus supply on competition among picoplankton and nanoplankton in an oligotrophic lake. Canadian Journal of Fisheries and Aquatic Sciences 50:936–945. Wehr, J. D., Descy, J.-P. 1998. Use of phytoplankton in large river management. Journal of Phycology 34:741–749. Wehr, J. D., Thorp, J. H. 1997. Impacts of navigation dams, tributaries and littoral zones on phytoplankton communities in the Ohio River. Canadian Journal of Fisheries and Aquatic Sciences 54:378–395. Weisse, T. 1993. Dynamics of autotrophic picoplankton in marine and freshwater ecosystems. Microbial Ecology 13:327–370. Wettstein, R. 1924. Handbuch der systematischen Botanik. LeipzigWien. Whelden, R. M. 1941. Some observations on freshwater algae of Florida. Journal of the Elisha Mitchell Science Society 57:261–272. Whelden, R. M. 1947. Algae, in: Polunin, N., Ed., Botany of Canadian eastern Arctic, II. Thallophyta and Bryophyta. Bulletin of the National Museum of Canada 97:13–127. Whitford, L. A., Schumacher, G. J. 1969. A manual of the freshwater algae in North Carolina. North Carolina Agricultural Experimental Station Technical Bulletin 188:1–313. Whitton, B. A. 1987. The biology of Rivulariaceae, in: Fay, P., Van Baalen, C., Eds., The cyanobacteria, Elsevier, Amsterdam, pp. 513–534. Whitton, B. A. 1992. Diversity, ecology, and taxonomy of the cyanobacteria, in: Mann, N. H., Carr, N. G., Eds., Photosynthetic prokaryotes. Plenum Press, New York, pp. 1–51. Whitton, B. A. 2002. Phylum Cyanophyta (Cyanobacteria). in: John, D. M., Whitton, B. A., Brock, A. J., Eds., The freshwater algal flora of the British Isles. An identification guide to freshwater and terrestrial algae. Cambridge University Press, pp. 25–122. Whitton, B. A., Carr, N. G. 1982. Cyanobacteria: Current perspectives, in: Carr, N. G., Whitton, B. A., Eds., The biology of cyanobacteria. Blackwell, Oxford, pp. 1–8. Whitton, B. A., Potts, M. 1982. Marine littoral, in: Carr, N. G., Whitton, B. A., Eds. 1982. The biology of cyanobacteria. Blackwell, Oxford, 515–542. Whitton, B. A., Potts, M., Eds. 2000. The ecology of cyanobacteria: Their diversity in time and space, Kluwer, Boston. Wilmotte, A., Stam, W. T. 1984. Genetic relationships among cyanobacterial strains originally designated as “Anacystis nidulans” and some other Synechococcus strains. Journal of General Microbiology 130:2737–2740. Wojciechowski, I. 1971. Die Plankton-Flora der Seen in der Umgebung von Sosnowica (Ostpolen). Annals of the University M. Curie-Skl⁄ odowska, Lublin, 26:233–263. Wood, H. C. 1869. Prodromus of a study of the fresh water algae of eastern North America. Proceedings of the American Philosophical Society 11:119–145. Wood, H. C. 1872. A contribution to the history of the fresh-water algae of North America. Smithsonian Contributions to Knowledge 19(241):1–262. Zalessky, M. M. 1926. Sur les nouvelles algues découvertes dans le sapropélogéne du lac Beloe. Revue Générale de Botanique 38:31–42. Zohary, T., Breen C. M. 1989. Environmental factors favoring the formation of Microcystis aeruginosa hyperscums in a hypertrophic lake. Hydrobiologia 178:179–192. Zohary, T., Robarts, R. D. 1990. Hyperscums and the population dynamics of Microcystis aeruginosa. Journal of Plankton Research 12:423–432.

4

FILAMENTOUS CYANOBACTERIA Jirˇí Komárek

Jaroslava Komárková

Institute of Botany Academy of Sciences of the Czech Republic Faculty of Biological Sciences University of South Bohemia CZ-37982 Trˇebonˇ, Czech Republic

Hydrobiological Institute Academy of Sciences of the Czech Republic Faculty of Biological Sciences University of South Bohemia CZ-37005 Cˇeské Budeˇ jovice Czech Republic

Hedy Kling Freshwater Institute Winnipeg, Manitoba Canada R3T 2N6 I. Introduction II. Morphology A. Cytology and Morphology B. Specialized Cells C. Reproduction III. Ecology IV. Methods

I. INTRODUCTION Filamentous cyanobacteria (blue–green algae, cyanoprokaryotes) include some of the most widely recognized and important freshwater algae in the world, many of which produce surface blooms, fix atmospheric nitrogen, and are important components of global carbon fixation (Fogg et al., 1973; Fay and Van Baalen, 1987; Whitton and Potts, 2000). Basic information on the biology and ecology of cyanobacteria is presented in Chapter 3 on coccoid forms; this chapter contains the information concerning the unique morphology and biology of filamentous blue–greens. The system that has been used for several decades for distinguishing genera of filamentous cyanobacteria is based on phenotypic characters that recently have been supported by ultrastructural and molecular data in many instances (Rippka et al., 1979; Anagnostidis and Komárek, 1988; Komárek and Anagnostidis, 1989). Current molecular Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

V. Key and Descriptions of Genera A. Key B. Descriptions of Genera Note Added in Proof VI. Guide to Literature for Species Identification Literature Cited

analyses support the separation of non-heterocystous and heterocystous genera1; however, we can expect further changes on both the generic and the species levels for many of these organisms (Castenholz, 1992; Li and Watanabe, 1998; Rudi et al., 1998, 2000). The present chapter recognizes three orders of filamentous cyanobacteria—Oscillatoriales, Nostocales and Stigonematales — which can be clearly defined by several diagnostic features (see Chap. 3, Sect. V.A). Several comprehensive reviews on filamentous cyanobacteria of North America have been published in the past. An early but thorough study on the distribution of blue–greens in North America is the monograph by Tilden (1910). Although largely using figures from other authors, with the list of exsiccates and very old records, it is a valuable view of the knowledge 1

The term “heterocyte” is used in this chapter to refer to cells commonly known as “heterocysts”; they are not truly cysts.

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of the North American cyanobacterial (= cyanoprokaryotic) flora from the beginning of this century. Several later studies present information on specific regions, including the upper Midwest (Wisconsin; Smith, 1920, 1950), Puerto Rico (Gardner, 1927), Yellowstone National Park (Copeland, 1936), the western Great Lakes area (Prescott, 1962), North Carolina (Whitford and Schumacher, 1969), and Illinois (Tiffany and Britton, 1952). Many of these past studies are still important sources of information. For example, in an account of Poulin’s collections from Arctic Canada, Whelden (1947) recorded 108 cyanobacterial taxa (following Geitler, 1932), comprising 15 genera, with 65 species that were filamentous. A number of more recent studies in many parts of North America document a particularly rich cyanobacterial flora (e.g., Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). The taxonomic status of many species and consequently their nomenclature have changed over the years, which may affect estimates of cyanobacterial diversity in future studies (Rippka et al., 1979; Anagnostidis and Komárek, 1988; Komárek and Anagnostidis, 1989; Castenholz, 1992). Nonetheless, previous published reports generally represent a useful source of information for current studies of the North American cyanobacterial microflora. In contrast, studies by Drouet and co-workers (Drouet and Daily, 1956; Drouet, 1968, 1973, 1981a, b) greatly oversimplified the taxonomy and classification of cyanobacteria and thus do not correspond to the true diversity in cyanobacterial assemblages in nature.

II. MORPHOLOGY A. Cytology and Morphology The internal cellular structure of filamentous cyanobacteria does not differ substantially from coccoid forms (Chap. 3, Sect. II.A and II.B). One important distinction is the arrangement of thylakoids inside cells, which differ from coccoid forms. Although there is wide variation in thylakoid patterns within the Chroococcales suggesting heterogeneity within that group, only three main types (with consistent variations) occur in the simplest filamentous order, the Oscillatoriales. These arrangements are parietal (family Pseudanabaenaceae; Figs. 1–3), radial (family Phormidiaceae; Figs. 12 and 15), and irregular (family Oscillatoriaceae; Fig. 9) (Anagnostidis and Komárek, 1988; Komárek and ˇ áslavská, 1991). Further, only one thylakoid type, irregC ular (similar to members of the family Oscillatoriaceae), exists in the Nostocales and Stigonematales. In all representatives of the filamentous orders, cells are arranged in so-called trichomes, forming one

physiological entity. Neighboring cells are connected through pores in their cross walls (microplasmodesms), the number and position of which are characteristic of the genera, and similar pores can sometimes be found on the outer cell walls, especially near the cross walls (Guglielmi and Cohen-Bazire, 1982a). Trichomes are capable of fragmentation (forming hormogonia) or complete disintegration into separate cells (Geitler, 1942, 1960; Watanabe and Komárek, 1989). Fragmentation proceeds after the formation of fine mucilaginous lamella between neighboring cells or via so-called necridic (sacrificial) cells (or necrids). These cells die off, after which the fragments separate (Geitler, 1960, 1982; Kondrateva, 1961; Anagnostidis and Komárek, 1988). Trichomes may be uniseriate, with cell division exclusively perpendicular to the trichome axis, or multiseriate in more differentiated types (Stigonemataceae), where cells also have the ability to divide longitudinally or in more directions. Cells of many genera produce slimy, colloidal substances external to the cell wall, composed of hydrated polysaccharides (Drews, 1973; Martin and Wyatt, 1974; Jürgens and Weckesser, 1985; deVecchi and Grilli-Caiola, 1986). They are recognizable in the form of mucilaginous envelopes or homogeneous or layered sheaths of various kinds (e.g., fine, firm, diffuse, homogeneous or lengthwise, perpendicular or funnel-like lamellated, colorless or variously colored) surrounding the trichomes. Trichomes with sheaths are traditionally termed filaments. The presence or absence of sheaths, or the ability to produce them under special conditions, is probably a genetically determined characteristic. The trichomes of various species may be morphologically uniform or exhibit a diversity of cell types. Species of some genera have the ability to form differentiated cells of various types at the ends of the trichome, which may have ecophysiological importance and may be used as a characteristic feature for identification of genera. Apical cells (one or more) may be narrower or wider (without thylakoids) or may be elongated. Apical cells in the family Rivulariaceae and in several genera of Nostocaceae (Nostocales) may become attenuated, elongated, and hyaline. These hairlike formations have been shown to be produced (or accentuated) under periods of inorganic phosphorus deficiency, and they may also be sites of alkaline phosphatase production (Livingstone et al., 1983; Whitton, 1987). The morphological diversity of trichomes is particularly noticeable in polarized (e.g., Calothrix and Rivularia) and branched types (e.g., Scytonematopsis and Fischerella; Figs. 20B, 25, and 27), where the basal parts of the filament are morphologically distinct from the apical parts or branches. In contrast to hairs, other

4. Filamentous Cyanobacteria

taxa have wider, shorter or spherical cells at the end of the filament, probably with a similar function (e.g., Trichodesmium and Scytonema). Facultative trichome motility is common in more simple forms or in segments of trichomes (motile hormogonia) that are liberated from the sheaths. Modifications of phototactic and other movements have been described in many species (Drews, 1959; Häder, 1974; Nultsch, 1974; Halfen, 1979; Whale and Walsby, 1984). For example, the genera Geitlerinema (motile) and Jaaginema (immotile) differ only in their motility (Anagnostidis and Komárek, 1988). Two basic types of branching occur, depending on the type of cell division and filament (+ sheath) morphology. In false branching, trichomes divide (often with necridic cells or at heterocytes) within the sheath and one or both ends of the divided trichome diverge, breaking out the sheath. The resulting filament (e.g., Tolypothrix and Scytonema; Figs. 20–23) consists of trichomes, diverging singly or in pairs from a common sheath, giving the appearance of branches. Modifications of this branching type are characteristic of certain genera (e.g., Oscillatoriales: Pseudophormidium, Blennothrix, and Plectonema; Nostocales: Tolypothrix, Hassallia, Scytonema, and Coleodesmium). True branching occurs in more complex forms (Stigonematales), in which the cells are also capable of longitudinal division (Figs. 35–40). The laterally dividing cell alters its growth polarity (direction of cell division), and grows more or less perpendicular to the original trichome, forming a lateral branch that is physiologically connected to the original trichome. Branching type appears to be a stable character in several genera (e.g., Brachytrichia, Hapalosiphon, and Stigonema; Desikachary, 1959; Bourrelly, 1970; Martin and Wyatt, 1974; Golubic´ et al., 1996); this morphology may also be related to filament polarity. Distinctly polarized filaments attached by one end to the substrate and with clearly polarized apical growth occur within several groups of filamentous cyanobacteria (e.g., Schizothrix and Rivularia). Filaments of some genera are often organized into different arrangements within colonies, and mucilage plays an important role in colony formation. Filaments of several planktonic and free-living metaphytic species can be arranged in clusters or fascicles, which are large enough to be recognizable in the field. Colonies with fasciculated (clustered) filaments are typical of several planktonic species in the genera Trichodesmium and Aphanizomenon and are radially arranged in spherical colonies in the genus Gloeotrichia. Periphytic species are capable of forming a large variety of mats and tufts on submerged as well as subaerial surfaces. Several species from the orders Nostocales and Stigonematales occur as slimy, irregular, or almost spherical macro-

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scopic colonies, including well-known taxa in the genus Nostoc. Other types of macroscopic brushlike fascicles occur in water or subaerial habitats, such as members of the genus Symploca or several species of Schizothrix and Symplocastrum. Characteristic stromatolite and travertine formations are produced by species precipitating calcium carbonate in their sheaths and within colonial mucilage. Several species from the genera Leptolyngbya, Schizothrix (subgenus Inactis), Calothrix, and Rivularia belong to the most important travertine-forming filamentous cyanobacteria (Pentecost and Whitton, 2000; Stal, 2000).

B. Specialized Cells Several members of orders within the filamentous cyanobacteria exhibit the greatest level of morphological and cellular differentiation. In the two most diversified orders, Nostocales and Stigonematales, two important types of specialized cells develop: heterocytes (heterocysts in old literature; see Komárek and Anagnostidis, 1989) and akinetes. Heterocytes develop from vegetative cells and may be solitary, in pairs, or several in a row (Fogg, 1949; Bahal and Talpasayi, 1972; Stewart, 1972; Wolk, 1982; Komárek and Anagnostidis, 1989). They produce cell walls, which are thick, multilayered, and apparently gas tight (Lang and Fay, 1971; Golecki and Drews, 1974; Stewart, 1980). During development, the thylakoid apparatus degrades and specific DNA rearrangements (e.g., nif genes) occur (Golden et al., 1985; Haselkorn, 1986). Heterocytes synthesize the enzyme nitrogenase, which enables fixation of gaseous nitrogen (N2) from the atmosphere (or dissolved in water) under anaerobic conditions, which are maintained within the heterocyte (Winkenbach and Wolk, 1973; Wolk, 1973, 1982). The morphology (shape) of the heterocytes and their position in the trichomes are apparently genetically predetermined, but their frequency in the trichomes in the populations depends on the nitrogen supply in the environment, with frequencies of these cells along the trichome declining with greater levels of NH4+ or NO3– (Bahal and Talpasayi, 1972; Stewart, 1972; Kohl et al., 1987). In some genera, heterocytes develop exclusively from the apical (terminal) vegetative cells (e.g., Cylindrospermum, Fig. 30) or intercalary (e.g., Anabaena, Fig. 28) at roughly regular distances from one another (metameric), or occasionally in pairs (e.g., Anabaenopsis, Fig. 29A) or in short rows. In polarized filaments, they have an obligatory position forming basal cells (e.g., Calothrix and Gloeotrichia, Figs. 25 and 26). Akinetes are resting cells that develop from solitary cells or after the fusion of two or more neighboring

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cells in several members of the Nostocales and Stigonematales (Geitler, 1932; Komárek and Anagnostidis, 1989; Anagnostidis and Komárek, 1990; Hindák, 2001). They often arise close to the end of the vegetative growth period, although their production is not obligatory (Rother and Fay, 1977). Akinetes possess thick cell walls, and an accumulation of photosynthetic assimilates and DNA increases during their development (Sutherland et al., 1985a, b). These cells function as resting stages to survive harsh conditions, such as winter temperatures, drought, or frost, and can germinate to form new filaments when conditions improve (Huber, 1984; Cmiech et al., 1986). The morphology of akinetes, their shape (spherical, ellipsoidal, cylindrical), size, position within the trichome, cell wall (epispore) characteristics (color, sculpture), and mode of germination apparently are genetically determined features (Komárek and Anagnostidis, 1989) and have been used as critical characters for distinguishing genera and species (e.g., Geitler, 1932; Desikachary, 1959). In some species in the genera Anabaena, Aphanizomenon, and Anabaenopsis (subfamily Anabaenoideae), akinetes develop in a defined configuration (up to five in a row) and in a regular position in relation to the heterocytes. Such akinetes can occur directly adjacent to the heterocytes or distant, within one to several vegetative cells from them. When mature, they are often much larger than the vegetative cells. In most species of Nostoc, Trichormus, and Nodularia (subfamily Nostocoideae), each vegetative cell may become an akinete, which develop in rows between the heterocytes and do not reach a size any larger than the vegetative cells (Komárek and Anagnostidis, 1989). Specialized cells with the same function as akinetes appear in the order Stigonematales; however, their morphology and position in the filaments are not as distinct as in true akinetes of Nostocales (Martin and Wyatt, 1974; Rippka et al., 1979; Anagnostidis and Komárek, 1990).

C. Reproduction In principle, cell division in filamentous cyanobacteria proceeds in the same way as it does in coccoid forms, usually perpendicular to the trichome axis. Among members of the Stigonematales, division can proceed in several planes. Reproduction can occur through solitary cells, which are liberated by several types of filamentous species, or by fragmentation of a filament and of a thallus. However, the most frequent mode of reproduction is fragmentation, forming distinct segments of trichomes, termed hormogonia. Fragmentation into hormogonia is simple in more primitive genera in the Pseudanabaenaceae, where

trichomes disintegrate between neighboring cells via formation of thin lamella without necridic cells. In more complex filamentous taxa, hormogonia separate at the site of necridic cell production (Phormidiaceae and Oscillatoriaceae). Hormogonia can be of varying length, ranging from short filaments of several cells to fragments of single cells. Different types of hormogonia formation and their reproductive function are described in Anagnostidis and Komárek (1988). Longer hormogonia are sometimes motile and may contain gas vesicles (e.g., Fischerella). Fragments of ensheathed filaments are called hormocytes. Akinetes may also serve for reproduction (see Sect. II.B).

III. Ecology The ecological requirements and importance of filamentous blue–greens and their geographical distribution were discussed in Chapter 3 (Sect. III.A–III.E) and are thoroughly reviewed in Whitton and Potts (2000). Their range of environments is as wide as any aquatic or terrestrial organisms on Earth. Many of the filamentous taxa are important members of lake and river phytoplankton, whereas others form thick, dense mats in a wide variety of benthic habitats. Still others form conspicuous floating masses on the water surface of various water bodies, or grow attached to stones, as epiphytes on aquatic angiosperms and other algae, epipelic and epipsammic on sediments or sand, respectively, and frequently colonize wet or moist soils. Some planktonic species are capable of forming water blooms in mesotrophic and eutrophic water bodies throughout the world. Important reviews that discuss ecological features of filamentous cyanobacteria are summarized in several important manuals (Jaag, 1945; Anagnostidis, 1961; Fogg et al., 1973; Carr and Whitton, 1973; Golubic´, 1980; Whitton and Potts, 2000). Aside from the general discussion of cyanobacterial ecology in Chapter 3, the most important ecological aspects of filamentous cyanobacteria in nature are as follows: • Many are major components of autotrophic biomass production, forming a basis for aquatic food webs, with picoplanktonic, nanoplanktonic and bloom-forming species, particularly from the genera Cyanobium, Planktothrix, Anabaena, Aphanizomenon, Nodularia, and Cylindrospermopsis (Komárek, 1958, 1999; Kohl et al., 1985; Watanabe, 1971; Kondrateva, 1972; Komárek et al., 1993; Oliver and Ganf, 2000; Stockner et al., 2000; Komárková, 2001). • Filamentous cyanobacteria are often major components of attached communities in


submerged (littoral and benthic) habitats, in both standing and flowing waters, as well as on moist subaerial surfaces and on tree bark in tropical forests (Desikachary, 1959; Golubic´, 1967a; Whitton, 1984; Rott and Pipp, 1999). • Cyanobacteria are frequently the sole or dominant autotrophic organisms in extreme environments, including thermal, desert, and polar environments (Copeland, 1936; Anagnostidis, 1961; Golubic´, 1980; Broady, 1984; Vincent, 2000; Ward and Castenholz, 2000; Wynn-Williams, 2000). • Certain species produce toxins and allergic compounds, particularly in dense planktonic populations (Jackim and Gentile, 1968; Mahmood and Carmichael, 1986; Prinsep et al., 1992; Codd, 1995; Carmichael, 1997; Falconer, 1998; Chorus and Bartram, 1999; Dow and Swoboda, 2000). • A number of species is responsible for a large portion of global nitrogen fixation in aquatic and terrestrial ecosystems (e.g., tundra), and several species are used as “natural fertilizers” in rice fields (Fogg et al., 1973; Wolk, 1973; Stewart, 1980; Fay, 1983; Van Baalen, 1987; Whitton, 2000).

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• Particular species are responsible (at least in part) for lithogenic processes (travertine, stromatolites) and, in contrast, boring into limestone substrata (Prát, 1929; Carr and Whitton, 1973; Golubic´ et al., 1975, 1981; Pentecost and Whitton, 2000; Stal, 2000). • Cyanobacteria are key organisms in the colonization of soils and rocks during primary succession in terrestrial communities (Golubic´, 1967a). • Many cyanobacteria are used in biotechnology and in mass cultures (Seshadri and Jeeji-Bai, 1992; Belay et al., 1994; Vonshak, 1997). • Many cyanobacterial strains are important model laboratory organisms in the study of photosynthetic processes and physiological processes (Carr and Whitton, 1982; Fay and Van Baalen, 1987; Komagata, 1987; Sugawara and Miyazaki, 1999).

IV. METHODS Methods of collecting, preparing and culturing filamentous cyanobacteria do not differ substantially from those described for coccoid and colonial forms (see Chap. 3).

V. KEY AND DESCRIPTIONS OF GENERA (FIGS. 1–40) A. Key Note: Many genera and species exhibit variable morphology without distinct morphological limits. Additional features that distinguish some genera exist only for ultrastructural and molecular characters, which cannot be used in the key. 1a.

Heterocytes2 and/or akinetes never occur in trichomes: order Oscillatoriales……........................……........................……........….…2

1b.

Heterocytes and/or akinetes develop commonly or occasionally in trichomes (if heterocytes are lacking, trichomes are morphologically complex with true branching or with akinetes) ………………………………………………………........................….……….…35

2a.

Trichomes (without sheaths) very narrow and cylindrical; ≤ 3 µm wide (rarely up to 6 µm); cells sometimes with separated centroplasma and chromatoplasma (parietal arrangement of thylakoids) ………………………………………….............................….…….3

2b.

Trichomes broader; width > 3 µm, usually 4–16 µm (rarely to 60 µm); trichomes cylindrical to moniliform; cell content homogeneous or variably structured (thylakoids arranged radially or irregular) ………………………………........................…….…………16

3a.

Trichomes without sheaths or within simple, thin sheaths (when present, always one trichome per sheath), solitary or in mats; trichomes (or filaments) isopolar (both poles with same morphology) with exception (see Figs. 11b and 14): family, Pseudanabaenaceae..................…………………..……............................................................................................................4

3b.

Sheaths wide, containing one or two or more trichomes, at least in a part of a filament; filaments mainly heteropolar: family Schizotrichaceae...............................................................................................................................................................................15

4a.

Trichomes without individual sheaths, but may possess wide or diffuse, mucilaginous envelopes……...............................................5

4b.

Trichomes with distinct, thin, fine or firm sheaths……..……………........................……........................…….......…...……..….……11

5a.

Trichomes straight, wavy, or irregularly coiled …………………………………........................……........................…….....................6

5b.

Trichomes in regular, screwlike coils ………………………….…………………………........................…….......................................10

2

Also termed “heterocyst” (see Komárek and Anagnostidis, 1989).

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6a.

Trichomes mainly short, curved or irregularly coiled, usually only few celled, disintegrating, sometimes enveloped by an indistinct, wide mucilaginous envelope; neighboring cells occasionally disorganized (Fig. 1A) ………………………..........................….Romeria

6b.

Trichomes more or less cylindrical, multicelled, usually without wide mucilaginous envelopes ………………………......……..……..7

7a.

Trichomes solitary or in fine colonies, sometimes constricted at cross walls; cells sometimes with polar aerotopes (groups of gas vesicles) ………...………........................……........................……........................……........................…….........…….......................8

7b.

Trichomes in larger clusters or in mats, cylindrical, usually not constricted at cross walls; cells always without aerotopes (occasionally with scattered or polar granules) ….…….……........................……........................……........................……........................…...9

8a.

Trichomes constricted at cross walls (sometimes distinctly), solitary or in small clusters; cells usually without polar aerotopes (Fig. 1B) …….……........................……........................……........................……........................……...................……Pseudanabaena

8b.

Trichomes unconstricted at cross walls, solitary, cylindrical; cells with polar and/or central aerotopes (Fig. 2A)................Limnothrix

9a.

Trichomes nonmotile, wavy, in clusters or mats (Fig. 2B)……........................……........................…….............……...........Jaaginema

9b.

Trichomes motile, straight or slightly coiled, in thin mats (Fig. 3)……........................……........................….............…..Geitlerinema

10a.

Coils free, long, and wide; trichomes always solitary (Fig. 4A)…………........................……............................................Glaucospira

10b.

Coils usually joined one to another; trichomes in mats or among other algae (Fig. 4B)……........................…………….........Spirulina

11a.

Trichomes (filaments) isopolar (both poles with same morphology), free living (planktonic, metaphytic, or creeping on substratum), in clusters, or forming mats……........................…….................................................……........................……........................….…12

11b.

Trichomes (filaments) heteropolar, with one end attached to the substratum……........................……........…........................…..…14

12a.

Filaments solitary in plankton and metaphyton, more or less short, straight, or irregularly or screwlike coiled (Fig. 5A) ……………………………........................……........................…….......……..........................……........................…...Planktolyngbya

12b.

Filaments creeping on the substratum, in clusters or forming mats……........................……........................……...........……………13

13a.

Filaments solitary or in small groups, epiphytic or creeping on substratum (Fig. 5B) ……...……………………………..……Leibleinia

13b.

Filaments in clusters or in mats, irregularly coiled (Fig. 6A)……........................……........................……..................…Leptolyngbya

14a.

Well-developed trichomes narrowed toward ends (terminal hairs may be present, sometimes separating) (Fig. 7A) ……........................……........................……........................……........................…….................……........................…Homoeothrix

14b.

Filaments and trichomes cylindrical, never tapering toward ends (Fig. 6B) ……………................……................…….Heteroleibleinia

15a.

Wide sheaths conically narrowed and closed at ends, individual trichomes occasionally enveloped by fine sheaths (Fig. 7B) …………………...…………………........................……........................……..........................……........................….……..Schizothrix

15b.

Wide sheaths open at the ends, trichomes without own sheaths, usually tightly joined into fascicles or clusters (Fig. 8A) …………..………………………………........................……........................……........................…….......................…….Trichocoleus

16a.

Trichomes distinctly moniliform (deeply constricted at cross walls), more or less short, without sheaths, isopolar; cells barrel shaped or subspherical: family Borziaceae ……………………………………………........................………………………………..............…17

16b.

Trichomes more or less cylindrical, long, sometimes constricted at the cross walls, but cells not barrel shaped or subspherical …………….………………………........................……........................……........................……........................……....................….18

17a.

Trichomes very short; with up to 8 (very rarely up to 16) cells (Fig. 8B)……........................……........................……...........…Borzia

17b.

Trichomes long (up to over 600 µm); consisting commonly of more than eight cells (Fig. 9A).…………...……………..Komvophoron

18a.

Cell length not less than one half width to ± isodiametric, or slightly longer than wide; protoplasts homogeneous or very finely striated or reticulate (radial thylakoid arrangement); trichomes 4–14 (18) µm wide: family Phormidiaceae …………….……...............19

18b.

Cells very short, always shorter than one half cell width; protoplast slightly granulated (irregular arrangement of short, coiled thylakoids); trichomes 6–35 (60) µm wide: family Oscillatoriaceae …………………………………..…….........................……….…32

19a.

Trichomes typically in screwlike, free coils, always without sheaths; few species with aerotopes (groups of gas vesicles) in cells (Fig. 9B) ...…….……………………........................……........................……........................……..................................….Arthrospira

19b.

Trichomes typically not in screwlike coils, with or without sheaths; individual genera always with or always without aerotopes........….................................................................................................................................................................20

20a.

Trichomes solitary, planktonic, or forming fine mats on submersed substrata, usually pale reddish with finely but distinctly reticulate (keritomized) protoplasts, with typical widened thylakoids, lacking aerotopes (Fig. 11A)……............................…….Tychonema

20b.

Trichomes solitary, in clusters or in mats; if planktonic, then with aerotopes; protoplasts sometimes irregularly striated, but not distinctly reticulate…………........................……........................……........................……........................……...................…..……21

21a.

Filaments (trichomes) always without sheaths, cells with aerotopes, only planktonic species; trichomes solitary or in fascicles........22


123

21b.

Filaments usually with sheaths, cells without aerotopes and not planktonic (exception: few lyngbya species), in various habitats (metaphytic, benthic, subaerial); trichomes (filaments) solitary, clusters or mats……........................……........................…….........23

22a.

Filaments (trichomes) solitary (Fig. 10A)……........................……........................…….......................……….……..…….Planktothrix

22b.

Filaments (trichomes) arranged in more or less parallel, microscopic fascicles (Fig. 10B) …………...............…………Trichodesmium

23a.

Always only one trichome in a sheath ………………………………..…………………........................……....................................…24

23b.

Several to many trichomes (rarely one in early stages) per sheath, sheaths sometimes slightly widened......……………….............…28

24a.

Filaments with obligatory (usually common) false branching……........................……........................……........................………...25

24b.

Filaments without, or only exceptionally with very rare false branching……........................……........................……........…..…..26

25a.

All trichomes cylindrical up to apex, without attenuated ends (Fig. 11B)……........................…………………...…Pseudophormidium

25b.

Trichomes (branches) distinctly attenuated toward the ends (Fig. 18A)......................…………………………………….. Ammatoidea

26a.

Sheaths thick, lamellated, usually colored with sheath pigments (Fig. 13A)……........................…………………….…Porphyrosiphon

26b.

Sheaths firm or thin, not distinctly lamellated or colored……........................……........................……........................…...………..27

27a.

Filaments in older mats joined (in fascicles) forming erect tufts (Fig. 12B)……........................……........................…….......Symploca

27b.

Filaments form flat (usually slimy) mats, not in fascicles arranged in tufts (Fig. 12A)……........................………………..Phormidium

28a.

Sheaths sometimes anastomozing (joining), thin and firm; when joined, two trichomes per sheath in parallel arrangement (Fig. 13B) ………………........................……........................……................................……........................……........................……Lyngbyopsis

28b.

Sheaths not anastomozing, sometimes widened, enclosing occasionally one, usually two to many trichomes, sometimes arranged in dense fascicles; trichomes within sheaths sometimes distant one from another……........................……........................…………....29

29a.

Widened sheaths contain (1) 2–several trichomes, which are usually slightly distant one from another (Fig. 14B)...............Dasygloea

29b.

Trichomes within sheaths are usually tightly arranged in fascicles……........................……........................……...........................…30

30a.

Trichomes within a common sheath and with own individual sheaths; filaments in fascicles, often forming tufts in older mats (Fig. 14A)……........................……........................……........................……........................……................…………..Symplocastrum

30b.

Trichomes within common, widened sheath, but without individual sheaths……........................……........................…….........….31

31a.

Sheaths hyaline, not stratified (Fig. 15A)……........................……........................……........................………........………Microcoleus

31b.

Sheaths finely lengthwise striated (Fig. 15B)……........................……........................…….........................…….……..…Hydrocoleum

32a.

Trichomes in vegetative state always without sheaths (if formed, only under stress; desiccation, hypersaline conditions) (Fig. 16) .………………........................……........................……........................……............................……........................………Oscillatoria

32b.

Trichomes in vegetative state always within distinct sheaths (only hormogonia and reproductive trichome segments can be without sheaths)……........................……........................……........................……........................……...…........................…………………33

33a.

Filaments with typical “scytonematoid” false branching (common paired branches) (Fig. 18B)……........................……...Plectonema

33b.

Filaments without paired false branching……........................……........................……........................………………………………34

34a.

Filaments contain one trichome per sheath; forming mats (rarely solitary filaments in plankton with cells with aerotopes) (Fig. 17) ...……………..……........................……........................……........................…….................................……........................…Lyngbya

34b.

Filaments typically arranged in parallel groups forming fascicles in common sheath; in fascicle-forming colonies or mats (Fig. 19)……........................……........................……........................……........................…….....…........................…….Blennothrix

35a.

Trichomes never branched or only with false branching: order Nostocales……........................……........................…….............…36

35b.

Trichomes (at least a portion) with true branching: order Stigonematales……........................……........................……........………60

36a.

Filaments with lateral false branches……........................……........................……........................……...................………………...37

36b.

Filaments always without any branching: family Nostocaceae……........................……........................……...............................….49

37a.

Filaments isopolar, solitary, entangled in clusters, or forming wooly mats with common lateral branches in pairs (V shape), or rarely with single (Y shape) branch (Fig. 20): family Scytonemataceae .………………………………………......................…………………38

37b.

Filaments typically heteropolar, solitary, forming fascicles or arranged parallel into flat or spherical colonies................……………39

38a.

Trichomes cylindrical up to the ends, apical ends of the trichomes (branches) not attenuated, end cells more or less rounded (Fig. 20A)......................…......................…......................…......................…......................…....................………………..Scytonema

38b.

Trichomes (branches) attenuated toward the ends, sometimes tapering into long, multicellular hairs (Fig. 20B).……Scytonematopsis

39a.

Trichomes cylindrical, not (or slightly) attenuated toward ends, apical cells rounded; heterocytes basal and intercalary: family Microchaetaceae......................…......................…......................…......................……………….......................…......................…..40

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39b.

Trichomes distinctly attenuated toward ends, may be elongated into long, thin, multicellular hairs; heterocytes mostly basal (rarely intercalary before false branching): family Rivulariaceae......................…………………………………………………….………........45

40a.

Filaments long, lateral branches common and typically solitary, rarely (in older filaments) in twos (“scytonematoid”); heteropolar growth sometimes indistinct in old trichomes with numerous intercalary heterocytes......................………………..…………..........41

40b.

Filaments short, with or without solitary lateral branches; heterocytes basal or occasionally intercalary.......................…………….44

41a.

Branches diverge distinctly from the main filament......................….............................................…………………………….………42

41b.

Branches remain partly inside main sheath, forming fascicles with several trichomes arranged in parallel; branches diverge only at filament ends (Fig. 23)......................…......................…......................…......................…......................…...........……Coleodesmium

42a.

Cells more or less uniform along length of trichome; sheaths thin or slightly widened and lamellated, branching common........…..43

42b.

Cells of variable length (shorter, longer than wide, or isodiametric), sometimes cylindrical, but barrel shaped near ends; sheaths very wide, funnel-like lamellated at ends; branching rare (Fig. 21B)......................…......................…......................……....…..Petalonema

43a.

Cells mostly shorter than wide, false branches mainly at one side of filament; trichomes usually distinctly constricted at cross walls (Fig. 21A)......................…......................…......................…......................…......................…...........................……………Hassallia

43b.

Cells isodiametric to longer than wide, false branching in several directions; trichomes cylindrical, constricted or unconstricted at cross walls (Fig. 22)......................…......................…......................…......................…......................…...........................Tolypothrix

44a.

Simple trichomes cylindrical or slightly attenuated toward ends (Fig. 24B)......................…......................………………..Microchaete

44b.

Simple trichomes widened distinctly toward the ends (Fig. 24A)......................…......................…......................…........………Fortiea

45a.

Heteropolar filaments simple; occur singly or in groups, but always distinctly separated one from another (Fig. 25A) ......................…......................…......................…......................……....................................................................................Calothrix

45b.

Heteropolar filaments distinctly branched (false branches); usually unified into mucilaginous colonies with parallel or radial arrangement......................…......................…......................…......................…..........…......................………………………………46

46a.

Filaments distinctly branched, in fascicles or clusters......................…......................…......................…......................………..……47

46b.

Filaments unified more or less in parallel, forming flat or spherical mucilaginous colonies......................…......................…...…….48

47a.

Sheaths distinctly broader than trichome width, often lamellated and usually closed at the ends (Fig. 26B).................…...Sacconema

47b.

Sheaths not distinctly wider than the trichome, firm or gelatinous, lamellated and funnel-like, frayed at the ends (Fig. 25B) ……..……......................…......................…......................…......................….............................……………………………Dichothrix

48a.

Colonies flattened or (young) hemispherical, sometimes large (millimeters to several square centimeters) with firm, individual sheaths; filaments arranged in parallel; akinetes absent (Fig. 27)......................…......................…………..…………………..Rivularia

48b.

Colonies spherical with radially arranged filaments; sheaths confluent, gelatinizing; basal akinetes often form near end of growth period (Fig. 26A)......................…......................…......................………………….………………………………………….Gloeotrichia

49a.

Heterocytes absent; akinetes may develop at the end of growth season......................…......................…......................…………….50

49b.

Heterocytes present (absent rarely in some species with high nitrogen supply)......................….....................................…………….51

50a.

Akinetes (when present) solitary, distinctly larger than vegetative cells; cells more or less cylindrical; planktonic (Fig. 31A) …………………......................…......................…......................…...................…......................…......................……….Raphidiopsis

50b.

Akinetes develop serially in rows, only slightly larger than the vegetative cells; cells mainly barrel shaped; planktonic, metaphytic, or in mats (Fig. 32B)......................…......................…......................…......................…......................…...............................….Isocystis

51a.

Heterocytes terminal, single pored; if intercalary, occur in pairs with trichomes fragmenting between them......................…….......52

51b.

Heterocytes mainly intercalary (terminal are exceptional), typically with pores on either side......................……………….........…..54

52a.

Heterocytes develop primarily intercalary in pairs; trichomes soon disintegrate between them, heterocytes then appear terminal; mainly planktonic species (Fig. 29A)......................…......................……………………………………………….......……Anabaenopsis

52b.

Heterocytes develop primarily from terminal cells; planktonic and/or forming mats......................…………………..........………….53

53a.

Trichomes forming benthic colonies, mats, epiphyton or metaphyton; akinetes always adjacent to terminal heterocytes; trichomes typically cylindrical (Fig. 30B) .………………………………………………......................…………...………………..Cylindrospermum

53b.

Trichomes planktonic and solitary; akinetes adjacent or slightly distant from heterocytes; trichomes slightly attenuated toward both ends or cylindrical (Fig. 30A) ……………………………………………………......................………….……...……Cylindrospermopsis

54a.

Trichomes somewhat asymmetrical in structure, with elongated and sometimes narrowed one to few terminal cells; akinetes elongated, cylindrical to long oval, rarely spherical; three species (from about 20 described) may form fascicles or clumps on the water surface (Fig. 29B)......................…......................…......................…........................……………………………………Aphanizomenon


125

54b.

Trichomes with more or less regularly spaced heterocytes along trichome (metameric); apical cells undifferentiated from other vegetative cells; akinetes of various shapes ……………………………......................…......................……………………....………..55

55a.

Akinetes adjacent to heterocytes or distant, solitary or up to five in a row, often formed after fusion of several vegetative cells; mature akinetes several times larger than vegetative cells ......................…......................…......................…………..………………56

55b.

Akinetes positioned more or less between heterocytes, in long rows; mature akinete size only slightly larger than vegetative cells ……..……………………......................…......................…..................................…......................…......................………………….57

56a.

Solitary planktonic trichomes, often with aerotopes (groups of gas vesicles), coiled or straight, or growing metaphytic and periphytic forming mats on substrata (Fig. 28)......................…......................……………….………………………………………..…Anabaena

56b.

Trichomes joined in macroscopic, benthic colonies; more or less spherical, saccate thallus with defined, firm surface (Fig. 31B) ……………………......................…......................…......................…......................….............................…......................…….Wollea

57a.

Cells (and akinetes) always shorter than wide; planktonic species in solitary trichomes (with aerotopes) or benthic species forming mats (Fig. 33)......................…......................…......................…......................…......................….............………….………Nodularia

57b.

Cells and akinetes always longer than wide......................…......................…......................…......................………………………..58

58a.

Trichomes in thin, firm sheaths (Fig. 31C)......................…......................…......................…........................…………………Aulosira

58b.

Trichomes in colonies without firm sheaths, but sometimes within wide, mucilaginous envelopes in marginal parts of colonies…..59

59a.

Trichomes more or less cylindrical, with constrictions at the cell walls, unified in fine, gelatinous, amorphous mats with diffuse surface (Fig. 32A)....................................…......................…......................…..................…......................….………..……Trichormus

59b.

Trichomes mainly moniliform, unified into firm, slimy colonies (often macroscopic) with a distinct, defined margin (Fig. 34) …………………………..............................…......................…......................…......................…......................………………...Nostoc

60a.

Portions of filament or thallus multiseriate; composed of cell agglomerations or filaments dividing lengthwise and laterally; uniseriate portions moniliform......................…......................…......................…......................…......................…........................……….61

60b.

Filaments only uniseriate; ± cylindrical, with distinct true branching......................…......................…......................…...………….64

61a.

Thallus formed from basal filaments (or cell agglomerations) and erect branches that are densely arranged and parallel (Fig. 35B) ……….…………......................…......................…......................…......................…......................….................……Stauromatonema

61b.

Thallus formed from filaments with distinct lateral branches (sometimes only short) not arranged in dense parallel series; basal or main trichomes multiseriate or uniseriate; at least some cells dividing laterally......................…......................…….………………...62

62a.

Basal filaments usually multiseriate, with rare true branching; long, uniseriate branches with false branching (like Tolypothrix): family Borzinemataceae (Fig. 37B)......................…......................…………………………………………………….…...Schmidleinema

62b.

Filaments only with (usually frequent) true branching......................…......................…......................…......................…………….63

63a.

Basal filaments mainly multiseriate, true branches (mainly uniseriate) differ morphologically (Fig. 37A).........................…Fischerella

63b.

All trichomes (basal and erect) more or less of the same usually multiseriate character, often regularly narrowing toward ends; basal system multiseriate; some forming macroscopic masses (Fig. 36) ......................…......................……………………........…Stigonema

64a.

Thallus shrublike, with distinct heteropolar growth from base to apex; filaments successively branched into typically pseudodichotomous branches......................…......................…...................…......................…......................…......................…........…...65

64b.

Thallus composed of creeping, uniseriate filaments; filaments branched laterally: family Mastigocladaceae ........................……….68

65a.

Heterocytes develop very occasionally intercalary, or (in several genera) absent: family Loriellaceae......................………….……...66

65b.

Heterocytes well developed, mainly lateral, at ends of short lateral branches or intercalary: family Nostochopsaceae (Fig. 39A)….......................…........…......................…......................…................................…Nostochopsis

66a.

Sheaths gelatinous, intensely encrusted with aragonite crystals, usually open at ends; inhabits caves (Fig. 38A).................…Geitleria

66b.

Sheaths firm and smooth on the surface, without encrustations, closed at the ends; inhabits thermal waters.....................…………67

67a.

Sheaths wide, with distinct, funnel-like lamellation (Fig. 38C)......................…......................….........................………..Colteronema

67b.

Sheaths thin, simple, fine, distinct, but not lamellated (Fig. 38B)......................…......................…........................………....Albrightia

68a.

Sheaths, particularly in older filaments, thick and confluent, with trichomes in parallel arrangement (Fig. 40B)....………Thalpophila

68b.

Sheaths simple, thin, fine or firm but never thick; forming confluent mass; trichomes not in parallel arrangement..................…….69

69a.

Cells in main trichomes and branches isomorphic, more or less cylindrical or barrel shaped (Fig. 39B)...................…...Hapalosiphon

69b.

Cells of different shapes in same thallus (e.g., short or elongated barrel shaped or cylindrical) (Fig. 40A).........………Mastigocladus

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B. Descriptions of Genera3 Oscillatoriales: Pseudanabaenaceae: Pseudanabaenoideae Geitlerinema (Anagnostidis et Komárek) Anagnostidis (Fig. 3) The thallus is thin, delicate, mostly bright blue– green, rarely violet or brown, diffuse, sometimes fascicle-like, usually forming thin macroscopic mats; occasionally isolated trichomes occur. Trichomes are (0.6) 1–4 (6.5) µm wide, usually in parallel arrangements, cylindrical, straight, sometimes slightly flexuous or (rarely) irregularly screwlike coiled (especially at the ends), without sheaths, mostly not constricted at the cross walls, rarely (more or less slightly) constricted, more or less gradually attenuated and bent or coiled at ends, rarely not attenuated and straight. Trichomes are motile, with intense gliding in the direction of the longitudinal axis, may be accompanied by distinctive clockwise or counterclockwise rotation, waving (oscillation) and circling. Cells are usually longer than wide, before division occasionally several times longer than wide; the cell content usually contains dispersed large cyanophycin granules or localized (apical) carotenoid bodies, without aerotopes; thylakoids are concentrically arranged, peripheral and parallel to longitudinal cell walls. Apical cells are conical, hooked, or bent, but mostly acuminate or rounded, occasionally sphericalcapitate, or straight, cylindrical, and rounded. Reproduction occurs through disintegration of trichomes into motile hormogonia, without necridic cells. Species of Geitlerinema (over 30 have been described) grow mostly in mats, on soils, or in various aquatic habitats on macrophytes or other substrata (mainly in unpolluted waters); they rarely occur as solitary filaments. Geitlerinema splendidum is the most common species in the periphyton of oligotrophic to mesotrophic waters in the north-temperate region; it is common in soft-water (granitic) lakes in eastern and western Canada (Bourrelly, 1966; Stein and Borden, 1979, as Oscillatoria splendida). G. amphibium (= Oscillatoria amphibia) is another widely distributed species (Prescott, 1951; Whitford and Schumacher, 1969; Stein and Borden, 1979). Other species are known from mineral waters and thermal springs: G. jasorvense and G. acus occur in Yellowstone National Park (Copeland, 1936); G. claricentrosum and G. earlei are reported from Puerto Rico (Gardner, 1927). The 3

Drawings in figures are based on North American literature; however, documentation of several species was selected from non–North American papers. Localities are cited only in figures from North America. Figures of trichome structures and reproductive processes were selected according to various authors.

metaphytic species are likely to occur elsewhere in North America. Jaaginema Anagnostidis et Komárek (Fig. 2B) Trichomes are usually flexuous, solitary or entangled in clusters, or forming thin, membranaceous thalli without sheaths (exceptionally with very fine mucilaginous layers around the trichomes); they are always nonmotile. Trichomes are up to 3 µm wide; they are usually not constricted at the cross walls, sometimes slightly attenuated at the ends, and not capitate. Cells are cylindrical, mostly longer (up to 10⫻) than wide, rarely almost isodiametric; the cell content is homogeneous, without aerotopes; thylakoids are probably parietal. Apical cells are mostly rounded, occasionally conical, without calyptra, rarely with a thickened outer cell wall. Reproduction occurs by fragmentation of trichomes, without necridic cells. Jaaginema is a little-known genus with more than 20 species, mainly benthic, in shallow water on sediments or aquatic plants in various types of water or in metaphyton. Several species are known from thermal or mineral springs in North America. Jaaginema filiforme (= Oscillatoria filiforme) was described from rivers in Yellowstone National Park by Copeland (1936). Limnothrix Meffert (Fig. 2A) Trichomes are solitary or in small, irregular fascicles or clusters, isopolar (both poles with the same morphology), usually free living, straight, slightly bent or flexuous, consisting of numerous cylindrical, mainly elongated cells. Trichomes are not constricted or very slightly constricted at indistinct, thin cross walls, 1–6 µm wide, cylindrical, not attenuated at ends, without or facultatively with fine, hyaline sheaths, without false branching, nonmotile or with reduced motility (slight trembling or gliding), often disintegrating. Cells are isomorphic, cylindrical, sometimes long, rarely slightly inflated, more or less isodiametric or frequently longer than wide, pale blue–green, blue–gray, yellowish, reddish, or pink; all cells are capable of dividing, with localized apical or central aerotopes, which may be lacking (depending on environmental conditions); thylakoids are mostly parietal; variable ratios of phycocyanin and phycoerythrin (recognizable mainly in cultures; Kohl and Nicklisch, 1981; Komárek, 1994). Apical cells are cylindrical, rounded or roundly flattened at ends, rarely conically attenuated and/or with pointed conical plasmatic protrusions, usually with one or few terminal aerotopes, without calyptra, not capitate. Cell division is perpendicular to the longitudinal axis, sometimes (rarely) asymmetrical. Daughter cells reach the original size before the next division. Repro-


FIGURE 1 (A) Romeria: (a) R. leopoliensis (original, population from Europe); (b) R. nivicola (after Kol from Smith, 1950, from snow in Yellowstone National Park); (c) R. leopoliensis, crosswise and lengthwise sections of cells with characteristic position of thylakoids. (B) Pseudanabaena: (a) P. catenata (after Lauterborn from Geitler, 1932); (b) P. limnetica (bar = 10 µm; after Komárek, 1958); (c) P. lonchoides (after Anagnostidis, 1961, possible occurrence in thermal springs); (d) P. galeata (after Anagnostidis, 1961); (e) Pseudanabaena sp., lengthwise section (bar = 1 µm; after Guglielmi and Cohen-Bazire, 1984).

127

128


FIGURE 2 (A) Limnothrix: (a) L. redekei (after Van Goor and Skuja from Anagnostidis and Komárek, 1988); (b) L. redekei (after Meffert, 1988); (c) Limnothrix sp. (original by Komárková, from Florida); (d) L. redekei, lengthwise section (after Kalina from Anagnostidis and Komárek, 1988). (B) Jaaginema: (a) J. neglecta (bar = 10 µm; after Komárek, 1975); (b) J. subtilissima (after Böcher from Starmach, 1966); (c) Jaaginema sp. (after Copeland, 1936, from Yellowstone National Park, sub Phormidium geysericola).


duction occurs by disintegration of the trichomes into nonmotile hormogonia, without necridic cells. Species of this genus occur in the plankton of mesotrophic to eutrophic lakes and reservoirs, sometimes developing metalimnetic maxima, mostly in temperate and northern areas (Whitton and Peat, 1969; Gibson, 1975; Meffert and Krambeck, 1977; Kohl and Nicklisch, 1981; Meffert, 1987, 1988; Kling, personal observation). There are about 20 revised species, but all Pseudanabaena species with polar aerotopes possibly belong to the genus Limnothrix. The most common species (L. redekei) is distributed throughout the temperate zone, where it occurs mainly in colder seasons. L. redekei was also one of the first species to respond to the artificial eutrophication of Lake 227 in the Experimental Lake Area in northwestern Ontario, Canada (recorded as Oscillatoria redekei by Kling and Holmgren, 1972). In large mesotrophic lakes, L. redekei may occur during spring and fall during turnover periods (Kling, personal observation). L. vacuolifera occurs in northern lakes in Scandinavia and Canada; the benthic L. guttulata was recorded from the United States (authors’ records). Pseudanabaena Lauterborn (Fig. 1B) Trichomes are solitary or in fine mats, straight or curved, less frequently wavy, cylindrical; they are usually short, consisting of a very few to several cells, or long with many cells, usually with conspicuous constrictions at the cross walls, 1–3.5 µm wide, rarely unconstricted. Trichomes lack firm sheaths, but sometimes have wide, fine, diffuse mucilage. Apical cells are not differentiated, without calyptra or thickened outer cell wall. Motility is lacking or facultative, usually slow gliding, occurring also in separated unicells, probably without rotation. Cells are usually cylindrical with rounded ends, but sometimes almost barrel shaped, always longer than wide, rarely almost isodiametric (after division), with or without polar aerotopes (groups of gas vesicles—types with gas vesicles probably belong to the genus Limnothrix); thylakoids are concentric and parietal, parallel to the long axis; with one central perforation in the cross walls and/or multiple pores (300–500) near the cell poles. Cell division is exclusively by binary fission in one plane, perpendicularly to the long axis, sometimes asymmetrical. Reproduction occurs through production of one-celled to multicelled hormogonia or trichome fragmentation, without necridic cells. Over 30 species have been described, several occurring in planktonic, metaphytic, periphytic, or benthic habitats in waters of different trophic status; a few occur on soil or within the mucilage of other algae or colonial rotifers (endogloeic). A few species are known from mineral waters and hot springs. Most reports of

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planktonic Pseudanabaena species are recorded as Oscillatoria, from which several species are also known from oligotrophic to eutrophic waters in North America (O. limnetica = P. limnetica, P. mucicola, P. catenata; authors’ results). P. thermalis occurs in alkaline, thermal waters (Oscillatoria amphigranulata sensu Castenholz, 1976). Romeria Koczwara in Geitler (Fig. 1A) The thallus is microscopic, very fine, and filamentous or pseudofilamentous. Trichomes are solitary, usually short, fine, irregular and fragile, up to 3.5 µm wide, 1–8 (18–32) celled, rarely with more cells, usually semicircular curved, flexuous or irregularly screwlike coiled, with one or two or more (up to eight) helices, constricted at the cross walls, without a distinctive sheath, but with fine, more or less thick mucilaginous envelopes. The ends of neighboring cells are sometimes slightly shifted one from another. Usually one, rarely few irregularly localized nonmotile trichomes are present in a fine mucilaginous envelope. The mucilage is colorless, diffuse, and indistinct. All cells are of the same morphology, elongated, long cylindrical or barrel shaped; terminal cells are rounded at the apex. Thylakoids are parietally arranged. Cell division is transverse, symmetric, or slightly asymmetric. Reproduction occurs by trichome fragmentation into hormogonia or solitary cells. Romeria species are mainly planktonic, living in clear oligotrophic to mesotrophic lakes and ponds, rarely in hypertrophic systems. Nineteen species have been described, most from the northern temperate zone; one species is marine (described from the Gulf of Mexico). From North America, Smith (1950) mentioned only the cryosestic R. elegans var. nivicola (= R. nivicola) from Yellowstone National Park, but several other U.S. species are known from the temperate zone (R. alascense from Alaska, R. elegans, R. leopoliensis, and marine R. mexicana) and from tropical regions (R. heterocellularis and R. hieroglyphica; Komárek, 2001). Several species are probably more widely distributed.

Oscillatoriales: Pseudanabaenaceae: Spirulinoideae Glaucospira Lagerheim (Fig. 4A) Filaments (trichomes) are solitary, short or slightly elongated, thin (up to 3 µm wide), without sheaths, regularly loosely screwlike coiled with wide and more or less long spirals, usually intensely motile (rotation), sometimes slightly flexible, cylindrical, not attenuated at ends, and not constricted at slightly visible cross walls. Cells are pale blue–green or yellowish, with homogeneous content, sometimes with few fine granules, probably always longer than wide. Cell division is per-

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FIGURE 3 Geitlerinema: (a) G. unigranulatum, lengthwise section with polar cyanophycin granules and parietally arranged thylakoids (after Komárek and Azevedo, 2000); (b) G. splendidum (bar = 10 µm; after Komárek, 1975); (c) G. lemmermannii (after Tavera et al., 1994, from Mexico); (d) G. earlei (after Gardner, 1927, from Puerto Rico; sub Oscillatoria earlei); (e) G. claricentrosa (after Gardner, 1927, from Puerto Rico, sub Oscillatoria claricentrosa).


FIGURE 4 (A) Glaucospira: (a) G. laxissima (after G. S. West, 1907); (b) Glaucospira sp. (after Copeland, 1936, from Yellowstone National Park; sub Spirulina caldaria var. magnifica); (c) Glaucospira sp. (after Drouet, 1937, from Massachusetts; sub Spirulina stagnicola). (B) Spirulina: (a) S. weissii (after Drouet, 1942, from the United States); (b) S. major (after Geitler, 1932); (c) Spirulina sp. (from culture collection, the United States, original photo by J. D. Wehr, with permission); (d) S. meneghiniana (after Komárek, 1989, from Cuba); (e) S. subsalsa (original by Komárková, from the Everglades, Florida).

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pendicular to the trichome axis. Reproduction probably occurs through trichome disintegration. Glaucospira is a very poorly known and problematic genus; the cyanobacterial character of some of the described (more than five) species must be confirmed. The known species are mainly planktonic or metaphytic. Copeland (1936) described G. yellowstonensis, a slightly thermophilic species, from the surface of gelatinous bottom sediments in Yellowstone Lake. Spirulina laxa (which corresponds to the genus Glaucospira) was described from ponds in the Piedmont and central plains regions of North Carolina (Whitford and Schumacher, 1969) and from lakes in Ontario and British Columbia (Duthie and Socha, 1976; Stein and Borden, 1979). Spirulina stagnicola, described by Drouet (1937) from brackish waters in Massachusetts, also belongs to this genus. Spirulina Turpin ex Gomont (Fig. 4B ) Trichomes are cylindrical, screwlike coiled, 0.3– 7.5 µm wide, of variable length, solitary or forming fine, mucilaginous mats. Their color ranges from blue– green, olive green, gray–green, brownish to reddish or violet. Regular coiling occurs in all species, but ranges from loosely to tightly screwlike or helical; coils may be compressed or may have spaces between them. Rarely irregularities in coiling, variable curves or occasionally circle-like coils and straight portions, can occur. Spirulina is intensely motile; trichomes glide with rapid clockwise or counterclockwise rotation. Sheaths or mucilaginous envelopes are usually lacking; a fine slime is produced in some mats. Cells are not constricted at scarcely visible cross walls (not seen without staining in light microscope); end cells are usually not attenuated. Cells are typically isodiametric or longer than wide; the cell content is homogeneous, without aerotopes; a special pore and a perforation pattern is apparent in the cell walls (i.e., several rows on the concave side); thylakoids are arranged parallel to the longitudinal cell axis. Apical cells are rounded, hemispherical, without calyptra or thickened outer cell wall. Reproduction occurs via fragmentation into motile hormogonia, without necridic cells. About 50 species have been described (about 20 revised; Anagnostidis and Golubic´, 1966; Anagnostidis and Komárek, 2003). A widely conceived genus (Geitler, 1932) has been divided according to the morphology of filaments, ultrastructure and molecular sequencing into Spirulina and Arthrospira (Phormidiaceae); the existence of the separated genus Arthrospira has been confirmed by both morphological and genetic analyses (Tomaselli et al., 1996; Mühling et al., 1997). Several typical Spirulina species sensu stricto are benthic and occur in metaphyton, sometimes in heavily polluted

habitats or among detritus. Other species grow in thermal and mineral springs or in saline lakes and ponds. Tilden (1910) recorded nine species of Spirulina from the United States (with Arthrospira). Smith (1920, 1950) noted three species, S. maior, S. subtilissima and S. labyrinthiformis; Whitford and Schumacher (1969) reported these plus S. weissii. North American records also include S. gigantea, S. nordstedtii, S. major, and S. princeps from northern and temperate lakes (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982).

Oscillatoriales: Pseudanabaenaceae: Leptolyngbyoideae Leibleinia (Gomont) L. Hoffmann (Fig. 5B) Filaments are solitary, waved or curved, 1.5–11 µm wide, with unique epiphytic habitat being attached to substrata along their length, or by a portion of it, later having both free ends, usually epiphytic, with a sheath, and very rarely with false branching. Sheaths are firm, thin, and colorless; trichomes are nonmotile. Cells are cylindrical, mainly pale blue–green or gray–blue, without aerotopes; they are usually longer than they are wide, with peripherally arranged thylakoids (?). Cells divide perpendicularly to the longitudinal axis; daughter cells approach the original size before subsequent division. Reproduction occurs via nonmotile hormogonia and hormocytes, separating without necridic cells, adhering to substrata lengthwise along their horizontal axis and growing upward at both poles. The majority of the 14 species described grow epiphytically on filamentous algae and aquatic macrophytes. Both freshwater and marine species are known. The filament morphology is very simple and may often be mistaken for Heteroleibleinia or Leptolyngbya. Many more species of Leibleinia probably exist, but the genus is poorly known from North American habitats with many species likely identified as species of Lyngbya, Phormidium, or Oscillatoria, such as L. calotrichicola from Yellowstone National Park (Copeland, 1936) and marine L. aeruginea from Puerto Rico (Gardner, 1927). Leptolyngbya Anagnostidis et Komárek (Fig. 6A) Filaments are rarely solitary, usually loosely arranged in flakelike clusters or mats, free floating or attached to substrata, seldom in fascicles or forming compact colonies (thallus); they are more or less flexuous, finely undulating, occasionally nearly straight and long. Ends are usually neither attenuated nor capitate. Facultative, firm, thin, hyaline sheaths are present; branching is rare but may occur with occasional pseudobranches. Sheath frequency is species specific, possibly dependent on environmental factors.


FIGURE 5 (A) Planktolyngbya: (a) P. limnetica (after Kondrateva, 1968; Hindák and Moustaka, 1988); (b) P. contorta (after Smith, 1950; Kondrateva, 1968); (c) P. tallingii (after Komárek and Kling, 1991). (B) Leibleinia: (a) Leibleinia sp. (original photo by Komárková, from the Everglades, Florida); (b) L. calotrichicola (after Copeland, 1936, from Yellowstone National Park).

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FIGURE 6 (A) Leptolyngbya: (a) Leptolyngbya sp. (typically coiled filaments in cluster; photos original by Komárková, from the Everglades, Florida); (b) L. foveolarum (after Komárek, 1988); (c) L. nostocorum (after Komárek, 1988); (d) L. cartilaginea (after Copeland, 1936, from Yellowstone National Park). (B) Heteroleibleinia: (a) H. kuetzingii (after Fott and Komárek, 1960); (b) H. minor (after Gardner, 1927, from Puerto Rico); (c) H. pusilla (bar = 10 µm; after Whelden, 1947, from Canadian Arctic); (d) H. profunda (reddish trichomes; after Tavera and Komárek, 1996, from volcanic lakes in Mexico).


Trichomes are 0.5–3.5 µm wide, motile (producing hormogonia) or nonmotile, or with slightly noticeable trembling. Cells are cylindrical, usually longer than wide, less frequently isodiametric or shorter than wide, usually with homogeneous contents, but often with recognizable chromatoplasma and centroplasma (= position of peripherally arranged thylakoids), without gas vesicles. Reproduction occurs via trichome fragmentation, with or without sacrificial cells (two subgenera); hormogonia range from nonmotile to barely motile, with trichome disintegration occurring from the apical portions of the trichomes. The genus Leptolyngbya is one of the most common (and taxonomically most difficult) cyanobacterial genera, containing numerous morphotypes and ecotypes (species), which are very common in soils and in periphyton and metaphyton in a variety of freshwater and saline (marine) environments (Anagnostidis and Komárek, 1988; Albertano and Kovácˇik, 1994). Several species are known from thermal and mineral waters or grow subaerially on wet rocks. Identification of species (more than 140 have been described) is difficult due to indistinct morphological differences. North American species have not been well documented (usually under the names Lyngbya or Phormidium). Whitford and Schumacher (1969) reported L. lagerheimii, L. subtilis, L. angustissima, and L. tenuis under the generic name of Phormidium. L. bijahensis, L. cartilaginea, L. geysericola, L. rubra, L. subterranea, L. vesiculosa and L. yellowstonensis were recorded from mineral and thermal waters in Yellowstone National Park; Schizothrix thermophila also probably belongs in this genus (Copeland, 1936). A survey and revision of North American species in this genus is necessary. Planktolyngbya Anagnostidis et Komárek (Fig. 5A) Filaments are free living, solitary, free floating, straight, flexuous, wavy, or more or less spirally screwlike or irregularly coiled, with firm, thin, colorless sheaths, and very rarely false branched. Trichomes are nonmotile, cylindrical, isopolar (both poles with the same morphology), uniseriate, unconstricted or slightly constricted at the cross walls, not attenuated toward ends, with rounded apical cells (not capitate). Cells are cylindrical, up to 3 (5) µm wide, usually longer than wide, rarely more or less isodiametric, with peripherally arranged thylakoids and without aerotopes or with facultative polar solitary aerotopes and rounded end cells. All cells are capable of division. Reproduction occurs by fragmentation, without necridic cells (short hormogonia). More than 15 species have been described from the plankton of large mesotrophic reservoirs and lakes, mainly from northern temperate zones (e.g., P. limnetica

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and P. contorta; Hindák, 1985; Cronberg and Komárek, 1994), but several species are specific to tropical and subtropical lakes (P. tallingii and P. regularis; Komárek and Kling, 1991; Komárková-Legnerová and Tavera, 1996; Komárek and Cronberg, 2001). Some Planktolyngbya species occur in North American reservoirs, but were previously recorded as the genus Lyngbya. Smith (1950) reported “Lyngbya contorta” from lakes in the United States. Planktolyngbya capillaris, P. contorta, P. bipunctata, and P. limnetica were reported in North America (authors’ unpublished data). A review of North American species in this genus is necessary.

Oscillatoriales: Psedudanabaenaceae: Heteroleibleinioideae Heteroleibleinia (Geitler) L. Hoffmann (Fig. 6B) Filaments are solitary or in groups, heteropolar, individually attached by one end to substratum, rarely forming membranaceous, tuftlike layers with parallel sessile filaments; filaments are typically short, usually up to 100 µm long, rarely longer. Sheaths are thin, firm, and colorless. Trichomes may be constricted or not constricted at the cross-walls. Cells are usually isodiametric or slightly shorter or longer than wide. Apical cells are rounded, without calyptra or thickened outer cell wall. Reproduction occurs by disintegration of trichomes into motile hormogonia and nonmotile hormocytes, separating particularly from the apical parts of the trichomes; necridia present (?). Heteroleibleinia does not differ substantially from thin forms of Leibleinia species except that the filaments are heteropolar, attached by one end to the substratum (main intergeneric feature). Almost all of the 30 species described are known from aquatic habitats worldwide, growing attached to different substrata in both marine and freshwater environments. Identification of species is difficult due to the small number of morphological characters. The genus is present in North American waters, but mainly mentioned under the previous generic names (usually Lyngbya); all species need to be accurately documented and their revisions are necessary, based on the current taxonomic criteria (Anagnostidis and Komárek, 1988). H. pusilla and H. versicolor are recorded (under Lyngbya) from several North American localities (Tilden, 1910; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). Homoeothrix (Thuret) Kirchner (Fig. 7A) Filaments are solitary or forming mats, simple, not branched, or very rarely laterally branched (false branching), erect, solitary or in small, loose fascicles, attached to substrata basally, sometimes radially oriented

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FIGURE 7 (A) Homoeothrix: (a) H. janthina (after Starmach, 1966); (b) H. crustacea (after Komárek and Kann, 1973); (c) H. varians (after Komárek and Kalina, 1965). (B) Schizothrix: (a) S. violacea (after Drouet, 1937, from the United States); (b) S. constricta (after Copeland, 1936, from Yellowstone National Park).


with bases in the center of the colony. Sheaths are thin, firm, hyaline or rarely slightly widened and lamellated, and yellowish in color. Trichomes are thin, straight or coiled, 3 (5, exceptionally to 7) µm wide, cylindrical, constricted or not constricted at the cross walls, tapering at ends, and sometimes elongated into thin, hyaline hairs (well-developed trichomes). Vegetative cells are short and cylindrical, bases of filaments of some species are enlarged (with wider and shorter cells); pale blue–green, olive green, or grayish. Reproduction occurs via hormogonia liberated from the upper part of trichomes after separation of the terminal hair. All species (about 25 are valid) are known from submersed aquatic habitats, especially from stones or macrophytes in streams, rivers, and lakes; a few species inhabit the mucilage of other algae (endogloeic). Several taxa are reported from North America, for example, H. varians and H. janthina from eastern and western mountainous locations (Prescott, 1951; Stein and Borden, 1979; authors’ results). Whitford and Schumacher (1969) recorded H. janthina (as Amphithrix) in rapids in the Piedmont region of North Carolina. However, H. janthina never forms encrustations, as it is sometimes reported, and also the marine habitats of this species (Tilden, 1910; Smith, 1950) are unlikely and may represent other species, e.g., (H. crustacea (?) and H. violacea (?). Whitford and Schumacher (1969) reported two other species (H. stagnalis and H. crustacea) from the pebbles of North Carolina streams (as Leptochaete).

Oscillatoriales: Schizotrichaceae Schizothrix Kützing ex Gomont (Fig. 7B) Filaments are solitary, free living in fascicles or attached and densely entangled forming a thallus; they are initially microscopic, but later form erect and polarized fascicles, with usually rich pseudobranched filaments attached by one end to stony substrata, or large, soft, thin or thick, fine or membranaceous layers. Two subgenera are recognized: In subgenus Schizothrix, fascicles are not encrusted and may be free floating or in mucilage of other algae; in subgenus Inactis, the thallus becomes firm, hard, sometimes spongy, later encrusted with calcium carbonate or calcified in basal parts. Encrusted colonies are usually lamellated, forming crustlike, hemispherical cushions or flat layers on stony substrata, 5 mm or more thick; the outside is warty, variously colored, gray, gray–brown, olive green, black–green, blue–green, or rusty reddish. Filaments of both subgenera are usually long, nearly straight or curved, sometimes densely aggregated, usually erect, with filaments parallel or occasionally radially arranged, sometimes forming ropelike tangles; filaments are rarely unbranched or sparsely, in tufts sometimes nearly pseu-

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dodichotomously pseudobranched, particularly at the ends. Sheaths are usually thickened and wide, rarely thin, soft to firm, often lamellated, delimited at the margin, rarely slightly diffused; they range from colorless to yellowish, gold–yellow, brownish, olive green or brownish, red or (rarely) violet or blue. The sheath surface ranges from smooth to uneven, rarely fibrous, ends attenuated or pointed, at free ends usually conically closed, rarely attenuated or funnel-like (facultative). Sheaths contain rarely one, usually several to numerous thin trichomes (facultatively changing in one species), often with more trichomes at the base, but with a single trichome in the upper part. Trichomes are nonmotile, distinctly, slightly or not constricted at the cross walls, and usually each with its own sheath. Cells are usually longer than they are wide, rarely almost isodiametric, with thylakoids probably only parietal. Apical cells are rounded, conically rounded, obtuse, or acutely conical, without a thickened outer cell wall or calyptra. Reproduction occurs via fragmentation into motile (?) hormogonia, without necridic cells. More than 70 species are currently recognized. Many grow in the littoral zone of lakes or in streams, attached to substrata in wave-swept areas or in metaphyton among plants in wetlands and swamps. Several species form encrusted hemispherical colonies or layers on stones or rocks; other species colonize saline environments. There are a few subaerial species, some of which create characteristic brownish fascicles on wood or soil. Several species are recorded from Canada, the United States, the West Indies, Mexico, and Alaska, but few were recently taxonomically reclassified into other genera (e.g., Symplocastrum). Smith (1950) noted 30 species in the United States; several were also reported from British Columbia (Stein and Borden, 1979). Whitford and Schumacher (1969) reported 13 species from North Carolina, several of them growing on moist soil or rocks; Schizothrix aikenensis was collected from the epipelon of a small pool, whereas several others were found on rocks or bottom substrata in streams. Whelden (1947) and Sheath and Steinman (1982) reported several species from arctic and subarctic freshwater habitats. Tilden (1910) listed seven species in the United States, the West Indies, and Mexico. Copeland (1936) recorded several encrusting species from Yellowstone National Park and described a new species: S. constricta. Trichocoleus Anagnostidis (Fig. 8A) Filaments are mainly solitary and grow among other algae or cyanobacteria; they are rarely densely aggregated, forming prostrate thalli (mats). Filaments are either not or very rarely divaricated (spread apart like branches), containing a few to numerous cylindri-

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FIGURE 8 (A) Trichocoleus: (a) T. erectiusculus (bar = 10 µm; after Kondrateva, 1968); (b) T. acutissimus (after Gardner, 1927, from Puerto Rico); (c) T. purpureus (after Gardner, 1927, from Puerto Rico). (B) Borzia: (a) B. trilocularis (life cycle according to Bicudo, 1985, from Brazil); (b) B. trilocularis (after Gomont from Smith, 1950).


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cal trichomes, arranged usually in parallel and forming dense fascicles. Sheaths are more or less cylindrical or (rarely) narrowed toward the ends, not or rarely lamellated, firm or mucilaginous, often diffuse, colorless, usually slightly or clearly distant from the trichome fascicles. The trichome width ranges from 0.5 to 2.5 (3) µm. Cells are always longer than they are wide. Apical cells are acutely conical, obtuse, or rounded, without calyptra or thickened outer cell wall. Reproduction occurs via hormogonia, which separate after disintegration from the ends of the trichomes. The genus was separated from Microcoleus based on cell morphology and cell structure (Anagnostidis, 2001). Trichocoleus species occur in the periphyton or metaphyton in fresh and saline waters, although some species prefer alkaline substrata. About 16 species belong to this genus, almost all originally described as Microcoleus species. In North America, T. sociatus occurs mainly in freshwater environments (Whitford and Schumacher, 1969); T. acutissimus, T. minor, and “Microcoleus purpureus” were described from Puerto Rico (Gardner, 1927); T. acutissimus was recorded from wet rocks in Arizona, Florida, Jamaica, and Puerto Rico (Gardner, 1927). One species is benthic in shallow brackish pools in Arctic region.

motile, straight or slightly flexuous, moniliform, simple, usually short or slightly elongated, up to 650 µm long, without firm sheaths. Cells are spherical or barrel shaped, up to 10 µm wide; thylakoids (known only in one species) are typically arranged in parallel arrays perpendicular to the cell walls. Aerotopes, heterocytes, and akinetes are absent. Reproduction occurs via fragmentation, without the formation of necridic cells, into nonmotile, rarely (occasionally) motile hormogonia. About 15 species have been described, most of which are benthic and living in solitary trichomes or forming thin mats on the bottom (sandy, stony, epipelic) of unpolluted lakes, pools, reservoirs, and streams. Two species are known from thermal and mineral springs. Komvophoron is relatively new genus (Anagnostidis and Komárek, 1988) not reported by early authors or identified as Pseudanabaena. Komvophoron specimens were recently collected in North America from benthic habitats in several northern temperate lakes (Manitoba and Ontario, Canada, the authors, personal observation). K. jovis was described from thermal springs in Yellowstone National Park (Copeland, 1936), K. groenlandicum from an oligotrophic lake in Greenland near Narssaq (Nygaard, 1984, under Isocystis sp.). More detailed work is required.

Oscillatoriales: Borziaceae

Oscillatoriales: Phormidiaceae: Phormidioideae

Borzia Cohn ex Gomont (Fig. 8B) Trichomes are solitary or aggregated in small groups, simple, very short, few celled, usually nonmotile, rarely motile or trembling, and constricted or unconstricted at the cross walls. Sheaths are usually lacking or sometimes present as a fine mucilage. Cells are cylindrical to barrel shaped, more or less isodiametric or slightly shorter or longer than wide. Thylakoids are coiled and probably spread over the cell volume. Apical cells are rounded. Reproduction occurs via fragmentation into nonmotile, few-celled hormogonia, without separation discs or necridia. Seven species were recorded from the periphyton and metaphyton of clear, mostly small reservoirs. Various species grow among macrophytes, are benthic in lakes, are subaerial on calcareous substrata, or occur within the mucilage (endogloeic) of other cyanobacteria and algae (Anagnostidis and Komárek, 1988, 2001). Borzia trilocularis is fairly cosmopolitan, but distribution data are sparse; this species is reported from several localities in the United States (Daily, 1943; Smith, 1950; Taft and Taft, 1970). The distribution of other species has not been verified in North America.

Arthrospira Stizenberger ex Gomont (Fig. 9B) Trichomes are solitary and free floating in the plankton or united into a fine, mostly slimy (diffuse margins), blue–green, olive-green, or reddish-brown thallus in the benthos. Trichomes are cylindrical, isopolar, regularly or rarely somewhat irregularly loosely spirally (screwlike) coiled. Trichomes are long or short, usually with relatively large spirals (width and height), sometimes attenuated at the ends, with variable coil width, not constricted or slightly constricted at the cross walls, usually nonmotile or rarely motile (gliding with clockwise or counterclockwise rotation). Sheaths are absent or facultatively present, fine and colorless. Cells are more or less isodiametric or shorter than wide, with visible cross walls in the trichomes, sometimes with aerotopes (groups of gas vacuoles, in planktonic species), with special pore and perforation patterns in the cell walls (one row of pores around cell) and whirl-like or radially arranged thylakoids. Apical cells are rounded or conical, occasionally with calyptras or thickened outer cell walls. No toxic strains are known. Reproduction occurs via trichome fragmentation into hormogonia or hormocytes with necridic cells. Cells divide perpendicularly to the horizontal axis. About 16 species are known, several of which are freshwater benthic (A. jenneri and A. platensis); others are planktonic in subtropical and tropical saline lakes

Komvophoron Anagnostidis et Komárek (Fig. 9A) Trichomes are solitary or agglomerated in small colonies enveloped by a fine mucilage, motile or non-

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FIGURE 9 (A) Komvophoron: (a) K. minutum (after Skuja, 1948); (b) K. schmidlei (after Jaag, 1938); (c) K. jovis (after Copeland, 1936, from Yellowstone National Park); (d) Komvophoron sp. (original by Komárková, from the Everglades, Florida). (B) Arthrospira: (a) A. jenneri (photo original by Komárková from the Everglades, Florida); (b) A. platensis (after Komárek and Lund, 1990; redrawn from type material from La Plata, Argentina); (c) A. skujae (after Magrin et al., 1997, from Brazil); (d) A. maxima (after Setchell and Gardner, 1917, from California); (e) A. khannae (after Drouet, 1942, from the United States).


(A. maxima) in southern California and Mexico. Two species, A. maxima and A. fusiformis, are used in mass cultivation, but usually designated as “Spirulina platensis.” Tilden (1910) reported two species from the United States, whereas Smith (1950), Prescott (1962), Whitford and Schumacher (1969), and Stein and Borden (1979) reported the benthic species A. jenneri and/or A. gomontiana from several locations. Phormidium Kützing ex Gomont (Fig. 12A) The thallus is usually expanded, more or less fine, thin or cohesive, gelatinous, mucilaginous, cartilaginous, membranaceous, feltlike to leathery, attached to substrata or (secondary) free floating in masses, sometimes forming clusters, penicillate tufts, or (rarely) living in solitary filaments. Filaments vary in curvature; they are not pseudobranched, usually entangled, slightly to strongly waved or loosely and irregularly screwlike coiled. Sheaths occur facultatively (under unfavorable conditions) or almost obligatorily (frequency depending on environmental conditions); they are firm or thin, colorless, adherent to the trichome, not lamellated, sometimes slightly to intensely diffuse (rarely thickened and lamellated when old; generic revision may be needed). Trichomes are cylindrical, mostly long, (1.8) 2.5–11(15) µm wide, unconstricted or slightly to distinctly constricted at the cross walls, clearly motile inside the sheaths and outside the sheath (gliding, creeping, waving, trembling, with or without oscillation and rotation). Cells are typically isodiametric or shorter or longer than wide, without aerotopes (clusters of gas vesicles). Apical cells are pointed, narrowed, or rounded, with or without calyptra. Thylakoids are typically radially oriented within cells (cell content may appear netlike or striated). Cells divide by transverse fission, and each cell reaches its original size before the next division. Reproduction occurs via trichome disintegration into short or long, motile hormogonia, sometimes with biconcave necridic cells. Phormidium is a very common genus, distributed worldwide, with nearly 200 described species. It forms mats on wet soil, mud, wetted rocks and macrophytes, and in standing and running waters. Several species are known from extreme environments (thermal springs and desert soils) a few species form travertine in springs, marl lakes, and streams (Prát, 1929). Tilden (1910) listed 27 species throughout North America from Alaska to Mexico, from Newfoundland to Florida. Smith (1950) mentioned 25 species in the United States. Of the most common species, Phormidium inundatum occurs on mud or rocks in lakes and ponds. P. retzii was the most commonly recorded macroscopic alga in North American stream sites (Sheath and Cole, 1992). Prescott (1962) reported 14 species in the western Great

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Lakes area; eight species were recorded from British Columbia (Stein and Borden, 1979). Other frequently reported species are P. autumnale and P. uncinatum, which colonize stones in streams and rivers, and P. fonticolum from cold stenotherm alpine and subarctic streams and seepages (authors’ results); P. minnesotense and P. favosum are known from the periphyton and metaphyton among the aquatic plants in rivers and standing waters (Whitford and Schumacher, 1969). Planktothrix Anagnostidis et Komárek (Fig. 10A) Trichomes are solitary, free floating, more or less straight or slightly irregularly waved, isopolar (both poles with the same morphology), cylindrical, not constricted or slightly constricted at the cross walls, usually planktonic (rarely metaphytic), in massive blooms arranged into small, disintegrating irregular clusters, or diffuse in tight clumps, more or less long (up to 4 mm), (2.3)3–12(15) µm wide, nonmotile, but occasionally with inconspicuous trembling or gliding, rarely oscillation. Trichomes are slightly attenuated or not attenuated at the ends, sometimes with terminal calyptra. Mucilaginous envelopes or sheaths are usually lacking, occasionally (e.g., in culture) with fine, visible sheaths (one species has obligatory sheaths in nature as well); false branching is absent. Cells are cylindrical, rarely slightly barrel shaped, usually slightly shorter than wide or up to more or less isodiametric, rarely longer than wide. Thylakoids are typically radially arranged. Aerotopes (groups of gas vesicles) are distributed throughout the cells. Apical cells (when fully developed) are widely rounded or narrowed conical, sometimes with calyptra or with thickened outer cell wall. Segments of trichomes (with several cells) without aerotopes are a feature of the genus; these appear less pigmented than other parts of the trichome (probably diazocytes according to Bergman, 2002). PC:PE ratio was found stable in various species (without photoacclimation). Characteristic carotenoids include myxoxanthophyll and oscillaxanthin. Geosmin and toxins are present in several strains (Skulberg and Skulberg, 1985). Cells divide perpendicularly to the trichome. Reproduction occurs via disintegration into nonmotile hormocytes (with necridia). Hormogonia without gas vesicles probably overwinter in sediments. Most species (about 15 have been described) are planktonic; a few form surface blooms and may be toxic (Skulberg and Skulberg, 1985; Skulberg et al., 1993). Only Planktothrix cryptovaginata is known from the metaphyton of unpolluted pools (Skácelová and Komárek, 1989). P. agardhii probably has a cosmopolitan distribution (Prescott, 1951; Komárek, 1958; Duthie and Socha, 1976; Stein and Borden, 1979; Skulberg and Skulberg, 1985; Niiyama et al., 1993;

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FIGURE 10 (A) Planktothrix: (a) P. rubescens (after Gomont, 1892); (b) P. agardhii (after Komárek, 1958); (c) P. mougeotii (after Skuja, 1948; Komárek, 1984). (B) Trichodesmium: (a) T. lacustre (after Nygaard, 1977); (b) T. iwanoffianum (after Nygaard, 1977); (c) Trichodesmium sp. (after Smith, 1920, from Wisconsin; sub Trichodesmium lacustre).


FIGURE 11 (A) Tychonema: (a) T. bourrellyi (after Lund, 1955). (B) Pseudophormidium: (a) P. flexuosum (after Gardner, 1927, from Puerto Rico; sub Plectonema flexuosum); (b) P. murale (after Gardner, 1927, from Puerto Rico; sub Plectonema murale); (c) P. batrachospermi (bar = 20 µm; after Starmach, 1957; sub Plectonema batrachospermii); (d) P. edaphicum (after Kondrateva, 1968, sub Plectonema edaphicum).

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Komárek and Cronberg, 2001). In North America, several Planktothrix species are reported as species of Oscillatoria. Whitford and Schumacher (1969) reported P. agardhii, and P. prolifica. Prescott (1951) recorded P. agardhii, P. prolifica and P. rubescens. P. agardhii and P. mougeotii are also known from Mexico and Brazil (authors’ unpublished records). Porphyrosiphon Kützing ex Gomont (Fig. 13A) Filaments are solitary among other algae or developed into an expanded and sometimes stratified blue– green, brown, or red thallus on various substrata. Filaments are contorted or undulating, rarely with single pseudobranches, containing one or, rarely, two trichomes in the sheaths. Sheaths are thick, firm, lamellated, and usually colored, red, reddish brown, purple, yellow, yellow–brown; they are rarely colorless. Young filaments have attenuated ends with closed sheaths; old sheaths have open ends, often characteristically widened or coiled and modified after fragmentation and hormogonia release. Trichomes are nonmotile, 6–10 µm wide, and constricted or not constricted at the cross walls. Cells are isodiametric or longer than wide. Apical cells are usually rounded or conical, without calyptra. Reproduction occurs by hormogonia released from the sheaths. Most of the 20 species described are terrestrial or subaerial on wet rocks, often in mountains; others occur on tree bark. Some species colonize mud or form periphyton in clear freshwaters. Porphyrosiphon notarisii was recorded from North Carolina (Whitford and Schumacher, 1969; Tilden, 1910) and Ellesmere Island, Northwest Territories (Sheath and Steinman, 1982); it is probably widely distributed throughout North America (Smith, 1950), but the identity of different populations should be revised. Tilden (1910) listed P. notarisii from the West Indies; two other species probably occur in North America (P. versicolor and P. fuscus). Pseudophormidium (Forti) Anagnostidis et Komárek (Fig. 11B) Filaments are long or short and are usually aggregated into expanded mats, tufts, or clusters, often densely entangled, sometimes radially arranged, rarely solitary among other algae and cyanobacteria. Filaments vary in curvature and are heavily (obligately) pseudobranched; they commonly disintegrate into numerous trichome segments with single branches overlapping in a common sheath. Sheaths are always present; they are firm, rarely mucilaginous and diffuse, colorless or colored, and lamellated or not. Trichomes are up to 10 µm in width, usually distinctly constricted at the cross walls. Cells are more or less isodiametric or shorter or longer than wide; cells lack aerotopes. Apical cells are usually rounded to obtuse conical, without

calyptra or thickened outer cell wall. Reproduction occurs via trichome fragmentation into nonmotile hormocytes or also possibly motile hormogonia (?). Pseudophormidium contains mostly periphytic species, commonly growing on soil or creeping on stone surfaces or other substrata in unpolluted streams (Anagnostidis and Komárek, 1988, 2001). One species (P. batrachospermi) is endogloeic in the mucilage of algae (Starmach, 1957, as Plectonema batrachospermi). Although not yet recorded, some of the 14 described species likely occur in American freshwaters and soils (a species similar to P. tenue was recorded from Massachusetts, North Carolina, and Wisconsin; Tilden, 1910). Symploca Kützing ex Gomont (Fig. 12B) The thallus is composed of entangled or parallel filaments, forming compact, more or less wooly, often terrestrial or subaerial masses. Filaments are at first prostrate, irregularly curved, later mainly united into numerous, partly pseudobranched, erect (rarely prostrate) fascicles that arise from the thallus as erect, conical, often confluent tufts. Sheaths are thin or thick, firm, distinct, in the fascicles often mucilaginous and laterally slightly confluent or somewhat gelatinized, at the ends straight and slightly attenuated, containing one trichome, always open at the end. Trichomes are straight, often weakly attenuated, thin or up to 8 (14) µm wide. Cells are isodiametric or either shorter or longer than wide, probably with a radial arrangement of thylakoids. Apical cells are never capitate, but often with a thickened outer cell wall. Reproduction occurs by motile hormogonia. Of the 70 species described, about 30 have been revised (Anagnostidis and Roussomoustakaki, 1985). Most species are terrestrial on wet soil or subaerophytic on rocks and mosses; some are marine. Smith (1950) listed S. muscorum on damp soils and moist cliffs, along with 11 other species from various locations in the United States. S. thermalis, S. nemecii, and S. ciliata were recorded from the edge of hot springs in Yellowstone National Park (Prát, 1929; Copeland, 1936). Whitford and Schumacher (1969) recorded S. borealis, S. dubia, and S. muralis on rocks and wet soil and at the bottom of drying pools in North Carolina. Other species reported from North America include S. cartilaginea (Gomont, 1892; Geitler, 1932), S. cavernarum (Copeland, 1936), and S. kieneri (Drouet, 1943a). Tilden (1910) listed nine species in the United States, Mexico, Greenland, and the West Indies. Trichodesmium Ehrenberg ex Gomont (Fig. 10B ) Trichomes are planktonic, free floating, more or less straight, rarely solitary, usually in parallel or radial


FIGURE 12 (A) Phormidium: (a) P. formosum (after Komárek, 1989, from Cuba); (b) P. richardsii (after Drouet, 1942, from the United States); (c) P. autumnale (after Komárek, 1988; Smith, 1950; microphoto originally by R. G. Sheath, with permission); (d) thylakoid arrangement in Phormidium cells (after Wolf from Komárek, 2001). (B) Symploca: (a) fascicles; (b) S. muscorum (bar = 10 µm; after Smith, 1950; Anagnostidis and Roussomoustakaki, 1985); (c) S. hydnoides (after Frémy, 1930).

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FIGURE 13 (A) Porphyrosiphon: (a) end of filament; (b) P. versicolor (after Frémy ex Geitler, 1932); (c) P. notarisii (Smith, 1950); (d) P. robustus (after Gardner, 1927, from Puerto Rico); (e) P. fuscus (after Frémy ex Geitler, 1932). (B) Lyngbyopsis: (a) L. willei (after Gardner, 1927 from Puerto Rico); (b–c) L. willei (after Komárek, 1989, from Cuba).


arrangements in colonies that form fascicles or flocculent masses, joined by diffuse mucilage. Trichomes occur without individual sheaths; they are straight or curved, rarely irregularly spirally twisted, slightly motile (inconspicuous gliding), 6–22 µm wide, cylindrical or with slightly tapering ends. Cells are typically isodiametric or slightly longer or shorter than wide, with homogeneous or finely granular content and with obligate aerotopes arranged irregularly throughout the blue–green or reddish protoplast. Apical cells are rounded or slightly capitate. Another distinguishing feature (from Planktothrix) is in the composition of fatty acids (Umezaki, 1974). Cell division occurs mainly in the meristematic zones in the middle of the trichomes; reproduction occurs via trichome fragmentation. Trichodesmium is primarily a pelagic species in oceans, capable of forming extensive water blooms, but several species occur in freshwaters (sometimes considered more related to Planktothrix); 11 exclusively planktonic species have been described, but freshwater populations need taxonomic revision (Niiyama et al., 1993). The freshwater T. lacustre (as Oscillatoria lacustris) is widespread but not abundant in North Carolina (Whitford and Schumacher, 1969) and in scattered locations in British Columbia (Stein and Borden, 1979). Smith (1920, 1950) mentioned the genus as being widely distributed throughout the United States. Tilden (1910) listed three marine species, two in the region of the West Indies and Central America. Tychonema Anagnostidis et Komárek (Fig. 11A) Trichomes are solitary or organized into fine mats, which may be benthic, tychoplanktonic, or planktonic. Trichomes are cylindrical, pale grayish–pinkish, purplish, reddish, or olive green, up to 5 mm long, 2–16 µm wide, without sheaths or with fine facultative mucilaginous sheaths, nonmotile (?) or with reduced motility (slightly trembling, gliding, or rotating), without false branching, usually not constricted at the cross walls. Filaments are straight or, occasionally, slightly irregularly coiled or more or less curved, not attenuated at ends. Cells are identical in morphology, cylindrical, more or less isodiametric, or shorter or longer (up to twice) than wide. Gas vesicles (and aerotopes) are always absent, but cells often contain prominent granules; cell content is pale and “alveolar” with keritomized chromatoplasma (seemingly vacuolated, but with radially arranged, widened thylakoids, and almost colorless; Komárek and Albertano, 1994). Apical cells are rounded, sometimes with thickened cell walls or narrow calyptras. Phycobilin content is variable (chromatic acclimation), mainly containing phycoerythrin (olive green to reddish brown); several characteristic

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carotenoids are present. Toxic strains are not known (Skulberg and Skulberg, 1985). Reproduction occurs via disintegration of trichomes into nonmotile hormocytes (motile hormogonia?). Tychonema species are planktonic or benthic (altogether eight species have been described, but only T. bourrellyi and T. bornetii are well known and their generic characters proved). Tychonema is possibly a cold-stenotherm genus of northern temperate areas. T. bourrellyi is known to form metalimnetic maxima in plankton, mainly recorded in northern Europe (Lund, 1955; Skulberg and Skulberg, 1985), but also in Canada (Kling and Holmgren, 1972; Findlay and Kling, 1979).

Oscillatoriales: Phormidiaceae: Microcoleoideae Dasygloea Thwaites ex Gomont (Fig. 14B) Dasygloea occurs as solitary, creeping filaments growing among other algae or in mucilaginous mats on substrata. Sheaths are distinctly irregularly widened, colorless or colored, distinctly outlined, with a smooth surface, usually clearly or indistinctly lamellated (stratified), sometimes funnel-like widened, irregularly branched or unbranched, usually closed at the ends. Sheaths contain one or two (rarely several) parallel, distant trichomes. Trichomes are cylindrical, usually wavy or almost straight, constricted or unconstricted at the cross walls, with noncapitate ends, (3) 4–10 (12) µm wide, with isodiametric cells or slightly shorter or longer than wide. Reproduction occurs by hormogonia, separated by necridic cells. The nine species described live on muddy sediments or are metaphytic in swamps and pools (Senna and Komárek, 1998). D. lamyi and D. amorpha are known from Jamaica and the United States (Pennsylvania) (Geitler, 1932; Starmach, 1966). D. calcicola and D. yellowstonensis were recorded from Yellowstone National Park from the periphyton on calcified encrustations near thermal springs (Copeland, 1936). Tilden (1910) listed other species from Pennsylvania. Hydrocoleum Kützing (Fig. 15B) Hydrocoleum is composed of solitary filaments or thallus usually composed of filaments joined in small, smooth, caespitose, spherical or cushion shaped, microscopic to macroscopic thalli, or forming extended amorphous, membranaceous, flat or cushion-like mats, rarely compact, often encrusted with CaCO3. Filaments are usually straight, sometimes of variable curvature, rarely sparsely pseudobranched, more or less parallel (rarely radial), or joined to form tufts or erect ropelike fascicles. Sheaths are thick, mucilaginous, and smooth, colorless or yellowish, initially firm, usually limited, when older diffuse, with longitudinal striation (strati-

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FIGURE 14 (A) Symplocastrum: (a) Symplocastrum sp. (after Drouet, 1939, from the United States; sub Schizothrix wollei); (b) S. purpurascens (after Gomont, 1892); (c) S. parciramosa (after Gardner, 1927, from Puerto Rico; sub Hypheothrix parciramosa). (B) Dasygloea: (a) D. yellowstonensis (after Copeland, 1936, from Yellowstone National Park); (b) D. brasiliense (after Senna and Komárek, 1998); (c) D. lamyi (after Gomont from Starmach, 1966).


FIGURE 15 (A) Microcoleus: (a) M. vaginatus (after Smith, 1950, from the United States); (b) M. chthonoplastes (after Gomont from Kondrateva, 1968); (c) thylakoid position in cells (cross section in Microcoleus chthonoplastes; after Hernandéz-Mariné, 1996). (B) Hydrocoleum: (a) H. groesbeckianum (after Drouet, 1943, from eastern California); (b) H. homeotrichum (after Gomont, 1892; Smith, 1950).

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fied), containing several (2–10) to many parallel and tightly fasciculated trichomes. Trichomes are straight, usually densely (rarely loosely) aggregated, more or less gradually attenuated at the ends. Cells are isodiametric or slightly shorter or longer than wide. Apical cells are capitate, often with a thickened outer cell wall or calyptra. Reproduction occurs via hormogonia. Of the several species described, about 12 are periphytic in clear streams and rivers or clear cold lakes. Some are present in marine littoral areas or in the littoral areas of marl lakes (with precipitations in the sheaths). Whitford and Schumacher (1969) observed H. homoeotrichum on wet soil in cultivated fields, whereas Prescott (1962) noted one species from the western Great Lakes region. Smith (1950) mentioned that H. homoeotrichum appears in the United States in both aerial (?) and aquatic habitats. Tilden (1910) listed eight species from the Caribbean, the east coast of the United States, Pennsylvania, and Texas, but a few of them were later reclassified into the genus Blennothrix (Oscillatoriaceae). Lyngbyopsis Gardner (Fig. 13B) The thallus occurs in the form of macroscopic, semiglobose or flattened mats (in strata) on submersed stony substrata or with filaments grouped into prostrate layers. The sheaths are thin or slightly thickened, colorless, firm, and membranaceous, containing one or few parallel trichomes, each developing an individual sheath. Filaments are sometimes anastomozed by their sheaths and linked together. Pseudobranching occurs in one or both directions toward the ends of the longitudinal axis. Trichomes are isopolar, with straight, not capitate, ends. Reproduction occurs via hormogonia, after trichome disintegration through the formation of necridic cells. Lyngbyopsis is a monotypic genus with the single species Lyngbyopsis willei. Gardner (1927) described Lyngbyopsis from Puerto Rico. It is also known from warm-water creeks in Cuba (Komárek, 1989). Microcoleus Desmazières ex Gomont (Fig. 15A) Filaments are solitary or in flat mats, usually prostrate on various substrata, and unbranched or sparsely pseudobranched. Sheaths are homogeneous, broad, occasionally indistinctly and irregularly longitudinally striated, mostly colorless, regularly cylindrical, firm, delimited or sometimes (when old) diffuse, occasionally transversely wrinkled, tapering and usually open at the ends, and containing numerous to many trichomes. The trichomes within sheaths are densely aggregated or ropelike contorted, nearly parallel and in tight fascicles, often extending beyond the sheath ends, without constrictions or slightly constricted at the cross walls,

straight, and mostly attenuated at the ends without individual sheaths. Cells are cylindrical, more or less isodiametric, with radially arranged thylakoids and granular contents. Apical cells are usually subconical to acutely conical, less frequently capitate, sometimes with calyptra. Reproduction occurs via trichome disintegration and hormogonia production. Several common species have been recorded from the littoral of clear lakes or slightly polluted pools; some species are benthic (in sediments or sand); other species are known from wet soils of different kinds, deserts, but also rivers, estuaries, and marine (haloplilic) environments, on wet rocks or mineral springs. More than 30 species were revised, after the genus Trichocoleus (Schizotrichaceae) was separated from Microcoleus (Anagnostidis and Komárek, 1988; Anagnostidis, 2001). Whitford and Schumacher (1969) mentioned seven species of which only M. lacustris grows in shallow freshwater; the others are known from wet soils and drying mud, very often in saline habitats. Smith (1950) listed a dozen freshwater species in the United States, and Tilden (1910) noted seven species from various areas of North America, including Alaska and the West Indies. Several other freshwater and marine species were described from locations in the United States (Setchell and Gardner, 1918, 1924; Drouet, 1943a) and Puerto Rico (Gardner, 1927). Symplocastrum (Gomont) Kirchner ex Engler et Prantl (Fig. 14A) The thallus is feltlike, in tufts, expanding to form velvet-like layers; it is composed of fascicles of parallel and tightly linked filaments. Trichomes have individual sheaths. Fascicles are mostly erect (rarely creeping), long (up to 3 cm), acutely pointed, and usually in closed fascicles enveloped by a common sheath. Enveloping sheaths are wide, colorless or colored, usually firm, sometimes lamellated, occasionally diffuse, variably branched, containing initially a few, later numerous trichomes. Trichomes are usually constricted at the cross walls. Cells are usually isodiametric or longer than wide. Reproduction occurs via hormogonia and thallus disintegration. Symplocastrum contains species that resemble the members of the genus Schizothrix by their filament and thallus morphology; however, the trichomes are wider and their structure corresponds to the Phormidiaceae. Eleven revised species (usually referred to under the name Schizothrix) were recorded from North America, including the terrestrial S. friesii and S. purpurascens, S. sauterianum epiphytic on Cladophora, and S. muelleri in standing waters (Prescott, 1962; Sheath and Steinman, 1982). Tilden (1910) listed four species in the United States, Canada, and the West Indies. Whitford and


Schumacher (1969) recorded S. penicillatum growing on submerged concrete in waters in the coastal plains of North Carolina. Within Symplocastrum probably also belong taxa referred to as Schizothrix acuminata, S. giuseppei, S. parciramosa, and S. telephoroides from Puerto Rico (Gardner, 1927), and S. acutissima, S. californica, S. chalybea, S. constricta, S. hancockii, S. mexicana, S. richardsii, S. rivularis, S. stricklandii, and S. taylori, recorded from various locations in the United States and Mexico (Tilden, 1910).

Oscillatoriales: Phormidiaceae: Ammatoideoideae Ammatoidea W. West et G. S. West (Fig. 18A) Filaments occur in clusters, in tufts, or in indefinite strata, or solitary among other algae, often growing prostrate with ascending branches and tips, or curved in the middle with ascending ends, more or less irregularly coiled, isopolar, narrowed toward both ends, sometimes falsely branched. Sheaths are firm, thin or thick, usually lamellate (at least in parts), colorless or yellowish to brown, open at both ends. Trichomes are isopolar, rarely (young trichomes) heteropolar, usually distinctly gradually narrowed toward both ends and with elongated cells, usually forming colorless cylindrical hairs, without constrictions or slightly constricted at the cross walls in old parts and in hormogonia. Meristematic zones with dividing cells are present. Heteropolarity and akinetes are absent. Reproduction occurs via fragmentation of the trichomes with necridia, forming motile hormogonia. Hormogonia separate from both ends of the trichomes (rarely central) after separation of terminal hairs. Filaments of A. normannii and A. yellowstonensis were observed by Copeland (1936) attached to the surface of gelatinous envelopes of other algae in Yellowstone National Park (see also Smith, 1950; Stein and Borden, 1979, under Hammatoidea).

Oscillatoriales: Oscillatoriaceae Blennothrix Kützing ex Anagnostidis et Komárek (Fig. 19) The thallus is mucilaginous, expanded or fasciculate and lengthened, cylindrical, filamentous or tufted, flaky, rarely hemispherical and cushion-like, up to 2 cm high, occasionally free floating, forming olive-green to black–green, bright to dull blue–green, blackish, rarely red–brown or black–violet masses. Filaments are straight or slightly undulating and entangled, in divaricated (spread apart like branches) or sparsely falsely branched fascicles or tufts, often with a special type of branching (“coleodesmoid,” not “plectonematoid”; Watanabe and Komárek, 1989; compare Figs. 18B and 19). Sheaths are always present, thin or thick, mucilagi-

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nous, firm or diffuse, sometimes lamellated, colorless, frequently with transverse lamellation and constrictions, open at apex, containing one to several trichomes, tightly aggregated (rarely loose). Trichomes are 8–40 µm wide, usually cylindrical or attenuated and straight up to capitate ends, and usually unconstricted to slightly constricted at the cross walls. Cells are very short and discoid and divide by perpendicular cleavage in rapid succession. Apical cells are present, often with calyptra or thickened outer cell wall. Reproduction occurs via trichome disintegration and formation of hormogonia with necridic cells. About 20 species have been described (usually under the name Hydrocoleum), which belong to two ecological groups: One colonizes clear mountain springs and streams; the other occurs in brackish and saline environments. Of North American species, B. ganeshii is common in mountain streams in central Mexico (Gold-Morgan et al., 1994; Gold-Morgan and Montejano, personal communication); B. ravenelii, B. heterotricha, and calcified B. groesbeckiana are known from clear standing and flowing waters (Tilden, 1910; Drouet, 1943a; Smith, 1950); B. coerulea was collected from mountain streams in Puerto Rico (Gardner, 1927). B. glutinosa, B. majus, and B. mirifica are known from marine habitats. Several other species occur in marine littoral and saline swamps in central America (e.g., B. cantharidosma and B. comoides in Florida, the Antilles, Bermuda, and Mexico; Geitler, 1932; Humm and Wicks, 1980, under Hydrocoleus). Lyngbya C. Agardh ex Gomont (Fig. 17) Filaments are straight or slightly undulating (several species are finely screwlike or coiled), rarely (a few free-floating species) solitary, mainly arranged in thin or thick, flat, compact, large, layered, leathery prostrate mats on the substrate, and very rarely false branched; they are usually wider than 6 µm. Sheaths are always present; only hormogonia and trichomes under extreme conditions leave the sheaths. Sheaths are attached to the trichome or slightly distant, firm, thin or thick, colorless or slightly yellow–brown or reddish (very rarely bluish), sometimes slightly lamellated, and containing one motile trichome. Trichomes are cylindrical and may or may not be constricted at the cross walls. Cells are short and discoid, always shorter than wide, and most without aerotopes (a few planktonic species have aerotopes). Apical cells usually have a thickened outer wall or calyptra. Reproduction occurs via trichome disintegration into usually short motile hormogonia, which often separate necridia formation. Over 60 species have been revised and confirmed, several of which are cosmopolitan. Most form periphytic and benthic mats on different submersed

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FIGURE 16 Oscillatoria: (a) O. limosa (after Smith, 1950, from the United States); (b) O. obtusa (after Gardner 1927 from Puerto Rico); (c) O. refringens (after Gardner, 1927, from Puerto Rico); (d) O. jenensis (after Komárek, 1989, from Cuba); (e) O. sancta (original photo by Kling, from central Canada); (f) Oscillatoria sp. (original photo by Komárková, from the Everglades, Florida); (g) O. princeps (original photo by R. G. Sheath, with permission).


FIGURE 17 Lyngbya: (a) L. splendens (after Gardner, 1927, from Puerto Rico); (b–c) Lyngbya sp. (original photo by Komárková, from the Everglades, Florida); (d) L. magnifica (after Gardner, 1927, from Puerto Rico); (e) L. intermedia (after Gardner, 1927, from Puerto Rico); (f–g) L. birgei (after Smith, 1920; original photo by Kling, from Canadian lakes).

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FIGURE 18 (A) Ammatoidea: (a) A. normanii (after W. West and G. S. West from Smith, 1950); (b) A. yellowstonensis (after Copeland, 1936, from Yellowstone National Park). (B) Plectonema: (a) P. tomasinianum (after Drouet, 1934, from Missouri; sub P. tomasinianum var. gracile); (b) P. tomasinianum (after Gomont from Kondrateva, 1968).


substrata in freshwater and brackish and marine waters. Some species are terrestrial or subaerial occurring on wet rocks; several of them are cosmopolitan. A few species (e.g., L. birgei, described from the United States) occur as solitary filaments in metaphyton and plankton. Lyngbya maior, L. martensiana, and L. spirulinoides are widely collected from periphyton in pools and streams (Tilden, 1910; Prescott, 1951; Whitford and Schumacher, 1969; Stein and Borden, 1979); L. aestuarii, L. meneghiniana, L. salina, and L. confervoides prefer saline waters (Geitler, 1932; Kosinskaja, 1948). Tilden (1910) listed 35 species, 29 of which are distributed over the North American continent; Smith (1950) recorded 35 species from the United States. However, many of these species now belong to other genera (e.g., L. contorta = Planktolyngbya contorta). Drouet (1934, 1942) described L. hahatonkensis from an artesian spring in the United States, along with periphytic L. giuseppei and L. patrickiana. The North American species require further research and revision. Oscillatoria Vaucher ex Gomont (Fig. 16) The thallus is usually flat, macroscopic, smooth, layered, rarely leathery, arranged in mats, less frequently in solitary trichomes. Trichomes are straight or slightly irregularly undulating, cylindrical, sometimes screwlike coiled at the ends, motile, gliding or oscillating in left-handed or right-handed rotation, usually wider than 6.8 µm (up to 70 µm), not constricted or constricted at the cross walls. Sheaths are usually absent, although they may occur under suboptimal conditions. Cells are short and discoid, always with lengths less than one half to one eleventh that of their widths; cell contents without aerotopes are homogeneous or sometimes contain large prominent granules. Cell division occurs in a rapid sequence transversely to the trichome axis. Reproduction occurs via trichome disintegration (sometimes completely) into short motile hormogonia, with the aid of necridia. Almost 70 widely distributed, mainly benthic species have been described (revised; traditionally planktonic species with thinner trichomes and different ultrastructure now classified under the genera Planktothrix, Pseudanabaena, or Limnothrix). Oscillatoria occurs in mats on different substrata (mud, plants, stones, sand) in shallow water bodies or marshes and swamps. With greater biomass, parts of the mats dislodge from the substratum and form floating clusters on the water surface in shallow water bodies. Of the numerous Oscillatoria species, the most frequently mentioned are the cosmopolitan O. limosa, O. sancta, O. princeps, and O. proboscidea; these species grow epipsammic or among other algae in lakes, ponds, and pools and in slowly flowing rivers (Prescott, 1962;

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Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). O. ornata, O. curviceps, and O. anguina have also been reported (Tilden, 1910; Prescott, 1951; Whitford and Schumacher, 1969). O. rhamphoidea was collected from drying pools with water plants in Winnipeg, Canada (authors’ results). O. limosa and O. tenuis were recorded on rock and mud habitats in arctic freshwater lakes and streams (Whelden, 1947). Several species grow in marine coastal waters (O. funiformis and O. margaritifera). Copeland (1936) described O. depauperata from Yellowstone National Park. Smith (1950) mentioned 40 species definitely known from the United States and Tilden (1910) listed 43 species widely distributed widely throughout the North American continent; however, many of these species cited by previous authors have been classified into other genera (Anagnostidis and Komárek, 1988). Plectonema Thuret ex Gomont (Fig. 18B) The thallus forms expanded and usually compact tufts up to 2 cm high, attached to various substrata, or as free clusters with coiled filaments. Free-floating masses also occur, composed of variously coiled, usually densely entangled, or rarely almost straight and parallel filaments. Filaments are obligatorily falsely branched (sparsely to frequently), up to 120 µm wide; false branches occur singly or in clusters, sometimes Scytonema-like. Sheaths are firm, thin to thick, up to 4 µm wide, initially colorless, later sometimes yellow– brown, homogeneous or distinctly lamellated. Trichomes are 8–25 (up to 72?) µm wide and are nonmotile. Cells are short and discoid; apical cells are rounded, without (rarely with) calyptra. Heterocytes and akinetes are absent. Reproduction occurs mainly by hormogonia, separating from apical portions of the trichome by the help of necridia, rarely by whole trichome fragmentation. Hormogonia separate from both trichome ends (isopolar). Of the many species described, only five have been confirmed (several have been transferred to other genera), occurring in the periphyton and metaphyton of clear, oligotrophic, and well-oxygenated ponds and pools; they are found less frequently in small streams or springs. Tilden (1910) and Smith (1950) recorded several species from the United States, most of which were attached to various submerged substrata and located on damp rocks or among mosses and liverworts; however, some were later reclassified into other genera. Whitford and Schumacher (1969) mentioned four species, with two freshwater species (P. wollei and P. tomasinianum). Recently, P. wollei from some North American lakes was found to contain toxins (Carmichael et al., 1997).

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FIGURE 19 Blennothrix: (a) false branching; (b) position of thylakoids in cells; (c) B. ganeshii (after Watanabe and Komárek, 1989, from Himalayas; occurring also in Mexico mountains); (d) B. cantharidosma (after Gomont ex Geitler, 1932).


FIGURE 20 (A) Scytonema: (a–c) bipolar germination of hormogonia; (d) S. capitatum (after Gardner, 1927, from Puerto Rico); (e) S. longiarticulatum (after Gardner, 1927, from Puerto Rico). (B) Scytonematopsis: (a) S. hydnoides (after Copeland, 1936, from Yellowstone National Park); (b) S. fuliginosa (after Kosinskaja, 1926).

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Nostocales: Scytonemataceae Scytonema Agardh ex Bornet et Flahault (Fig. 20A) The thalli form wooly mats or irregular clusters, yellow to brownish in color. Filaments are free floating or in fascicles, curved, sometimes densely coiled, and creeping on substratum. False branching is simple, usually into two branches (rarely in one branch). Branching often occurs after the formation of necridic cells, less frequently at heterocytes following disintegration of the trichome; the free ends of the trichomes squeeze through a rupture in the sheath and form new branches. Trichomes are isopolar (both poles with the same morphology), cylindrical, at their terminal parts cylindrical or sometimes widened, constricted or unconstricted at the cross walls, pale blue–green, olive green, or brownish, rarely violet. Sheaths are firm, lamellated or without visible structure, sometimes wide, often yellow–brown in color (scytonemin; Jaag, 1945; Geitler, 1960). Cells are cylindrical to barrel shaped, sometimes elongated in the central part of the trichome; apical cells are rounded. Heterocytes are solitary and intercalary. Akinetes have occasionally been reported, but not confirmed. Reproduction occurs via isopolar hormogonia released at both ends of the trichomes. Germination of hormogonia at both ends. Scytonema species are mainly aerial or subaerial on alkaline substrata, wet rocks, wood, and soil, sometimes encrusted with calcium carbonate; some species grow in the periphyton in lakes (mainly calcareous) and at sea coasts. Scytonema is a common, variable genus, with over 100 described species, many of which are known only in tropical regions. Tilden (1910) reported 36 species from North America. Whelden (1947) noted Scytonema crustaceum embedded in calcareous precipitates inside the hollow nodes of a Nostoc colony, S. myochrous in a freshwater tundra pool (typically S. myochrous grows in travertine habitats), and S. ocellatum from the edge of an Arctic lake. Twenty species of Scytonema were reported by Smith (1950) primarily from subaerial habitats, but also from damp walls and rocky cliffs; one species has been found in algal mats in marshes in the Florida Everglades. Diplocolon heppii, which is considered synonymous with the genus Scytonema (Geitler, 1932, 1942), was recorded from Niagara Falls (Smith, 1950). Whitford and Schumacher (1969) reported 19 species in North Carolina, three of which (S. arcangelii, S. crispum, and S. dubium) were periphytic in freshwaters, whereas the rest were subaerial on wet soils and rocks. Several taxa were also reported from several locations across Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982); S. cincinnatum, S. ocellatum, and S. tolypothrichoides were reported from the central United States (Tiffany and Britton, 1952).

Scytonematopsis E. Kiseleva (Fig. 20B) The thalli are brownish, low, prostrate, and closely attached to the substrata or in clusters. Filaments are falsely branched, heteropolar when young, later usually isopolar, forming clusters with parallel or a bristle-like arrangement; old trichomes are isopolar. Branching (usually two) starts at necridic cells. Trichomes are usually constricted at the cell walls. Sheaths are firm, sometimes parallel lamellated and telescopic, hyaline or yellowish-brown in old filaments. Cells vary from short to long cylindrical or barrel shaped; several terminal cells are distinctly elongated, often forming a thick tapering hair, with the terminal cell bluntly or sharply pointed. Heterocytes are mostly intercalary and solitary. Akinetes have been described, but are rarely found. Reproduction occurs via hormogonia separated from filaments, which germinate from both ends. This genus contains both periphytic and metaphytic species occurring in temperate mountain streams; one species (S. hydnoides) was described from thermal springs in Yellowstone National Park (Copeland, 1936); the type species S. fuliginosa was described from the Hawaiian islands (Tilden, 1910; Copeland, 1936).

Nostocales: Microchaetaceae: Tolypotrichoideae Coleodesmium Borzì ex Geitler (Fig. 23) Filaments are united into highly branched, mucilaginous, polarized blue–green or brownish fascicles, up to 1 cm high. Sheaths are firm, thin or thick, and often lamellated and opened at the apex, colorless or yellow–brown, containing one to several trichomes (with own sheaths). Trichomes are heteropolar with basal, elliptical heterocytes, parallel orientation, false branching, cylindrical, uniseriate, constricted at the cross walls, and sometimes slightly attenuated or widened toward the ends. False branching is often initiated at intercalary heterocytes. Cells are short, barrel shaped to cylindrical, with widely rounded end cells. Aerotopes are absent. Akinetes are rare. Cell division is perpendicular to the longitudinal axis of the trichome. Reproduction takes place by the production of hormogonia (separated by necridic cells), liberated from the sheaths. Coleodesmium species are epiphytic on macrophytes or epilithic on stones in clear, unpolluted streams, especially in mountainous and northern areas. Coleodesmium floccosum (originally identified as C. wrangelii) has been recorded from Connecticut (Geitler, 1932; Komárek and Watanabe, 1990) and British Columbia (Stein and Borden, 1979); typical C. wrangelii occurs in subarctic to arctic waters (Elster et al., 1997) and (as Desmonema wrangelii; Smith, 1950) in several other localities of North America. Whitford and Schumacher (1969) collected C. wrangelii (also as Desmonema) on concrete in a swift stream in North Carolina.


FIGURE 21 (A) Hassallia: (a) Hassallia false branching; (b) H. discoidea (after Gardner, 1927, from Puerto Rico); (c) H. granulata (after Gardner, 1927, from Puerto Rico); (d) H. byssoidea (after Frémy ex Kondrateva, 1968). (B) Petalonema; (a) P. alatum (after Komárek ex Fott, 1956; Kondrateva, 1968); (b) P. involvens (after Frémy ex Geitler, 1932).

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FIGURE 22 Tolypothrix: (a) branching; (b) T. willei (after Gardner, 1927, from Puerto Rico); (c) T. robusta (after Gardner, 1927, from Puerto Rico); (d) T. papyracea (after Gardner, 1927, from Puerto Rico); (e) T. amoena (after Gardner, 1927, from Puerto Rico); (f) T. tenuis (after Smith, 1950; original microphoto Komárková from the Everglades, Florida); (g) T. penicillata (after Geitler, 1932).


Hassallia Berkeley ex Bornet et Flahault (Fig. 21A) The thalli are filamentous, forming low, bristle-like bundles or thin or wooly mats, or they are solitary, creeping along substrata. Rich false branching is due to many simple, solitary, short lateral branches, each starting at a heterocyte. Filaments are heteropolar, cylindrical, and curved. Sheaths are firm, thick and lamellated, sometimes yellowish brown in color. Trichomes are usually constricted at the cross walls, with end cells sometimes slightly attenuated or rounded, with granular content. Cells are barrel shaped to cylindrical when old, mostly shorter than wide to discoid. Heterocytes are mostly hemispherical or discoid, sometimes spherical, mostly single pored, situated at the trichome base or intercalary. Akinetes are unknown. Cells divide in meristematic zones at the ends of the trichomes, perpendicular to the main axis. Reproduction occurs via separation of entire branches and hormogonia. Species occur in aerial or subaerial habitats on stones or tree bark or submersed. About 10 species are known; several occur in the United States and Canada, but are recorded mainly under Tolypothrix (e.g., subaerial Hassallia byssoidea; Tilden, 1910; Smith, 1950; Stein and Borden, 1979). Petalonema Berkeley ex Kirchner (Fig. 21B) Filaments are solitary or in small groups, heteropolar, often with a basal heterocyte, forming a creeping thallus on substrata, slightly curved or bent; false branching occurs next to heterocytes or after filament disintegration. Trichomes are cylindrical and narrow in comparison with the thick sheaths. Sheaths are very wide, firm, distinct, limited, funnel-like, lamellated, colorless to yellow–brown. Cells are elongated cylinders in the central parts of the trichomes, short and barrel shaped at the ends, with rounded apical cells. Heterocytes are spherical, solitary, situated at the trichome base, intercalary or at the basal point of the branch. Akinetes are absent. Cells divide in meristematic zones. Reproduction occurs via hormogonia from the end of the trichomes. A common representative, Petalonema alatum, is known from subaerial or submersed habitats on calcareous substrata, particularly in temperate zones. About eight species were recorded from North America (under Scytonema; e.g., Tilden, 1910; Stein and Borden, 1979), but some require revision. Tolypothrix Kützing ex Bornet et Flahault (Fig. 22) Thalli form wooly mats, tufts, or caespitose colonies; they are grayish blue–green to yellowish or dark brownish in color. Young filaments are heteropolar with basal heterocytes and free apical ends, often very long. False branching begins next to the heterocyte, diverging singly from the main filament or form-

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ing two morphologically identical branches. Trichomes are uniseriate, with one or more heterocytes at the base, constricted or unconstricted at the cell walls, sometimes slightly widened toward the apex. Cells range from long cylindrical to short barrel shaped, sometimes with a few granules, blue–green, olive green, yellow to reddish. Cells in the subapical meristematic zones divide transversely to the trichome axis. Heterocytes are spherical, cylindrical, or discoid with one or two pores, situated intercalary, originally singly or in pairs, often at the base of branches. Akinetes are rarely seen. Reproduction occurs via hormogonia that separate from the filament ends or from a disintegrating part of a filament. Hormogonia germinate from one or both ends; growth later becomes heteropolar. Species occur in unpolluted freshwaters attached to stones, macrophytes, and other algae, or forming mats in mineral springs, streams, and alkaline swamps or subaerial habitats. Many species are known from tropical and subtropical habitats (Gardner, 1927; Geitler, 1932; Desikachary, 1959). Forty species have been described, but there are difficulties with their identification (Hoffmann and Demoulin, 1985). Tolypothrix is more commonly encountered in aquatic habitats than the mainly terrestrial and subaerial Scytonema. Colonies originally formed in benthic habitats may later separate and float to the surface. Tilden (1910) listed 10 species distributed throughout the Arctic, the United States, and the West Indies. Whelden (1947) reported T. bouteillei, T. distorta, T. penicillata, T. tenuis, and T. tenuis f. minor (noted as a new taxon found on the bottom of a stream) from the Arctic. Smith (1950) reported T. distorta, T. setchellii, T. lanata, and T. penicillata from various areas in the United States, as have many other studies (e.g., Tiffany and Britton, 1952; Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). Eleven species are recorded from North Carolina, mostly subaerial or in submersed periphyton (T. limbata and T. rupestris). Tolypothrix is mainly distinguished from Hassallia by its diverse thallus structure (Komárek and Anagnostidis, 1989).

Nostocales: Microchaetaceae: Microchaetoideae Fortiea De Toni (Fig. 24A) The thallus is filamentous and epiphytic or periphytic on macrophytes or stones. Filaments are solitary or in groups, heteropolar, characterized by basal heterocytes and free, widened apical ends, cylindrical, simple, solitary or in small groups. Filaments always contain only one trichome. Sheaths are firm, colorless, occasionally thick and lamellated length- and crosswise. Trichomes are cylindrical, constricted or unconstricted at the cross walls, widened toward ends, with the last

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FIGURE 23 Coleodesmium: (a) false branching; (b) C. wrangelii (after Frémy, 1927; Bourrelly, 1970); (c) Coleodesmium sp. (after Smith, 1950, from the United States; sub Desmonema wrangelii); (d) C. floccosum (after Komárek and Watanabe, 1990, from Connecticut).


FIGURE 24 (A) Fortiea: (a) trichome; (b) F. salinicola (after Komárek, 1984, from Cuba); (c) F. monilispora (after Komárek, 1984, from Cuba). (B) Microchaete: (a) M. tenera (after Smith, 1950; Kondrateva, 1968); (b) M. robinsonii (bar = 20 µm; after Komárek, 1994, from southern Manitoba).

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cell of the trichome widely rounded or spherical. Cells are cylindrical or barrel shaped and more elongated in the center of the trichome. Heterocytes are spherical, hemispherical, or cylindrical, obligatory at the base of the trichome and facultatively also intercalary, with one or two pores. Trichomes disintegrate near the heterocytes. Akinetes, if any, develop basally in a row. Reproduction occurs via hormogonia, which separate from the trichome with necridic cells, and by akinetes. Most species are epiphytic on macrophytes or other algae in tropical alkaline pools and swamps and in moors in temperate zones; some are epilithic in streams and in waterfalls (in limestone regions). Several species (F. bossei, F. monilispora, and F. salinicola) occur in Central America (Komárek, 1984). Species within Fortiea resemble those in Microchaete, but they have widened apical ends. Data from North America need revision. Microchaete Thuret ex Bornet et Flahault (Fig. 24B) Thalli are filamentous, with filaments solitary or in small groups, attached to substrata or forming a thin mat on stones, heteropolar, cylindrical throughout the filament length. Trichomes, with a basal heterocyte, are constricted or unconstricted at the cross walls, cylindrical or slightly tapering toward ends, with meristematic zones at the ends. False branching is infrequent at intercalary heterocytes. Sheaths are distinct, firm, colorless, sometimes lamellated, and form open tubes. Cells are cylindrical or slightly barrel shaped, varying in length/width ratio; end cells are hemispherical or rounded. Heterocytes are ovoid, spherical, or cylindrical, in basal or intercalary position, with one or two pores. Akinetes develop facultatively. Reproduction occurs via hormogonia separating from the apical ends; germination of hormogonia is heteropolar and isopolar. Most of about 20 species are epiphytic on algae, mosses, and plants in swamps and ponds, several are epilithic in streams, and a few occur only in tropical or marine environments (Geitler, 1932; Desikachary, 1959). Tilden (1910) listed four species from Canada, the United States, and the West Indies. Smith (1950) reported three species of Fremyella (synonymous with Microchaete; F. diplosiphon, F. robusta, and F. tenera) in standing waters in the United States; others also reported from various regions across North America (e.g., Whitford and Schumacher, 1969; Stein and Borden, 1979). Komárek (1994) described Microchaete robinsonii from a river in central Canada.

Nostocales: Rivulariaceae Calothrix Agardh ex Bornet et Flahault (Fig. 25A) The thallus is filamentous, attached to substratum basally forming bristle-like groups or thin mats.

Filaments are heteropolar, with a wider basal part (with heterocytes and occasionally an associated akinete and/ or with enlarged basal vegetative cells) and an apical portion forming an elongated, tapering, hairlike form. Heterocytes develop basally; false branching occurs occasionally with the formation of a separated trichome inside of its own sheath. Trichomes are constricted or unconstricted at the cross walls and always taper terminally. Sheaths are always present, firm, in some species lamellated or enlarged at the end, forming funnelformed collars yellow to brownish in color (e.g., in C. fusca) or colorless. Depending on their position in the trichome, cells may be barrel shaped, cylindrical, or narrowly elongated toward the ends (= hairs), especially with nutrient limitation (Whitton, 1987). Aerotopes are absent from vegetative cells but may be present in hormogonia. Meristematic zones are known in several species. Heterocytes are ellipsoidal, spherical to hemispherical, mainly basal, sometimes intercalary (especially near false branches). Akinetes are ellipsoidal to cylindrical, appearing above basal heterocytes and developing from a vegetative portion of the trichome. Reproduction occurs via hormogonia (sometimes with aerotopes), which are released from the end of the filament after the end hair has separated. Calothrix consists primarily of aquatic epilithic and epiphytic species (approximately 60); some species occur in marine littoral zones. Calothrix is taxonomically difficult genus. Tilden (1910) listed 39 species, 36 of which were found in the United States, Greenland, and the West Indies. Three species (C. kawraiskii, C. stagnalis, and C. parietina) were reported from freshwater habitats in Illinois (Tiffany and Britton, 1952). Smith (1950) referred to more than a dozen species growing attached to rocks or woods in flowing and standing waters in the United States. Whitford and Schumacher (1969) recorded 11 species from North Carolina. Calothrix fusca and C. parietina-complex are apparently widespread, whereas C. braunii, C. donnellii, C. ascendens, C. elenkinii, C. epiphytica, C. juliana, C. scytonemicola, C. rivularis, and C. stellaris were recorded less frequently. Whelden (1947) reported several species from the Canadian Arctic: C. borealis (from shallow streams), C. braunii (on rotting Carex leaves), C. contarenii (on rocks in a tidal pool), C. parietina (on stream boulders), and C. pulvinata (marine near the high-tide mark). Many other locations are known in Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). Dichothrix Zanardini ex Bornet et Flahault (Fig. 25B) The thallus is composed of filaments forming bristle-like groups attached to various substrata.


FIGURE 25 (A) Calothrix: (a) trichome; (b) C. tenella (after Gardner, 1927, from Puerto Rico); (c) C. fusca (after Starmach from Kondrateva, 1968); (d) C. simplex (after Gardner, 1927, from Puerto Rico); (e) Calothrix sp. (original photo by J. D. Wehr from the United States). (B) Dichothrix: (a) false-branched thallus; (b) D. willei (after Gardner, 1927, from Puerto Rico); (c) D. orsiniana (after Frémy, 1930); (d) Dichothrix sp. (original microphoto by J. D. Wehr, with permission).

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Filaments are heteropolar with differentiated basal and apical portions and with strong lateral false branching beside the heterocytes. Branches form individual sheaths and diverge from the basal filament in parallel (at least in part) with the old sheath. Trichomes with basal heterocytes are constricted or unconstricted at the cross walls and taper toward hairlike ends. Sheaths are always present, firm, colorless and unlamellated or lamellated and yellow to brownish. Cells are short barrel shaped or short cylindrical; the apical region is composed of long, narrow, hyaline cells. Basal heterocytes are almost spherical to hemispherical, with single pores, occasional intercalary heterocytes with two pores; akinetes are unknown. Meristematic zones occur in several species. Reproduction occurs via hormogonia that separate from the filament by the help of necridic cells, following the separation of a terminal hair. About 30 species have been described from lake periphyton on stones or macrophytes and in rapidly flowing streams (Geitler, 1932; Golubic´, 1967a). Several species are known from the marine littoral zone. Tilden (1910) listed 13 freshwater species widely distributed throughout North America: Dichothrix calcarea, D. hosfordii, and D. inyoensis were described from the United States; other species (D. baueriana, D. compacta, D. gypsophila, and D. orsiniana) were found several times. Some of these species likely have a cosmopolitan distribution (Smith, 1950), but several are reported in various taxonomic concepts. Whelden (1947) noted six species of Dichothrix in a survey of marine and freshwater habitats in the Canadian Arctic: D. baueriana and D. compacta occurred in fast-flowing streams, D. gypsophila in lime-encrusted masses on a rock in a stream and in a freshwater pool, D. hosfordii in freshwater, D. orsiniana in marshy soil, and D. rupicola on rocks in a tidal pool. Villeneuve et al. (2001) reported an interesting species from benthic cyanobacterial mats in an Arctic lake (Ward Hunt Lake, Ellesmere Island); other sites are also known (Stein and Borden, 1979; Sheath and Steinman, 1982). Whitford and Schumacher (1969) reported five periphytic or epiphytic species (D. compacta, D. gypsophila, D. meneghiniana, D. orsiniana, and D. spiralis) and one subaerial species (D. baueriana) from North Carolina. Gloeotrichia Agardh ex Bornet et Flahault (Fig. 26A) Thalli typically occur as mucilaginous, ball-like or hemispherical colonies, enveloped by more or less firm, distinctly limited mucilage. Filaments are heteropolar with basal heterocytes and apical hairs, radially arranged within the colony inside a mucilaginous envelope. Colonies range from microscopic to several centimeters in diameter; they are free floating or attached to substrata, mainly aquatic plants. Sheaths

are always present and distinct, but in some species they are gelatinized and confluent. Trichomes are constricted or unconstricted, tapering toward their ends to a narrow hair, straight, curved, or bent. False branching is rare. Branches grow parallel to the original trichome, reaching its full length, and are radially oriented in rows in colonies with the basal part of the filaments in the center. Cells are barrel shaped or cylindrical to hairlike, blue–green or reddish to dark brownish in color, with or without aerotopes. Heterocytes are spherical to hemispherical or ellipsoid, positioned basally (center of colony) or sometimes intercalary. Akinetes are cylindrical, elongated with rounded ends, adjacent to basal heterocytes, but inside the sheath. Cell division occurs mainly in meristematic zones in the vegetative portion of the trichome. Reproduction occurs via disintegration of trichomes, with hormogonia, which are released after the separation of apical hairs. Of the 16 species described, two are planktonic with aerotopes (clusters of gas vesicles) in the cells, several are epiphytic on submerged water plants in pools, ponds, and littoral zones of lakes; they occur less frequently on stones, especially in mesotrophic and eutrophic waters (Kondrateva, 1968; Barbiero and Welch, 1992). All are exclusively freshwater species except planktonic G. echinulata, which can also occur in slightly brackish waters (eastern Baltic; Pankow, 1976). Gloeotrichia echinulata is a common species in many (mainly eutrophic) lakes and ponds in North America (e.g., Smith, 1950; Whitford and Schumacher, 1969; Duthie and Socha, 1976; Stein and Borden, 1979). Tiffany and Britton (1952) reported only two epiphytic species (G. pisum and G. natans). Smith (1950) mentioned several species, including G. echinulata (= Rivularia planctonica), G. natans, G. pilgeri (the latter from alkaline subtropical and tropical swamps), and G. pisum from localities in the United States. Rivularia (Roth) Agardh ex Bornet et Flahault (Fig. 27) The thallus is composed of filaments arranged in parallel, attached basally to substrata, forming small or large hemispherical colonies (sometimes hollow) when young, later becoming macroscopic (up to several centimeters) layered strata (hemispherical, flat, or irregular) and several millimeters thick; it is gelatinous or encrusted with CaCO3. Filaments are heteropolar, with basally widened ends and apical hairlike sections. Sheaths are firm, mucilaginous, remaining inside the parental sheath after branching, often not reaching the end of the filaments. Trichomes are usually less cylindrical, constricted or unconstricted at the cross walls, with basal heterocytes, separating and branching at intercalary heterocytes, and hairlike narrowed toward the ends. Cells are barrel shaped or cylindrical,


FIGURE 26 (A) Gloeotrichia: (a) planktonic G. echinulata (after Komárek, 1958); (b) G. pisum (after Kondrateva, 1968). (B) Sacconema: (a) S. rupestre (after Smith, 1950, from the United States); (b) S. rupestre (after Borzì, 1882).

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FIGURE 27 Rivularia: (a) trichome disintegration and false branching within colony; (b) Rivularia sp., colonies on stony substrata (original photo by J. D. Wehr, with permission); (c) R. dura (after Smith, 1950; Kondrateva, 1968); (d) R. aquatica (after Kondrateva, 1968, original microphoto by J. D. Wehr, with permission; probably young colony of R. aquatica).


without aerotopes, elongated and narrow at the apex. Heterocytes range from spherical to hemispherical, positioned basally or occasionally intercalary; akinetes are unknown. Meristematic zones are known in several species. Reproduction occurs via hormogonia, which often remain in the upper parts of the colonies, creating upper layers in the strata. Of the 20 species described, most are epilithic or epiphytic, especially on calcareous substrata, occasionally forming travertine. Many species colonize in clear unpolluted standing and running waters, several in marine rocky littoral; some are CaCO3 encrusted. Rivularia is widely reported genus in many floras and ecological studies. Smith (1950) reported several species (R. biasolettiana, R. compacta, R. dura, R. globiceps, R. haematites, and R. minutula) from the United States. Whelden (1947) reported four epiphytic and periphytic species (R. biasolettiana, R. compacta, R. dura, and R. minutula with a possible new variety) from High Arctic freshwater habitats, whereas R. biasolettiana and R. globiceps were also recorded on submersed freshwater substrata in North Carolina (Whitford and Schumacher, 1969). Tilden (1910) reported 19 species (five uncertain) distributed throughout the United States and in the West Indies; several species were reported from Ontario, British Columbia, and the Northwest Territories (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). Several species are known from marine habitats. The calcified R. dura is mainly distributed in tropical and subtropical limestone regions, but Weldon (1947) also reported this species from the eastern Arctic (revision is needed). Sacconema Borzì (Fig. 26B) The thallus is composed of radially arranged or creeping filaments. Filaments are heteropolar, irregularly falsely branched, attached at the base to substrata or creeping among detritus and other algae, forming loose shrublike colonies. Trichomes are solitary or in pairs within a single sheath, uniseriate, heteropolar, constricted at the cross walls, with basal heterocytes. In the apical portion, cells are attenuated into elongated hair cells, which are cylindrical and hyaline. Sheaths are very thick, swollen, and widened at the apex (up to 10⫻ broader than the trichome width). The sheath is initially closed, but later opened at the apex, intensely lamellated, funnel-like, and yellowish brown to brown in color. Heterocytes are basal, mainly spherical, but not well known. Cell division proceeds perpendicularly to the longitudinal axis of the trichome; meristematic zones are not well known. Reproduction most likely occurs via hormogonia. Sacconema is most commonly subaerial on rock wall seepages and in rocky littoral communities. Tilden

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(1910) reported one species from Massachusetts; S. rupestris was reported from Kootenay Lake, British Columbia (Stein and Borden, 1979). The genus is rarely reported in the literature but has recently been observed in rock scrapings from the splash zones in Lake of the Woods, Ontario (Komárek, Kling, and Komárková, unpublished observations). Sacconema is likely present in more locations in North America; reports will probably increase with increasing studies of the algal flora of the littoral zones of northern lakes.

Nostocales: Nostocaceae Anabaena Bory ex Bornet et Flahault (Fig. 28) The thallus, or with solitary filaments, or is arranged in free clusters, in macroscopic mats, or in the tissues of aquatic plants. Trichomes are straight, curved, or regularly coiled, rarely in bundles with parallel-oriented filaments. Sheaths are never firm, in form of mucilaginous, hyaline and colorless, diffuse envelopes, occurring only in some species. Trichomes are constricted or unconstricted at the cross walls, uniseriate, isopolar (both poles with the same morphology). Cells are spherical, ellipsoidal, short or long cylindrical, sometimes bent (reniform), pale to bright blue–green or yellow–green; the planktonic species have aerotopes (subgenus Dolichospermum), occasionally with granular contents. Heterocytes are intercalary, solitary, at fairly regular intervals along filament (metameric), up to nine (rarely more) per filament. Akinetes are spherical, ellipsoidal, cylindrical, curved, intercalary, solitary or in groups of two to five, sometimes with a yellowishcolored epispore, in some species adjacent to heterocytes. Cells divide perpendicularly to trichome axis; meristematic zones are not noted. Reproduction occurs via trichome fragmentation and akinete production. Anabaena is a common genus worldwide; about 110 species have been described, a large part of them planktonic (subgenus Dolichospermum). Some form dense surface blooms and some are recorded as toxic (Carmichael, 1992; Skulberg et al., 1993; Codd, 1995; Chorus and Bartram, 1999). Species without aerotopes (clusters of gas vesicles) form mats in the littoral zone or cover sediments or aquatic plants in freshwater pools and ponds and in saline lakes. North American records include both benthic and planktonic species. Duthie and Socha (1976), Stein and Borden (1979), and Sheath and Steinman (1982) recorded about 18 or 19 taxa in the Canadian flora from Ontario, British Columbia, and the Northwest Territories, respectively. Tiffany and Britton (1952) reported nine species of Anabaena from Illinois. Smith (1950) mentioned 36 species in the United States, some of which produce water blooms. Whitford and Schumacher (1969)

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FIGURE 28 Anabaena: (a) origin of metameric trichomes in Anabaena (development of proheterocytes and heterocytes in regular distances); (b) planktonic A. perturbata (after Nygaard, 1949; original microphoto by Komárková); (c) planktonic A. viguieri (after Nygaard, 1949; original microphoto by Komárková); (d) benthic A. oblonga (after Komárek, 1989).


referred to 14 species both with and without aerotopes in North Carolina. Hill (1976a–c) described three new planktonic species from Minnesota lakes. Tilden (1910) listed 15 species (four “not well understood”) as widely distributed throughout North America. Several interesting planktonic species from the United States (e.g., A. mendotae, and A. perturbata, which are common in the temperate zone) were described by Trelease (1889) and Hill (1976a, b, c). Several nonplanktonic Anabaena species were recently transferred into the genus Trichormus (see Komárek and Anagnostidis, 1989, and the Trichormus description in this chapter), which differs from Anabaena substantially by mode of akinete formation. Anabaenopsis (Wo l⁄ oszyn´ska) Miller (Fig. 29A) The thallus is filamentous. Filaments are solitary or in free clusters, free floating, with or without constrictions at the cross walls; they are frequently bent, spirally coiled, or sigmoid, rarely straight. They lack sheaths, but have a mucilaginous, diffuse envelope. Trichomes have a characteristic pattern of metameric (regularly repeating) origin of pairs of heterocytes in the trichome: Trichomes break between mature heterocytes leaving filaments with “apical” heterocytes at both ends. Several pairs of young heterocytes can be situated at regular intervals over a filament. Cells are almost spherical, barrel shaped to cylindrical, bent or straight, pale blue–green to brownish, with facultative aerotopes. Heterocytes are spherical to hemispherical, intercalarly in pairs after asymmetrical division of two neighboring vegetative cells; they may be smaller or larger than the vegetative cells. Akinetes are spherical or ellipsoidal, bent or straight, solitary or several together, intercalary, often distant from heterocytes. All cells in the trichome are capable of division; division is asymmetric before heterocyte formation. Reproduction occurs via akinete production and trichome fragmentation. Most of the 16 species described are planktonic, forming water blooms. The spirally coiled filaments occur in mesotrophic to eutrophic lakes or in alkaline and saline waters, sometimes with epizooic protozoans (e.g., Vorticella sp.; Canter-Lund and Lund, 1995). Few species live in the metaphyton of small temperate water bodies in tropical and subtropical regions. Several species were reported from Midwestern or higher conductivity U.S. reservoirs (Tilden, 1910; Smith, 1950; Whitford and Schumacher, 1969). This genus is rare in low-conductivity, low-nutrient boreal waters and apparently absent from arctic and subarctic waters (Duthie and Socha, 1976; Sheath and Steinman, 1982). However, species of the genus were reported in prairie lakes in central Canada (Kling, 1975).

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Aphanizomenon Morren ex Bornet et Flahault (Fig. 29B) The thallus is filamentous. Filaments are free floating, solitary, or gathered in small or large fasciculated colonies with filaments in characteristically parallel arrangements (may appear as macroscopic flakes or like grass clippings on lake surface). Trichomes are isopolar, subsymmetrical, straight, sometimes with slightly curved or bent ends, cylindrical or narrowing toward the ends, with or without constrictions at the cross walls. Firm sheaths are absent, but the trichomes of several species are covered with fine, diffuse, mucilaginous, colorless envelopes. Cells are cylindrical or long barrel shaped, with quite variable width/length ratio, pale blue–green, with aerotopes (sometimes facultative). The end cells are often much longer than the central cells, and hyaline. In some species, they are cylindrical and rounded at the end without visible cell contents; in others, narrowed and pointed. Heterocytes are almost spherical, ellipsoidal, or cylindrical with two pores, always intercalary and solitary, usually only few (1–3) per trichome (rarely more). Akinetes are long or short cylindrical with rounded ends, or elliptical, rarely almost spherical, solitary or groups of two or three, adjacent to heterocytes or distant. Cell division occurs along the whole trichome with the exception of the end cells. Reproduction occurs via trichome disintegration and akinete germination. About 20 species have been described, but some are not clearly defined; the position of species with narrowed and pointed apical cells is unresolved. Almost all are planktonic, often forming dense water blooms in temperate zones (Aphanizomenon flos-aquae in eutrophic lakes, reservoirs, regulated fish ponds, and cattle ponds). One species prefers brackish conditions or seawater (Baltic Sea). Few species are known only from tropical regions (Cronberg and Komárek, in press). The most noted species in North America are A. flos-aquae and A. gracile, although several other species have been reported (A. issatschenkoi, A. skujae, and A. aphanizomenoides). One species, A. schindleri, has only been identified from mesotrophic to eutrophic Canadian Shield lakes in northwestern Ontario (Kling et al., 1994). Early authors (e.g., Tilden, 1910; Smith, 1950; Prescott, 1962), as well as Whitford and Schumacher (1969) and others (e.g., Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982) mentioned only one species, A. flosaquae, which is widely distributed throughout North America. Aulosira Kirchner ex Bornet et Flahault (Fig. 31C) The thallus is filamentous. Filaments are solitary or grouped in clusters, rarely in mats. Filaments are slightly

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FIGURE 29 (A) Anabaenopsis: (a) origin of metameric trichomes in Anabaenopsis (development of two neighboring heterocytes in regular distances); (b) A. elenkinii (after Smith, 1950). (B) Aphanizomenon: (a) subsymmetrical trichomes; (b) A. flos-aquae (after Komárek, 1958; original microphoto by J. D. Wehr, with permission); (c) A. schindleri (after Kling et al., 1994, from central Canadian lakes).


FIGURE 30 (A) Cylindrospermopsis: (a) development of terminal heterocytes; (b) C. raciborskii (after Horecká and Komárek, 1979). (B) Cylindrospermum: (a) development of symmetrical trichomes; (b) C. stagnale (bar = 20 µm; after Komárek, 1975); (c) C. longisporum (bar = 10 µm; after Komárek, 1975); (d) C. minutissimum (after Komárek, 1989, from Cuba).

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irregularly coiled, sometimes oriented in parallel rows, with firm, distinct colorless sheaths enveloping one or occasionally two trichomes (young trichomes have no sheath), open at the end. Trichomes are of uniform width throughout their length, not attenuated at ends, with constrictions at the cross walls, uniseriate, isopolar, metameric (regularly repeating pattern), with several solitary heterocytes regularly arranged along trichome. Cells are cylindrical or barrel shaped, typically isodiametric, but becoming longer than wide with age, blue–green, pale grayish–blue, olive green, or reddish, occasionally (rarely) with aerotopes, often with prominent granules. Terminal cells are rounded. Heterocytes are spherical, oval or cylindrical, generally same width or slightly wider than vegetative cells. Akinetes are usually elongated, oval to cylindrical (rarely spherical), apoheterocytic (arise in rows between two heterocytes and develop successively toward a heterocyte), or sometimes irregularly situated. Cell division occurs perpendicularly to the trichome axis, with cells growing to the original size and shape before subsequent divisions; all cells are capable of division, with no meristematic zones. Reproduction is by hormogonia emerging from sheaths. The species of the genus appear benthic or in the metaphyton of unpolluted lakes, ponds, and reservoirs, generally among or on submersed water plants or on sticks. Most species are geographically delimited, many to tropical regions (Frémy, 1930b; Geitler, 1932; Desikachary, 1959). Several species are important members of the microflora of rice paddies (e.g., A. fertilissima). Twenty species have been described but only six are well known. Smith (1920) noted the genus in widely separate localities, also reported from Wisconsin and British Columbia (Prescott, 1962; Stein and Borden, 1979). It is most likely present in tropical and subtropical regions as well (marshes in central America; authors’ data). Cylindrospermopsis Seenaya et Subba Raju (Fig. 30A) The thallus is filamentous. Filaments are solitary, straight, bent, or coiled, free floating, narrowed toward the ends (in some species). Trichomes are isopolar or secondary heteropolar, subsymmetrical, with or without constrictions at the cross walls. Cells are cylindrical or barrel shaped, pale blue–green or yellowish, facultatively with aerotopes. End cells are often conical or bluntly or sharply pointed. Heterocytes are ovoid or conical, sometimes slightly curved in a droplike form, single pored, and always terminal; they develop after asymmetrical division of the end cells in one or in both ends of the filament. Akinetes are ellipsoidal or cylindrical, developing among vegetative cells, distant, or occasionally adjacent to apical heterocytes. Reproduction occurs via akinetes and trichome fragmentation.

The nine species described are planktonic, bloom forming, and common in tropical eutrophic freshwaters (pantropical) (Komárková, 1998; Komárek, 2001). The common C. raciborskii has been shown to be invasive into the warm regions of the temperate zone in Europe (Horecká and Komárek, 1979; Padisák, 1990, 1997; Dokulil and Mayer, 1996). C. raciborskii is a wellknown species forming water blooms in tropical and subtropical water bodies (Komárková et al., 1999) and producing toxins (Skulberg et al., 1993). Occurrences of members of this genus apparently have increased in south-temperate and subtropical lakes (e.g., Florida ) in the last few years (authors’ data). However, Hill (1970a) lists an Anabaenopsis (= Cylindrospermopsis) raciborskii from southern Minnesota. Also his Raphidiopsis mediterranea (Hill, 1970b) could be a part of the developmental stage of C. raciborskii with only akinetes. A new species (C. catemaco) has been described from Mexico (Komárková-Legnerová and Tavera, 1996). Cylindrospermum Kützing ex Bornet et Flahault (Fig. 30B) The thallus is composed of fine or compact mucilaginous mats. Filaments are slightly curved or irregularly coiled, typically cylindrical throughout their length. Trichomes are without a firm sheath, but with a fine, colorless, homogeneous, and diffuse mucilaginous envelope. Trichomes are symmetrical, usually slightly constricted at the cell walls. Cells are cylindrical, more or less isodiametric, pale or bright blue–green or grayish, without aerotopes, occasionally granulated. Heterocytes are always terminal, developing from the end cells, ellipsoidal, ovoid, or conical, single pored, situated at one or both ends of the filaments. Akinetes are ellipsoidal, rarely spherical, developing near both ends of the filament at heterocytes, solitary or in rows up to seven, often with sculptured epispore. Cell division occurs throughout entire trichome; there areno meristematic zones. Reproduction occurs via akinete germination and fragmentation of trichomes into hormogonia. Most of the 50 species described are benthic, epiphytic, or epilithic in unpolluted or slightly eutrophic waters, or from soils. Tilden (1910) noted eight common species, whereas Smith (1950) reported 11 species from terrestrial and aquatic habitats in the United States. Whitford and Schumacher (1969) distinguished 11 species based on differences in akinete shape. One aquatic species, C. catenatum, occurred in waters across North Carolina; others were subaerial. Tiffany and Britton (1952) reported three species of this genus from Illinois. A few species have been reported in scattered locations across Canada (Duthie and Socha,


FIGURE 31 (A) Raphidiopsis: (a) R. curvata (after Fritsch and Rich from Geitler, 1932). (B) Wollea: (a) W. saccata, macroscopic colonies and solitary trichomes (after Wolle from Geitler, 1932, and Smith, 1950). (C) Aulosira: (a) Aulosira sp. (original photo by Komárková, from the Everglades, Florida); (b) A. implexa (after Geitler, 1932); (c) A. laxa (after Kondrateva, 1968).

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FIGURE 32 (A) Trichormus: (a) apoheterocytic akinete development between two heterocytes; (b) T. variabilis (after Frémy ex Geitler, 1932; sub Anabaena variabilis); (c) T. luteus (after Gardner, 1927, from Puerto Rico; sub Anabaena lutea); (d) T. fertilissimus (after Desikachary, 1959; sub Anabaena fertilissima); (e) T. subtropicus (after Gardner, 1927, from Puerto Rico; sub Anabaena subtropica). (B) Isocystis: (a) I. planctonica (after Starmach, 1966); (b) I. infusionum (after Hollerbach et al., 1953).


1976; Stein and Borden, 1979; Sheath and Steinman, 1982). Several interesting species are known from swampy habitats in the Caribbean (Komárek, 1989). Isocystis Borzì ex Bornet et Flahault (Fig. 32B) Isocystis is filamentous, growing in solitary filaments, in small macroscopic fascicles, or in indistinct microscopic gelatinous colonies attached to substrata. Filaments are enveloped by a fine, diffuse, unstructured, colorless, barely visible gelatinous envelope. Trichomes are uniseriate, simple curved, or coiled, isopolar, generally slightly attenuated toward both ends, and clearly constricted at the cross walls. The gelatinous sheaths are fine, diffuse, unstructured, colorless, and barely visible. Cells are barrel shaped, isodiametric, longer or shorter than wide, sometimes with solitary granules. Heterocytes are absent. Akinetes have thick cell walls, are slightly larger than vegetative cells, and are situated in apoheterocytic fashion (in rows in the middle of trichomes). All cells are capable of cell division, perpendicular to the trichome axis. Reproduction occurs via short hormogonia, which sometimes disintegrate into solitary cells, and akinete germination. Species of Isocystis appear in the metaphyton or are free floating. Several species are periphytic in different freshwater habitats (springs and littoral areas of standing waters), one species is subaerial, colonizing wet walls, one is planktonic (Starmach, 1966), and one was described from soil (Geitler, 1932). Eight species have been described, of which five have been accepted as valid (Starmach, 1966; Komárek and Anagnostidis, 1989). The genus is not well known and probably rare (but present) in North America (not yet published). Nodularia Martens ex Bornet et Flahault (Fig. 33) This is a filamentous genus occurring as solitary filaments or in groups or clusters, occasionally (several benthic species) in mats. Filaments are isopolar (both poles with the same morphology), unbranched, more or less straight, curved, coiled, or irregularly spirally coiled with fine, diffuse, two-layered sheaths, open at both ends. Trichomes are uniseriate, cylindrical, occasionally (rarely) with slightly attenuated trichome ends, constricted at the cross walls, metameric (several heterocytes situated at regular intervals). Cells are short, barrel shaped, with length never exceeding width. Two groups of species (subgenera) differ cytologically and ecologically from each other; facultative gas vesicles occur only in planktonic species. Cell contents are yellowish, pale olive green, blue–green, or pinkish; thylakoids are irregularly coiled and distributed throughout the cell, sometimes peripheral. Heterocytes are typically identical in shape to vegetative cells. Akinetes are short, barrel shaped (shorter than wide) or

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spherical, but often with irregularities. All cells are capable of division, transverse to the trichome axis; trichomes lack meristematic zones. Reproduction occurs via hormogonia, disintegration of trichomes, and akinete germination. After revision of the genus, only 9 of the 24 species (27 taxa) described were well defined (Nordin and Stein, 1980; Komárek et al., 1993; Hayes et al., 1997; Barker et al., 2000). Four species with aerotopes (groups of gas vesicles) are planktonic (including type species N. spumigena), but other planktonic taxa occur in localities worldwide, including North and Central America. Species in this group sometimes form dense water blooms in inland lakes or reservoirs with higher salinity, in brackish coastal lakes and lagoons, or in the sea or estuaries of rivers (Huber, 1984; Blackburn and Jones, 1995; Tavera and Komárek, 1996; Perez et al., 1999). Five species always lack aerotopes and are benthic; the most common, N. harveyana (cosmopolitan?), occurs in coastal or inland saline and mineral pools, as well as in marshes and lakes. N. sphaerocarpa (temperate) occurs in alkaline streams and in the littoral areas of lakes and pools, and N. willei (pantropical) occurs in rice fields, supplying an important source of nitrogen; it is highly probable that several species also colonize soils. Blooms of Nodularia have been noted in lakes in western North America, as well as in saline lakes from North American prairies (Nordin and Stein, 1980) and in Nevada (Galat et al., 1990). Tilden (1910) recorded six species of the genus Nodularia in North America, Tiffany and Britton (1952) reported two species, and Smith (1950) mentioned all three species recognized prior to 1950 in the United States. Prescott (1962) reported two species from the western Great Lakes area, and several species have been reported from across Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). Nostoc Vaucher ex Bornet et Flahault (Fig. 34) The thallus is microscopic or macroscopic, gelatinous, spherical, or irregular gelatinous mats or flat colonies, smooth or warty at the surface, often with superficial mucilaginous exterior. Filaments are typically coiled, forming irregular, loose, or dense clusters, concentrated near the colony surface. The sheath is mucilaginous, firm, wide, and sometimes yellow to brownish, but visible only in young colonies and confluent with common mucilage. The mucilage of the colony is sometimes colored, yellowish green or brownish. Trichomes are isopolar, sometimes very long, curled inside the colony, often moniliform. Cells are barrel shaped or spherical, a uniform shape and size along trichome, pale to bright blue–green or olive green. Heterocytes are barrel shaped or spherical, soli-

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FIGURE 33 Nodularia: (a–d) N. cf. spumigena (after Pérez et al., 1999, from coastal lakes in Uruguay); (e) N. litorea (after Bornet and Thuret from Geitler, 1932); (f) N. baltica (after Smith, 1950, from the United States, sub N. spumigena var. minor); (g) N. harveyana (after Geitler, 1932); (h) N. willei (after Gardner, 1927, from Puerto Rico).


FIGURE 34 Nostoc: (a) Nostoc sp., typical initial stages (after Smith, 1950; original photo by R. G. Sheath, with permission); (b) typical position of trichomes in old colony (after Smith, 1950); (c) N. paludosum (after Komárek, 1975); (d) N. edaphicum (after Kondrateva, 1968); (e) N. commune (after Kondrateva, 1968).

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tary, developing at ends of trichomes or intercalary. Akinetes are ellipsoidal, only slightly larger than vegetative cells, arising in rows between heterocytes. Cells divide perpendicularly to the trichome axis; meristematic zones are unknown. Colony morphology changes during development, and reproduction is specific for each subgenus; motile hormogonia develop between heterocytes, by akinete germination or filament disintegration. Nostoc is common and widespread genus with more than 200 taxa described; a few species have been precisely revised (Mollenhauer, 1970). Species of Nostoc are mainly benthic, occurring in many epiphytic, epipelic, and epilithic habitats, in unpolluted lakes, ponds and pools, streams, and rivers, and on soils (including desert soils), some reaching 30 cm in diameter (Dodds et al., 1995). N. parmelioides, with spherical colonies, grows on stones in streams and rivers and produces ear-shaped colonies after being occupied by aquatic midge larvae (Ward et al., 1985; Dodds, 1989). A few Nostoc species are endophytic in fungi (Geosiphon), mosses, and ferns (Mollenhauer et al., 1996; Mollenhauer and Mollenhauer, 1996), and phycobionts in lichens. At the time of Tilden (1910), the genus Nostoc comprised 31 species, 29 of which were found in North America. Whelden (1947) recorded several taxonomically uncertain species from the Arctic: Nostoc aureum (freshwater), N. commune (soils or tundra seepages), N. kihlmanii (floating in freshwater), N. linckia (in a drying pool), N. minutum (lake edges), N. paludosum (in marshes), N. pruniforme (floating in lakes), N. sphaericum (small pools and on rotting leaves in freshwater), and N. sphaeroides (among mosses). Smith (1950) recorded some species as mostly terrestrial, others as strictly aquatic, and still others as free floating in pools or attached to submersed substrata. Prescott (1962) recorded 14 species of the genus Nostoc from the western Great Lakes area; several species are widely reported across Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982). Raphidiopsis Fritsch et Rich (Fig. 31A) The thallus is filamentous. Trichomes are solitary, free floating, without sheaths and mucilaginous envelopes, straight or screwlike coiled, unconstricted or slightly constricted at the cell walls. Cells are usually cylindrical, facultatively with aerotopes and granulations. Apical cells are conical, pointed, or bluntly pointed at the ends of the trichomes. Heterocytes are unknown. Akinetes are cylindrical to elongated ellipsoidal, intercalary, usually slightly off center in the trichomes. Reproduction occurs by trichome fragmentation and akinete germination.

The species are mainly planktonic. Hill (1970b) reported R. mediterranea from Minnesota, and the genus has been collected several times in Mexico (Komárková-Legnerová and Tavera, 1996). Smith (1950) and Taft and Taft (1970) mentioned only one species, R. curvata from Lake Erie. Whitford and Schumacher (1969) reported the same species in North Carolina, which has also been collected from standing and flowing water (irrigation ditches) in British Columbia (Stein, 1975; Stein and Borden, 1979). Trichormus (Ralfs ex Bornet et Flahault) Komárek et Anagnostidis (Fig. 32A) Thalli occur as macroscopic mats, in clusters and in strata; filaments are rarely solitary. Trichomes are uniseriate, straight, bent, or coiled, and constricted at the cross walls. Firm sheaths do not develop, but slimy envelopes are sometimes present. Cells are barrel shaped to cylindrical, pale blue–green, without aerotopes. Heterocytes are spherical to ellipsoidal, intercalary, in metameric (regularly repeating) configuration. Akinetes are oval to barrel shaped, rarely almost cylindrical, and develop in rows apoheterocytically between two heterocytes. One vegetative cell becomes one akinete. Reproduction occurs via hormogonia that separate from trichomes and via akinete germination. Thirty-two species have been described (mainly formerly as Anabaena, which differ substantially by the strategy of akinete formation; Komárek and Anagnostidis, 1989). The majority of species colonize soils (often in rice fields) or mud; some are periphytic in the littoral of different waterbodies or live endophytic (known as “Anabaena azollae,” “Anabaena cycadearum,” and “Nostoc gunnerae”). Several species are tropical (e.g., T. anomalus, T. doliolum, and T. fertilissimus). Trichormus has been revised recently (Komárek and Anagnostidis, 1989) and thus may not be noted as such by earlier authors of North American studies. However, several species, listed as species of Anabaena, and recently recombined into Trichormus, are known from North America (e.g., Trichormus [Anabaena] azollae from Ontario; Duthie and Socha, 1976), and including the species Anabaena = Trichormus variabilis from soil habitats, reported from the United States by Whelden (1947), but from atypical localities. Several species living in intercellular spaces of vascular plants are also known from North and Central America: T. (Anabaena) azollae is a common symbiont of the aquatic fern Azolla (our results) and another species colonizes the roots of cycads (Mercado, 1977). Wollea Bornet et Flahault (Fig. 31B) Thalli are gelatinous, macroscopic colonies, with smooth surface, forming 5- to 10-cm tubelike (saccate)


colonies or irregular balls, closed at apex, initially attached to a substrate and later free floating. Filaments are typically cylindrical. Trichomes are uniform along their length, more or less straight or slightly curved, uniseriate, unbranched, not attenuated or widened at the ends, highly constricted at the cross walls, with rounded end cells. Trichomes are irregular or almost parallel within a common diffuse mucilage. A fine pelicula (slimy but firm mucilage) is apparent on the surface of colonies, but sheaths around the trichomes are absent. Heterocytes are intercalary and solitary, and their position results in a metameric structure (regularly repeating pattern) along the trichomes. Spherical or oval akinetes arise in a short series, at both sides of the heterocytes. Cell division occurs by perpendicular binary fission. Reproduction is by hormogonia. Wollea colonizes benthic habitats in freshwaters. The type species, W. saccata, is infrequently reported but known from several localities (mainly standing waters) in North America, including the Great Lakes region, Massachusetts, Minnesota, New Jersey, North Dakota, Louisiana, North Carolina, British Columbia, and Panama (Tilden, 1910; Prescott, 1962; Whitford and Schumacher, 1969; Stein, 1975). A second species (W. bharadwajae) was described from benthic habitats of ponds and paddy rice fields in India. Only these two species are known.

Stigonematales: Capsosiraceae Besides the genus Stauromatonema, Smith (1950), Tilden (1910), and Whitford and Schumacher (1969) also mentioned a subaerophytic genus Capsosira (C. brebissonii) from New England and North Carolina. However, the original drawings do not correspond to the characteristics described by Frémy (Geitler, 1932), and North American records must be revised (see Fig. 35A). Stauromatonema Frémy (Fig. 35B) The thallus is firm and flat; the encrusting layer is composed primarily of basal creeping and coiled filaments and erect branches. Basal trichomes become multiseriate, sometimes disintegrating into coccoid-like masses from which grow secondary trichomes. Branches (true branching) are densely arranged and parallel, situated perpendicularly to substrata, uniseriate, and producing repeated V branches in successive layers. Sheaths are thin, firm, colorless, and closed at the apex of young filaments; they sometimes gelatinize and become slightly lamellated. Trichomes, constricted at the cell walls, are more or less cylindrical or slightly widened toward the ends, with rounded terminal cells. Cells are irregularly disclike or bluntly barrel shaped. Heterocytes are intercalary and barrel shaped. Akinetes

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are unknown. Cells divide in different planes in basal trichomes and in pseudoparenchymatous masses, but divide perpendicularly to the long axis in erect branches. Reproduction occurs via the liberation of monocytes (solitary reproduction cells) from the ends of branches after the sheaths are opened (rarely happens in intercalary cells). Hormogonia and hormocysts are unknown. Thalli colonize stones in springs, creeks, and waterfalls, and rarely, littoral in lakes. Three species are tropical. S. viride has been found on the stones in an outlet from Lake Catemaco, Veracruz, Mexico (Carmona-Jimenéz and Gold-Morgan, 1994).

Stigonematales: Stigonemataceae Stigonema Agardh ex Bornet et Flahault (Fig. 36) The thallus is filamentous, wooly or crusty, composed of free, coiled, true branched filaments, which are not diversified into morphologically different basal filaments and branches. Thalli are usually attached to substrata. Trichomes may be uniseriate or multiseriate (uniseriate mainly in young trichomes or ends of branches). Branches are thick, irregularly lateral, narrowed toward the ends, but with the apical cells often larger than others. Sheaths are thin or thick with a clear outer margin becoming wider with age, lamellated, and usually yellowish brown. Individual envelopes may develop around cells in the older parts of filaments. Trichomes disintegrate often in separated cells within the filaments. Cells are barrel shaped or irregularly rounded, usually connected by one pore (“pit connections”) with one another (pores disappear in segments of some trichomes). The cell content is blue–green or olive green, usually with prominent solitary granules. Heterocytes are intercalary, solitary, occasionally lateral, and similar in form to adjacent vegetative cells. Akinetes are unknown. Clusters of coccoid cells occur occasionally. Cell division occurs in all planes; horizontal (crosswise) fission is most common, and meristematic zones are only present in sections where hormogonia arise. Reproduction occurs via hormogonia liberated from the ends of trichomes and branches. Hormogonia are uniseriate, morphologically different from trichomes, usually more cylindrical, and consist of two to many cells, sometimes with aerotopes. Species of Stigonema attach to the substrata in standing and flowing waters or are subaerial or occur on soils (not common); they are distributed worldwide but apparently are more common in tropical regions (Gardner, 1927; Frémy, 1930b; Desikachary, 1959). Subaerial species colonize tree bark and rocks from lowlands to alpine regions (the common epilithic S. minutum, and S. informe from alpine wetted rocks are most known). Other species are known from the

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FIGURE 35 (A) Capsosira: (a) C. brebissonii (after Smith, 1950, from the United States); (b) C. brebissonii (after Frémy, 1930). (B) Stauromatonema: (a–b) S. viride (bar = 50 µm; after Carmona-Jimenez and GoldMorgan, 1994, from Mexico); (c) S. viride (after Geitler and Ruttner, 1935).


FIGURE 36 Stigonema: (a) thallus; (b) S. congestum (after Gardner, 1927, from Puerto Rico); (c) S. elegans (after Gardner, 1927, from Puerto Rico); (d) S. mamillosum (after Frémy and Kosinskaja from Kondrateva, 1968).

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metaphyton or attached to wood and stones in pools, swamps, and wetlands. In tropical habitats, several species are important soil organisms (Desikachary, 1959). In North America and northern Europe, species can be found attached to rocks in oligotrophic lakes and mountain streams. There are many reports of this genus throughout North America, including Tilden (1910), who recorded nine Stigonema species of the ten described at that time in North America. Several species have been noted from the Arctic (Whelden, 1947): Stigonema informe (small freshwater pools and subaerial on wet granitic rocks), S. mamillosum (boulders in swift flowing streams), S. minutum (seepage rocks), S. ocellatum (stream), and S. turfaceum (peaty brook, tundra pool). In addition, Smith (1950) recorded S. hormoides, S. panniforme, and S. thermale. Prescott (1962) reported five species from the western Great Lakes area, whereas Whitford and Schumacher (1969) recorded 11 species in North Carolina; S. hormoides was the most common species in aquatic habitats with S. ocellatum. S. mesentericum, S. minutum var. saxicola, and S. mirabile were found on dripping rocks. At least nine taxa have been recorded from across Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982).

Stigonematales: Fischerellaceae Fischerella (Bornet et Flahault) Gomont (Fig. 37A) The thallus is composed of morphologically diverse, uniseriate or multiseriate, usually creeping filaments and more or less erect uniseriate branches. Most species produce feltlike, rarely compact mats. Creeping trichomes with barrel-shaped cells sometimes occur within a gelatinous matrix. Trichomes are generally moniliform and enveloped by thick, sometimes from outside wavy or slightly lamellated, colored sheaths. Erect true branches (with T-type branching), usually unilateral, arise after longitudinal cell division in basal trichomes. Branches, primarily cylindrical, are generally composed of cylindrical elongated cells in colorless sheaths. Cell contents are slightly granular, with thylakoids distributed irregularly. Heterocytes are intercalary, subspherical in basal trichomes, and cylindrical in branches. Akinetes (known only in a few species) occur occasionally and irregularly in basal trichomes. Cell division occurs mainly via horizontal (crosswise), perpendicular fission. Reproduction occurs via uniseriate hormogonia separating from the ends of branches. Hormogonia liberated under wet or humid conditions, and are usually morphologically distinct from other cells in branches and contain aerotopes (gas vacuoles). Most species are subaerial, growing on wet (often acidic, peaty) soils and rocks. Several species occur only in the tropical forests on mosses and the bark of

trees. Two species, F. ambigua and F. thermalis, are reported in Tilden (1910). Fischerella ambigua is known from the United States, Canada, Mexico, the West Indies, and Hawaii as intermingled with large algae on moist rocks, wet earth, or the trees. F. thermalis is known from New Hampshire on damp stones and granite rocks and from thermal water in Hawaii and British Columbia (e.g., Stein and Borden, 1979). Prescott (1962) reported two species from the western Great Lakes region, and three were recorded from British Columbia (Stein and Borden, 1979). Whitford and Schumacher (1969) observed six species among the material collected from North Carolina, and three of these (F. ambigua, F. letestui, and F. major) were found on submerged substrata whereas the others were from subaerial habitats.

Stigonematales: Borzinemataceae Schmidleinema De Toni (Fig. 37B) Thalli consist of mats on various substrata or mixed with other algae, composed of basal prostrate creeping filaments and more or less erect branches. Basal creeping trichomes are torulous (thickened slightly), uniseriate or multiseriate, sometimes with true branching. Branches are cylindrical, uniseriate, erect, and usually repeatedly falsely branched (like Tolypothrix). Sheaths are usually (in parts) yellow–brown, distinctly outlined, thick and lamellated, closed or funnel-like or sometimes with a telescopic opening at the apex. Cells are barrel shaped or nearly cylindrical, pale blue–green to olive green, slightly or generally granular, sometimes with solitary distinct granules. Intercalary heterocytes are solitary or in pairs, usually single pored (prior to false branching), cylindrical, hemispherical or almost spherical. Akinetes are absent. Cell division occurs in several planes in older creeping filaments, perpendicular to the longitudinal axis in young trichomes and in branches; meristematic zones occur in the apical parts of branches. Reproduction occurs via motile hormogonia formed from the ends of branches and ensheathed hormocysts, differentiated in the basal or apical parts of the thallus. Hormogonia and hormocysts germinate on both sides of the heterocytes that develop in their center, becoming basal during later development. Schmidleinema occurs in freshwater habitats, mainly tropical to subtropical, or subaerial. The type species, Schmidleinema indicum, is known from humid subaerial habitats such as on liverworts, tree trunks, and humid walls in India. Two other taxa (S. cubanum and S. roberti-lamyi) are known from alkaline swamps in the Antilles (Cuba and Guadeloupe; Komárek, 1989); S. cubanum is also known from the Everglades, Florida, and Belize.


FIGURE 37 (A) Fischerella: (a) F. thermalis (after Frémy from Geitler, 1932); (b) F. ambigua (after Frémy from Geitler, 1932). (B) Schmidleinema; (a) thallus; (b) S. cubanum (after Komárek, 1989, from Cuba).

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FIGURE 38 (A) Geitleria: (a–d) G. calcarea (after Friedmann, 1955, 1979; Couté, 1985); (B) Albrightia: (a) A. tortuosa (after Copeland, 1936 from Yellowstone National Park). (C) Colteronema: (a) C. funebre (after Copeland, 1936, from Yellowstone National Park).


Stigonematales: Loriellaceae Albrightia Copeland (Fig. 38B) The thallus consists of small clusters of free filaments. Filaments and branches are of the same morphology, cylindrical, single or true branched, flexuous, and irregularly and loosely tangled. Sheaths are firm, thick, homogeneous or layered. Trichomes are uniseriate and cylindrical and composed of moniliform rows of cells with deep constrictions at the cross walls. Growth is terminal, with branches of the same width and morphology as the main filaments and trichomes. True branching is either T type or pseudodichotomous V type. Cells are barrel shaped to cylindrical, narrowed at both ends, usually longer than wide, blue–green in color, with a granular content and sometimes prominent solitary granules. Terminal cells are more cylindrical and slightly elongated; end cells are rounded. Coccoid cells (two-celled groups) sometimes appear. Heterocytes and akinetes are unknown. Cell division is by horizontal (crosswise) or perpendicular fission. Reproduction occurs via few-celled hormogonia differentiated terminally at the ends of branches. Filaments disintegrate into persisting chroococcoid groups. The type species Albrightia tortuosa is the only member of this genus. It grows in slightly thermal, alkaline springs in Yellowstone National Park among other cyanobacteria or on the surface of their thalli (Copeland, 1936; Smith, 1950). It may be present in other thermal streams in North America. Colteronema Copeland (Fig. 38C) The thallus is leathery or fibrous, membranous, and macroscopic, up to 1 mm in height, composed of true branched filaments. Filaments are usually crooked and roughly cylindrical. Trichomes are of identical morphology, more or less free, growing horizontally, densely crowded, slightly flexible, more or less cylindrical, torulous, uniseriate, distinctly constricted at the cross walls, and enclosed in colorless to yellow–brown, thick, firm, lamellated sheaths. Trichome branching is true (T and V types) with pseudodichotomous branches, arising after longitudinal division of the apical cells; lateral branching from the intercalary cells sometimes also occurs. Growth is apical. Cells are barrel shaped to ellipsoidal, subcylindrical, and longer than wide. End cells are rounded, sometimes slightly to noticeably enlarged. Heterocytes and akinetes have not been reported. Cell division occurs by horizontal (crosswise) perpendicular fission. Reproduction occurs via fewcelled hormogonia separating as the ends of branches. The single species (Colteronema funebre) is still known only from terrestrial, atmophytic (influenced by hot steam) habitats near thermal springs in Yellowstone National Park (Copeland, 1936).

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Geitleria Friedmann (Fig. 38A) The thallus is filamentous, loosely tufted, up to 2 mm thick, consisting of coiled creeping or erect filaments with irregular lateral and pseudodichotomous true branches (V and T types), with no prostrate basal system and no morphological differentiation into the main and lateral branches. Sheaths are firm, colorless, intensely lime encrusted, containing single trichomes. Trichomes are uniseriate, moniliform, and indistinctly to strongly constricted at the cross walls. Cells are generally barrel shaped or cylindrical, occasionally with narrow individual envelopes. Cell contents are granular, pale grayish–green in color with distinct chromoplasma and solitary granules. End cells are sometimes slightly widened and rounded. Thylakoids are distributed throughout the cell. Heterocytes and akinetes are absent. Cell division is horizontal (crosswise) or longitudinal (before true branching). Reproduction occurs via hormogonia (fragmented filament segments). The habitat is aerial, subaerial, and epilithic, especially on calcareous rocks in caves. The type species, Geitleria calcarea, is probably distributed worldwide (e.g. Israel, France, Romania, Spain, ancient Yugoslavia, the United States, and the Cook Islands), but its occurrence is patchy; thus far, it has only been collected under specific ecological conditions, such as in aragonite caves (Friedmann, 1955, 1979). A second species, G. floridana, growing in similar habitats, is known only from Florida (Friedmann, 1979).

Stigonematales: Nostochopsaceae Nostochopsis Wood ex Bornet et Flahault (Fig. 39A) Thalli are gelatinous and attached to substrata, irregularly spherical or lobed with generally smooth mucilaginous surface, hollow center, up to 3.5 cm in diameter, bluish, olive green, or yellow–green. Common mucilage is usually homogeneous, colorless to yellowish brown. Relatively radially oriented filaments are present within the thallus; they are commonly true branched, slightly coiled, with fine, gelatinous, and diffuse (at margins), the colorless to brownish-yellow sheaths. Trichomes are always uniseriate, with typically isodiametric, barrel shaped or elongated (up to twice as long as wide), ellipsoid, blue–green cells. True lateral (T type or V type) branching is present, with long, cylindrical, multicelled branches terminating in slightly narrowed, rounded apical cells, or very short branches (with one to several cells) terminated by apical heterocytes. Heterocytes are always present, intercalary (bipored), lateral (single pored), or terminal (single pored). Akinetes are unknown. Cell division is horizontal (crosswise), sometimes in meristematic region, but primarily in apical trichome parts. Reproduction occurs via bipolar germinating hormogonia with short barrel-shaped cells.

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FIGURE 39 (A) Nostochopsis: (a) structure of thallus; (b–d) N. lobata (after Bornet and Frémy from Geitler, 1932). (B) Hapalosiphon: (a) true branching; (b) H. hibernicus (after Frémy from Geitler, 1932); (c) H. welwitschii (after Hirose and Hirano, 1981).


Only one freshwater species of Nostochopsis is known. It grows attached to mosses, stones, and submersed wood (rarely free floating when old) in unpolluted creeks and streams in temperate, subtropical, and tropical regions. One variable species, N. lobata, is referred by Smith (1950) to several findings in the eastern United States; Tilden (1910) listed it from Vermont. It has also been collected from several localities in the Greater Antilles (Cuba; Augusto Comas, personal communication).

Stigonematales: Mastigocladaceae Hapalosiphon Nägeli ex Bornet et Flahault (Fig. 39B) The thallus is filamentous, composed of free, coiled filamentous clusters, initially attached to the substratum, later floating free in the metaphyton. Filaments are irregularly curved with uniseriate trichomes (basal trichomes and branches), only rarely with some longitudinally dividing cells. True branching is usually lateral (T type), with no morphological differentiation in the main and lateral trichomes (branches rarely slightly narrower). Sheaths are colorless, thin, firm, and rarely indistinctly lamellated. Heterocytes are intercalary. Cells divide horizontally (crosswise) without meristematic zones. Reproduction occurs via hormogonia separated by necridic cells, usually at the ends of trichome branches. Most species (about 15 have been described) occur in the littoral metaphyton of lakes and in swamps in tropical and temperate zones, usually with macrophytes. Several species prefer moors and peat bogs (H. fontinalis and H. tenuis). One species from the United States occurs in thermal waters (similar to H. fontinalis; Copeland, 1936), and two are subaerial. Smith (1950) recorded H. aureus, H. flexuosus, H. fontinalis, H. hibernicus, H. pumilus, and H. welwitschii in the United States. Prescott (1962) added H. brasiliensis and H. confervaceus to this list; most of these species (seven of eight) were also recorded from British Columbia, along with H. delicatulus and H. intricatus (Stein and Borden, 1979). Whitford and Schumacher (1969) recorded five species from freshwaters on submerged plants and wood and two from subaerial habitats. Tilden (1910) listed seven species from the United States and the West Indies. Mastigocladus Cohn ex Kirchner (Fig. 40A) The thallus is composed of usually densely tangled filaments forming soft, spongy mats, sometimes containing small carbonate crystals, varying from smooth or gelatinous at the surface to compact, occasionally layered, dirty blue–green to olive-green mass. Solitary filaments occur commonly among other cyanobacteria. Trichomes are uniseriate, irregularly coiled with thin,

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distinct, colorless sheaths becoming diffuse with age. Branching is true T type or less frequently reverse Y type, often unilateral (in some stages branching is rare or almost absent). Branches continually taper toward the ends. False branches are facultatively present but uncommon. Heterocytes are intercalary, solitary or in pairs (rare). Akinetes are rare, solitary in older parts of the trichomes. Polymorphic species sometimes occur in unbranched, moniliform filaments. Cell division is horizontal (crosswise) or longitudinal before branching. Reproduction occurs via hormogonia or disintegration of trichomes. The type species, Mastigocladus laminosus, is a thermophilic species from hot springs throughout the world, including Yellowstone National Park (Castenholz and Wickstrom, 1975), British Columbia (Stein and Borden, 1979), Oregon (Jackson and Castenholz, 1975), and Antarctica (Broady, 1984, Broady et al., 1987). It is most likely dependent on the following environmental conditions: 45–60°C, pH >7.5, low oxygen content, and low salinity. M. laminosus shows an extremely large degree of morphological variability, perhaps suggesting several morphotypes and genotypes. Thalpophila Borzì (Fig. 40B) The thallus is filamentous, forming mats on substrata, composed typically of parallel oriented or slightly coiling and creeping true branched filaments. Trichomes are slightly diversified, uniseriate, initially cylindrical, thin and constricted at the cross walls, not attenuated toward the ends, with rounded terminal cells, becoming torulous with age. Lateral T-type branching occurs with branches bending immediately in the direction of the original trichome. Sheaths are thick, lamellated, gelatinizing, from outside becoming confluent or merging with adjacent sheaths. Cells are barrel shaped or cylindrical with a fine granular texture. Heterocytes are uncommon, intercalary, barrel shaped, and solitary. Akinetes are small, forming rows in old trichomes, and embedded with a thick wall. Cell division is horizontal (crosswise). Reproduction occurs via solitary cells (monocytes), liberated from open sheaths. Hormogonia or hormocytes are unknown. The type species, Thalpophila cossyrensis, was described from volcanic rocks near thermal springs (atmophytic) in Italy. Another species, T. imperialis, was described by Copeland (1936) from the calcareous substrata of thermal springs in Yellowstone National Park.

NOTE ADDED IN PROOF A new genus of heterocytous filamentous cyanobateria (Nostocales, Scytonemataceae) was described from United States in 2002.

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FIGURE 40 (A) Mastigocladus: (a–f) M. laminosus (bar = 10 µm; after Anagnostidis 1961). (B) Thalpophila: (a) true branching; (b) T. imperialis (after Copeland, 1936, from Yellowstone National Park).


Spirirestis Flechtner et Johansen (Figure 41) Filaments heteropolar, free, forming tight, regular coils, more or less creeping, densely arranged in colonies, without external mucilage production, 6–20 µm in diameter. Sheats firm, thin or thick, sometimes lamellated. False branches double or single. Trichomes cylindrical, solitary within sheaths, without hairs, rounded at the ends. Cells shorter than broad. Heterocytes basal and intercalar, spherical to oval or appressed. Akinetes not observed. Genus supported by molecular analyses. The single species, S. rafaelensis, was described from soils of a semi-arid juniper community in Utah, USA (Flechtner et al. 2002).

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VI. GUIDE TO LITERATURE FOR SPECIES IDENTIFICATION See Chapter 3, page 110 (this volume) for guide.

LITERATURE CITED Albertano, P., Kovácˇik, L. 1994. Is the genus Leptolyngbya (Cyanophyte) a homogeneous taxon? Archiv für Hydrobiologie/Algological Studies 75:37–51. Anagnostidis, K. 1961. Untersuchungen über die Cyanophyceen einiger Thermen in Griechenland. Institut für Systematische Botanik und Pflanzengeographie Thessaloniki, 322 p.

a

b

c

d

FIGURE 41 Spirirestis rafaelensis (a) Appearance of spiraling filaments; (b) filaments ends in relaxed coils, showing sheath, constrictions at crosswalls, and hormogonia; (c) heterocytes forming; (d) variability in heterocyte shape and false branching in relaxed spirals. Reprinted with permission from Flechtner et al., 2002. © 2002 E. Schweizebart’sche Verlagsbuchlandlung. (mail@schweizbartide, www.schweizerbartide). All scales = 10 µm.

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5

RED ALGAE Robert G. Sheath Office of Provost and Vice President for Academic Affairs California State University, San Marcos San Marcos, California 92096 I. Introduction II. Diversity and Morphology A. Diversity B. Vegetative Morphology C. Reproduction III. Ecology and Distribution A. Streams and Rivers B. Other Inland Habitats

The freshwater red algae in form, in physiology and in habitat may be considered as comprising an elite group of plants, long neglected by American botanists and thus all the more enchanting as representing a research frontier rich with the promise of happy days in the field and laboratory (Flint, 1970, p. 18).

IV. Collection and Preparation for Identification V. Key and Descriptions of Genera A. Key B. Descriptions of Genera VI. Guide to Literature for Species Identification Literature Cited

1932), caves (Nagy, 1965; Hoffmann, 1989), and even sloth hair (Wujek and Timpano, 1986). Freshwater red algae in North America represent a widespread division, with members that range from the high Arctic to the tropical rainforest (Sheath and Hambrook, 1990). The number of species increases from the tundra to the tropics (6 to 25), similar to the trend seen for marine species of Rhodophyta.

I. INTRODUCTION The Rhodophyta, the red algae, constitute a division of organisms that share the following combination of attributes: eukaryotic cells, lack of flagella, floridean starch, phycobiliprotein pigments (red and blue), unstacked thylakoids, and chloroplasts lacking an external endoplasmic reticulum (Woelkerling, 1990). They are primarily marine in distribution, with only approximately 3% of the over 5000 species occurring in truly freshwater habitats (Sheath, 1984). Most of the inland species of Rhodophyta are restricted to streams and rivers (lotic forms), although a few species are found in lakes and ponds (lentic forms) (Sheath and Hambrook, 1990). A small number of inland rhodophytes occurs in habitats other than typical freshwaters, such as hot springs (Doemel and Brock, 1971), soils (Geitler, Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

II. DIVERSITY AND MORPHOLOGY A. Diversity Freshwater rhodophytes have a relatively low diversity compared to other major groups of algae. Analysis of the literature reveals a flora with 65 infrageneric taxa and 25 genera from North America (Table I). Among these taxa, 14 infrageneric taxa are known only from North America. The subclass Florideophycidae accounts for 52 infrageneric taxa, whereas the Bangiophycidae consists of 13 species. Batrachospermum is the most diverse genus, with 26, or 40%, of the total infrageneric taxa. I am proposing in this chapter that there are three potential origins of the predominantly lotic forms: (1) 197

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TABLE I Taxa, Habitats, Forms, Chloroplast Types and Reproduction of Inland Rhodophyta in North America Taxon Bangiophycidae Porphyridiales Chroodactylon ornatuma Chroothece mobilis Cyanidium caldarium Flintiella sanguinaria Kyliniella latvica Porphyridium purpureum P. sordidum Rufusia pilicolab Bangiales Bangia atropurpureaa Compsopogonales Boldia erythrosiphonb Compsopogon coeruleus C. prolificus Compsopogonopsis leptocladus Florideophycidae Acrochaetiales Audouinella eugenea A. hermannii A. macrospora A. pygmaea A. tenellab Balbianiales Rhododraparnaldia oregonicab Batrachospermales Batrachospermaceae Batrachospermum Section Contorta B. ambiguum B. globosporum B. intortum B. louisianaeb B. procarpum Section Setacea B. androinvolucrumb B. atrum Section Virescentia B. elegans B. helminthosum Section Aristata B. macrosporum Section Turfosum B. turfosum Section Hybrida B. virgato-decaisneanum Section Batrachospermum B. anatinum B. arcuatum B. boryanum B. carpocontortumb B. carpoinvolucrumb B. confusum B. gelatinosum B. gelatinosum forma spermatoinvolucrum B. heterocorticumb B. involutumb B. pulchrumb

Habitat

Formc

Chloroplast typed

Reproductive disseminatione

Lake, stream Cool spring Hot spring Cool spring Stream Soil Soil Sloth hair

pf pf u u pf u u pf

cs cs pd pl pmd cs cs pd

ms cd cd cd cd cd cd cd, es

Lake, stream

ff

cs

ms

Stream Stream, cool spring Stream Stream, cool spring

tu ff ff ff

pmd pmd pmd pmd

ms ms ms ms

Stream Stream Stream Stream Stream

tf tf tf tf tf

pmd pmd pmd pmd pmd

ms ms, cs, ts ms ms ts

Stream

m

pmd

cs

Stream Stream Stream Stream Stream

gf gf gf gf gf

pmd pmd pmd pmd pmd

cs cs cs, ms cs cs

Stream Stream

gf gf

pmd pmd

cs cs

Stream Stream

gf gf

pmd pmd

cs cs

Stream

gf

pmd

cs

Stream, pond, bog

gf

pmd

cs, ms

Stream

gf

pmd

cs

Stream, cool spring Stream Stream, cool spring Stream Stream, cool spring Stream, cool spring Stream, cool spring Stream Stream Stream, cool spring Stream

gf gf gf gf gf gf gf gf gf gf gf

pmd pmd pmd pmd pmd pmd pmd pmd pmd pmd pmd

cs cs cs cs cs cs cs cs cs cs cs (Continues)

5. Red Algae

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TABLE I (Continued) Taxon

Habitat

Formc

Chloroplast typed

Reproductive disseminatione

B. skujae B. trichocontortumb B. trichofurcatumb Other genera Sirodotia huillensis S. suecica Tuomeya americanab Lemaneaceae Lemanea borealis L. fluviatilis L. fucina var. parvab Paralemanea annulata P. catenata P. mexicanab Thoreales Nemalionopsis tortuosa Thorea hispida T. violacea Hildenbrandiales Hildenbrandia angolensis Ceramiales Ballia prieuriia Bostrychia moritzianaa B. radicansa B. tenellaa Caloglossa leprieuriia C. ogasawaerensisa Polysiphonia subtilissimaa

Stream Stream Stream

gf gf gf

pmd pmd pmd

cs, ms cs cs

Stream, cool spring Stream Stream

gf gf pp

pmd pmd pmd

cs cs cs

Stream Stream Stream Stream Stream Stream

pp pp pp pp pp pp

pmd pmd pmd pmd pmd pmd

cs cs cs cs cs cs

Stream Stream, cool spring Stream, cool spring

ff ff ff

pmd pmd pmd

ms ms, cs? ms

Stream, cool spring

cr

pmd

g

Stream stream Stream Stream Stream Stream Stream, cool spring

ff ff ff ff pp pp ff

pmd pmd pmd pmd pmd pmd pmd

f? st st st f? f? f?

a

Potential invader from brackish/marine habitats. Unique to North America. c pf, pseudofilament; u, unicell; ff, free filament; tu, tube; tf, tuft; m, mat; gf, gelatinous filament; pp, pseudoparenchymatous form; cr, crust d cs, central stellate with pyrenoid; pd, single peripheral disc; pl, peripheral, intricate lamellate; pmd, peripheral, multiple disc. e ms, monosporangia; cd, cell division; es, endospores; cs, carpospores; fs, tetraspores; g, gemmae; f, fragmentation; st, stichidia. b

specialists that evolved early within the stream environment and are absent in other habitats, (2) generalists that occur in a wide range of other freshwater bodies, such as lakes and ponds, and (3) upstream migrants from estuaries. This proposal is an expanded version of that originally given by Skuja (1938). Of the 60 stream-inhabiting taxa, 50 appear to be specialists, whereas three are generalists and nine are potentially brackish/marine invaders (Table I). Note that there is overlap in two of the species, Chroodactylon ornatum and Bangia atropurpurea, in the last two categories. The other generalist species is Batrachospermum turfosum (Sheath et al., 1994c; Müller et al., 1998).

B. Vegetative Morphology The red algae occurring in typical freshwater habitats tend to be macroscopic and benthic (as defined in Chapter 2) (Sheath and Hambrook, 1990). None-

theless, these algae exhibit a smaller size range than do marine species with the majority (80%) of freshwater rhodophytes having a length range of 1–10 cm. Among the forms occurring in North America, there are 28 gelatinous filaments, 12 free filaments (individual filament without a gelatinous matrix) (e.g., Figs. 2B, 6H, I, and 7B, F, G), nine pseudoparenchymatous forms (tissue-like but composed of compacted filaments) (Figs. 4H, 5A–F, and 7C–E), five tufts (short radiating filaments without a common matrix) (Fig. 2F), two pseudofilaments (loose chains of cells held tegether with a common gelatinous matrix) (Fig. 1C–E), and one each of unicells (Fig. 1A), tubes (Fig. 1H), mats (flat plant body composed of tightly interwoven filaments), and crusts (flat thallus composed of compacted tiers of cells) (Fig. 6E, Table I). Species distributed in hot springs or soils are unicellular and Rufusia on sloth hair is pseudofilamentous. Among the 58 forms that have a filamentous construction, only three have

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multiaxial growth, the members of the Thoreales (Figs. 5G, H and 6A, B) (Sheath et al., 1993b); the rest are uniaxial, except for Bangia, which has a uniaxial base and multiaxial apex at maturity (Fig. 1F). Uniaxial filaments may be corticated with one or more layers of smaller cells covering those of the main axis (Figs. 6I and 7A, B, F, G). The various morphological forms encounter the stress caused by flow in riverine habitats in various ways according to Sheath and Hambrook (1988, 1990). Crusts and short tufts occur within the boundary layer or at least in a region of reduced current velocity and hence avoid much of the flow-related stress. The remaining macroscopic species can be regarded as semierect, experiencing bending, tensile, and compressive forces and perhaps torsional stresses in flowing waters (Vogel, 1984). This group includes mucilaginous and nonmucilaginous filaments and pseudoparenchymatous forms and tubes. It would be expected that the semierect forms possess adaptive mechanisms to tolerate flow, such as branch reconfiguration and extension of thalli in high water motion (Sheath and Hambrook, 1988,1990). Of the 65 infrageneric taxa of Rhodophyta in inland habitats in North America, 20 are reddish, while 45 are largely blue to olive in color. This trend contrasts with that of marine red algae, in which the great majority of species appear red in color. Nonetheless, there are a number of chloroplast morphologies among freshwater taxa: all members of the Florideophycidae have multiple discoidal or ribbon-like chloroplasts without pyrenoids (e.g., Fig. 2G); the bangiophycidean taxa have this type as well as central stellate chloroplasts with a pyrenoid (Fig. 1A), a peripheral lamellate structure, and a single peripheral disc (Table I). However, some of the species, which appear to have multiple discoidal or ribbon-like chloroplasts, may actually contain a complex, interconnected single chloroplast, such as Compsopogon coeruleus (Gantt et al., 1986). Like other red algae, the chloroplasts of freshwater species contain single thylakoids with phycobilisomes (granules consisting of the accessory pigments) on both sides (Sheath, 1984). The phycobilisomes of blue-colored species, such as Porphyridium aerugineum and Compsopogon coeruleus, tend to be hemidiscoidal in shape and predominated by the blue pigment phycocyanin (Gantt et al., 1986). In contrast, the phycobilisomes of the red-colored Porphyridium purpureum are larger and hemispherical and composed mostly of the red pigment phycoerythrin. Phycoerythrin and photosystem (PS) II activity appear to be absent from the pyrenoid of P. purpureum whereas PS I and ribulose1,5-bisphosphate carboxylase/oxygenase (RuBisCO) activities can be detected in this structure (McKay

and Gibbs, 1990). Chloroplasts of Batrachospermum gelatinosum develop from proplastids, which have a double-membraned envelope and a parallel outer photosynthetic thylakoid (Brown and Weier, 1968). This outer thylakoid functions in the production of additional ones to the interior. Thylakoids have been observed to be coiled in serial sections of Cyanidium, Compsopogon, and Batrachospermum (Pueschel, 1990). They can fragment and form dilated tubular units in some freshwater species, such as Batrachospermum gelatinosum and Sirodotia suecica, when subjected to reduced illumination (Sheath et al., 1979). Another characteristic that is useful in analyzing the morphology of freshwater Rhodophyta is the external covering. Unicells and pseudofilaments typically have a gelatinous matrix surrounding the cells (e.g., Fig. 1C, D, G) which varies in thickness, depending on the age and physiological state of the organisms (Sheath, 1984). The gelatinous filamentous members of the Batrachospermales, such as Batrachospermum and Sirodotia, have distinct cell walls as well as an overall matrix surrounding the filament. The free filaments and pseudoparenchymatous forms have only cell walls. The gelatinous matrices of Porphyridium and Batrachospermum are complex mixtures of a variety of monomeric sugars, including galactose, glucose, and xylose (Craigie, 1990). The cell walls of a freshwater isolate of Bangia atropurpurea are similar to those of marine collections in having repeating water-soluble dissacharide units of agarose and porphyran and insoluble residues of galactose and mannose (Youngs et al., 1998). The cell walls of Paralemanea annulata have xylan as the major polysaccharide as well as cellulose in small quantities as the fibrillar components (Gretz et al., 1991). The amorphous component consists of a glucuronogalactan. Water-soluble cell wall polymers of freshwater Bostrychia moritziana are composed of a complex mixture, including methyl agarose and methyl porphyran (Youngs et al., 1998). The insoluble residues contain a mixture of galactose and glucose. Many freshwater red algal species exhibit differential staining of external coverings with Alcian Blue, particularly of mucilaginous layers, rhizoids, sporangia and spermatangia (Sheath and Cole, 1990).

C. Reproduction Freshwater red algal species exhibit a diversity of reproductive types, particularly in terms of dissemination (Table I). Cell division is the major mode of population increase among the unicellular forms. During mitosis of Porphyridium purpureum and Flintiella sanguinaria, the nuclear envelope remains intact with polar openings, the spindle apparatus is composed of

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interdigitating half spindles, and the nuclear-associated organelle (NAO) is an electron-dense bipartite structure (Broadwater and Scott, 1994). Cell division in the other forms is generally the mechanism by which the thallus is expanded. Mitosis in Batrachospermum anatinum is similar to that of the unicells but also includes perinuclear endoplasmic reticula and a bipartite NAO that is composed of a small ring within a large one (Scott, 1983). Monosporangia formation is the major form of asexual reproduction among the pseudofilamentous and filamentous taxa (Table I) and typically involves the formation of single spores that germinate back into the life history phase that produced them. In Chroodactylon ornatum and Bangia atropurpurea, spores are released by localized digestion of the common filamentous matrix (Fig. 1G) (Sheath, 1984). The order Compsopogonales is characterized by its method of monospore production, which involves the cleavage of a relatively small cell from a larger vegetative cell by oblique cell division (Fig. 2A, C, D) (Garbary et al., 1980). Monosporangia of the Acrochaetiales and Batrachospermales are specialized, enlarged, and obovoid cells typically produced at the apices of vegetative branches (Figs. 2G and 3I) (Sheath, 1984). Monosporangia can be regenerated after spore release by protrusion and cleavage of cytoplasm from the subtending cell in Audouinella hermannii (Hymes and Cole, 1983) and Batrachospermum intortum (Sheath et al., 1992). Monospores are also a mechanism by which certain life history phases perpetuate themselves in complex life history alternation, such as the “chantransia” phase (see below) of the Batrachospermales (Sheath, 1984). Another form of asexual spore reported from inland rhodophytes is the endospore in Cyanidium caldarium and Rufusia pilicola (Wujek and Timpano, 1986; Seckbach, 1991). Sexual reproduction and life history alternation are known for many species of freshwater red algae, although these phenomena have not been conclusively demonstrated for freshwater members of the Bangiophycidae or the Hildenbrandiales of the Florideophycidae (Table I). In the freshwater Audouinella species for which the life history has been analyzed, the free-living gametophyte and tetrasporophyte are isomorphic, both having the same tuftlike morphology (Fig. 2F) (Drew, 1935; Necchi et al., 1993a). The haploid gametophyte produces the gametangia. The female gametangium, the carpogonium, is a colorless cell with an inflated base and narrow tip, the trichogyne (Fig. 2I). The male gametangium, the spermatangium, is also colorless, obovoid in shape, and releases one spermatium at a time (Fig. 2H). Spermatia attach to the trichogyne and one eventually fertilizes the carpo-

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gonium. The zygote divides into a microscopic diploid phase, the carposporophyte, which remains attached to the gametophytic stage until its deterioration (Fig. 2J). Carpospores germinate into the diploid tetrasporophyte, which, at maturity, forms tetrasporangia at the branch tips (Fig. 3A). Haploid tetraspores are formed by meiosis and germinate into the gametophytic stage, thereby completing the life history. In a small stream in Rhode Island, both the tetrasporangia and the carpogonia of A. hermannii are formed in a brief period of time, February and May, respectively (Korch and Sheath, 1989). It would appear that production of these structures in a short period of time is common in North America because gametangia and carposporophytes were observed in only 7 out of 75 collections from throughout the continent (Necchi et al., 1993a). Audouinella tenella, which has only been collected in California, has only been found to contain tetrasporangia (Necchi et al., 1993a). The blue-colored Audouinella species have not been observed to contain gametangia, carposporophytes, or tetrasporangia in 34 collections in North America (Necchi et al., 1993b). This finding may indicate that they are not, in fact, true Audouinella species, but rather one of the life history stages of the Batrachospermales, the “chantransia” (see below). Further substantiating this possibility, Pueschel et al. (2000) have demonstrated that isolates formerly classified as the blue-colored Audouinella macrospora were positioned in an 18S rRNA gene tree with samples of Batrachospermum and not with the freshwater red-colored Audouinella hermannii. In addition, the pit plugs of A. macrospora were also like those of the Batrachospermales. Rhododraparnaldia oregonica has characteristics that are intermediate between the Acrochaetiales and the Batrachospermales, including reproductive structures (Sheath et al., 1994d). The carpogonia and spermatangia are similar in morphology to those of Audouinella (Fig. 3C) and fertilized carpogonia form microscopic carposporophytes (Fig. 3D). However, unlike the Acrochaetiales, the spermatangia are formed on specialized colorless stalks, rather than at the apices of vegetative branches (Fig. 3C). What is similar between Rhododraparnaldia and the Batrachospermales is the fact that the free-living life history stages are heteromorphic, being quite different in morphology. The semierect gametophytes are composed of a main axis with barrelshaped axial cells that are distinctly larger in diameter than the more elongate lateral branch cells (Fig. 3B). The microscopic to small macroscopic “chantransia” stage (so named because it was originally thought to be a separate genus) contains simple branched filaments with no difference in diameter and no obvious main axis (Fig. 3E) (Sheath et al., 1994d). The latter stage is

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diploid and formed from the germination of carpospores. Where it differs from the tetrasporophyte of the Acrochaetiales described above is in the process of meiosis. In the “chantransia” stage, tetraspores are not produced but rather the haploid gametophyte is formed directly attached to this stage (Fig. 3E) (Sheath, 1984; Sheath et al., 1994d). In the freshwater red algal species for which this process has been studied, it appears that meiosis takes place in an apical cell of the “chantransia” filament; in each division, a residual nucleus is extruded into a lateral protrusion, which is then separated by wall formation (Sheath, 1984). The one remaining haploid nucleus forms the gametophyte. This life history alternation is typical of the Batrachospermales. Fifty-nine percent of the freshwater Rhodophyta in North America belong to the order Batrachospermales or Thoreales (Table I). Like Rhododraparnaldia, the gametophyte is macroscopic and semierect, while the “chantransia” stage is often microscopic or composed of tiny tufts (Fig. 4C). The latter forms are morphologically similar to the bluish forms of Audouinella noted above. Sheath (1984) proposed that the “chantransia” stage of the Batrachospermales may be an evolutionary adaptation for population maintenance in the upper portions of drainage basins. This stage is typically perennial, seasonally producing the attached gametophyte (Yoshida, 1959; Sheath, 1984; Necchi, 1993). Therefore, the population can continue to proliferate upstream while colonizing downstream with the release of carpospores (Sheath, 1984). If these algae possessed the typical red algal life history, as exhibited by the Acrochaetiales, release of both tetraspores and carpospores would result in a gradual shift of populations downstream until they were solely in the larger trunk river, which is too deep, turbid, and sedimented to support the growth of most autotrophs. Raven (1993) demonstrated that the photosynthetic rates in situ of the “chantransia” stage of Lemanea mamillosa are one twentieth of those of the gametophyte and the former phase has a negligible role in provisioning the growing gametophyte. He also concluded that the key role of the “chantransia” stage is to occupy space throughout the year, including possible exposure during summer drawdown. This stage may also act in population dispersal through the production of monospores (Sheath, 1984; Raven, 1993). In North American temperate streams, gametophytes are typically present from late fall to late spring (Sheath and Hambrook, 1990). Another key feature pertaining to the reproduction of the Batrachospermaceae of the Batrachospermales is the formation of relatively enlarged and persistent trichogynes of the carpogonia, compared to those of red algae from other orders (Figs. 3K–M and 4E, I) (Sheath, 1984). The larger surface area and longevity

would enhance the probability of spermatia contact. Hambrook and Sheath (1991) demonstrated that the mean percentage fertilization rate for various species of Batrachospermum was 45–72%, including dioecious taxa. This rate may be obtained because spermatia are released into turbulent eddies downstream of rocks, where they are carried through the female plants numerous times as the water is moving back and forth (Sheath and Hambrook, 1990). Carpogonia are borne on carpogonial branches that may be little to highly differentiated from adjacent vegetative branches in the Batrachospermales (Fig. 3K, M). In the undifferentiated carpogonial branch of Batrachospermum involutum, the cells are uninucleate with abundant starch granules and several well-developed peripheral chloroplasts (Sheath and Müller, 1997). In contrast, the short carpogonial branch cells of B. helminthosum have no visible starch, chloroplasts are highly reduced, and cross walls break down among cells. Freshwater populations of Hildenbrandia of the order Hildenbrandiales are typically vegetative and reproduce by gemmae, dense aggregations of cells formed in cavities in the thallus (Fig. 6F, G) (Starmach, 1969; Seto, 1977; Sherwood and Sheath, 2000b). The gemmae are eventually released from the thalli and germinate into new crusts, presumably of the same ploidy level. Most of the freshwater members of the Ceramiales in North America have only been observed in their vegetative stages, including Ballia, Caloglossa, and Polysiphonia (Sheath et al., 1993c). They probably proliferate through fragmentation of the thallus and subsequent growth of the fragments. In contrast to Ballia in North American streams, inland populations from South America and Malaysia have been observed to contain monosporangia (Kumano, 1978; Couté and Sarthou, 1990; Necchi, 1995). In addition, brackish water populations of Caloglossa ogasawaraensis have been observed to contain gametangia and carposporophytes or tetrasporangia (Tanaka and Kamiya, 1993). A few freshwater populations of the genus Bostrychia form stichidia which are inflated, multichambered structures at the tips of vegetative branches (Fig. 6I). These chambers may form the tetrasporangia, but this has only been observed conclusively in collections of B. moritziana from Venezuela and Brazil (D’Lacoste and Ganesan, 1987; Kumano and Necchi, 1987; Sheath et al., 1993c).

III. ECOLOGY AND DISTRIBUTION A. Streams and Rivers Because 94% of the inland rhodophytes of North America occur in streams or rivers (Table I), this

5. Red Algae

chapter will concentrate on this habitat. Much of the ecology of riverine red algae has been summarized by Sheath and Hambrook (1990) and I will synthesize the trends and give updated information here.

1. Patterns of Distribution In North America, 51% of 1000 first- to fourthorder stream reaches surveyed contain red algae and 24% have two or more species (Sheath and Hambrook, 1990; Sheath and Cole, 1992). The maximum number of red algal species found per reach is six. The most widespread species is Batrachospermum gelatinosum, which occurs in about 13% of the streams examined. This species occurs from the polar desert on Ellesmere Island, Northwest Territories (80°N) to the southeastern coastal plain in Louisiana (37°N) (Vis et al., 1996a; Vis and Sheath, 1997). Other widespread species include Audouinella hermannii (North Slope of Alaska to Georgia) (Necchi et al., 1993a), Batrachospermum helminthosum (Washington and Maine to central Mexico) (Sheath et al., 1994a), B. turfosum (North Slope of Alaska to central Mexico) (Sheath et al., 1994d; Müller et al., 1997), and Lemanea fluviatilis (central Alaska to Arkansas) (Vis and Sheath, 1992). Among the taxa of lotic Rhodophyta in North America with more restricted patterns of distribution, there are some interesting trends. Members of the Compsopogonaceae, Thoreales, Hildenbrandiales, Ceramiales, and Batrachospermum section Contorta are largely restricted to warmer waters from south temperate to tropical streams (Vis et al., 1992; Sheath et al., 1992, 1993a, b). There are also taxa that are mostly in north temperate to tundra habitats, such as Batrachospermum gelatinosum forma spermatoinvolucrum and Lemanea borealis (Vis and Sheath, 1992, 1996, 1998). Boldia and Tuomeya have been collected only in eastern North America, ranging from Quebec to the southeastern United States (Howard and Parker, 1980; Sheath and Hymes, 1980; Kaczmarczyk et al., 1992). There are a number of species that appear to be localized only in southwestern spring-fed streams, such as Flintiella sanguinaria (Ott 1976), Chroothece mobilis (Blinn and Prescott, 1976), and three members of Batrachospermum section Batrachospermum (B. carpocontortum, B. carpoinvolucrum and B. involutum) (Vis and Sheath, 1996; Sherwood and Sheath, 1999a). Some of the warm-water groups noted above are also distributed in spring-fed streams of southwestern North America.

2. Physical Factors Riverine red algae exhibit a wide range of occurrence with respect to current velocity (Sheath and Hambrook, 1990). Nonetheless, most species are

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found in moderate flow regimes (mean 29–57 cm s–1). Moderate flow enhances various aspects of metabolism, including productivity and pigment content (Thirb and Benson-Evans, 1982), growth (Whitford, 1960), respiration rate (Schumacher and Whitford, 1965), and phosphorus uptake level (Schumacher and Whitford, 1965). In addition, it has a positive influence on the ecology of these organisms, such as washout of loosely attached competitors (Whitton, 1975), constant replenishment of gases and nutrients (Hynes, 1970), and reduction of the boundary layers of depletion around the algal thallus (MacFarlane and Raven, 1985). Few taxa are typically localized at high current velocities (>1 m s–1), the exceptions being Lemanea and Paralemanea of the Lemaneaceae (e.g. Everitt and Burkholder, 1991; Vis et al., 1991). The morphology of some species, such as Sirodotia delicatula, can be altered under different flow regimes (Necchi, 1997). At high current velocities (132 cm s–1), plants are denser, having shorter internodal lengths. Sheath and Hambrook (1988) calculated mean potential velocities (in cm s–1) at which various morphological forms of red algae would break: tufts, 80; mucilaginous filaments, 160; and cartilaginous and pseudoparenchymatous thalli, 580. The light regime, which includes changes in intensity, quality, and photoperiod, is one of the key factors affecting the distribution and seasonality of riverine Rhodophyta (Sheath and Hambrook, 1990). Illumination affects algal growth via photosynthesis, by processes indirectly related to photosynthesis, and by those processes unrelated to photosynthesis. In the case of freshwater red algae, distribution within a drainage basin and seasonality are determined by the photoregime established by the surrounding tree canopy. In a headwater Rhode Island stream containing Batrachospermum boryanum, the total illumination reaching the water surface is reduced by 90–99% on both sunny and cloudy days in a shaded reach compared with a nearby open segment (Kaczmarczyk and Sheath, 1991). There is a slight but significant increase in green light under the canopy and a corresponding increase in the red pigment phycoerythrin compared to the blue pigment phycocyanin. The action spectrum of collections from the canopied and open sites are similar and quite broad. Nonetheless, the populations of Batrachospermum mostly disappear during periods of peak canopy shading (Hambrook and Sheath, 1991). Likewise, many species of stream-inhabiting Rhodophyta exhibit a positive correlation to light and a negative one to temperature (Kremer, 1983; Sheath, 1984; Leukart and Hanelt, 1995). In addition, they tend to exhibit low saturating levels of illumination for photosynthesis (35–400 µmol photons m–2 s–1).

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Temperature regime influences the latitude, elevation, drainage basin distribution, as well as the seasonality of freshwater red algae (Sheath and Hambrook, 1990). Latitudinal patterns have been discussed above. Kremer (1983) concluded that some of the geographic patterns of riverine Rhodophyta are based on photosynthetic response to temperature. For example, the concentration of members of the Compsopogonaceae in warm waters can be explained by a maximum photosynthesis rate at 30–35°C. In large drainage basins, elevation and basin distribution patterns are interrelated; mean temperatures tend to increase from the source to the mouth, although the amplitude of diurnal fluctuations in temperature become less (Whitton, 1975). Israelson (1942) reported that most rhodophytes in Sweden were restricted to elevations less than 900 m above sea level. From our surveys of North America, we have observed a similar trend. Exceptions include the Lemaneaceae and some members of the Acrochaetiales, such as Audouinella hermannii and A. tenella, which can be abundant in montane streams (e.g., Necchi et al., 1993a; Vis and Sheath, 1992). In temperate regions, most freshwater red algae exhibit maximum biomass, growth, and reproduction between late fall and early summer (Sheath and Hambrook, 1990), but in many cases this seasonality is more related to light penetration to the stream surface than to temperature (e.g., Hambrook and Sheath, 1991). Necchi (1993) noted a similar seasonality for batrachospermalean species in a tropical drainage basin in southeastern Brazil where a combination of lower temperature and reduced turbidity during the dry winter months promoted growth of macroscopic gametophytes. In contrast, Compsopogon coeruleus was present throughout the year and distribution was not related to temperature, but to current velocity in these Brazilian streams.

3. Chemical Factors The interaction between pH and the form of inorganic carbon can greatly influence the productivity and distribution of freshwater Rhodophyta (Sheath and Hambrook, 1990). Although widespread species are found in a wide range of pH values, the majority occur in mildly acidic waters between pH 6 and 7. However, there are exceptions to this pattern, including Bangia, Chroodactylon, Hildenbrandia, and members of the Compsopogonaceae, Thoreales, and Ceramiales, which may be considered to be alkalophiles (Sheath, 1987; Sheath et al., 1993a–c; Vis et al., 1992; Vis and Sheath, 1993). The effect of pH can be attributed to the form of inorganic carbon available; some taxa, such as Lemanea mamillosa, have been shown to use only free CO2 as a carbon source for photosynthesis,

which is the predominant form at mildly acidic pH values (e.g., Raven et al., 1994). Above pH 8, the proportion of free CO2 drops below 2–5% and species occurring in these waters would require flow replenishment or use of alternative sources of inorganic carbon (Sheath and Hambrook, 1990). One species commonly distributed in high-pH waters is the crustose Hildenbrandia rivularis, which also utilizes CO2 as a carbon source but may also use HCO3–, although this possibility has not been confirmed (Raven et al., 1994). Specific conductance and pH are related in that alkaline waters are high in ions, such as carbonates, and are buffered strongly above pH 8 (Sheath and Hambrook, 1990). In contrast, waters draining igneous rock catchment areas are less well buffered and usually more acidic. Four common freshwater red algal species in North America, Audouinella hermannii, Batrachospermum gelatinosum, Lemanea fluviatilis, and Tuomeya americana, exhibit a negative frequency distribution in relation to specific conductance (total ions); the greatest frequencies occur below < 100 µS cm–1 (Sheath and Hambrook, 1990). This pattern is due in part to the form of inorganic carbon. Those taxa that typically occur at higher pH values, as noted above, are also distributed at high conductance ranges. Likewise, species typically localized in hardwater streams constitute the same list given above for high pH values. Freshwater red algae are found in a wide range of oxygen concentrations (0.2–21 mg L–1), but there tends to be an increase in frequency of occurrence with higher concentrations (Sheath and Hambrook, 1990). To some extent, this relationship results from the occurrence of many species in the cooler months from late fall to late spring when oxygen solubility is highest. Nevertheless, freshwater Rhodophyta are not commonly associated with stagnant, organic-rich waters with very low oxygen contents. Freshwater rhodophytes occur over a broad range of nutrient values, but they are more typically found in low to moderate nutrient regimes (e.g., PO43– below detection to 100 µg L–1) (Sheath and Hambrook, 1990). The common occurrence of red algae at low nutrient levels is partially due to flow replenishment and reduction of the boundary layer of depletion in riverine systems. In addition, many species form colorless hair cells that may be produced in response to nutrient deficiency, as is the case for some green algal filaments (Gibson and Whitton, 1987). Some researchers have employed rhodophytes for classification of streams; for example, in Austria Hildenbrandia is typical of lowland rivers with relatively high nutrients, whereas Lemanea is regarded as indicative of high-altitude streams with low nutrients (Pipp and Rott, 1994).

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In general, freshwater red algae are localized in reasonably unpolluted waters and are infrequent to absent in streams and rivers that are organically enriched, greatly silted, or very high in inorganic nutrients (Sheath and Hambrook, 1990). However, Lemanea fluviatilis and Bangia atropurpurea appear to be tolerant of some heavy-metal pollution (Lin and Blum, 1977; Harding and Whitton, 1981). For example, L. fluviatilis can occur at aqueous concentrations of zinc up to 1.16 mg L–1.

4. Biotic Factors Thirty-eight riverine animals to date have been observed to ingest freshwater red algae, based on gut content or feeding studies (Sheath and Hambrook, 1990; Sheath et al., 1995). These grazers include two amphipods and the larvae of six mayfly, thirteen caddisfly, six stonefly, six chironomids, one beetle, as well as two snails and two cyprinoid fish. Most animals remove small pieces of 5–20 cells and digest the cytoplasm, leaving the empty walls intact. The majority of these animals are polyphagous, consuming a wide variety of food matter, including detritus, leaf fragments, and other algal taxa. In grazing experiments done by Hambrook and Sheath (1987), it was observed that the preference of consumption was Audouinella hermannii, Batrachospermum helminthosum, followed by Tuomeya americana; this trend was likely based on increasing toughness of the thallus and reduction of protein content. Rosemond (1993) noted that irradiance, nutrients, and herbivore grazing simultaneously limited algal biomass, including Audouinella sp., in a small forested stream. Cases of larvae and pupae from six caddisfly species have been observed to contain pieces of freshwater rhodophyte thalli in North America (Sheath et al., 1995). Seven genera of Rhodophyta (Batrachospermum, Bostrychia, Compsopogon, Compsopogonopsis, Lemanea, Paralemanea, and Tuomeya), representing 13 species and 35 populations, have been observed in this association. Strips of the algae are fit together in a transverse, concentric, or spiralled fashion. In some of the associations, pieces of the rhodophyte are also found in the gut of the caddisfly. In addition, some of the algal strips remain viable in these cases. Three genera of chironomid larvae have also been observed to incorporate pieces of red algae in their cases in North America (Sheath et al., 1996a). Five genera of Rhodophyta are used in this process, Audouinella, Batrachospermum, Lemanea, Paralemanea, and Sirodotia. The cases are tubular in shape with longitudinally oriented strips of algae held together with silken threads.

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Competition for suitable substrata can occur among species of freshwater red algae or with other benthic algae at various levels (Sheath and Hambrook, 1990). Unicellular forms, such as Flintiella, microscopic stages, and low-growing forms, including the “chantransia” stage of the Batrachospermales, Audouinella and Hildenbrandia, are common components of the stream epilithic community. As such, they compete with a complex association of microalgae, usually dominated by diatoms during early colonization stages (e.g., Steinman and McIntire, 1986). Fritsch (1929) noted that Hildenbrandia thalli are often overgrown by diatoms and cyanobacteria in British streams. In later stages of succession, filamentous and stalked species can form a canopy, providing a competitive advantage for light and nutrient replenishment. The semierect forms fit into this category and are subjected to competition with macrophytes and other macroalgae. Bryophytes are frequent dominants in the upper reaches where red algae occur (e.g., Sheath et al., 1986). The macrophytes are subjected to removal by flooding events, allowing new periphyton colonization. Therefore, lotic communities containing red algae are generally in a nonequilibrium state, consisting of most successional stages (Sheath and Hambrook, 1990). On a geographic scale, certain freshwater rhodophyte species have distributional patterns that are correlated to that of other species (Sheath and Hambrook, 1990). For example, in tropical streams in North America, Bostrychia, Compsopogon, and Hildenbrandia frequently cohabit the same stream reaches. In temperate and boreal regions, Audouinella hermannii and members of the Lemaneaceae are frequently found together; the former taxon can be both epiphytic on Lemaneaceae and epilithic in these situations.

B. Other Inland Habitats Soft-water ponds and bogs represent the second most common habitat to encounter inland Rhodophyta (Sheath, 1984). In particular, Batrachospermum turfosum is common in these habitats, and Yung et al. (1986) noted its broad occurrence in the northeastern regions of Canada and the United States. Müller et al. (1997) studied the phenology of a population of B. turfosum in a boreal pond in Newfoundland. The gametophyte is perennial with a peak cover in summer, which is correlated to water temperature and day length. Carpogonia and spermatangia are present throughout the year except for October and November when monospore production is predominant. Two species, which can be classified as brackish/ marine invaders of freshwaters, Chroodactylon ornatum and Bangia atropurpurea, are common species in

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hard-water sections of the lower Great Lakes (Sheath and Morison, 1982; Sheath, 1987; Müller et al., 1998). These species are absent from the low-ion waters of Georgian Bay, the North Channel, and Lake Superior (specific conductance < 200 µS cm–1) (Sheath et al., 1988). Collections of Bangia from Lakes Ontario, Erie, Huron, Michigan, and Simcoe have identical DNA sequences for the nuclear gene coding for the small subunit of ribosomal RNA (18S rDNA), the chloroplast gene coding for the large subunit of RuBisCO (rbcL) and the spacer unit between rbcL, and the gene coding for the small subunit of RuBisCO (rbcS) (Müller et al., 1998). These sequences are also nearly identical to those from freshwater collections of Bangia from Europe, the Thames and Shannon Rivers, United Kingdom, and Lake Garda, Italy. Hence, it would appear that the North American freshwater Bangia arose by a single invasion from a European freshwater population, possibly by vector-assisted transport (e.g., ballast water of ships), rather than by migration from the Atlantic Ocean. Cyanidium caldarium is an unusual species restricted to several acid hot springs in North America, including those in the United States, Mexico, and El Salvador (DeLuca et al., 1979). These springs exhibit the following range of conditions: pH 1–2.6 and temperature 25–44°C. Doemel and Brock (1971) examined growth properties of Cyanidium isolated from Yellowstone National Park and observed that the optimum pH was 2–3 (range 0.5–5.0) and the optimum temperature was 45°C (range 25–56°C). This species also forms surface and subsurface crusts in hot, acidic soils and river banks near thermal springs (Smith and Brock, 1973). Species of Porphyridium can form gelatinous crusts on moist soils and decaying wood (Geitler, 1932; Ott, 1972). In these habitats, these species are reasonably desiccation resistant and shade tolerant (Hoffmann, 1989).

IV. COLLECTION AND PREPARATION FOR IDENTIFICATION Because 94% of inland red algal taxa occur in streams or rivers, I will concentrate on collecting in this environment. While macroalgae can be distributed

throughout a major drainage basin, they tend to be more common in the smaller channels of first- to fourth-order reaches (Sheath and Cole, 1992). Because only approximately 5% of stream channels examined in North America have 50% or more of the stream bottom covered by one or more species of red algae (Sheath and Hambrook, 1990), it is often necessary to actively search for these taxa. A view box, composed of a glass bottom and Plexiglas sides, is of great help in being able to observe the stream bottom during this search process. To attain a representative sampling of species, at least a 20-m length should be carefully examined using the view box, including a variety of flow regimes and substrata. For example, in a large, fast-flowing stream, members of the Acrochaetiales and Lemaneaceae may be found attached to rocks in more rapidly flowing portions in the midchannel, whereas members of the Batrachospermaceae may be localized in quiet side channels and pools, attached to a variety of substrata, such as logs and tree roots. Long forceps are quite useful for grabbing specimens in deep waters, particularly gelatinous filaments of the Batrachospermaceae. Razor blades are necessary to remove crustose forms. Other collecting equipment for consideration includes hip waders, diver’s gloves for winter collecting, and various portable meters, such as pH, specific conductance, temperature, turbidity, and current. Red algal specimens are best viewed shortly after collection in a live state using a combination of dissecting and compound microscopes. If they cannot be examined quickly, then they should be fixed in 2.5% histological-grade glutaraldehyde, buffered with a pinch of CaCO3, and stored in a dark and cool environment. Under these conditions, they will maintain their morphology and pigmentation for several years. Other fixatives, such as formalin or Lugol’s, cause more distortion, and drying of herbarium specimens results in considerable morphological damage. Generally, samples are sorted and initially viewed at low power with a dissecting microscope. Then a reproductive piece is removed, mounted on a microscope slide with cover slip, finely chopped with a sharp razor blade and squashed to obtain flat images, and then viewed at 200⫻ or 400⫻ in a compound microscope. It is necessary to find all of the key vegetative and reproductive features noted in the section below to achieve a proper identification.

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V. KEY AND DESCRIPTIONS OF GENERA A. Key 1a.

Thallus unicellular (Fig. 1A)...............................................................................................................................................................2

1b.

Thallus multicellular...........................................................................................................................................................................4

2a.

Reproduction by endospores, hot springs............................................................................................................................Cyanidium

2b.

Reproduction by cell division, other inland habitats...........................................................................................................................3

3a.

Parietal chloroplasts without pyrenoids..................................................................................................................................Flintiella

3b.

Axial chloroplast with pyrenoid.....................................................................................................................................Porphyridium

4a.

Thallus pseudofilamentous (Fig. 1C–E)..............................................................................................................................................5

4b.

Thallus crustose, filamentous, or pseudoparenchymatous..................................................................................................................8

5a.

Axial chloroplast with pyrenoid.........................................................................................................................................................6

5b.

Parietal chloroplasts without pyrenoids..............................................................................................................................................7

6a.

Few-celled colony, no branching (Fig. 1B)..........................................................................................................................Chroothece

6b.

Multiple celled with false branching (Fig. 1C)...............................................................................................................Chroodactylon

7a.

Discoidal base (Fig. 1D), occurs in streams and rivers...........................................................................................................Kyliniella

7b.

No discoidal base, in sloth hair.................................................................................................................................................Rufusia

8a.

Thallus crustose (Fig. 6E)..............................................................................................................................................Hildenbrandia

8b.

Thallus a monostromatic, hollow sac (Fig. 1H)..........................................................................................................................Boldia

8c.

Thallus filamentous or pseudoparenchymatous..................................................................................................................................9

9a.

Thallus filamentous..........................................................................................................................................................................10

9b.

Thallus pseudoparenchymatous (tissue-like but with compacted filamentous construction).............................................................21

10a.

Multiaxial filaments (Figs. 5G, H and 6A, B)...................................................................................................................................11

10b.

Uniaxial filaments (Figs. 2G and 3B)................................................................................................................................................12

11a.

Monosporangia on short branches at base of assimilatory filaments (Fig. 6C)..........................................................................Thorea

11b.

Monosporangia on long branches at periphery of assimilatory filaments (Fig. 5I).........................................................Nemalionopsis

12a.

Filament uniaxial at base and multiaxial at reproductively mature apices (Fig. 1F)...................................................................Bangia

12b.

Filament uniaxial throughout..........................................................................................................................................................13

13a.

Filaments non-corticated (Figs. 2G, I and 3B)..................................................................................................................................14

13b.

Filaments corticated (Figs. 2B, C and 7B).........................................................................................................................................16

14a.

No obvious main axis or reduction in diameter in lateral branches...................................................................................Audouinella

14b.

Distinct main axis and lateral branches reduced in diameter.............................................................................................................15

15a.

Lateral branches not compacted, spermatangial stalks (Fig. 3C)............................................................................Rhododraparnaldia

15b.

Lateral branches compacted and pinnate, no spermatangial stalks (Fig. 6H)...............................................................................Ballia

16a.

Reproduction by monosporangia, which are cleaved obliquely from cortical cells (Fig. 2C).............................................................17

16b.

Reproduction by transversely cleaved monosporangia and/or sexual only........................................................................................18

17a.

Rhizoidal cells confined to filament base, cortical cells cuboidal (Fig. 2C).....................................................................Compsopogon

17b.

Rhizoidal cortication scattered throughout filament (Fig. 2E)................................................................................Compsopogonopsis

18a.

Thallus with whorls of determinate lateral branches (Figs. 3F–H and 4D).......................................................................................19

18b.

Thallus with alternate or opposite lateral branches (Fig. 2G)...........................................................................................................20

19a.

Carposporophytes a distinct mass of determinate gonimoblast filaments (Fig. 4A, B)..............................................Batrachospermum

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19b.

Carposporophytes indistinct, indeterminate gonimoblast filaments (Fig. 4F).........................................................................Sirodotia

20a.

Trichoblasts (hair cells) from cortical cells, no stichidia (Fig. 7F).....................................................................................Polysiphonia

20b.

No trichoblasts, stichidia at branch tips (Fig. 6I).................................................................................................................Bostrychia

21a.

Thallus a branched flat blade with a distinct midrib (Fig. 7C, D)........................................................................................Caloglossa

21b.

Thallus a cartilaginous tube with no midrib.....................................................................................................................................22

22a.

Whorled lateral branches appearing like beads in a row at low magnification (Fig. 4H)........................................................Tuomeya

22b.

Thallus tubular, sometimes with distinct nodal swellings (Fig. 5A)...................................................................................................23

23a.

Main axis corticated, spermatangia in rings around the diameter of the thallus (Fig. 5D)................................................Paralemanea

23b.

Main axis uncorticated, spermatangia in patches (Fig. 5A)....................................................................................................Lemanea

B. Descriptions of Genera (for a list of species in each genus, see Table I)

Porphyridiales: Unicellular Forms Cyanidium L. Geitler Cyandidium is composed of spherical unicells, rarely united in a common mucilaginous matrix, 1.5–6 µm in diameter. Each cell contains one blue– green, parietal, spherical to cuplike chloroplast without a pyrenoid, a mitochondrion, and a nucleus, but no vacuole. The chloroplast contains single, parallel, concentric thylakoids with phycobilisomes and predominant C-phycocyanin. Reproduction occurs by four endospores (2–3 µm). Sexual reproduction has not been reported. This genus is widespread in acidic thermal areas. The temperature range in situ is 25–44°C; the pH range in situ 0.05–5.0 (Doemel and Brock, 1971; DeLuca et al., 1979). Flintiella F. D. Ott in Bourelly Flintiella is composed of spherical cells united in a gelatinous matrix, each with a massive, parietal, reddish chloroplast without a pyrenoid. The cell diameter ranges from 9 to 20 (45) µm. Reproduction occurs by cell division. Sexual reproduction is unknown. This genus is found in Barton Springs, Austin, Texas, in autumn (Ott, 1976). Total dissolved solids, 513 mg L–1; NO3–1, 301 mg L–1; pH, 6.9. Porphyridium Nägeli (Fig. 1A) Porphyridium is composed of spherical to ovoid unicells with a stellate chloroplast and prominent central pyrenoid. The cell diameter is 5–10 µm in the exponential phase, 7–16 µm in the stationary phase. Cells are solitary, but often grouped into irregular colonies with an ill-defined mucilaginous matrix. Species are distinguished by chloroplast color. Reproduction occurs by cell division.

Porphyridium forms gelatinous coatings on various surfaces; it is widespread in freshwaters, brackish environments, and moist soils.

Porphyridiales: Multicelluar Forms Chroodactylon Hansgirg (Fig. 1C) Chroodactylon is composed of false-branched pseudofilaments of globose or elliptical cells enclosed in a broad, gelatinous sheath and arranged in an irregular uniseriate manner. Each cell contains a blue– green, stellate, axial plastid and a prominent pyrenoid. The cell diameter is 3–17 µm; the length is 6–28 µm. There is a linear correlation between the false branch number and the filament length (approximately 1 branch per 200 µm length). The maximum filament length is 1240 µm. Reproduction occurs by monospores and fragmentation. No sexual reproduction has been observed. Stylonema sp. has been reported from streams in central Mexico (Jimenez, 1999) based on multiaxial pseudofilaments, but most of the figures in this presentation appear to be Chroodactylon (Jimenez’s figures 1b–d, g, h). This genus is epiphytic on Cladophora and Rhizoclonium in the Laurentian Great Lakes and scattered streams from Ontario to Arizona (Vis and Sheath, 1993). Chroodactylon is epilithic on limestone cave walls in Kentucky (Nagy 1965). It occurs in freshwaters, largely hardwater, with a specific conductance of 170–540 µS cm–1 and a pH of 7.8–8.5. Chroothece Hansgirg in Wittrock et Nordstedt (Fig. 1B) Chroothece is composed of ellipsoidal to cylindrical cells, each with a broad, firm gelatinous envelope. The cell diameter is 20–30 µm; the length is 30–45 (50) µm. Cells contain an axial, stellate, blue–green to yellow–brown or orange chloroplast with a prominent pyrenoid. Cells are solitary or joined pole to pole into a few-celled colony. The basal pole, with a gelatinous sheath, extends into a lamellated stalk. Reproduction

5. Red Algae

FIGURE 1 Freshwater bangiophycidean algae. (A) Porphyridium purpureum, unicellar form with axial stellate chloroplast with peripheral lobes (arrowheads). (B) Chroothece sp., elongated cells enclosed by a layered gelatinous stalk (arrowheads). (C) Chroodactylon ornatum, pseudofilamentous form with false branches (arrowheads). (D, E) Kyliniella latvica: (D) Pseudofilament extending from a discoid base (arrowhead). (E) Rhizoidal outgrowths (arrowheads) from upright cells. (F, G) Bangia atropurpurea: (F) Mature filament, which is biaxial toward the base and multiaxial at the apex (arrowhead). (G) Release of monospores (arrowhead) by localized digestion of matrix at apex. (H) Boldia erythrosiphon, macroscopic view showing saccate thallus. C–H reprinted from Sheath (1984) with permission.

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occurs by cell division; the pyrenoid divides and then the cell undergoes transverse invagination. This species also forms resting akinetes. No sexual reproduction has been reported. Chroothece is a rare component of freshwater streams, moist soils, and peat bogs. Populations occur in an Arizona spring at a temperature of 13°C, a pH of 7.2–7.8, and a specific conductance of 570 µS cm–1 (Blinn and Prescott, 1976). Kyliniella Skuja (Fig. 1D, E) Kyliniella is composed of unbranched pseudofilaments growing in gray–green clusters from a discoid, pseudoparenchymatous base. The maximum filament length is 2–3 cm. The cells are 18–19 µm in diameter and 5–17 µm long, surrounded by a mucilaginous envelope up to 16 µm wide. Cells are contiguous or separate, containing several parietal, blue–green discoid chloroplasts. Rhizoidal outgrowths [17–25 (150) µm long] occur at points of contact. Vegetative reproduction occurs by release of small fragments (hormogonia). Asexual reproduction occurs by monospores shed by expulsion through the sheath. Presumptive sexual reproduction has been reported in a New Hampshire population with small colorless spermatia and large, pigmented carpogonia with tubular projections. The fate of the zygote is unknown. This genus is epiphytic on macrophytes in softwater streams in the northeastern United States (Rhode Island and New Hampshire), mostly in the summer and fall (Vis and Sheath, 1993). Rufusia Wujek et Timpano Rufusia is composed of branched pseudofilaments, composed of spherical or elliptical cells, each with several reddish–violet, parietal, discoid to band-shaped chloroplasts without pyrenoids. Cells are 5.5–15 ⫻ 3.5–10 µm. Cell division is apical and intercalary. Asexual reproduction occurs by endospores; vegetative propagation occurs by fragmentation. Sexual reproduction has not been observed. Rufusia grows within the hair tissues and furrows of the two-toed sloth (Choloepus) and three-toed sloth (Bradypus) in Panama and Costa Rica (Wujek and Timpano, 1986). It is not found on adjacent vegetation.

Bangiales Bangia Lyngbye (Figs. 1F, G) Bangia is composed of filiform, unbranched cylinders of cells embedded in a firm gelatinous matrix. It is attached by down-growing rhizoids, usually in dense purple–black to rust-colored clumps. The initial uniaxial filament (diameter 10–30 µm) becomes largely multiaxial at maturity (the diameter is 60–6180 µm for

freshwater filaments). The cell number and filament length are highly correlated in uniaxial filaments; filament lengths range from 0.2 to 35 cm. Vegetative cells contain a large, axial, stellate chloroplast with prominent pyrenoid. The apical region differentiates into packets of cells, the monosporangia. This genus occurs in hard-water lakes in North America, particularly Lakes Ontario, Erie, Huron, Michigan, and Simcoe, as well as the upper St. Lawrence River (Müller et al., 1998).

Compsopogonales Boldia Herndon (Figs. 1H and 2A) This genus occurs as a mauve–pink to reddishbrown (rarely olive green), hollow, monostromatic sac or tube, 1–20 (40–75) cm long and 0.1–2.0 cm in diameter. Vegetative cells are rectangular, 5–20 (45) µm in diameter, containing several peripheral, ribbon-like chloroplasts and a large central vacuole. Secondary filaments arise as outgrowths from vegetative cells, elongating between and above vegetative cells and eventually dividing to form monospores, 5–9 µm in diameter. Monospores germinate into a prostrate, monostromatic disc or an aggregation of creeping filaments. The disc or aggregation produces a cushionlike mound of cells that functions as a perennial holdfast, producing seasonally macroscopic thalli. Sexual reproduction has not been observed. Boldia is localized in scattered streams in eastern North America, extending from central Alabama to Ontario and Québec. The range of ecological factors is as follows: current velocity 3–71 cm s–1;, pH, 6.1–8.5; specific conductance, 18–290 µS cm–1, dissolved oxygen, 4.5 mg L–1 (saturation); temperature, 12–25°C. In the southern range, it is often associated with snails of the family Pleuroceridae with high manganese content in the shells, appearing in late winter and usually disappearing by early summer. In northern streams, it is largely epilithic, occurring throughout the summer (Howard and Parker, 1980; Sheath and Hymes, 1980). Compsopogon Montagne in Bory and St. Vincent et Durieaux (Fig. 2B–D) Compsopogon is composed of a branched, bluish to violet–green, uniaxial filament with older portions corticated. The small-celled cortex is produced by vertical division of axial cells into one to five layers. Plants are up to 20–50 cm long and 250–2000 µm in diameter. Axial cells are enlarged and are evident by slight constrictions in the older portions. Axial cells may break down, leaving hollow cylinders. This genus may be free floating or benthic. If attached, rhizoids form by outgrowths of lower cortical cells or a basal disc. Cortical cells 7–22 ⫻ 10–48 µm, contain several

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FIGURE 2 Freshwater bangiophycidean and acrochaetalian algae. (A) Boldia erythrosiphon, thallus surface with large vegetative cells and smaller monosporangia (arrowheads) between and above them. (B–D) Compsopogon coeruleus: (B) Branched uniaxial filament covered by small cortical cells. (C) Transverse section of mature thallus with hollow center resulting from deterioration of axial filament stained with Toluidine Blue O. Monosporangia are cleaved from the cortical cells and are evident as small, external cells (arrowheads). (D) Surface view with smaller, more densely pigmented monosporangia (arrowheads). An empty monosporangium (double arrowhead) that has released its contents is also included. (E) Compsopogonopsis leptocladus, portion of branched filament showing rhizoidal cortication (arrowheads). (F–J) Audouinella: (F) A. hermannii, macroscopic view of tufts (arrowheads) that are epiphytic on Lemanea. (G) A. eugenea with monosporangium (arrowhead) on short lateral branch. (H–J) A. hermannii: (H) Spermatangial clusters (arrowheads) at the tips of short lateral branches. (I) Carpogonium with cylindrical base (arrowhead), narrow trichogyne (double arrowhead), and attached spermatium (triple arrowhead). (J) Carposporophyte, a dense mass of gonimoblast filaments with obovoid carposporangia of branch tips (arrowheads). A, B, and F reprinted from Sheath (1984) with permission. C courtesy of T. Rintoul. G, I, and J reprinted from Necchi et al. (1993a) with permission.

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peripheral, discoid chloroplasts. Reproduction occurs by fragmentation or monosporangia (9–28 µm long), which are cleaved from cortical cells by oblique, unequal cell division. The monospore divides into creeping, branched filament; a central cell eventually elongates vertically and divides to form the erect stage. Microaplanospores reported may represent spermatia but this is not confirmed. Compsopogon is largely distributed in tropical to warm temperate streams; in North America, welldocumented collections range from Virginia and Texas to Belize and the Caribbean islands (Vis et al., 1992). Streams tend to be warm (13–27°C) and alkaline (pH, 7.3–8.6; specific conductance, 46–1880 µS cm–1). Height, diameter, monosporangium number, branching, and basal disc presence are affected by current velocity. It is usually epilithic but can be epiphytic. C. coeruleus is epizoic on the parasitic copepod Lernaea, where it is attached to cyprinid fish in Mud River, Kentucky (Camburn and Warren, 1983). Compsopogon was recently found at a depth of 21 m in central Lake Huron of the Laurentian Great Lakes (Manny et al., 1991). Compsopogonopsis Krishnamurthy (Fig. 2E) Compsopogonopsis is composed of a branched, bluish to olive-colored, uniseriate filament with older parts corticated. The cortex is initiated by rhizoid-like outgrowths, generally one layer but occasionally two or more. Thalli are 20–40 cm long and 0.2–1.0 mm in diameter. Cortical cells, 22–51 ⫻ 19–55 µm, contain several peripheral, discoid chloroplasts. Monosporangia are produced by cortical and uncorticated axial cells. Monospores (16–23 ⫻ 15–23 µm) germinate into a few-celled mass, one cell of which divides into the erect filament and the others contribute to the holdfast. Sexual reproduction has not been observed. The genus is considered by some researchers to be synonymous with Compsopogon (e.g., Rintoul et al., 1999). Compsopogonopsis occurs in scattered freshwater streams, which are typically warm (18–24°C) and alkaline (pH, 7.6–7.7; specific conductance, 1760–1880 µS cm s–1). North American collections are restricted to New Mexico and Puerto Rico (Vis et al., 1992).

Acrochaetiales Audouinella Bory de St. Vincent (Figs. 2F–J and 3A) Audouinella is composed of short, branched, uniaxial filaments, which typically grow in dense tufts, usually less than 1 cm in diameter but up to 2–3 cm. The filaments may be composed of erect and prostrate axes. Apices of erect axes often terminate with colorless hair cells. Cells contain either reddish or bluish, parietal, ribbon-like chloroplasts. The cell diameter is 6–26 µm. Filaments occur most commonly with mono-

sporangia (5–38 µm in diameter) at the branch tips. Only reddish species have been observed to be sexual or tetrasporic. Colorless spermatangia (4–5 µm in diameter) occur in clusters at the branch tips; carpogonia have a cylindrical base and thin trichogyne (30 µm in length). Carposporophytes are spherical, compact mass of short gonimoblast filaments; carposporangia are obovoid (10 ⫻ 13 µm). Tetrasporangia are also formed at the branch tips (9 µm in diameter). Audouinella is a widespread genus in streams, ranging from the North Slope of Alaska to Costa Rica (Necchi et al., 1993a, b). A. hermannii, the most common species in North America, tends to occur in cool waters (11°C), with a low ion content (104 µS cm–1) and mildly alkaline pH (7.5). A. eugenea and A. pygmaea are typical of warm streams of high ion content.

Balbianiales Rhododraparnaldia Sheath, Whittick, et Cole (Fig. 3B–E) This genus is composed of crimson-colored filaments up to 15 cm long with barrel-shaped axial cells that have a distinctly larger diameter (17.3–30.1 µm) than that of the lateral branches (4.3–8.5 µm). Unique spermatangial stalks produce two types of spermatangia at their tips. The carpogonium is borne on an undifferentiated branch and has a swollen, cylindrical base and thin trichogyne. The carposporophyte consists of a spherical, compact mass of short gonimoblast filaments. The carposporangia are spherical up to 8 µm in diameter. Carpospores germinate into a “chantransia” phase with cells 5–7 ⫻ 16–38 µm; this phase produces gametophytes. DAPI-relative fluorescence values are twice as high for gonimoblast cells, carposporangia, and “chantransia” cells as for the gametophyte vegetative cells and gametangia. The single species R. oregonica combines characteristics of both the Acrochaetiales and the Batrachospermales. Rhododraparnaldia is found in two mountain streams in Oregon; the type locality has moderate current velocity (35–61 cm s–1), temperatures of 8–11°C, a pH of 8.3, and a specific conductance of 30 µS cm–1 (Sheath et al., 1994d).

Batrachospermales: Batrachospermaceae Batrachospermum A.W. Roth (Figs. 3F–M and 4A–C) Batrachospermum is composed of gelatinous gametophyte filaments, up to 40 cm long, with beaded appearance, varying from blue–green, olive, violet, gray to brownish. It contains a uniaxial central axis with large cylindrical cells; four to six pericentral cells produce repeatedly branched fascicles of limited growth. Rhizoid-like cortical filaments typically develop from

FIGURE 3 Freshwater members of the Acrochaetiales, Balbianiales, and Batrachospermaceae. (A) Audouinella hermannii, tetrasporangium (arrowhead) at the tip of a vegetative branch. (B–E) Rhododraparnaldia oregonica: (B) Branched filament with lateral branches, both opposite and alternate, arising from large barrel-shaped axial cells. (C) Carpogonia with cylindrical bases, thin trichogynes (arrowhead), and attached spermatia (double arrowhead). Spermatangia (triple arrowheads) are formed at the apex of colorless stalk cells. (D) Carposporophyte, consisting of compact gonimoblast filaments and carposporangia (arrowheads) at branch tips. (E) “Chantransia” phase (arrowhead) producing an attached gametophyte (double arrowhead). (F–M) Batrachospermum: (F) B. gelatinosum, branched filament with barrel-shaped whorls containing several carposporophytes (arrowheads). (G) B. ambiguum, branched filament with obovoid whorls and single axial carposporophytes (arrowheads). (H) B. louisianae, branched filament with confluent whorls and large carposporophytes (arrowheads) extending beyond whorls. (I) B. intortum, with monosporangia (arrowheads) at tips of vegetative branches. (J) B. gelatinosum forma spermatoinvolucrum, spermatangia (arrowhead) and hair cells (double arrowheads) at the tips of vegetative branches. (K) B. anatinum, with carpogonium with inflated trichogyne (arrowhead) with four attached spermatia (double arrowhead). The carpogonial branch is little differentiated from typical vegetative branches. (L) B. helminthosum with immature carpogonium (arrowhead). The carpogonial branch (double arrowhead) is composed of cells that are substantially smaller than those of adjacent vegetative branches. (M) B. intortum with immature carpogonium (arrowhead). The carpogonial branch (double arrowhead) is differentiated and twisted. A reprinted from Necchi et al. (1993a) with permission. B–E reprinted from Sheath et al. (1994d) with permisson of the International Phycological Society. F reprinted from Vis et al.(1996a) with permission. G–I and M reprinted from Sheath et al. (1992) with permission of the Journal of Phycology. J reprinted from Vis and Sheath (1996) with permission of the International Phycological Society. K reprinted from Vis et al. (1996b) with permission. L reprinted from Sheath et al. (1994a) with permission of the Journal of Phycology.

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FIGURE 4 Members of the Batrachospermales. (A–C) Batrachospermum: (A) B. gelatinosum with a dense, spherical carposporophyte (arrowhead). (B) B. globosporum with loose carposporophyte with carposporangia (arrowheads) at tips of gonimoblast filaments. (C) B. gelatinosum “chantransia” stage (arrowhead) producing attached gametophyte (double arrowhead). (D–F) Sirodotia suecica: (D) Branched filament with obovoid whorls. (E) Carpogonium with base (arrowhead) with a protuberance, cylindrical trichogyne (double arrowhead), and attached spermatium (triple arrowhead). (F) Carposporophyte with indeterminate gonimoblast filament (arrowhead), and carposporangia (double arrowhead) at the tips of short lateral branches. (G–J) Tuomeya americana: (G) Macroscopic view of a mature plant, which is a well-branched, cartilaginous tube. (H) Branch apex showing confluent whorls. (I) Carpogonium with small base (arrowhead) and large trichogyne (double arrowhead) perpendicularly attached to a stalk. (J) Carposporophyte with prominent carposporangia (arrowheads) at the tips of short gonimoblast filaments. (K) Lemanea fluviatilis, macroscopic view showing thallus, which is tubular, with inflated nodes, branches, and a stalk. A reprinted from Vis et al. (1996a) with permission. B reprinted from Sheath et al. (1992) with permission of the Journal of Phycology. D–F reprinted from Necchi et al. (1993c) with permission of the Journal of Phycology, G, H, and K reprinted from Sheath (1984) with permission. I and J reprinted from Kaczmarczyk et al. (1992) with permission of the Journal of Phycology.

5. Red Algae

the lower side of the pericentral cells. Cortical filaments grow downward and ensheath axial cells, often producing secondary fascicle branches. Each fascicle cell contains several, ribbon-like, parietal chloroplasts with no pyrenoid. Few species form monosporangia in the gametophyte stage. Spermatangia bud off from terminal primary and secondary fascicle cells or in some species from involucral filaments of the carpogonial branch; they are spherical, colorless, and 4–8 µm in diameter. Carpogonial branches range from little modified to well differentiated with a twist in the section Contorta. Carpogonia with broad trichogynes are sometimes stalked on a small base containing the nucleus. Carposporophytes are generally a spherical or semispherical, compact or loose mass of gonimoblast filaments; carposporangia form at the apices. Carpospores germinate into the “chantrasia” stage, a crustose growth consisting of large basal cells and erect,

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the sparsely branched filaments. Filaments can form monosporangia or divide meiotically, producing an attached gametophyte and two residual cells. Batrachospermum is a cosmopolitan genus occurring in moderately flowing, reasonably unpolluted streams (Sheath, 1984). B. turfosum is also common in bogs. Both euryphotic (e.g., B. gelatinosum) and shade or brown-water (e.g., B. turfosum) species exist. Filament fragments are common in the guts of grazing amphipods, insect larvae, and snails. Batrachospermum section Contorta is mostly concentrated in tropical to subtropical regions. B. gelatinosum, the only widespread species in tundra, tolerates a large range of conditions: temperature, 0–24°C; current velocity, 7–181 cm s–1; pH, 4.1–8.2; specific conductance, 10–360 µS cm–1; PO43– < 1–4900 µg L–1 (Sheath and Hambrook, 1990).

Key to the North American Sections of Batrachospermum 1a.

Carpogonial branch with little or no differentiation from fascicles (Fig. 3K)...........................................................Batrachospermum

1b.

Carpogonial branch well differentiated..............................................................................................................................................2

2a.

Carpogonial branch almost as elongate as fascicles................................................................................................................Aristata

2b.

Carpogonial branch much shorter than fascicles (Fig. 3L, M)............................................................................................................3

3a.

Carpogonial branch with one or more well-developed twists (Fig. 3M).................................................................................Contorta

3b.

Carpogonial branch straight or with slight curves..............................................................................................................................4

4a.

Carpogonial branches a combination of straight and curved in same filament.......................................................................Hybrida1

4b.

Carpogonial branch always straight (Fig. 3L).....................................................................................................................................5

5a.

Mean mature whorl (with carposporophytes) diameter < 170 µm, mean fascicle cell number, 300 µm; mean fascicle cell number, ≥ 7........................................................................................................6

6a.

Stalked trichogyne...............................................................................................................................................................Virescentia

6b.

Sessile trichogyne......................................................................................................................................................................Turfosa

1

Vis and Entwisle (2000) proposed sinking this section into Contorta based on the positioning of Batrachospermum virgato-decaisneanum in rbcL gene trees.

Sirodotia Kylin (Fig. 4D–F) Sirodotia is composed of attached, gelatinous gametophytic filaments, up to 17 cm long, with a beaded appearance varying from blue–green to yellow–green. It contains a uniaxial central filament with large, cylindrical cells; four to six pericentral cells produce repeatedly branched fascicles of limited growth. In most species, rhizoid-like cortical filaments develop from the lower side of the pericentral cells. Each fascicle cell contains several, ribbon-like, parietal chloroplasts with no pyrenoid. Spermatangia bud off from terminal fascicle cells; they are spherical, colorless, and 4–7 µm in diameter. Carpogonial branches are somewhat differentiated with small cells. Carpogonia with broad trichogyne are attached off-center to the base,

the latter structure having a definite protrusion. The carposporophyte occurs as a branched indeterminant filament creeping along the main axis; carposporangia form at branch apices. Carpospores germinate into the “chantransia” stage, composed of branched, uniaxial filaments. Meiosis and monosporangia have not been observed. Sirodotia was considered to be a section of Batrachospermum by Necchi and Entwisle (1990), but this was questioned by Necchi et al. (1993c). Sirodotia occurs in scattered, small, typically softwater (pH, 5.7–7.6; specific conductance, 10–140 µS cm–1) streams in boreal to tropical environments; in North America, species range from northern Quebec and Newfoundland to central Mexico (Necchi et al., 1993c). The most widespread species, S. suecica, has

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little increase in drag with increasing current velocity (20–80 cm s–1) due to branch reconfiguration. The second species, S. huillensis, is apparently restricted to arid, southwestern habitats (Vis and Sheath, 1999). Tuomeya W. H. Harvey (Fig. 4G–J) Tuomeya has a densely branched, cartilaginous, and cylindrical gametophytic thallus, 1–5 (6.5) cm long. It ranges in color from blue–green to olive to black. The uniaxial filament is covered by two or three layers of cortical filaments; dense laterals arising from approximately six pericentral cells are of limited growth with outer cells fused. Axial cells are evident in mature branches by constrictions. Spermatangia form at the tips of the laterals. Carpogonia are asymmetrical with an irregularly broadened trichogyne attached obliquely or perpendicularly to a stalk and borne on a curved carpogonial branch derived from a pericentral cell. The carposporophyte occurs as a globular mass of filaments. Carpospores germinate into the branched, uniseriate “chantransia” stage in culture, but in situ gametophytes develop from an undifferentiated mass of cells. Tuomeya was considered to be a section of Batrachospermum by Necchi and Entwisle (1990), but this was questioned by Kaczmarczyk et al. (1992). Tuomeya occurs in scattered freshwater streams in eastern North America, from Florida to Newfoundland, from fall to early summer. The range of conditions is as follows: temperature, 5–26°C; current velocity, 16–125 cm s–1; pH, 4.7–7.6; specific conductance, 10–124 µS cm–1 (Kaczmarczyk et al., 1992). It tolerates considerable stress before breaking (1780 ± 850 kN m–2 ), stretching 22% in the process. Tuomeya is common in the guts of grazing amphipods and insect larvae; its cartilaginous structure and low protein content make it a little preferred food source.

Batrachospermales: Lemaneaceae Lemanea Bory de St. Vincent (Figs. 4K and 5A, B) Lemanea is composed of tufts of cartilaginous, tubular, pseudoparenchymatous gametophytic thalli, lacking cortical filaments, around a central uniseriate axis. T- or L-shaped ray cells are closely applied to the outer cortex. It is 1–40 cm long and 0.2–2.0 mm in diameter. It is blue–green to olive when young, becoming rusty-brown to black at maturity. Species are characterized by the presence of branching, by their diameter, and by the degree of basal constriction. Several parietal discoid chloroplasts occur in the outer cells only. Spermatangia develop as yellowish circular patches. Nearby small carpogonial branches are entirely internal except for a thin trichogyne that protrudes beyond the outer cell layer. Carposporophytes are microscopic, spherical masses of filaments forming

large, ellipsoidal carpospores within the central cavity; they are released by thallus deterioration and germinate into branched, uniseriate “chantransia” filaments. The “chantransia” stage produces attached gametophytes seasonally after meiosis. This genus is widespread in temperate and boreal streams and rivers with typically high current velocities (up to 2 m s–1) (Sheath and Hambrook, 1990). Species have adapted to flow by developing dense turfs closely adherent to rocks and high breaking stress (910 ± 430 kN m–2 for L. fluviatilis). Lemanea is common at high elevations up to 1200 m. L. fluviatilis tolerates a temperature of 4–25°C, a pH of 4.1–8.2, and a specific conductance of 10–300 µS cm–1. It occurs at low nutrient concentrations. In a Rhode Island river, growth and reproduction of L. fluviatilis gametophytes are confined to April–August, after which the thallus deteriorates and carpospores are released; between September and March, remnants persist. Paralemanea Vis et Sheath (Fig. 5C–F) Paralemanea is composed of tufts of cartilaginous, tubular, pseudoparenchymatous gametophytic thalli, with cortical filaments, around a central, axial filament and simple ray cells not abutting outer cortical cells. The mean length is 4.3–9.5 cm and the diameter is 0.5–0.7 mm. Species are characterized by the presence of branching, by their length, and by their diameter. Several parietal discoid chloroplasts occur in the outer cells only. Spermatangia develop as yellowish to brownish nodal rings. Small carpogonial branches are entirely internal except for a thin trichogyne that protrudes beyond the outer layer. Carpsoporophytes are small, spherical masses of filaments forming large, ellipsoidal carpospores in the central cavity; they are released by thallus deterioration and germinate into branched, uniaxial “chantransia” filaments. The “chantransia” stage produces attached gametophytes seasonally after meiosis. Most populations of Paralemanea in North America are from the southeastern United States and northern California, but extend from central Mexico to New York (Vis and Sheath, 1992). This genus occurs under a wide range of conditions: mean current velocity, 18–110 cm s–1; temperature, 4–17°C; pH, 5.5–8.6; specific conductance, 42–500 µS cm–1. It is predominant in cool seasons in a North Carolina stream (Everitt and Burkholder, 1991).

Thoreales Recent molecular sequence analyses have demonstrated that this family is not closely aligned with the Batrachospermaceae and Lemaneaceae and should be removed from the Batrachospermales (e.g., Vis et al.,

5. Red Algae

FIGURE 5 Members of the Lemaneaceae and Thoreales. (A, B) Lemanea: (A) L. fluviatilis with nodes obvious as a series of swellings will have patches of spermatangia (arrowheads). (B) Longitudinal Toluidine Blue O–stained longitudinal section showing outer cortical layer (arrowhead) and a central strand of carpospores (double arrowhead). (C–F) Paralemanea: (C) P. mexicana, macroscopic view with whorled branching (arrowhead), rebranching (double arrowhead), and tubular construction. (D) Paralemanea sp. with an obvious spermatangial ring (arrowhead). (E) P. mexicana, Toluidine Blue O–stained longitudinal section showing outer cortical layer (arrowhead), axial filament (double arrowhead), which is surrounded by inner cortical filaments (triple arrowhead). (F) P. mexicana, Toluidine Blue O–stained section showing carposporophyte (arrowhead) and large carpospores (double arrowhead). (G–I) Nemalionopsis tortuosa: (G) Branched, multiaxial filament. (H) Section of thallus showing colorless, central medulla (arrowhead) and photosynthetic, assimilatory branches (double arrowhead). (I) Monosporangia (arrowheads), which are formed at the tips of assimilatory filaments. A reprinted from Sheath (1984) with permission. B, E, and F reprinted from Sheath et al. (1996b) with permission. C and D reprinted from Vis and Sheath (1992) with permission of the International Phycological Society. I reprinted from Sheath et al. (1993b) with permission.

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1998). We have proposed that the Thoreaceae be placed in a new order, the Thoreales (Sheath et al., 2000), Müller et al., 2002. Nemalionopsis Skuja (Fig. 5G–I) This genus contains a sparsely branched, cordlike thallus, which is burgundy to yellow–brown in color; it is composed of a central medullary region of interwoven, colorless filaments and an outer pigmented cortex of branched laterals of limited growth. The length is 5–30 (50) cm, the diameter is 700–1000 µm. It may be flattened or coiled in some portions. Monosporangia form at the outer tips of lateral filaments, 7–12 ⫻ 8–14 µm. The spore-bearing branch-tovegetative branch length ratio is greater than 0.64. Monospores germinate into prostrate filaments forming monostromatic discs from which erect filaments arise. Sexual reproduction has not been observed. Nemalionopsis is known from four freshwater streams in North America in Florida, Louisiana, and North Carolina with temperatures ranging from 13 to 22°C, current velocity of 29 cm s–1, pH of 7.1–8.3, and specific conductance of 220 µS cm–1 (Sheath et al., 1993b). Thorea Bory de St. Vincent (Fig. 6A–D) Thorea is composed of branched gametophytic filaments, up to 20–200 cm long and 0.5–3 mm in diameter. It is composed of interwoven, colorless medullary filaments and dense, photosynthetic laterals of limited growth. It is olive green to reddish to black. Chloroplasts in assimilatory filaments are parietal and ribbon-like. Monosporangia are solitary or in clusters, formed at the base of the assimilatory filaments (spore-bearing branch-to-vegetative lateral length ratio < 0.3). Sexual reproduction is known for a few species. In T. violacea, spermatangia are borne on specialized branches near the base of assimilatory filaments, colorless, elliptical or obovoid, 8–10 ⫻ 4–7 µm. Carpogonia are conical with an elongated trichogyne 5–7 µm wide; the carpogonial branch is short and located at the base of assimilatory filaments. Carposporophytes are sparsely branched and compact. Carposporangia are terminal, 9–13 ⫻ 17–25 µm; carpospores germinate into branched, uniseriate “chantransia” filaments. This genus is widespread in tropical to warm temperate, freshwater streams; in North America, it ranges from New York to Grenada (Sheath et al., 1993b). It typically occurs in alkaline waters (e.g., in North America, pH, 7.5–8.2; specific conductance, 180– 500 µS cm–1). An exception to this distribution pattern is a population in the Hudson River, New York, which occurs in temperatures ranging from 15 to 19°C,

specific conductance of 44–136 µS cm–1 and pH of 6.6–7.2 (Pueschel et al., 1995).

Hildenbrandiales Hildenbrandia Nardo (Fig. 6E–G) This genus contains a bright-red uncalcified crustose thallus, which is composed of a single basal layer that gives rise to vertical files of cells. The thallus height varies from 23 to 182 µm. The cell dimensions, 2–8 ⫻ 4–10 µm, indicate that a single species, H. angolensis, exists in freshwater habitats of North America [H. rivularis appears to be largely restricted to Europe (Sherwood and Sheath, 2000a)]. Reproduction is largely by gemmae, dense aggregation of cells formed in the thallus, which are eventually released and germinate into new crusts. Gemma production continues year round in two spring-fed streams in Texas (Sherwood and Sheath, 2000b). Although H. angolensis has been reported from Pennsylvania, well-documented collections occur in streams and springs from Texas in the north to Costa Rica in the south and throughout the Caribbean islands (Sheath et al., 1993a). These streams are mostly warm (14–27°C), alkaline in pH (7.0–8.6), and variable in current velocity (5–67 cm s–1) and specific conductance (70–1558 µS cm–1). The freshwater populations of Hildenbrandia in North America form a monophyletic clade in 18S rRNA gene trees distinct from marine populations of the genus (Sherwood and Sheath, 1999b).

Ceramiales Ballia W. H. Harvey (Fig. 6H) Ballia is composed of reddish filaments with a distinct main axis with hexagonal-shaped cells and smaller, pinnate, determinate lateral branches that may rebranch. The apical cell is typically quite long in B. prieurii (43–89 µm in length). Plants typically are small (3–15 mm). They can reproduce by the production of monosporangia. Choi et al. (2000) have proposed that marine species of this genus from Australia are not members of the Ceramiales, based on the presence of two pit plug cap layers and positioning in 18S rRNA gene trees. However, they were unable to resolve the status of the freshwater species, B. prieurii. Three freshwater collections have been made in North America from Belize and Costa Rica, with slow to moderate current velocities (1–65 cm s–1), pH of 7.6–7.8, specific conductance of 50–100 µS cm–1, and temperatures of 19–22°C (Sheath et al., 1993c). Bostrychia Montagne (Figs. 6I and 7A, B) Bostrychia is composed of dark-reddish filaments with tiers of pericentral cells around the axial cells; the

FIGURE 6 Freshwater members of the Thoreales, Hildenbrandiales, and Ceramiales. (A–D) Thorea: (A) T. hispida, branched, multiaxial filament, with an obvious outer, assimilatory layer (arrowhead) and inner medullary layer (double arrowhead). (B) T. violacea, transverse section showing colorless, central medulla (arrowhead) and outer, assimilatory filaments (double arrowhead). (C) T. violacea with monosporangium (arrowhead) on short branch at the base of assimilatory filament. (D) T. violacea carpogonium with thin, elongate trichogyne (arrowhead). (E–G) Hildenbrandia angolensis: (E) Toluidine Blue O–stained transverse section showing rows of erect filaments. (F) Scanning electron micrograph of thallus surface with gemmae (arrowhead) formed in cavities. (G) Released gemma with basal rhizoids for attachment (arrowhead). (H) Ballia prieurii, apex showing hexagonal axial cells and pinnate lateral branches that rebranch (arrowhead). (I) Bostrychia moritziana, apex with uniaxial branch tip, mature axis covered with pericentral cells (double arrowhead) and stichidium (triple arrowhead). A and D reprinted from Sheath et al. (1993b) with permission. B and C reprinted from Sheath (1984) with permission. E and F courtesy of A. Sherwood. H and I reprinted from Sheath et al. (1993c) with permission of the Journal of Phycology.

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FIGURE 7 Freshwater members of the Ceramiales. (A, B) Bostrychia: (A) B. moritziana hapteron, specialized branch with colorless rhizoids (arrowhead) at tip. (B) B. tenella with an additional layer of cortication (arrowhead) outside of pericentral cells. (C–E) Caloglossa: (C) C. leprieurii, macroscopic view of flat blades with subdichotomous branching, constrictions at nodes (arrowhead), and obvious midrib (double arrowhead). (D) C. leprieurii showing cortical filaments of midrib area (arrowhead) and rhizoids (double arrowhead). (E) C. ogasawaerensis with new branch (arrowhead) and rhizoids arising at node. (F, G) Polysiphonia subtilissima: (F) Apex with axis surrounded by pericentral cells and trichoblast (arrowhead). (G) Rhizoid (arrowhead) arising from pericentral cell. A–G reprinted from Sheath et al. (1993c) with permission of the Journal of Phycology.

thallus may become additionally corticated to the outside of the pericentral cells (e.g., B. tenella). Specialized rhizoidal branches, the haptera, attach filaments to the substrata. Vegetative branching is bilateral; branches near the apex tend to incurve and there may be both long and short shoots. Only tetrasporangia have been observed in freshwater collections, which are formed in inflated, multichambered structures termed stichidia. Freshwater populations in North America appear to be restricted to streams in the Caribbean islands which are warm (21–26°C), alkaline (pH 7.0–8.4), and range in specific conductance (56–440 µS cm–1) (Sheath et al., 1993c).

Caloglossa J. Agardh (Fig. 7C–E) Caloglossa is composed of flat, dichotomously branched reddish blades with constrictions. A prominent midrib is evident and is composed of a broad axial row of cells surrounded by a cortex of elongated cells. The outer portions of the blade are monostromatic with an oblique series of hexagonal cells. Rhizoids arise at constrictions, either from the midrib area or from the peripheral layer of cells. Population spread in freshwaters is vegetative, although gametophytic and tetrasporic plants have been collected in brackish waters. Two species, C. lepreurii and C. ogasawarensis, have been collected from streams in Puerto Rico and

5. Red Algae

Costa Rica, respectively. Current velocities are moderate (33–43 cm s–1), temperature warm (23–24°C), pH alkaline (7.6–8.4), with a specific conductance of 100–200 µS cm–1 (Sheath et al., 1993c). Polysiphonia Greville (Fig. 7F, G) Polysiphonia is composed of dark-reddish filaments with a single tier of pericentral cells around the axial cell. No freshwater collections have an additional layer of cortication but a few marine species do. Delicately branched hairs (trichoblasts) are formed in upper portions of the plant. Rhizoidal branches arise from pericentral cells. Freshwater samples have not been observed as being either sexual or tetrasporic. This genus is common in marine and brackish habitats; only two populations of P. subtilissima have been collected in North American freshwaters in Florida and Jamaica (Sheath et al., 1993c). These streams have moderate flow (25 cm s–1), warm temperature (22–26°C), alkaline pH (7.7–7.8), and high specific conductance (1150–1840 µS cm–1).

VI. GUIDE TO LITERATURE FOR SPECIES IDENTIFICATION The following is a list of key North American references, each of which contains citations to older literature and those from other continents: 1. Audouinella—Necchi et al. (1993a, b) 2. Ballia—Sheath et al. (1993c) 3. Bangia—Sheath and Cole (1984), Müller et al. (1998) 4. Batrachospermum—Sheath et al. (1992, 1993d, 1994a–c), Sheath and Vis (1995), Vis et al. (1996a, b), Vis and Sheath (1996, 1997, 1998), Müller et al. (1997) 5. Boldia—Howard and Parker (1980), Sheath and Hymes (1980), Rintoul et al. (1999) 6. Bostrychia—Sheath et al. (1993c) 7. Caloglossa—Sheath et al. (1993c) 8. Chroodactylon—Vis and Sheath (1993) 9. Chroothece—Blinn and Prescott (1976) 10. Compsopogon - Vis et al. (1992), Rintoul et al. (1999) 11. Compsopogonopsis—Vis et al. (1992), Rintoul et al. (1999) 12. Cyanidium—Seckbach (1991) 13. Flintiella—Ott (1976) 14. Hildenbrandia—Sheath et al. (1993a), Sherwood and Sheath (1999b, 2000b) 15. Kyliniella—Vis and Sheath (1993) 16. Lemanea—Vis and Sheath (1992), Sheath et al. (1996b)

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17. Nemalionopsis—Sheath et al. (1993b) 18. Paralemanea—Vis and Sheath (1992), Sheath et al. (1996b) 19. Polysiphonia—Sheath et al. (1993c) 20. Porphyridium—Ott (1972) 21. Rhododraparnaldia—Sheath et al. (1994d) 22. Rufusia—Wujek and Timpano (1986) 23. Sirodotia—Necchi et al. (1993c), Vis and Sheath (1999) 24. Thorea—Sheath et al. (1993b), Vis et al. (1998), Müller et al. (2002) 25. Tuomeya—Kaczmarczyk et al. (1992)

ACKNOWLEDGMENTS I thank everyone who has worked on freshwater red algae in my laboratory and contributed to much of the knowledge summarized in this chapter, including JoAnn Burkholder, Lesley Campbell, Tracey Carlson, Wayne Chiasson, Dana Couture, Julie Hambrook, Bev Hymes, Don Kaczmarczyk, Judith Korch, Mollie Morison, Kirsten Müller, Orlando Necchi, Jr., Tara Rintoul, Troina Shea, Alison Sherwood, Al Steinman, Stacey Thompson, Kathy Van Alstyne, and Morgan Vis. In addition, collaborators from other laboratories and institutions have been very helpful in the various studies reported, such as Murray Colbo, Dave Larson, Tim Entwisle, Gary Saunders, and Alan Whittick. A special thank you to Kay Cole whose long-time collaboration has been an inspiration. Many collecting trips were made more enjoyable with the support and help of Mary Koske. Assistance in manuscript preparation from Toni Pellizzari and helpful reviews by Orlando Necchi, Jr., Morgan Vis, and John Wehr are also appreciated. Research support from NSERC, NSF, Universities of Guelph and Rhode Island, PCSP, and the Northern Training Program is gratefully acknowledged.

LITERATURE CITED Blinn, D. W., Prescott, G. W. 1976. A North American distribution record for the rare Rhodophyceae, Chroothece mobilis Pascher and Petrova. American Midland Naturalist 96:207–210. Broadwater, S. T., Scott, J. L. 1994. Ultrastructure of unicellular red algae, in: Seckbach, J., Ed., Evolutionary pathways and enigmatic algae: Cyanidium caldarium. Kluwer Academic, Dordrecht, pp. 215–230. Brown, D. L., Weier, T. E. 1968. Chloroplast development and ultrastructure in the freshwater red alga Batrachospermum. Journal of Phycology 4:199–206. Camburn, K. E., Warren, M. L., Jr. 1983. Epizoic occurrence of Compsopogon coeruleus (Rhodophyta) on Lernaea (Copepoda) from Kentucky fishes. Canadian Journal of Botany 61:3545–3548.

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Choi, H.-G., Kraft, G. T., Saunders, G. W. 2000. Nuclear smallsubunit rDNA sequences from Ballia spp. (Rhodophyta): Proposal of the Balliales ord. nov., Balliaceae fam. nov., Ballia nana sp. nov. and Inkyuleea gen. nov. (Ceramiales). Phycologia 39:272–287. Couté, A., Sarthou, C. 1990. Révision des espèces d’eau douce du genre Ballia (Rhodophytes, Céramiales). Cryptogamie Algologie 11:265–279. Craigie, J. S. 1990. Cell walls, in: Cole, K. M., Sheath, R. G., Eds., Biology of the red algae. Cambridge Univ. Press, Cambridge, UK, pp. 221–257. D’Lacoste, V., Ganesan, E. K. 1987. Notes on Venezuelan freshwater algae I. Nova Hedwigia 45:263–281. DeLuca, P., Gambardella, R., Merola, A. 1979. Thermoacidophilic algae in North and Central America. Botanical Gazette 140:418–27. Doemel, W. N., Brock, T. D. 1971. The physiological ecology of Cyanidium caldarium. Journal of General Microbiology 67:17–32. Drew, K. M. 1935. The life history of Rhodochorton violaceum (Kütz.) comb. nov. (Chantrasia violacea Kütz.). Annals of Botany 49:439–450. Everitt, D. T., Burkholder, J. M. 1991. Seasonal dynamics of macrophyte communities from a stream flowing over granite flatrock in North Carolina, USA. Hydrobiologia 222:159–172. Flint, L. H. 1970. Freshwater red algae of North America. Vantage Press, New York, 110 p. Fritsch, F. E. 1929. The encrusting algal communities of certain fast-flowing streams. New Phytologist 28:165–196. Gantt, E., Scott, J., Lipschultz, C. 1986. Phycobiliprotein composition and chloroplast structure in the freshwater red alga Compsopogon coeruleus (Rhodophyta). Journal of Phycology 22:480–484. Garbary, D. J., Hansen, G. I., Scagel, R. F. 1980. A revised classification of the Bangiophyceae (Rhodophyta). Nova Hedwigia 33:145–166. Geitler, L. 1932. Porphyridium sordidum n. sp., eine neue Süβwasserbangiale. Archiv für Protistenkunde 76:595–604. Gibson, M. T., Whitton, B. A. 1987. Hairs, phosphatase activity and environmental chemistry in Stigeoclonium, Chaetophora and Draparnaldia (Chaetophorales). British Phycological Journal 22:11–22. Gretz, M. R., Sommerfeld, M. R., Athey, P. V., Aronson, J. M. 1991. Chemical composition of the cell walls of the freshwater red alga Lemanea annulata (Batrachospermales). Journal of Phycology 27:232–240. Hambrook, J. A., Sheath, R. G. 1987. Grazing of freshwater Rhodophyta. Journal of Phycology 23:656–662. Hambrook, J. A., Sheath, R. G. 1991. Reproductive ecology of the freshwater red alga Batrachospermum boryanum Sirodot in a temperate headwater stream. Hydrobiologia 218:233–246. Harding, J. P. C., Whitton, B. A. 1981. Accumulation of zinc, cadmium and lead by field populations of Lemanea. Water Research 15:301–319. Hoffmann, L. 1989. Algae of terrestrial habitats. Botanical Review 55:77–105. Howard, R. V., Parker, B. C. 1980. Revision of Boldia erythrosiphon Herndon (Rhodophyta, Bangiales). American Journal of Botany 67:413–422. Hymes, B. J., Cole, K. M. 1983. The cytology of Audouinella hermannii (Rhodophyta, Florideophyceae). II. Monosporogenesis. Canadian Journal of Botany 61:3377–3385. Hynes, H. B. N. 1970. The ecology of running waters. Liverpool Univ. Press, Liverpool. Israelson, G. 1942. The freshwater Florideae of Sweden. Symbolae Botanicae Upsalienses 8(1):1–134.

Jimenez, J. C. 1999. Estudio floristico (taxonomico-ecologicobiogeografico) de las rodofitas de agua dulce en la region central de Mexico. Ph.D. thesis, Universidad Nacional Autonoma de Mexico. Kaczmarczyk, D., Sheath, R. G. 1991. The effect of light regime on the photosynthetic apparatus of the freshwater red alga Batrachospermum boryanum. Cryptogamie Algologie 12: 249–263. Kaczmarczyk, D., Sheath, R. G., Cole, K. M. 1992. Distribution and systematics of the freshwater genus Tuomeya (Rhodophyta, Batrachospermaceae). Journal of Phycology 28:850–855. Korch, J. E., Sheath, R. G. 1989. The phenology of Audouinella violacea (Acrochaetiaceae, Rhodophyta) in a Rhode Island stream (USA). Phycologia 28:228–236. Kremer, B. 1983. Untersuchungen zur Ökophysiologie einiger Süsswasserrotalgen. Decheniana (Bonn) 136:31–42. Kumano, S. 1978. Notes on freshwater red algae from West Malaysia. Botanical Magazine (Tokyo) 91:97–107. Kumano, S., Necchi, O., Jr. 1987. Studies on the freshwater Rhodophyta of Brazil-5: Record of Bostrychia radicans (Montagne) Montagne f. moniliforme Post in freshwaters. Revista Brasileira de Biologia 47:437–440. Leukart, P., Hanelt, D. 1995. Light requirements for photosynthesis and growth in several macroalgae from a small soft-water stream in the Spessart Mountains, Germany. Phycologia 34:528–532. Lin, C. K., Blum, J. L. 1977. Recent invasion of a red alga Bangia in Lake Michigan. Journal of the Fisheries Research Board of Canada 34:2413–2416. MacFarlane, J. J., Raven, J. A. 1985. External and internal CO2 transport in Lemanea: Interactions with the kinetics of ribulose bisphosphate carboxylase. Journal of Experimental Botany 36:610–622. Manny, B. A., Edsall, T. A., Wujek, D. F. 1991. Compsopogon cf. coeruleus, a new benthic red alga (Rhodophyta) in the Laurentian Great Lakes. Canadian Journal of Botany 69:1237–1240. McKay, R. M. L., Gibbs, S. P. 1990. Phycoerythrin is absent from the pyrenoid of Porphyridium cruentum: Photosynthetic implications. Planta 180:249–256. Müller, K. M., Vis, M. L., Chiasson, W. B., Whittick, A., Sheath, R. G. 1997. Phenology of a Batrachospermum population in a boreal pond and its implications for the systematics of section Turfosa (Batrachospermales, Rhodophyta). Phycologia 36:68–75. Müller, K. M., Sheath, R. G., Vis, M. L., Crease, T. J., Cole, K. M. 1998. Biogeography and systematics of Bangia (Bangiales, Rhodophyta) based on the Rubisco spacer, rbcL gene, 18S rRNA gene sequences and morphometric analyses. I. North America. Phycologia 37:195–207. Müller, K. M., Sherwood, A. R., Pueschel, C. M., Gutell, R. R., Sheath, R. G. 2002. A proposal for a new red algal order, the Thoreales. Journal of Phycology 38:807–820. Nagy, J. P. 1965. Preliminary note on the algae of Crystal Cave, Kentucky. International Journal of Speleology 1:479–490. Necchi, O., Jr. 1993. Distribution and seasonal dynamics of Rhodophyta in the Preto River Basin, southeastern Brazil. Hydrobiologia 250:81–90. Necchi, O. Jr. 1995. Occurrence of the genus Ballia (Ceramiaceae, Rhodophyta) in freshwater in Brazil. Hoehnea 22:229–235. Necchi, O., Jr. 1997. Microhabitat and plant structure of Batrachospermum (Batrachospermales, Rhodophyta) populations in four streams of São Paulo State, southeastern Brazil. Phycological Research 45:39–45. Necchi, O., Jr., Entwisle, T. J. 1990. A reappraisal of generic

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6

FLAGELLATED GREEN ALGAE Hisayoshi Nozaki Department of Biological Sciences Graduate School of Science University of Tokyo Hongo, Bunkyo-ku, Tokyo 113-0033 Japan

I. II. III. IV.

Introduction Diversity and Morphology Ecology and Distribution Collection and Preparation for Identification V. Key and Descriptions of Genera A. Key to Families of the Freshwater Green Flagellates B. Polyblepharidaceae C. Haematococcaceae

I. INTRODUCTION Green algae are organisms which are characterized by having chlorophylls a and b as the major photosynthetic pigments, starch located within the chloroplast as the major storage product and flagella of the whiplash (smooth) type (e.g., Bold and Wynne, 1985). They can be also distinguished from other eukaryotic algae in having two chloroplast membranes and stellate structure in the flagellar transition region (e.g., van den Hoek et al., 1995). Members of the flagellate forms of the green algae were traditionally assigned to the order Volvocales or Chlamydomonadales of the Chlorophyceae (e.g., Smith, 1950). However, recent ultrastructural studies and molecular phylogenetic analyses of the green plants (green algae and land plants) suggest that some of the flagellated green algae should be classified in the Prasinophyceae or Pedinophyceae (see Norris, 1980; Moestrup, 1991; Friedl, 1997). Therefore, members of the flagellated green algae described in this chapter belong to the Chlorophyceae Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

D. Chlamydiomonadaceae E. Phacotaceae F. Volvocaceae G. Goniaceae H. Tetrabaenaceae I. Spondylomoraceae VI. Guide to Literature for Species Identification Literature Cited

or Prasinophyceae, based on the current concepts of taxonomy/phylogeny of green algae (e.g., van den Hoek et al., 1995; Friedl, 1997). Although the flagellated chlorophycean algae can be divided into unicellular and colonial types of organization (Chlamydomonadales vs. Volvocales; see Mattox and Stewart, 1984), recent molecular phylogenetic analyses indicate that the colonial organisms represent polyphyletic status within the Volvocales (Buchheim et al., 1994). Volvox is the organism which was first observed with a light microscope by van Leeuwenhoek (1700) within the Volvocales, and the generic name Volvox was established by Linnaeus (1758).

II. DIVERSITY AND MORPHOLOGY About 100 genera with more than 1000 species are included within the freshwater green flagellates (Ettl, 1983). Among these taxa, 44 genera are known from North America. Chlamydomonas is the most diverse 225

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genus, with more than 400 described species worldwide (Ettl, 1983) and more than 20 species in North America. The Volvocales usually have cell walls, loricae, or gelatinous matrices. The main component of the cell walls is glycoprotein, rather than cellulose (Harris, 1989). Ultrastructure of the cell walls reveals their median, tripartite structure. Such a structure is fundamentally recognized in the gelatinous matrices of the green flagellates, such as the colonial alga Pandorina and Volvox (Fulton, 1978; Kirk et al., 1986). The vegetative cells have two or sometimes four or eight equal flagella of the whiplash type at the anterior end, a single nucleus in the center, and two contractile vacuoles generally located near the base of the flagella. The chloroplasts of most species are single and cupshaped or they may appear H-shaped, asteroid, lamellate, or be divided into discoidal units (Iyengar and Desikachary, 1981; Ettl, 1983). The chloroplast envelope consists of two layers without a chloroplast endoplasmic reticulum. The pyrenoid and stigma (eyespot) are generally contained within the chloroplast. The major photosynthetic pigments include chlorophylls a and b. However, colorless, heterotrophic genera also exist. The major storage product of photosynthetic species is starch (α-1, 4 and α-1, 6 polymer of glucose). which is localized within the chloroplast and stains positively with iodine. Asexual reproduction is accomplished by zoospore formation within the parental cell wall in the unicellular forms. In the colonial forms, all cells in a colony or some specialized (large) reproductive cells divide twice, three times or more (n) simultaneously to form four, eight or more (2n) daughter cells which soon become a daughter colony with the cell number and shape both characteristic for a given species (autocolony formation). After the daughter colonies are released from the parental colony, the new colonies increase their size with their cell number unchanged. Naked unicellular species usually undergo bipartition or cell division during swimming. Sexual reproduction is either isogamous, anisogamous, or oogamous. In isogamy, two conjugating gametes are essentially identical in size, whereas in anisogamy the cell size of male gametes is smaller than that of female gametes. When the female gametes lack flagella, such cells and the male gametes are called sperm and eggs, respectively, representing oogamy. Although evolution from isogamy to oogamy through anisogamy can be recognized in relation to increase in colonial complexity within the colonial Volvocales (Nozaki and Ito, 1994; Kirk, 1998; Nozaki et al., 2002), unicellular genera such as Chlamydomonas or Chlorogonium exhibit these three types of sexual reproduction (Ettl, 1983). After the conjugation,

the zygote secretes a heavy cell wall to become a hypnozygote (thick-walled dormant zygote). After a period of dormancy, the hypnozygote undergoes meiosis and germinates to give rise to four haploid nuclei (Stein, 1958b; Coleman, 1959). In most members of the green flagellates, therefore, four gone cells (reproductive cells from germinating zygote) of identical size are formed on zygote germination. However, only one of the four haploid nuclei survives to form a single gone cell in the colonial Volvocales (Volvocaceae) (Coleman, 1959; Goldstein, 1964; Nozaki et al., 1989). Buchheim et al. (1996) and Nakayama et al. (1996b), on the basis of ribosomal (r) RNA gene sequence data, resolved that the volvocalean algae constitute an ancestral and nonmonophyletic (paraphyletic) group with closely related derived groups including the chlorococcalean and tetrasporalean species. In addition, other DNA sequence data, such as the rbcL gene (large subunit of ribulose-1, 5-bisphosphate carboxylase/oxygenase) gene suggest that several genera (including Volvox) of the colonial forms are nonmonophyletic (Nozaki et al., 1995a, 1997a). Thus, further studies using both molecular and morphological data are needed to establish a natural taxonomic system for the flagellated green algae. Species of the volvocalean algae are generally delineated on their morphological differences. On the basis of intercrossings of heterothallic strains of the colonial species such as Pandorina morum (Coleman, 1959, 1977), multiple syngens (biological species) were demonstrated within the single morphological species. In addition, morphological species in the volvocalean algae may be nonmonophyletic or monophyletic depending upon the alga and the degree of morphological characterization of species (Nozaki et al., 1997b, c).

III. ECOLOGY AND DISTRIBUTION The green flagellates are found in a wide variety of freshwater habitats including lakes, ponds, rivers, rice paddies, and rainwater pools (e.g., Entwisle et al., 1998) as well as on ice or snow (Hoham, 1980; Ling, 1996). The majority are free living, but some strains are endosymbiotic (Lembi, 1980). Water bodies with elevated levels of nutrients are often especially rich in Volvocales (Lembi, 1980; Kirk, 1998), although Chlamydomonas has been reported in the nutrientpoor lakes (Lembi, 1980). Heterotrophic and photoheterotrophic algae (e.g., Polytoma, Astrephomene, and Pyrobotrys) grow in highly eutrophic waters, such as sewage oxidation ponds (Silva and Papenfuss, 1953), cow pasture ponds (Stein, 1958a), and rice paddies (Nozaki, 1983). Such photoheterotrophic algae

6. Flagellated Green Algae

generally require acetate for their growth (Pringsheim and Weissner, 1960; Brooks, 1972) and often exhibit plasticity of pyrenoids depending upon growth conditions (e.g., Nozaki et al., 1994b, 1995, 1998b). Three species of Pyrobotrys require anaerobic conditions for their growth (Nozaki, 1986). Some species, such as Tetrabaena socialis (= Gonium sociale), are distributed from the temperate zone to Antarctica and one Antarctic isolate does not grow at 25°C (Nozaki and Ohtani, 1992). Snow or ice algae are known in the unicellular genera Chloromonas, Chlamydomonas, Carteria, Chlainomonas, and Smithsonimonas (Hoham, 1980; Ling, 1996). Some species of Chlamydomonas grow in the extremely acidic water (Ettl, 1983). Although the volvocalean algae do not usually exhibit predominant species, Volvox and Haematococcus sometimes form water blooms in lakes or reservoirs. Many species are essentially worldwide in distribution (Iyengar and Desikachary, 1981; Ettl, 1983), although records from the tropics are sparse (Coleman, 1996). Extensive genetic and/or molecular analyses have been carried out on the colonial volvocalean species with worldwide distribution. Pandorina morum contains more than 20 sexually isolated groups (syngens) distributed throughout the world (Coleman, 1959, 1977) and their genetic differences are large (Coleman et al., 1994). Molecular phylogenetic analyses resolved that a large number of Gonium pectorale isolates from five continents are divided into six

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subclades, which are consistent with the interfertility between the isolates (Fabry et al., 1999). Although almost all of the well-studied genera of green flagellates are global in their distribution (Ettl, 1983), endemism can be considered in the colonial genus Platydorina, which has been reported only in the United States and Mexico (Kofoid, 1899; Harris and Starr, 1969; Ortega, 1984). Endemic distribution of species or infraspecific level is often recognized, particularly in Volvox (Smith, 1944; Nozaki, 1988).

IV. COLLECTION AND PREPARATION FOR IDENTIFICATION General methods for collection and preservation of freshwater algae have been described by Smith (1950) and Entwisle et al. (1998). Collection of the vegetative cells or colonies of the green flagellates can be carried out using plankton nets. These cells should soon be identified while they are alive, since some morphological features diagnostic for genus or species level (e.g., contractile vacuoles, flagella) are unstable after collection. When fixation is needed, the samples should be fixed with 1–2% glutaradehyde (rather than formaldehyde) soon after collection, and then stored at 4°C before identification. For snow algae, such as Chloromonas, cells fixed with 1–2% OsO4 will result in better cell morphology than those fixed with glutaradehyde.

V. KEY AND DESCRIPTIONS OF GENERA A. Key to Families of the Freshwater Green Flagellates 1a.

Cell with a depression at the flagellar base and/or lacking cell walls, gelatinous matrix, or lorica..........................Polyblepharidaceae

1b.

Cell without a depression at the flagellar base and enclosed by cell walls, gelatinous matrix, or lorica...............................................2

2a.

Cell with numerous protoplasmic strands radiating within the wall or gelatinous matrix.......................................Haematococcaceae

2b.

Cell without numerous protoplasmic strands radiating within the wall or gelatinous matrix.............................................................3

3a.

Unicellular..........................................................................................................................................................................................4

3b.

Colonial.............................................................................................................................................................................................5

4a.

Cell with a wall ...............................................................................................................................................Chlamydomonadaceae

4b.

Cell with a lorica...............................................................................................................................................................Phacotaceae

5a.

Colony surrounded by a gelatinous matrix.........................................................................................................................................6

5b.

Colony without a gelatinous matrix.......................................................................................................................Spondylomoraceae

6a.

Colony generally four-celled.........................................................................................................................................Tetrabaenaceae

6b.

Colony cell number eight or more......................................................................................................................................................7

7a.

Tripartite boundary of the gelatinous matrix surrounding each cell of the colony (cellular boundary); inversion absent during colony formation.............................................................................................................................................................................Goniaceae

7b.

Tripartite boundary of the gelatinous matrix surrounding the whole colony (colonial boundary); inversion occurs during colony formation..........................................................................................................................................................................Volvocaceae

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B. Polyblepharidaceae The flagellates of this family are unicellular and different from the typical green flagellates (e.g., Chlamydomonas) in that the cells lack cell walls (or lorica) and/or the protoplast exhibits depression at the flagellar base (Ettl, 1983). Asexual reproduction is generally affected by the bipartition of cells. Sexual reproduction is unknown in culture except for Nephroselmis (Suda et al., 1989). Based on the ultrastructure of cells, some genera traditionally assigned to

the Polyblepharidaceae have been classified in the separate class of green algae, namely, the Prasinophyceae (Micromonadophyceae) or Pedinophyceae (Norris, 1980; Moestrup, 1991). Recent molecular phylogenetic analyses essentially support this separation (e.g., Friedl, 1997). However, this chapter follows the traditional taxonomic style here because of the easy identification/recognition of the green flagellates using the light microscope. Eight genera are known:

1a.

Cell with a single flagellum (Fig. 1).................................................................................................................................Pedinomonas

1a.

Cell with two or more flagella............................................................................................................................................................2

2a.

Cell with eight flagella (Fig. 2).....................................................................................................................................Polyblepharides

2b.

Cell two or four flagella.....................................................................................................................................................................3

3a.

Cell with two flagella.........................................................................................................................................................................4

3b.

Cell with four flagella.........................................................................................................................................................................6

4a.

Flagella equal, inserted in the dorsal side of the platelike cell (Fig. 3)...............................................................................Mesostigma

4b.

Flagella unequal, inserted in the edge of the platelike cell..................................................................................................................5

5a.

Chloroplast lacking pyrenoid and stigma (Fig. 4)...............................................................................................................Scourfieldia

5b.

Chloroplast containing pyrenoid and stigma (Fig. 5)......................................................................................................Nephroselmis

6a.

Cell spindle-shaped and curved (Fig. 6)........................................................................................................................Spermatozopsis

6b.

Cell ovoid or ellipsoidal.....................................................................................................................................................................7

7a.

Cells containing photosynthetic pigments; cell wall or theca present..................................................................................................8

7b.

Cells lacking photosynthetic pigments; cell wall or theca absent (Fig. 7)............................................................................Polytomella

8a.

Lateral margins of cell not flattened or winged (Fig. 8).......................................................................................................Tetraselmis

8b.

Lateral margins of cell flattened and winged (Fig. 9)............................................................................................................Scherffelia

Mesostigma Lauterborn (Fig. 3) Disk-shaped cells are markedly compressed in the transapical axis, with two equal flagella inserted in the dorsal side of the cell. Cells without a wall are surrounded by gelatinous envelopes or numerous scales (Iyengar and Desikachary, 1981; Ettl, 1983). Two or more contractile vacuoles are present near the base of the flagella. Chloroplast is single and cup-shaped, with or without a pyrenoid and stigma. Asexual reproduction is by bipartition. Sexual reproduction has not been observed with certainty. Two species have been described. This genus is assigned to the Prasinophyceae (Norris, 1980), although recent phylogenetic analyses of actin genes show that Mesostigma is the earliest divergence within the Streptophyta (Charophyceae and land plants) (Bhattacharya et al., 1998; Karol et al., 2001). Based on the multiple chloroplast gene sequences, Mesostigma was resolved as the most basal organism

within the green plants (Streptophyta plus Chlorophyta) (Lemieux et al., 2000), but this phylogenetic position is questioned (Karol et al., 2001). M. viride and M. grande (Fig. 3) have been observed from the United States (e.g., FL, GA, IN, KY, OH) (Smith, 1950; Dillard, 1989). Nephroselmis Stein (Fig. 5) Cells are equatorially compressed, lacking a cell wall, with two unequal flagella inserted in the edge of the plate-like cell. A single contractile vacuole is located near the base of the flagella (Iyengar and Desikachary, 1981; Ettl, 1983). The chloroplast is single and cup-shaped, with a single basal pyrenoid and a stigma. The flagella and cell of this genus have a scaly covering (Suda et al., 1989). Asexual reproduction takes place by bipartition. Isogamous sexual reproduction has been confirmed in culture


2 1

3

5

6 4

7

8

9

FIGURE 1 Uniflagellate vegetative cell of Pedinomonas minor. (× 5500.) FIGURE 2 Polyblepharides singularis, showing anterior depression and eight flagella. (× 1400.) FIGURE 3 Two views of vegetative cell of Mesostigma grande. (× 1600.) FIGURE 4 Vegetative cell of Scourfieldia cordiformis, showing unequal flagella. (× 4500.) FIGURE 5 Vegetative cell of Nephroselmis olivacea, showing anterior depression. (× 3000.) FIGURE 6 Quadriflagellate vegetative cell of Spermatozopsis exsultans, showing twisted cell body. (× 7300.) FIGURE 7 Naked vegetative cell of Polytomella citrii showing four flagella. (× 2400.) FIGURE 8 Vegetative walled cell of Tetraselmis cordiformis, showing anterior depression. (× 2000.) FIGURE 9 Two views of vegetative cell of Scherffelia phacus. (× 2500.)

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(Suda et al., 1989). Two freshwater species have been described. Nephroselmis was recently assigned to the Prasinophyceae (Norris, 1980). This genus was designated as Heteromastix Korshikov by Smith (1950), who recorded North American H. angulata, a taxonomic synonym of Nephroselmis olivacea (Fig. 5). Pedinomonas Korshikov (Fig. 1) Cells are compressed, ellipsoidal to subcircular in front view, without a cell wall. A single flagellum is posteriorly directed and inserted at the anterior pole of the cell (Iyengar and Desikachary, 1981; Ettl, 1983). A single contractile vacuole is present near the base of the flagellum. Chloroplast is single and parietal, with a single pyrenoid and a stigma. Asexual reproduction is by bipartition (Ettl, 1983). Sexual reproduction has not been observed with certainty. Approximately ten species are recognized in freshwater habitats (Ettl, 1983). This genus was recently assigned to the Pedinophyceae on the basis of ultrastructure of the flagellar apparatus (Moestrup, 1991). P. maior, P. rotunda, and P. minor (Fig. 1) have been found in the United States (e.g., AL, FL) (Ettl, 1983; Dillard, 1989). This genus has also been collected in the Laurentian Great Lakes (Munawar and Munawar, 1981). Polyblepharides Dangeard (Fig. 2) Naked cells are ovoid to ellipsoidal, with eight equal flagella and two or four contractile vacuoles at the anterior pole. Chloroplast is single and massive, with a single pyrenoid surrounded by starch granules (Smith, 1950; Ettl, 1983). Asexual reproduction is by bipartition (Smith, 1950; Iyengar and Desikachary, 1981; Ettl, 1983). Although this genus is reported from various localities of the world, the existence of Polyblepharides has not been confirmed in culture (Bold and Wynne, 1985). P. singularis (Fig. 2) and P. fragariiformis are known from the United States (e.g., FL, NC) (Smith, 1950; Ettl, 1983). Polytomella Aragao (Fig. 7) Naked cells are ovoid, ellipsoidal, or spherical, with four equal flagella inserted in the anterior papilla of the cell (Iyengar and Desikachary, 1981; Ettl, 1983). Two or four contractile vacuoles are present near the base of the flagella. Chloroplasts degenerate to become colorless leucoplasts. A single stigma may be present depending upon the species (Ettl, 1983). Pyrenoids are lacking. Asexual reproduction is by bipartition. Isogamous sexual reproduction is reported. More than ten species have been described (Ettl, 1983). On the

basis of ultrastructure of flagellar apparatus and molecular phylogeny of 18S rRNA gene sequences, Polytomella was assigned to the Chlorophyceae (Nakayama et al., 1996a). P. citrii (Fig. 7) and P. agilis have been reported from the United States (Smith, 1950). Scherffelia Pascher (Fig. 9) Walled cells are ovoid or ellipsoidal, compressed equatorially, with two large chloroplasts and four equal flagella inserted in the anterior depression of the cell. Lateral margins of the cell are flattened and winged (Ettl, 1983). The cell wall, or theca, is formed by the fusion of cell body scales characteristic of the Prasinophyceae. Two contractile vacuoles are present near the base of the flagella. The chloroplast lacks pyrenoids. Asexual reproduction is via formation of four zoospores within the theca. Sexual reproduction is unknown. Eight species were recognized (Ettl, 1983). Although Smith (1950) and Ettl (1983) classified this genus in the Chlamydomonadaceae, 18S rRNA gene sequence data suggest that Scherffelia is closely related to Tetraselmis (Steinkötter et al., 1994). S. phacus Pascher (Fig. 9) is reported from the United States (e.g., OH, TN) (Smith, 1950). Scourfieldia G. S. West (Fig. 4) Naked cells are equatorially compressed, with two long unequal flagella inserted in the edge of the platelike cell (Iyengar and Desikachary, 1981; Ettl, 1983). Two contractile vacuoles are present near the base of the flagella. Chloroplast is single and cup-shaped, lacking pyrenoid and stigma. Asexual reproduction is by bipartition. Sexual reproduction has not been observed with certainty. Three freshwater species have been described (Ettl, 1983). Although flagella and cell body of this genus lack scaly covering, Scourfieldia was recently assigned to the Prasinophyceae on the basis of ultrastructure of flagellar apparatus (see Moestrup, 1991). S. cordiformis (Fig. 4) has been found in the United States (Dillard, 1989) and the Canadian province of Ontario (Duthie and Socha, 1976). The genus has also been reported from the arctic regions of Nunavut, Canada (Sheath and Steinman, 1982). Spermatozopsis Korshikov (Fig. 6) Naked cells are spindle-shaped and curved or twisted, with four equal flagella at the anterior end (Iyengar and Desikachary, 1981; Ettl, 1983). Two contractile vacuoles are located near the base of the flagella. A single chloroplast is linear, lying along the convex side of the cell, with a single stigma. Pyrenoid is lacking. Asexual reproduction is by bipartition. Sexual


reproduction has not been observed in culture. Two species have been described (Ettl, 1983; Preisig and Melkonian, 1984) On the basis of ultrastructure of flagellar apparatus and molecular phylogeny of 18S rRNA gene sequences, Spermatozopsis was assigned to the Chlorophyceae (Friedl, 1997). S. exsultans (Fig. 6) has been found in several location in of the United States (e.g., FL, KY, TN) (Smith, 1950; Dillard, 1989). Tetraselmis Stein (Fig. 8) Walled cells are ovoid or ellipsoidal, somewhat compressed equatorially, with four equal flagella inserted in the anterior depression of the cell. The cell wall or theca is formed by the fusion of cell body scales characteristic of the Prasinophyceae. Two contractile vacuoles are present near the base of the flagella in freshwater species (Ettl, 1983). The chloroplast is single and cup-shaped, with a single basal pyrenoid and a stigma. The pyrenoid may be absent depending upon the species (Ettl, 1983). Asexual reproduction is by bipartition within the theca. Sexual reproduction is

231

unknown. Although Tetraselmis is generally found in marine water, seven freshwater species were recognized (Ettl, 1983). T. cordiformis (Fig. 8) and T. subcordiformis is reported from the United States (e.g., FL, TN) (Smith, 1950; Ettl, 1983). Smith (1950) placed the latter species in the genus Platymonas (as P. subcordiformis) and he recorded North American P. elliptica, which is a taxonomic synonym of T. cordiformis (Ettl, 1983).

C. Haematococcaceae Cells of members of this family have numerous protoplasmic strands radiating within the cell wall or gelatinous matrix (Ettl, 1983). The family may be unicellular (Haematococcus) or colonial (Stephanosphaera). In Haematococcus, asexual reproduction is accomplished by zoospore formation, whereas daughter colony formation take place in Stephanosphaera. Sexual reproduction is effected by spindle-shaped small isogametes in both genera (Pocock, 1960; Ettl, 1983).

1a.

Unicellular (Fig. 10).....................................................................................................................................................Haematococcus

1b.

Colonial (Fig. 11).......................................................................................................................................................Stephanosphaera

10

11 FIGURE 10 Vegetative cell of Haematococcus pluvialis. (× 900.) FIGURE 11 Two views of vegetative colony of Stephanosphaera pluvialis. (× 590.)

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Haematococcus C. A. Agardh (Fig. 10) Cells are spherical, ellipsoidal, or pear-shaped, with two flagella and a cup-shaped chloroplast with a single or multiple pyrenoids (Pocock, 1960; Ettl, 1983; Thompson and Wujek, 1989). Protoplast is enclosed by a swollen, gelatinous cell wall and produces protoplasmic strands that extend through the wall. Accumulation of haematochrome often causes reddish color of the protoplasts. Sexual reproduction is by small, spindle-shaped isogametes. Six species are recognized (Ettl, 1983; Thompson and Wujek, 1989). H. pluvialis (= H. lacustris) (Fig. 10) and H. carocellus are reported widely from the United States (Smith, 1950; Thompson and Wujek, 1989). The latter species was collected at least twice from Minnesota, but has not been studied in culture (Thompson and Wujek, 1989). The genus is also widespread in Canada, including British Columbia (Stein and Borden, 1979), Nunavut (Sheath and Steinman, 1982) and Ontario (Duthie and Socha, 1976). H. pluvialis is also reported from Mexico (Ortega, 1984). Stephanosphaera Cohn (Fig. 11) Colonies are spherical to ellipsoidal, containing eight elongate cells arranged in a ring within a gelatinous matrix. Cells have two equal flagella, a stigma,

several contractile vacuoles, and a cup-shaped chloroplast (filling much of the cell) with several pyrenoids (Ettl, 1983). Protoplast exhibits numerous processes on the surface. Asexual reproduction occurs by autocolony formation by all the colonial cells. Sexual reproduction is isogamous (Ettl, 1983). Although this genus is sometimes classified in the family Volvocaceae (e.g., Bold and Wynne, 1985), a recent molecular phylogenetic study indicated that Stephanosphaera is closely related to Haematococcus, and separated from the volvocacean algae (Buchheim et al., 1994). Stephanosphaera is monotypic, with S. pluvialis (Fig. 11), which has been observed in the United States (Smith, 1950).

D. Chlamydomonadaceae The unicellular flagellates of this family have cell walls, chloroplasts, stigmata, and contractile vacuoles (Iyengar and Desikachary, 1981; Ettl, 1983). Asexual reproduction is accomplished by zoospore formation. Sexual reproduction is isogamous, anisogamous or oogamous (Ettl, 1983). Eight genera are known in North America (Figs. 12–19) and are distinguished by their flagellar number and cell form:

1a.


1b.

Cell with four flagella.........................................................................................................................................................................8

2a.

Cells lacking photosynthetic pigments (Fig. 15).....................................................................................................................Polytoma

2b.

Cells containing photosynthetic pigments...........................................................................................................................................3

3a.

Contractile vacuoles distributed throughout the cell surface (Fig. 16)............................................................................Chlorogonium

3b.

Contractile vacuoles positioned only near the base of the flagella......................................................................................................4

4a.

Two flagella remote from each other at the surface of the protoplast (Fig. 17).................................................................Gloeomonas

4b.

Two flagella in close proximity to each other at the surface of the protoplast....................................................................................5

5a.

Cell wall with protuberances or swollen............................................................................................................................................6

5b.

Cell wall without protuberances and not swollen...............................................................................................................................7

6a.

Cell wall with protuberances (Fig. 18)...............................................................................................................................Lobomonas

6b.

Cell wall without protuberances but swollen (Fig. 19)...................................................................................................Vitreochlamys

7a.

Chloroplasts containing pyrenoids (Fig. 12)...............................................................................................................Chlamydomonas

7b.

Chloroplasts lacking pyrenoids (Fig. 13).........................................................................................................................Chloromonas

8a.

Cell walls not swollen, tightly surrounding the protoplast (Fig. 14).........................................................................................Carteria

8b.

Cell walls swollen, separated from the protoplast surface..............................................................................................Chlainomonas


13 12

15 14

16

18

17 19 FIGURE 12 Vegetative cell of Chlamydomonas sonowiae, showing cell wall and two equal flagella. (× 2300.) FIGURE 13 Vegetative cells of Chloromonas minima. (From Pascher, 1927.) (× 1900.) FIGURE 14 Quadriflagellate vegetative cell of Carteria eugametos. (From Nozaki et al., 1994a, reproduced by permission of International Phycological Society.) (× 1500.) FIGURE 15 Vegetative cell of Polytoma uvella, lacking chloroplast. (× 1250.) FIGURE 16 A–C: Vegetative cells of Chlorogonium. A: Scanning electron microscopy of C. elongatum. (× 1600.) B: Optical section of C. euchlorum cell. (× 690) C: Surface view of C. capillatum cell, showing many contractile vacuoles. (× 690.) D: Two-celled stage of asexual reproduction in C. euchlorum. (From Nozaki et al., 1998b, reproduced by permission of the Journal of Phycology.) (× 690.) FIGURE 17 Vegetative cell of Gloeomonas ovalis (From Pascher, 1927). FIGURE 18 Vegetative cell of Lobomonas rostrata (× 2400.) FIGURE 19 Vegetative cell of Vitreochlamys fluviatilis, showing swollen cell wall (× 1150.)

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Carteria Diesing (Fig. 14) General features of the cells in this genus are essentially the same as in Chlamydomonas except for the typical four flagella (Ettl, 1983). Four contractile vacuoles and/or two stigmata are observed in some species. Asexual reproduction is by zoospore formation within the mother cell wall. Sexual reproduction is isogamous (Ettl, 1983). Aplanogamous sexual reproduction was recently reported in C. eugametos (Nozaki, 1994) (Fig. 14). More than 60 species have been described (Ettl, 1983). Lembi (1975), on the basis of the ultrastructure of flagellar apparatuses, recognized two distinct groups within the genus Carteria. Recent molecular phylogenetic analysis resolved such two clades of Carteria (Buchheim and Chapman, 1992; Buchheim et al., 1996, 2002). Nozaki et al. (1994a), on the basis of SEM observations, characterized the swastika-shaped anterior papillae of the cell wall in one of these two groups of Carteria. Twelve species of Carteria were observed in the United States (Smith, 1950; Dillard, 1989; Starr and Zeikus, 1993; Nozaki et al., 1994a) as well as British Columbia, Canada (Stein and Borden, 1979), the Laurentian Great Lakes (Munawar and Munawar 1981), and Mexico (Ortega, 1984). Chlainomonas Christen (not pictured) Cells of this genus have a swollen or gelatinous cell wall separated from the protoplast surface but otherwise similar to those of Carteria cells. Three species have been described (Ettl, 1983). Asexual reproduction is accomplished by zoospore formation (Ettl, 1983). Two species of this genus have been collected from snow in Canada and the United States; C. rubra in British Columbia, Canada, and Washington (Hoham, 1974a), and C. kollii in Oregon and Washington (Hoham, 1974b). Chlamydomonas Ehrenberg (Fig. 12) Cells are spherical, ovoid or ellipsoidal, with a cell wall, two equal flagella at the anterior pole and two contractile vacuoles at the base of the flagella (Ettl, 1983). The chloroplast is single and fills much of the cell, with a single or several pyrenoids. Stigma is generally single. Asexual reproduction is by zoospore formation within the mother cell wall. Sexual reproduction is isogamous, anisogamous, or oogamous. More than 400 species have been described (Ettl, 1983). It is very difficult to distinguish these morphological species, based on only field-collected materials. Analysis of rRNA gene sequences showed that Chlamydomonas was resolved as a basal, nonmonophyletic group within the Volvocales (Buchheim et al., 1996). Dillard (1989) listed 24 species of Chlamydomonas in the southeastern United States, such as C. sonowiae

(Fig. 12) [excluding four Chloromonas species and including Chlamydomonas tetragama (as Chlorogonium tetragamum)]. The genus is also reported from across Canada (e.g., Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982), as well as Mexico (Ortega, 1984), Guadelope (Bourrelly and Marguin, 1952), and Costa Rica (Haberyan et al., 1995). Chlorogonium Ehrenberg (Fig. 16) Cells are spindle-shaped, elongate-ovoid, ovoid, or ellipsoidal, with a cell wall and two equal flagella at the anterior end (Nozaki et al., 1998b). Contractile vacuoles are two or more, generally positioned in both of the anterior and posterior halves of the cell, but sometimes distributed in only the anterior portion of the cell. The chloroplast is parietal, lacking pyrenoids, or with one or more pyrenoids (Ettl, 1983). Asexual reproduction is by zoospore formation within the mother cell wall (Nozaki et al., 1996a). During asexual reproduction, the first division is transverse without cross rotation of the parental protoplast. This type of initial cell division distinguishes Chlorogonium from Chlamydomonas (Ettl, 1980; Nozaki et al., 1996a). Sexual reproduction may be isogamous, anisogamous, or oogamous. Recently, paedogamous sexual reproduction (conjugation of gametes within the gametangium) was observed in a culture of C. capillatum (Nozaki et al., 1995). Ettl (1983) used the presence or absence of pyrenoids in vegetative cells for distinguishing species of Chlorogonium and he recognized 12 pyrenoidlacking species and six pyrenoid-containing species. However, Chlorogonium strains completely lacking pyrenoids in the chloroplast of the vegetative cells have not previously been studied in culture (Nozaki et al., 1998b). On the basis of ultrastructure and rbcL gene sequences, the genus Chlorogonium was resolved for at least two phylogenetically separated groups (Nozaki et al., 1998b). Five species assignable to Chlorogonium have been found in the United States (e.g., FL, LA, TN, VA, WV) (Smith, 1950; Dillard, 1989; Nozaki et al., 1998b). This genus has been collected widely in Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982) and Mexico (Ortega, 1984). Chloromonas Gobi (Fig. 13) Chloromonas is distinguished from Chlamydomonas only by the lack of pyrenoids in the chloroplasts. More than 100 species have been described (Ettl, 1983). Complete absence of pyrenoids in the chloroplasts was demonstrated by light and electron microscopy in four species of Chloromonas (Morita et al., 1998). Buchheim et al. (1997), on the basis of 18S rRNA gene sequence data, showed that the genus


Chloromonas is nonmonophyletic within the Volvocales. Morita et al. (1999) and Nozaki et al. (2002a) resolved that Chloromonas and several species of Chlamydomonas constituted a closely related lineage. Pröschold et al. (2001) included several pyrenoid-containing species in the genus Chloromonas, mainly based on the 18S rRNA gene phylogeny. Smith (1950) reported North American Platychloris minima, which should be assigned to Chloromonas, as C. minima (Fig. 13). Four other species assignable to this genus (C. clathrata, C. platystigma, C. depauperata, and C. anglica) were observed from the United States (e.g., GA, NC, SC, TN) (Dillard, 1989). The genus is also reported from Canada (e.g., Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982) as well as Mexico (Ortega, 1984) (Haberyan et al., 1995). In addition, C. pinchiae, C. nivalis, C. brevispina, C. polypyera and C. granulosa have been collected in snow in the United States and Canada (e.g., Hoham et al., 1979, 1983). Gloeomonas Klebs (Fig. 17) Cells are spherical, ovoid, or ellipsoidal, with a thick cell wall or gelatinous envelope, two equal flagella at the anterior pole, and two contractile vacuoles near the base of the flagella (Iyengar and Desikachary, 1981; Ettl, 1983). The two flagella are remote from each other at the surface of the protoplast. Chloroplast is single and fills much of the cell or numerous discoid, lacking pyrenoids. A stigma may be present or absent depending on the species. Twelve species are recognized (Ettl, 1983). Asexual reproduction is by zoospore formation within the mother cell wall. Sexual reproduction is unknown. Vegetative cells of G. kupfferi have been examined by transmission electron microscopy (Domozych, 1989; Domozych and Nimmons, 1992). G. ovalis (Fig. 17) has been found in the United States (e.g., MA, ME) (Smith, 1950) and British Columbia, Canada (Stein and Borden, 1979). Lobomonas Dangeard (Fig. 18) Cells are spherical, ovoid, or ellipsoidal, with a cell wall, two equal flagella at the anterior pole, and two contractile vacuoles at the base of the flagella. The cell wall has many protuberances on the surface and may be swollen (Iyengar and Desikachary, 1981; Ettl, 1983). The chloroplast is single and massive, with a single pyrenoid and a stigma. Pyrenoid may be absent, depending on the species. Thirteen species are recognized (Ettl, 1983). Asexual reproduction is by zoospore formation within the mother cell wall. Sexual reproduction is unknown. L. rostrata (Fig. 18) has been found in the United States (e.g., NJ, VJ) (Smith, 1950; Dillard, 1989). This

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genus has also been reported from British Columbia, Canada (Stein and Borden, 1979). Polytoma Ehrenberg (Fig. 15) Cells of this genus are colorless, but otherwise similar to those of Chlamydomonas cells. More than 30 species have been described (Iyengar and Desikachary, 1981; Ettl, 1983). Asexual reproduction is via zoospore formation within the mother cell wall. Sexual reproduction is isogamous, anisogamous, or oogamous. On the basis of 18S rRNA gene sequence data,13 Polytoma strains were resolved as a nonmonophyletic group (Rumpf et al., 1996). Members of this genus occur in water rich in organic matter (Ettl, 1983). P. uvella (Fig. 15) and P. granuliferum have been found in the United States (Moewus, 1959; Ettl, 1983). Vitreochlamys Batko (Fig. 19) Cells are spherical, ovoid, or ellipsoidal, with a cell wall, two equal flagella at the anterior pole, and two or three contractile vacuoles at the base of the flagella (Ettl, 1983). The cell wall is swollen or gelatinous, which is characteristic of the genus (Ettl, 1983; Nakazawa et al., 2001). The chloroplast is single and fills much of the cell, with a single or multiple pyrenoids and a stigma. Pyrenoid may be absent, depending upon the species. Asexual reproduction is by zoospore formation within the mother cell wall. Sexual reproduction is unknown. Twenty-three species are recognized (Ettl, 1983). Although the name Sphaerellopsis Korshikov (1925) is generally used (e.g., Ettl, 1983), this name is a homonym of the fungus genus Sphaerellopsis M. C. Cooke (1883). Thus, Batko (1970) proposed Vitreochlamys as a nomen novum for Sphaerellopsis Korshikov (1925). The genus Vitreochlamys was resolved as three separate lineages basal to the Tetrasporales or the colonial Volvocales based on the rbcL gene phylogeny, representing the ancestral situation of these two orders (Nakazawa et al., 2001). V. fluviatilis (Fig. 19) and Sphaerellopsis (Vitreochlamys) gelatinosa have been collected in the United States (e.g., CA, KY, NC, TN, VA) (Smith, 1950; Dillard, 1989). The genus has been reported from Guadeloupe (Bourrelly and Manguin, 1952) and British Columbia, Canada (Stein and Borden, 1979), as Sphaerellopsis.

E. Phacotaceae The unicellular, biflagellate algae in this family are characterized by having nonliving investments surrounding the protoplast or loricae (Bold and Wynne, 1985). The loricae are sometimes impregnated with iron and/or manganese salts rendering them brown in color. In some genera, the lorica is composed of two

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parts that separate at reproduction (Hepperle and Krienitz, 1997). The vegetative cell has a stigma, two contractile vacuoles at the base of the flagella and a cup-shaped chloroplast with a single or more pyrenoids. Asexual reproduction is by zoospore formation (Hepperle and Krienitz, 1997). Isogamous or

anisogamous sexual reproduction is known in some species. Recent molecular phylogenetic analysis of the phatocacean algae indicates that Dysmorphococcus is separated from the clade composed of Phacotus and Pteromonas (Hepperle et al., 1998). Nine genera are distributed in North America.

1a.

Cell with four flagella (Fig. 20)...........................................................................................................................................Pedinopera

1b.


2a.

Lorica composed of two overlapping halves......................................................................................................................................3

2b.

Lorica not composed of two overlapping halves................................................................................................................................4

3a.

Lorica smooth and hyaline (Fig. 21)..................................................................................................................................Pteromonas

3b.

Lorica finely granulate and not hyaline (Fig. 22)....................................................................................................................Phacotus

4a.

Posterior portion of lorica narrowed or protruded.............................................................................................................................5

4b.

Posterior portion of lorica broad or rounded.....................................................................................................................................6

5a.

Compressed face of lorica with projections (Fig. 23).......................................................................................................Wislouchiella

5b.

Compressed face of lorica without projections (Fig. 24)................................................................................................Cephalomonas

6a.

Cells more or less compressed; lorica not broad (Fig. 25).............................................................................................Thoracomonas

6b.

Cells not compressed; lorica broad....................................................................................................................................................7

7a.

Lorica very thin and smooth except for numerous spines or granules distributed on the surface (Fig. 26).....................Granuochloris

7b.

Lorica thick and rough.......................................................................................................................................................................8

8a.

Two flagella projected through a common opening of lorica (Fig. 27)..............................................................................Coccomonas

8b.

Each flagellum projected through an individual opening of lorica (Fig. 28).............................................................Dysmorphococcus

Cephalomonas Higinbotham (Fig. 24) Loricae are compressed, rigid, and brittle, forming a broad anterior half and a narrowed posterior half. Cells have two equal flagella, two contractile vacuoles at the base of the flagella, a stigma, and a cup-shaped chloroplast with a single pyrenoid (Ettl, 1983). Asexual reproduction is by zoospore formation. Isogamous sexual reproduction is known. Cephalomonas is a monotypic genus with C. granulata (Ettl, 1983). C. granulata (Fig. 24) has been reported from the United States (e.g., FL, KY, MD) (Smith, 1950; Dillard, 1989). Coccomonas Stein (Fig. 27) Loricae are thick and not compressed, impregnated with lime (CaCo3) and iron compounds. Cells are ovoid to spherical in shape, with two equal flagella, two contractile vacuoles at the base of the flagella, a stigma, and a cup-shaped chloroplast with a single basal pyrenoid (Ettl, 1983). The two flagella are projected through a common opening of the lorica. Asexual reproduction is by zoospore formation. Sexual reproduction is unknown. Seven species are known in this genus (Ettl, 1983).

C. orbicularis (Fig. 27) has been recorded from the United States (e.g., FL, GA, KY, NC, SC, TN, VA) (Smith, 1950; Dillard, 1989), and from British Columbia, Canada (Stein and Borden, 1979). Dysmorphococcus Takeda (Fig. 28) Loricae are thick and with pores, sometimes becoming brown in color. Cells spherical or ovoid, with two equal flagella, two contractile vacuoles at the base of the flagella, a stigma, and a cup-shaped chloroplast with a single or multiple pyrenoids (Ettl, 1983). Each flagellum is projected through an individual opening of the lorica. Asexual reproduction is by zoospore formation. Sexual reproduction is unknown. Eight species are known in this genus (Ettl, 1983). D. variabilis (Fig. 28) and D. globosus have been observed from several locations in the United States (e.g., AL, KY, OH, TN, VA, WV) (Bold and Starr, 1953; Dillard, 1989). Granuochloris Pascher et Jahoda (Fig. 26) Loricae are thin and not compressed, with numerous granules or spines distributed throughout the surface. Cells have two equal flagella, two contractile


22 21 20

24 25 23

26

27

28

FIGURE 20 Two views of Pedinopera granulosa vegetative cell. (× 1000.) FIGURE 21 Two views of Pteromonas aculeata vegetative cell. (× 790.) FIGURE 22 Two views of vegetative cell of Phacotus lenticularis. (× 770.) FIGURE 23 Vegetative cell of Wislouchiella planctonica. (× 1370.) FIGURE 24 Vegetative cell of Cephalomonas granulata. (× 1800.) FIGURE 25 Vegetative cell of Thoracomonas phacotoides. (× 1900.) FIGURE 26 Vegetative cell of Granulochloris spinifera. (× 1700.) FIGURE 27 Vegetative cell of Coccomonas orbicularis, showing two flagella projecting through one anterior pore of the lorica. (× 1300.) FIGURE 28 Vegetative cell of Dysmorphococcus variabilis. (× 1500.)

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vacuoles at the base of the flagella, a stigma, and a cup- or H-shaped chloroplast with a single pyrenoid (Fott, 1963). Reproduction is not known in detail. Three species are recognized in this genus (Ettl, 1983). G. spinifera (Fig. 26) has been collected in Massachusetts (Fott, 1963). Pedinopera Pascher (Fig. 20) Loricae are compressed, with granulate surface, with or without longitudinal ridges. Cells are flattened and ovoid to pear-shaped, with four equal flagella and a cup-shaped chloroplast (Ettl, 1983). Pyrenoids/stigma may be present or absent, depending upon a given species. Although five species have been described, reproduction in this genus is unknown (Ettl, 1983). P. granulosa (Fig. 20) and P. rugulosa have been collected in the United States (e.g., FL, NC, TN) (Dillard, 1989). Phacotus Perty (Fig. 22) Loricae are highly compressed and lens-shaped, composed of two parts (sometimes called shells), without pores. The loricae are usually dark brown colored and impregnated with lime, exhibiting a mineralized surface composed of many calcite crystals (Hepperle and Krienitz, 1997). Cells are flattened, with two equal flagella, two contractile vacuoles at the base of the flagella, a stigma, and a cup-shaped chloroplast. Pyrenoids may be single, multiple, or lacking depending upon a given species (Ettl, 1983). Asexual reproduction is by formation of 2–16 zoospores within a gelatinous sporangial sheath after separation of the two shells of the parental lorica (Hepperle and Krienitz, 1996). Sexual reproduction is rare and isogamous (Ettl, 1983). Calcification of P. lenticularis requires external supersaturation of calcium (Hepperle and Krienitz, 1997). P. lenticularis (Fig. 22), P. angustus, P. glaber, and P. subglobosus have been collected from across in the United States (Smith, 1950; Dillard, 1989). The genus is also reported from Ontario and British Columbia, Canada (Duthie and Socha, 1976; Stein and Borden, 1979) and the Laurention Great Lakes (Munawar and Munawar, 1981). Pteromonas Seligo (Fig. 21) Loricae are compressed, forming projecting wings around the cell, composed of two hyaline shell-like portions joined at the wings (Iyengar and Desikachary, 1981; Ettl, 1983). Cells are flattened and pear-shaped, with two equal flagella, two contractile vacuoles at the base of the flagella, a stigma and a cup-shaped chloroplast. Pyrenoids may be single or multiple, which is species specific (Ettl, 1983). Asexual reproduction is by

formation of two or four zoospores within the parental lorica. Sexual reproduction is isogamous. Approximately 20 species have been described (Ettl, 1983). Five species of Pteromonas, P. aculeata (Fig. 21), P. cordiformis, P. angulosa, P. sinuosa, and P. cruciata have been collected in the United States (e.g., KY, NC, TN, WV) (Smith, 1950; Dillard, 1989). Thoracomonas Korshikov (Fig. 25) Loricae are somewhat compressed, verrucose, and thin. Cells have two equal flagella, two contractile vacuoles at the base of the flagella, a stigma, and a cupshaped chloroplast with a single or multiple pyrenoids (Ettl, 1983). Asexual reproduction is by zoospore formation. Sexual reproduction is not known in detail. Four species are recognized in this genus (Ettl, 1983). T. feldmanii and T. phacotoides (Fig. 25) have been observed in the United States (e.g., LA, TN) (Smith, 1950; Dillard, 1989). Wislouchiella Skvortzow (Fig. 23) Loricae are strongly compressed and finely verrucose, forming a broad wing-like expansion with two cylindrical projections in each compressed face (Ettl, 1983). Cells are ovoid in side view and rhomboidal in vertical view, with two equal flagella, two contractile vacuoles at the base of the flagella, a stigma, and a cup-shaped chloroplast with a single pyrenoid. Reproduction is unknown. Wislouchiella is a monotypic genus with W. planctonica (Ettl, 1983). W. planctonica (Fig. 23) has been reported from the United States (Smith, 1950; Dillard, 1989) and British Columbia, Canada (Stein and Borden, 1979).

F. Volvocaceae The coenobic colonial organisms (fixed number of cells) in which biflagellate Chlamydomonas-like cells are surrounded by a gelatinous matrix were traditionally assigned to the Volovocaceae and Astrephomenaceae (Bold and Wynne, 1985). However, the classification of these two families has recently become controversial. Nozaki and Kuroiwa (1992) removed the genus Gonium from the Volvocaceae and classified Astrephomene and Gonium in a single family, the Goniaceae, based on the vegetative ultrastructure of the extracellular (gelatinous) matrix. Subsequently, Nozaki and Ito (1994), on the basis of cladistic analysis of morphological data, resolved the four-celled species Tetrabaena socialis (Gonium sociale) as a sister group to the monophyletic group composed of other species in the Volvocaceae and the Goniaceae, and established a new family, the Tetrabaenaceae, for encompassing T. socialis. Furthermore, Nozaki et al. (1996b)


assigned another four-celled colonial alga Basichlamys sacculifera to the Tetrabaenaceae, on the basis of the further cladistic analysis based on morphological data. Although the author follows the taxonomic concept of Nozaki et al. (1996b), the phylogenetic relationships and status of these three colonial families are uncertain in the phylogenetic analyses based on the rbcL and/or atpB (ATP synthase beta-subunit) gene sequence data except for the robust monophyly of the Tetrabaenaceae (Nozaki et al., 1995a, 1997a, 1999). However, the recent molecular phylogenetic analyses using five chloroplast genes resolved the monophyly of the Volvocaceae as well as the Goniaceae (Nozaki et al., 2000). The volvocacean algae now can be characterized by having a tripartite colonial boundary of the extra-

239

cellular matrix in vegetative colonies (Nozaki and Kuroiwa, 1992). In asexual reproduction, all of cells in the colony or only large reproductive cells (gonidia) divide successively to form a miniature of the parental colony (autocolony). The volvocacean genera exhibit a unique phenomenon called inversion or eversion just after the successive divisions during colony formation (see Smith, 1955). Sexual reproduction is either isogamous, anisogamous, or oogamous. In anisogamous and oogamous genera, spindle-shaped male gametes are produced as they are grouped, forming a hemispherical or flattened coenobic colony called a “sperm packet.” This family includes seven genera (Figs. 29–35) which are distinguished by shape of the colony and cellular differentiation.

1a.

Colony flattened, with cells arranged in a single layer (Fig. 33)..........................................................................................Platydorina

1b.

Colony spheroidal, with cells arranged radially..................................................................................................................................2

2a.

Colony generally with up to 32 cells; differentiation of small somatic cells absent or incomplete.......................................................3

2b.

Colony generally with 64 or more cells; differentiation of small somatic cells complete.....................................................................6

3a.

Maximum colony cell number 16; cellular envelopes absent..............................................................................................................4

3b.

Maximum colony cell number 32; cellular envelopes present.............................................................................................................5

4a.

Cells keystone-shaped or pear-shaped; colony contiguous in the center (Fig. 29).................................................................Pandorina

4b.

Cells hemispherical or lenticular; colony hollow (Fig. 30).....................................................................................................Volvulina

5a.

Sexual reproduction isogamous (Fig. 31)........................................................................................................................Yamagishiella

5b.

Sexual reproduction anisogamous with sperm packets (Fig. 32)............................................................................................Eudorina

6a.

Colony generally 32- 64- or 128-celled, with 20–50% somatic cells (Fig. 34)....................................................................Pleodorina

6b.

Colony containing more than 500 cells, composed mostly of somatic cells (Fig. 35).................................................................Volvox

Eudorina Ehrenberg (Fig. 32) Colonies are ovoid, ellipsoidal, or cylindrical, containing 16 or 32 cells arranged radially in the periphery of a gelatinous matrix, forming a hollow sphere (Goldstein, 1964). Each cell of the colony is enclosed tightly by the fibrillar layer (cellular envelope) of the extracellular matrix of the colony (Nozaki and Kuroiwa, 1992). The matrix does or does not form individual sheaths, depending upon the species (Goldstein, 1964). Cells are ovoid or spherical, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with one (basal) or multiple pyrenoids (Goldstein, 1964). Stigmata of anterior cells are larger than in posterior cells. No differentiation between somatic and reproductive cells occurs except for E. illinoisensis, in which anterior four cells are small and facultatively somatic. Sexual reproduction is anisogamous with sperm packets, producing walled hypnozygotes (Goldstein, 1964). Upon germination, the zygote gives rise to a single or two biflagellate gone

cell. Eudorina is cosmopolitan and contains about seven species (Goldstein, 1964). Although Eudorina was often confused with Pleodorina, it is now distinguished based of the absence of obligately somatic cells (Nozaki et al., 1989). Molecular phylogenetic analyses resolved that the genus Eudorina is paraphyletic, exhibiting the ancestral situation of Pleodorina and Volvox (excluding section Volvox) (Nozaki et al., 1995a, 1997a, b, 1999, 2000). E. elegans is among the most frequently encountered species of green algae. Goldstein (1964), on the basis of cultured material originating from the United States and Canada, recognized five species assignable to Eudorina, E. elegans (Fig. 32), E. unicocca, E. illinoisensis, E. cylindrica, and E. conradii. Although Prescott (1955) described E. interconnexa from the Panama Canal Zone, Ettl (1983) questioned its existence. Several species have been recorded widely in the United States and Canada (Smith, 1950; Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982; Whitford and Schumacher, 1984).

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Hisayoshi Nozaki

30

29

32

31

33

34

35 FIGURE 29 Vegetative colony of Pandorina morum, showing 16 cells compactly arranged. (From Nozaki, 1995, reproduced by permission of Koudan-Sha Scientific.) (× 520.) FIGURE 30 Sixteen-celled vegetative colony of Volvulina steinii, showing pyrenoid (py) developing in the brim of the cup-shaped chloroplast. (From Nozaki, 1982, reproduced by permission of Journal of Japanese Botany.) (× 470.) FIGURE 31 Yamagishiella unicocca vegetative colony, showing 32 cells loosely arranged. (From Nozaki, 1981, reproduced by permission of Journal of Japanese Botany.) (× 270.) FIGURE 32 Vegetative colony of Eudorina elegans. (From Nozaki, 1995, reproduced by permission of Koudan-Sha Scientific.) (× 300.) FIGURE 33 Two views of vegetative colony of Platydorina caudata. (From Kofoid, 1899.) (× 200, × 130.) FIGURE 34 Light micrographs of Pleodorina californica. (× 85.) A: Vegetative colony. B: Daughter colony formation. FIGURE 35 Light micrographs of Volvox aureus. A: Asexual colony with divided gonidia. (× 290.) B: Daughter colonies within the parental colony. (× 110.) C: Surface view of asexual colony. Note delicate cytoplasmic bridges between cells. (× 340.)


Pandorina Bory de St.-Vincent (Fig. 29) Colonies are ovoid or ellipsoidal, containing eight or 16 cells compactly arranged radially in a gelatinous matrix (Ettl, 1983). Cells are keystone-shaped or ovoid, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with one basal or multiple pyrenoids (species dependent) (Nozaki and Kuroiwa, 1991). Stigmata in anterior cells are larger than in posterior cells. Sexual reproduction is isogamous, forming walled hypnozygotes (Coleman, 1959; Nozaki and Kuroiwa, 1991). Upon germination, zygotes give rise to single biflagellate gone cells. Pandorina is cosmopolitan in fresh waters (Coleman, 1959, 1977). Although this genus was frequently confused with Eudorina, it is now distinguished by the difference in structure of extracellualr matrix of the vegetative colony and in sexual reproduction (Nozaki, 1981; Nozaki and Kuroiwa, 1992). Recently, Pandorina unicocca was removed to the genus Yamagishiella, on the basis of its cellular envelopes within the colony (Nozaki and Kuroiwa, 1992). Although seven species were described, only P. morum and P. colemaniae seem reliable based on cultural studies (Coleman, 1959; Nozaki and Kuroiwa, 1991). P. morum (Fig. 29) is broadly distributed throughout the United States (Coleman, 1959). P. smithii has been found in Wisconsin (Ettl, 1983). P. morum has also been reported widely from Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982) as well as Mexico (Ortega, 1984) and Guadeloupe (Bourrelly and Marguin, 1952). P. charkowiensis, reported by Smith (1950), Thompson (1954) and Dillard (1989) should be assigned to Eudorina or Yamagishiella. Platydorina Kofoid (Fig. 33) Colonies are flattened, slightly twisted, and horseshoe-shaped with three to five posterior projections of a gelatinous matrix (Kofoid, 1899). Each colony contains 16 or 32 cells arranged in one layer and oriented in different directions in a gelatinous matrix. Cells are spherical or pear-shaped, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with a single basal pyrenoid (Kofoid, 1899). Colony formation involves inversion and subsequent intercalation (Harris and Starr, 1969). Sexual reproduction is anisogamous with sperm packets, forming walled hypnozygotes (Harris and Starr, 1969). Platydorina is a monotypic genus with P. caudata (Fig. 33) and collected from sites in the United States (e.g., AL, FL, GA, KY, SC, TN) (Kofoid, 1899; Harris and Starr, 1969). The genus has been reported from Mexico, but no species was given (Ortega, 1984).

241

Pleodorina Shaw (Fig. 34) Colonies are spherical, ovoid, or ellipsoidal, containing 32, 64, or 128 cells arranged radially at the periphery of a gelatinous matrix (Nozaki et al., 1989). Colonies have small, obligately somatic cells at the anterior pole, and large reproductive cells (gonidia) in the remaining portion. The matrix may or may not exhibit individual sheaths, depending upon a given species (Nozaki et al., 1989). Cells are spherical or ovoid, each with two equal flagella, a stigma, many contractile vacuoles on the cell surface, and a massive cup-shaped chloroplast. Chloroplasts of somatic cells have a single basal pyrenoid, whereas reproductive cells have multiple ones. Stigmata of anterior cells are larger than in posterior cells. Sexual reproduction is anisogamous with sperm packets, forming walled hypnozygotes (Nozaki et al., 1989). A single biflagellate gone cell is released from the germinating zygote. Pleodorina is cosmopolitan in freshwater and includes four species (Nozaki et al., 1989). Pleodorina and Eudorina are distinguished based on the presence or absence of obligately somatic cells respectively (Nozaki et al., 1989). Recent molecular phylogenetic analysis suggests that P. indica is separated from P. californica and P. japonica (Nozaki et al., 1997a). P. californica (Fig. 34) has been recorded from the United States (e.g., FL, GA, TN, VA, WV) and the Panama Canal Zone (Goldstein, 1964) as well as from Ontario, Canada (Duthie and Socha, 1976). P. indica, originating from Mexico, has been studied by light and electron microscopy (Nozaki et al., 1989; Nozaki and Kuroiwa, 1992). Volvox Linnaeus (Fig. 35) Colonies are spherical, subspherical, ellipsoidal, or ovoid, containing 500–50,000 cells arranged radially at the periphery of a gelatinous matrix, forming a hollow sphere. Several to approximately 50 large reproductive cells (gonidia) are situated in posterior 1/2 to 2/3 of colony (Smith, 1944). Each cell is enclosed by a gelatinous sheath which is distinct or confluent, depending upon the species (Smith, 1944). Somatic cells are spherical, ovoid, or star-shaped, each with two equal flagella, two contractile vacuoles at the base of the flagella, and a cup-shaped chloroplast with a single pyrenoid. Cytoplasmic strands between cells are thick, thin, or absent, and this trait is species dependent (Smith, 1944). Stigmata in the anterior cells are larger than in posterior cells. Sexual reproduction is oogamous (Nozaki, 1988); in monoecious species, the sexual colony has both sperm packets and eggs. In dioecious species, the male colony contains androgonidia which divide successively into sperm packets; such males may be markedly reduced in size (dwarf

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male) or nearly as large as asexual colonies. The female colony has eggs, whose number is nearly the same as that of gonidia in asexual colonies (facultative female) or much larger (special female). After fertilization zygotes develop a heavy cell wall that may be ornamented with reticulation or spines (Smith, 1944). Upon germination, the zygote gives rise to a single biflagellate gone cell (Nozaki, 1988). This genus contains approximately 20 species, which are classified into four sections based on the differences in gelatinous matrix and cytoplasmic strands (Smith, 1944). Ultrastructure of the flagellar apparatus of two species of Volvox (Hoops, 1984) and molecular phylogenetic analyses based rRNA sequences (Larson et al., 1992; Kirk, 1998) and internal transcribed spacer sequences (Coleman, 1999) indicate that Volvox is polyphyletic. Molecular phylogenetic analyses based on rbcL gene sequence data strongly suggest that section Volvox (=Euvolvox) is separated from the other three sections (Nozaki et al., 1995a, 1997a, 1999). Results of chloroplast multigene phylogeny indicate that the genus Volvox represents four separate lineages (Nozaki et al., 2002b). Volvox is cosmopolitan in fresh waters and V. aureus (Fig. 35) is a widely reported species. Eleven species of Volvox have been found in the United States, namely, V. aureus, V. globator, V. africanus, V. carteri, [including V. weismannia (= V. carteri f. weismannia)], V. perglobator, V. powersii, V. spermatosphaera, V. tertius, V. dissipatrix, V. prolificus, and V. rousseletii (Smith, 1950; Dillard, 1989). The genus has also been widely reported from Canada (Duthie and Socha, 1976; Stein and Borden, 1979; Sheath and Steinman, 1982), Mexico (Ortega, 1984), and Guadeloupe (Bourrelly and Manguin, 1952). Starr (1970) described V. pocockiae based on cultured material originating from Mexico. Volvulina Playfair (Fig. 30) Colonies are ovoid or spherical, containing eight or 16 cells embedded in the periphery of a gelatinous matrix, forming a hollow structure (Pocock, 1954; Thompson, 1954; Stein, 1958a; Starr, 1962). Cells are lenticular or hemispherical, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella or many contractile vacuoles scattered on the cell surface, a massive cup-shaped chloroplast without pyrenoids, or with one in the bottom or brim (Starr, 1962; Nozaki, 1982; Nozaki and Kuroiwa, 1990). Stigmata in anterior cells are larger than in posterior cells. Sexual reproduction is isogamous and walled hypnozygotes are formed (Stein, 1958a; Starr, 1962; Nozaki, 1982; Nozaki and Kuroiwa, 1990). Germinating zygotes give rise to a single biflagellate gone cell. Volvulina is cosmopolitan but rare. The current concept of Volvulina is based on the presence

of hollow colonies with lenticular cells (Nozaki and Kuroiwa, 1990). V. steinii is often collected from water rich in organic matter. Three reliable species were described (Nozaki and Kuroiwa, 1990). V. steinii (Fig. 30) has been observed in various localities across the United States (Thompson, 1954; Stein, 1958a; Carefoot, 1966). Starr (1962) described V. pringsheimii Starr based on cultured material originating from Texas. V. steinii has also been collected in British Columbia, Canada (Stein and Borden, 1979). Yamagishiella Nozaki (Fig. 31) Colonies are ovoid, ellipsoidal, or cylindrical, containing 16 or 32 cells arranged radially in the periphery of a gelatinous matrix, forming a hollow sphere (Rayburn and Starr, 1974). Each cell of the colony is enclosed tightly by the fibrillar layer (cellular envelope) of the extracellular matrix of the colony (Nozaki and Kuroiwa, 1992). Cells are ovoid or spherical, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with a single basal pyrenoid (Rayburn and Starr, 1974). Stigmata of anterior cells are larger than in posterior cells. No differentiation between somatic and reproductive cells occurs. Sexual reproduction is isogamous, producing walled hypnozygotes (Rayburn and Starr, 1974). Upon germination, the zygote gives rise to a single biflagellate gone cell. Yamagishiella is a monotypic genus with Y. unicocca (Fig. 31). This genus is distinguished from Pandorina by its cellular envelopes and 32-celled colonies (Nozaki and Kuroiwa, 1992). Although Yamagishiella differs from Eudorina by its isogamous sexual reproduction, the vegetative morphology and asexual reproduction characteristics of these two genera (especially Y. unicocca and E. unicocca) are indistinguishable. However, such a taxonomic problem may be resolved based on the analysis of rbcL gene sequence data (Nozaki et al., 1998a). Y. unicocca has been recorded from Indiana, Oregon, and Massachusetts (as Pandorina unicocca; Rayburn and Starr, 1974).

G. Goniaceae This family was revived by Nozaki and Kuroiwa (1992) to comprise the two genera Gonium and Astrephomene. These two genera (Figs. 36 and 37) exhibit essentially the same colony structure in which each vegetative cell is enclosed by a tripartite boundary of the extracellular matrix (cellular boundary). This situation is essentially different from that of the Volvocaceae (colonial boundary). Asexual reproduction is by daughter colony formation without inver-


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37

36

FIGURE 36 Light micrographs of Gonium pectorale. (× 340.) A: 16-celled colony. B: Eight-celled colony. C: Daughter colony formation. FIGURE 37 Vegetative colony of Astrephomene gubernaculifera. (From Nozaki, 1983, reproduced by permission of Journal of Japanese Botany.) (× 230.)

sion. All the vegetative cells in Gonium colonies undergo reproduction, whereas small somatic cells are present in the posterior pole of Astrephomene colonies.

Sexual reproduction is isogamous and three types of zygote germination are recognized in the Goniaceae (see Nozaki and Ito, 1994).

1a.

Colony flattened (Fig. 36).......................................................................................................................................................Gonium

1b.

Colony spherical (Fig. 37)..............................................................................................................................................Astrephomene

Astrephomene Pocock (Fig. 37) Colonies are ovoid or subspherical, containing 32, 64, or 128 cells arranged radially at the periphery of a gelatinous matrix (Pocock, 1954). Colonies have two to several small somatic cells (rudder cells) at the posterior pole. Cells are nearly spherical or lenticular, each with two equal flagella, a stigma, many contractile vacuoles on the cell surface, and a massive cup-shaped chloroplast that lacks pyrenoids or develops one in the brim (Nozaki, 1983). Stigmata in the anterior cells are larger than in posterior cells. Each protoplast is enclosed by a gelatinous sheath (cellular boundary) and constitutive cells attach or connect by the fusion or attachment of these sheaths, forming a hollow colony (Nozaki, 1983). Sexual reproduction is isogamous and walled hypnozygotes are formed. On germination, the zygote gives rise to a single biflagellate gone cell (Brooks, 1966; Nozaki, 1983). Astrephomene is rare and grows in freshwater rich in organic matter. This genus contain two species: A. gubernaculifera (Fig. 37) and A. perforata (Nozaki, 1983). The two species differ in character of the gelatinous cellular sheaths, pyrenoids, and number of somatic cells (Nozaki, 1983). Some authors recognize the family Astrephomenaceae for only a single genus Astrephomene, based on having spheroidal colonies but lacking inversion during colony formation (Pocock, 1954). A. gubernaculifera has been collected in the United States and Mexico (Stein, 1958a; Brooks, 1966).

Gonium O. F. Müller (Fig. 36) Colonies are flattened, containing eight, 16, or 32 cells arranged in one layer and oriented in the same direction (Stein, 1958b; Ettl, 1983). The eight-celled colony exhibits a characteristic cell arrangement for a given species (Nozaki, 1989a). Cells are ovoid to angular, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with one or multiple pyrenoids (Pocock, 1955; Stein, 1958b; Nozaki, 1989a). Each protoplast is enclosed by a gelatinous sheath (cellular boundary), and the cells attaching or connecting to one another by the union or attachment of the sheaths form a colony. Sexual reproduction is isogamous, forming hypnozygotes with smooth walls (Stein, 1958b; Nozaki, 1989a). Germinating zygote produces four biflagellate gone cells which are joined in a colony (germ colony) (Stein, 1958b; Nozaki, 1989a), except for G. multicoccum (Nozaki and Ito, 1994). Gonium is cosmopolitan in fresh waters (Fabry et al., 1999). G. pectorale (Fig. 36) is one of the most commonly encountered species of all chlorophytes. Five species have been studied in culture (Nozaki et al., 1997a). Although cladistic analysis based on morphological data indicates that Gonium is paraphyletic (Nozaki and Ito, 1994), recent molecular phylogenetic analyses using combined data set from multiple chloroplast protein-coding gene sequences resolved that the genus is monophyletic (Nozaki et al., 1999, 2000).

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Stein (1965) demonstrated the existence of 33 sexual populations of G. pectorale from the United States and Canada. Pocock (1955) described G. multicoccum and G. octonarium based on material collected in the United States. Prescott (1942) described G. discoideum from Louisiana. Although G. formosum has been frequently collected in the United States (Smith, 1950; Dillard, 1989) and British Columbia, Canada (Stein and Borden, 1979), this species has not been previously studied in culture.

H. Tetrabaenaceae The Tetrabaenaceae includes Basichlamys and Tetrabaena, both of which were sometimes assigned to the genus Gonium (e.g., Stein, 1959). However, this

family differs from the Goniaceae in having vegetative colony composed of only four cells and reticulate zygote or hypnospore walls (Nozaki et al., 1996b). All the cells of the colony divide into a daughter colony in asexual reproduction. Sexual reproduction is isogamous. Based on the combined data set from rbcL and atpB gene sequences, Basichlamys and Tetrabaena are resolved as a close clade separated from the monophyletic group composed of the genus Gonium (Nozaki et al., 1999). Such separation between the Tetrabaenaceae and the genus Gonium was supported by the occurrence of the xanthophyll loroxanthin within the colonial Volvocales (Schagerl and Angeler 1998). Recently, the Tetrabaenaceae was resolved as the most basal lineage within the colonial Volvocales on the basis of the multigene phylogeny (Nozaki et al., 2000).

39

38 FIGURE 38 Light micrographs of Basichlamys sacculifera. (× 650.) A: Upper view of vegetative colony. India ink preparation. B: Side view of vegetative colony. India ink preparation. C: Aplanospores with reticulate walls. FIGURE 39 Light micrographs of Tetrabaena socialis vegetative colonies. India ink preparation. (× 640.) A: Upper view of vegetative colony. B: Side view of vegetative colony.


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1a.

Colonial cells separated from one another, but attached to their parental cellular sheath (sac) (Fig. 38)............................Basichlamys

1b.

Colonial cells connecting to one another by the union or attachment of the cellular sheaths (Fig. 39)................................Tetrabaena

Basichlamys Skuja (Fig. 38) Colonies contain four cells attached to their parental cellular sheath (sac), forming a square (Stein, 1959). Cells are ovoid and somewhat asymmetrical, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with a single basal pyrenoid. Akinetes or hypnospores (thick-walled dormant cells) are sometimes formed (Stein, 1959). Sexual reproduction is isogamous. Basichlamys contains only a single species B. sacculifera (Fig. 38) and it is distinguished from Tetrabaena and Gonium in lacking connections of the cellular sheaths of the constitutive cells, which attach only to the sac. Some authors do not recognize Basichlamys and use Gonium sacculiferum (Stein, 1959). This genus is rarely abundant, but it is apparently cosmopolitan in freshwaters (Stein, 1959). Stein (1959) studied morphology and reproduction of B. sacculifera (as Gonium sacculiferum) based on the cultured materials originating from Indiana, Minnesota, and California; it has also been reported from British Columbia (Stein and Borden, 1979). Tetrabaena Fromentel (Fig. 39) Colonies contain four cells attached to each other by the protuberances of their cellular sheath, forming a square (Stein, 1959; Nozaki and Ohtani, 1992). Cells are ovoid and somewhat asymmetrical, each with two

equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with a single basal pyrenoid (Stein, 1959; Nozaki and Ohtani, 1992). Sexual reproduction is isogamous (Stein, 1959). This genus is rare but cosmopolitan in fresh water, occurring from temperate zones to the Antarctic (Nozaki and Ohtani, 1992). Tetrabaena contains only a single species, T. socialis (Fig. 39) and it is distinguished from Gonium in lacking colonies with more than four cells. Some authors do not recognize this genus and use Gonium sociale (Stein, 1959). T. socialis has been found in various localities of the United States (e.g., FL, NC, SC, TN, VA) (Smith, 1950; Stein, 1959; Dillard, 1989), and Canada (Duthie and Socha, 1976; Stein and Borden, 1979).

I. Spondylomoraceae Coenobic colonies of the Spondylomoraceae are composed of Chlamydomonas-like cells as in the Volvocaceae, the Goniaceae and the Tetrabaenaceae, but lack a gelatinous matrix surrounding the colony (Ettl, 1983; Bold and Wynne, 1985). Cells may be bior quadriflagellate (Ettl, 1983). Asexual reproduction takes place by autocolony formation by all the cells of the colony. Isogamous sexual reproduction is known for Pyrobotrys and Pascherina (Korshikov, 1928; Nozaki, 1986). Four genera in this family can be distinguished as follows:

1a.

Cells quadriflagellate (Fig. 40)....................................................................................................................................Spondylomorum

1b.

Cells biflagellate.................................................................................................................................................................................2

2a.

Colonial cells interconnected to each other by elongate protuberances of the cell walls (Fig. 41)......................................Chlorcorona

2b.

Colonial cells connected to each other by the direct attachment of cell walls......................................................................................3

3a.

Chloroplasts containing pyrenoids (Fig. 42)........................................................................................................................Pascherina

3b.

Chloroplasts lacking pyrenoids (Fig. 43).............................................................................................................................Pyrobotrys

Chlorcorona Fott (Fig. 41) The colonies contain eight cells arranged in two parallel, alternating rhomboid tiers of four cells each, without encompassing gelatinous matrix (Ettl, 1983; Hoops and Floyd, 1982). The cells have a cell wall and are interconnected to each other by elongate protuberances of the walls. Each cell is ovoid and has two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a cup-shaped chloroplast without pyrenoids (Ettl, 1983; Hoops and Floyd, 1982). Sexual reproduction is unknown. Chlorcorona

is a monotypic genus with C. bohemica (Fig. 41), and rarely found in freshwater habitats of Europe and the United States. This genus was originally described by Fott (1949) as Corone. However, this generic name is a homonym of Corone (Hoffmannseg ex H. G. L. Reichenbach) Fourreau (1868) and Corona Lefébure et Cheneviére (1938). Fott (1967) proposed Chlorcorona as a nomen novum for Corone Fott (1949). Recently, C. bohemica was collected in Ohio and its flagellar apparatus has been observed by electron microscopy (Hoops and Floyd, 1982).

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40

41

42 43 FIGURE 40 Vegetative colony of Spondylomorum quaternarium. (From Pascher, 1927.) (× 800.) FIGURE 41 Vegetative colony of Chlorcorona bohemica. (From Fott, 1949.) (× 1100.) FIGURE 42 Vegetative colony of Pascherina tetras. (From Korshikov, 1928.) (× 2100.) FIGURE 43 Vegetative colony of Pyrobotrys casinoensis. (From Nozaki, 1995, reproduced by permission of Koudan-Sha Scientific.) (× 950.)


Pascherina Silva (Fig. 42) Colonies are mulberry-shaped, containing four cells arranged in two alternating tiers of two cells each, without encompassing gelatinous matrix (Korshikov, 1928; Smith, 1950). Cells are walled and ellipsoidal to ovoid, each with two equal flagella, a stigma, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast with a single basal pyrenoid. Sexual reproduction is isogamous (Korshikov, 1928). Pascherina is infrequently collected from cool, eutrophic habitats and only P. tetras (Fig. 42) has been described. Silva (1959) published Pascherina as a nomen novum for Pascheriella Korshikov (1928), an illegimate homonym of Pascherella (Conrad, 1926). P. tetras has been observed in the United States (Smith, 1950; Dillard, 1989). Pyrobotrys Arnoldi (Fig. 43) Colonies are star- or mulberry-shaped, containing four, eight or 16 cells arranged in two or four tiers, without encompassing gelatinous matrix (Nozaki, 1986). Cells are spherical, subspherical, ovoid, ellipsoidal, pear-shaped or irregularly pear-shaped, each with two equal flagella, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast without pyrenoids. A stigma may be present in each cell of the colony. Cell walls are delicate with a papilla at the base of the flagella (Nozaki, 1986). In sexual reproduction, all the cells divide into four, eight, or 16 small, biflagellate isogametes that fuse to form planozygotes (Nozaki, 1986). Mature planozygotes are quadriflagellate with a large stigma and a speciesdependent form (Nozaki, 1986). Mature aplanozygotes are spherical with a heavy cell wall. On germination, the zygote gives rise to four biflagellate gone cells released separately (Nozaki, 1989b). Pyrobotrys is cosmopolitan and found in fresh water rich in organic matter. This genus contains 12 described species. Some species require anaerobic conditions for growth in culture (Nozaki, 1986). Pyrobotrys also appears in the literature under the names Uva Playfair (1914) or Chlamydobotrys Korshikov, (1924). Silva (1972), however, resolved this nomenclatural confusion and used the name Pyrobotrys Arnoldi (1916). P. casinoensis (Fig. 43) and P. stellata have been found in the United States (e.g., KY, VA) (Smith, 1950; Dillard, 1989). Planozygotes of P. casinoensis has been described as Chlorobrachis gracilima (e.g., Smith, 1950). Morphological description of Chlorobrachis gracilima by Smith (1950) seems to be incorrect; Figure 31 B is not “side view” of the alga (see Smith, 1950), but it is the immature stage of the alga (see Korshikov, 1925; Nozaki, 1986).

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Spondylomorum Ehrenberg (Fig. 40) Colonies are star- or mulberry-shaped, with eight or 16 cells arranged in four-celled tiers, without encompassing gelatinous matrix. Cells are walled and ovoid or pear shaped with a long posterior tail, each with four equal flagella, two contractile vacuoles at the base of the flagella, and a massive cup-shaped chloroplast without pyrenoids. Sexual reproduction is unknown. Spondylomorum is distinguished from other spondylomoraceaen genera by its quadriflagellate vegetative cells. This genus contains two described species and S. quaternarium (Fig. 40) has been recorded from various localities of the world (Huber-Pestalozzi, 1961). However, no culture studies have been carried out, and its existence has been questioned (Pringsheim, 1960). S. quaternarium has been found in the United States (Smith, 1950; Dillard, 1989) and Ontario, Canada (Duthie and Socha, 1976).

VI. GUIDE TO LITERATURE FOR SPECIES IDENTIFICATION Species identification of fixed materials is possible based on the taxonomic concepts of Smith (1950), Huber-Pestalozzi (1961), Ettl (1983), and Dillard (1989). Correct identification of species within the Volvocales often needs clonal cultured materials. Morphological characteristics of cultured materials observed under the controlled laboratory conditions provide stable and objective species diagnoses such as in Eudorina (Goldstein, 1964), Pyrobotrys (Nozaki, 1986), Carteria (Nozaki et al., 1994a), and Chlorogonium (Nozaki et al., 1998b). DNA sequence data, such as the rbcL gene, obtained from clonal cultures, may help confirm identification of species/genus within the volvocalean algae (see Nozaki et al., 1997a, 1998a, b). For general methods for clonal cultures of microalgae, see Stein (1975) and Starr and Zeikus (1993). The following is a list of key recent references, each of which contains citations to older literature and those from other continents: 1. Pedinomonas—Iyengar and Desikachary (1981), Ettl (1983) 2. Polyblepharides—Iyengar and Desikachary (1981), Ettl (1983) 3. Mesostigma—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 4. Scourfieldia—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 5. Nephroselmis-Iyengar and Desikachary (1981), Ettl (1983) 6. Spermatozopsis—Iyengar and Desikachary (1981), Ettl (1983), Preisig and Melkonian (1984)

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7. Polytomella—Iyengar and Desikachary (1981), Ettl (1983) 8. Tetraselmis—Ettl (1983) 9. Scherffelia—Iyengar and Desikachary (1981), Ettl (1983) 10. Haematococcus—Iyengar and Desikachary (1981), Ettl (1983), Thompson and Wujek (1989) 11. Stephanosphaera—Iyengar and Desikachary (1981), Ettl (1983) 12. Chlamydomonas—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 13. Chloromonas—Iyengar and Desikachary (1981), Ettl (1983), Pröschold et al. (2001) 14. Carteria—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989), Nozaki et al. (1994a) 15. Polytoma—Iyengar and Desikachary (1981), Ettl (1983) 16. Chlorogonium—Ettl (1983), Dillard (1989), Nozaki et al. (1998b) 17. Gloeomonas—Iyengar and Desikachary (1981), Ettl (1983) 18. Lobomonas—Iyengar and Desikachary (1981), Ettl (1983) 19. Vitreochlamys—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989), Nakazawa et al. (2001) 20. Pedinopera—Iyengar and Desikachary (1981), Ettl (1983) 21. Pteromonas—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 22. Phacotus—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 23. Wislouchiella—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 24. Cephalomonas—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 25. Thoracomonas—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 26. Granulochloris—Iyengar and Desikachary (1981), Fott (1963), Ettl (1983) 27. Coccomonas—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 28. Dysmorphococcus—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 29. Pandorina—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989), Nozaki and Kuroiwa (1991) 30. Volvulina—Iyengar and Desikachary (1981), Ettl (1983), Nozaki and Kuroiwa (1990) 31. Yamagishiella—Rayburn and Starr (1974), Nozaki and Kuoiwa (1992) 32. Eudorina—Iyengar and Desikachary (1981), Goldstein (1964), Ettl (1983)

33. Platydorina—Harris and Starr (1964) 34. Pleodorina—Iyengar and Desikachary (1981), Nozaki et al. (1989) 35. Volvox—Iyengar and Desikachary (1981), Smith (1944), Ettl (1983), Starr (1970), Dillard (1989) 36. Gonium—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989), Nozaki (1989) 37. Astrephomene— Nozaki (1983) 38. Basichlamys—Stein (1959), Iyengar and Desikachary (1981) 39. Tetrabaena—Stein (1959), Nozaki and Ohtani (1982) 40. Spondylomorum—Iyengar and Desikachary (1981), Ettl (1983) 41. Chlorcorona—Iyengar and Desikachary (1981), Ettl (1983) 42. Pascherina—Iyengar and Desikachary (1981), Ettl (1983), Dillard (1989) 43. Pyrobotrys —Iyengar and Desikachary (1981), Ettl (1983), Nozaki (1986)

ACKNOWLEDGMENTS I would like to thank Dr. Robert Sheath who kindly added many non-U.S. references to the distributions of the flagellated green algal genera and gave me valuable comments on the manuscript.

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NONMOTILE COCCOID AND COLONIAL GREEN ALGAE L. Elliot Shubert Department of Botany The Natural History Museum London SW7 5BD United Kingdom I. II. III. IV. V.

Introduction Diversity and Morphology Ecology and Distribution Collection and Preparation for Identification Key and Descriptions of Genera A. Key

The systematic investigation of the green algae has so far been largely a descriptive process; this applies not only to the nineteenth century when only the light microscope was in use but also to the twentieth century with the electron microscope. (Round, 1984, p. 9)

I. INTRODUCTION The nonmotile coccoid and colonial green algae belong to the division Chlorophyta. The Chlorophyta include a diversity of taxa (morphologically and ecologically), ranging from unicellular and freshwater taxa (e.g., Chlamydomonas; Chap. 6) to multicellular and marine taxa (e.g., Ulva). The nonmotile coccoid and colonial microscopic algae (hereafter referred to as “nonmotile greens”) represent a large subgroup currently distributed over five classes encompassing nine different orders. They are commonly called little green balls, because they often appear as aggregations or clumps of green cells. At first glance a novice might think that these organisms all look the same, but using careful and detailed observation reveals that there are stable and discrete morphological characters that separate many of the genera from each other (Prescott, 1978; Komárek and Fott, 1983; Dillard, 1999). NeverFreshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

VI.

B. Descriptions of Genera C. Addendum Guide to Literature for Species Identification Literature Cited

theless, numerous genera have been reassigned after detailed analysis with transmission electron microscopy (TEM) and/or molecular analysis (Watanabe and Floyd, 1989; Huss et al., 1999). Species identification is even more difficult because many species exhibit morphological variability or phenotypic plasticity, as well as motile stages in part of their life cycle. Although the taxa are nonmotile in the vegetative state (actively growing), some species produce flagellated stages or resting stages during part of their life cycle (Sect. II). Thus, for a definitive determination of species it may be necessary to culture the organism under controlled environmental conditions, to examine it at the ultrastructural level with TEM and/or scanning electron microscopy (SEM), or to use molecular methods. The nonmotile greens include many ubiquitous species, which are found worldwide in a variety of habitats (see Sect. III). Whether distributed by air currents, birds, or the feet of animals, they are effective colonizers of denuded soils (e.g., disturbed by human activities), new soils (e.g., volcanic lava), and newly formed water bodies (e.g., refilled potholes and depressions formed by mechanical disturbance). Thus, they play an important role in primary and secondary successional processes. 253

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There are approximately 200 recognized genera and thousands of described species in this group (Komárek and Fott, 1983; Ettl and Gärtner, 1995). Unfortunately, the proliferation of species names, such as in Scenedesmus (Hegewald and Silva, 1988), has created more confusion rather than a better understanding of the taxa. The problem arises because the classical taxonomic approach relies on differences in morphological characters of organisms collected from the field and the assumption is that these characters are stable. However, when these same organisms are isolated into axenic culture and grown under different environmental conditions (e.g., light, temperature, and nutrients) a range of morphological types can be expressed (Trainor, 1998). Phenotypic plasticity has been well documented for some taxa. The elucidation of the range of morphological types in taxa must be considered if we are to gain a better understanding of their systematics, phylogenetic relationships and eco-physiological roles. Some of the algae are considered to be “green weeds” (e.g., Chlorella, Chlorococcum Scenedesmus, and Desmodesmus), because they can be easily cultured in the laboratory or can produce bloom conditions in the field. Indeed, the first report of an alga cultured from field material was that of Chlorella (Beijerinck, 1890).

II. DIVERSITY AND MORPHOLOGY The vegetative cells of nonmotile greens are eukaryotic and either uni- or multinucleate (coenocytic). All are nonmotile in the vegetative state. When cell division occurs in coccoid and colonial algae, the progeny cells form their own walls. When the new cell or colony is released, the empty remains of the parent cell wall (“ghost cell”) can be observed (e.g., Desmodesmus and Pediastrum). The dead remains of Desmodesmus and Pediastrum may be persistent for a considerable time in sediments, due to decay resistant compounds, such as algaenans and sporopollenin, and are still readily identifiable (Pickett-Heaps and Staehelin, 1975; Graham and Wilcox, 2000). In some cases these algae can withstand degradation for millions of years, and are associated with mudstones and oil deposits (Fleming, 1989; Gelin et al., 1997). In some taxa the remains of the parent wall become part of the progeny cells (e.g., Dictyosphaerium) and are distinctive enough to be used as a taxonomic character. The vegetative cells of some taxa (uninucleate) are embedded in mucilage (homogenous or lamellated), and pseudoflagella (giving the appearance of flagella, but not functional) are often present and visible with the light microscope (LM). Other vegetative cells have a range of morphology, that is, unicells to colonies

(uni- or multinucleated), and mucilage may be present, but there are no pseudoflagella. Some colonial forms have a well-developed mucilaginous sheath (Dillard, 1989). Motile cells may be present in some genera (e.g., Bracetococcus, Characium, Chlorococcum, Myrmecia, and Tetracystis) and most are chlamydomonad in appearance (two whiplash flagella of nearly equal length, contractile vacuoles, basal bodies, persistent eye spot, and a cup-shaped chloroplast). The motile cells may function as zoospores (asexual) or gametes (sexual). If sexual reproduction occurs, it can be either isogamous (similar morphology) or anisogamous (dissimilar morphology). The nonmotile greens have a zygotic meiosis life cycle. The nonmotile greens share characteristics common to all eukaryotic green algae, such as wall structure, pigments, storage products, and motility. They contain at least one plastid and are autotrophic. Some taxa are facultative heterotrophs. The dominant pigments are chlorophyll-a and -b, which give the cell a grass-green appearance. The light harvesting pigment complex is associated with proteins, which bind chlorophyll-a and -b, and accessory pigments (e.g., carotenoids). The accessory pigments may function to harvest light, but more typically provide protection from photo-oxidation and dispersal of excess energy. The major accessory pigment is lutein or a derivative, and β-carotene is always present. When a cell ages in the absence of essential nutrients, it may appear yellow to orange in color. Chlorosarcinopsis appears bright orange (presumably carotenoid production) when the culture is deficient in nitrogen (Starks and Shubert, 1979). There may be one to several chloroplasts per cell. The shape of the chloroplast is often used to distinguish among genera that have similar cell shapes. For example, the spherical unicells Chlorococcum and Trebouxia are distinguishable from one another because the former has a cup-shaped chloroplast and the latter has a starshaped chloroplast (Dillard, 1989). The storage product for members of this group is true starch, amylose, and amylopectin (α-1,4-linked polyglucans), and is found inside the chloroplasts. The starch can be detected with a drop of I2KI, which turns dark blue–black. Staining for starch is a diagnostic tool for distinguishing green algae from morphologically similar algae that belong to other groups, but do not store starch. The starch (seen as whitish granules with the TEM) can often be observed surrounding the pyrenoid, a distinct spherical structure embedded in the chloroplast. There may be more than one pyrenoid or the pyrenoid is not always present (e.g., Ankistrodesmus and Tetraedron) or the pyrenoid is lacking (e.g., Bracteococcus and Cerasterias). It may be used as a

7. Nonmotile Coccoid and Colonial Green Algae

taxonomic character to distinguish between some genera (e.g., Palmodictyon and Tetraspora; Dillard, 1989). Cells collected from nature may be full of food reserve, obscuring the plastid. This may make it difficult to determine plastid type, which is a problem if this character is necessary for identification to genus or species. Archibald and Bold (1970) solved this problem by culturing isolates in high levels of nitrogen Bold’s Basal Medium (BBM). However, Blackwell et al. (1991) pointed out that “there are inherent dangers in assuming, a priori, that all members of a taxon will respond to particular growth conditions...” after they tried to grow C. submarinum in BBM and were unsuccessful. In the end, for an individual taxon, we have no data on possible modifications of plastid structure at low, intermediate, and high nitrogen levels. It is advisable to consider growth under standard conditions as a recommendation rather than an absolute requirement for determining a taxon. Nevertheless, culturing an organism may provide valuable information on its range of phenotypic plasticity (including the production of flagellated stages) and physiological tolerances under different environmental conditions. Chlorococcum is considered a type genus for the chlorococcalean algal group (Bold and Wynne, 1978). Although this group of algae is nonmotile in the vegetative state, some taxa produce motile cells (planospores). Planospores may be asexual zoospores or sexual gametes. To form motile cells, the vegetative cell undergoes multiple cleavages of the protoplasm and the nucleus. The resulting division together with associated organelles becomes surrounded by a cytoplasmic membrane and a cell wall (sometimes they are “naked” because the wall is absent). Trainor (1978) described a number of possible outcomes when planospore production occurs, depending on the way in which they are released (e.g., singly through a pore or breakdown of a wall, or released as a group within a flexible membrane, the vesicle). A single planospore would be a unicell such as Chlorococcum, whereas a group of planospores would be unicells or a colony, such as Hormotilopsis and Pediastrum, respectively. If the planospores are retained within the parent cell wall and attach to each other, the organism would be colonial such as Hydrodictyon. These distinctions can be very helpful when observing live algae collected from the field or mixed cultures in the laboratory with respect to identification and understanding the reproductive process. Alternatively, the method of formation of aplanospores (nonmotile cells) determines the morphological outcome (Trainor, 1978). Cells released singly result in a unicellular organism (e.g., Ankistrodesmus, Chlorella, and Chlorococcum), but if the progeny cells are

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retained and remain attached to each other, then a colony is produced (e.g., Coelastrum and Scenedesmus). Chlorococcum is found in both aquatic and terrestrial habitats, and can produce both planospores and aplanospores in culture. Trainor (1978) suggested that sufficient nutrients and water might “trigger” the formation of planospores in Chlorococcum, whereas in older cultures or in a dry soil, aplanospores would develop. “Aplanospores have the ontogenetic potentiality for being flagellate and motile” (Bold and Wynne, 1978). Isogamous sexual reproduction occurs in Chlorococcum and the gametes are morphologically similar to zoospores, but function as sex cells. In most representative taxa, the cells are surrounded by a cellulose cell wall. Some taxa may also have chitin or sporopollenin deposited on the wall. This gives added strength and is thought to help prevent desiccation (Graham and Wilcox, 2000). Some taxa have wall ornamentation, such as scales, a rough texture, thick walls with distinct layers, warts, ridges, and spines (Trainor, 1978). Some of these structures can be seen with the LM (e.g., ridges and spines), whereas others are visible only with TEM and SEM (e.g., scales and warts). Wall ornamentation is often used as a taxonomic character (e.g., number and position of spines); however, with some taxa this may not be a stable character, particularly at the species level. Trainor and Egan (1990a, b) demonstrated cyclomorphosis in an axenic clone of Desmodesmus armatus (as Scenedesmus armatus), which exhibited a variety of spine patterns that ranged from two spines on each of the terminal cells, to one spine on each of the terminal cells, to no spines on any cells (spineless). Some of these morphs could be identified as different species if spine number and pattern were assumed to be stable characters. Thus, caution is advised when making species identifications from field material, because some taxa (e.g., Chlorococcum, Desmodesmus, and Scenedesmus) have a variable morphology. To complicate matters, unicellular morphs of Desmodesmus have been misidentified as other genera: Lagerheimia (Hegewald and Schmidt, 1987, 1991; Trainor and Egan, 1990c; Trainor, 1991), Franceia, or Oocystis. This is unfortunate and is the consequence of relying solely on field material for identification (Smith, 1950) rather than using axenic cultures or considering cyclomorphosis and phenotypic plasticity (Hegewald and Silva, 1988). Subsequent investigations, beginning in the 1960s, by Trainor and co-workers (summarized in Trainor, 1998) unequivocally demonstrated that the colony form of Desmodesmus could produce unicells. The unicells are formed within the parent wall and are released as new progeny. Alternatively, unicells can produce more unicells or colonies, depending on the environmental conditions

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(Trainor, 1998). Shubert (1975) described a new species, Scenedesmus trainorii, based on a strain (UTEX 1588) that showed phenotypic plasticity. The alternation between colonies and unicells was mediated by modifying the concentration of phosphorus (Shubert and Trainor, 1974). Forma designation was used to differentiate between the unicellular stage, f. trainorii, and the colonial stage, f. quadricauda. Komárek and Fott (1983) dropped the forma designation because they considered that the ecomorphs had no taxonomic significance. In rejecting the forma designation, they are ignoring morphological plasticity in responseknown environmental variables. Hegewald and Silva (1988) considered f. quadricauda to be an invalid name because it shared the type strain with f. trainorii and was a typical form (e.g., four-celled stage), thus required no taxonomic recognition. The decisions by Komárek and Fott (1983) and Hegewald and Silva (1988) imply that forma is a redundant rank, but the International Botanical Code permits the use of forma to discriminate between contrasting morphs within a species. My opinion is that the genera Franceia and Lagerheimia require revalidation as distinct taxa using modern taxonomical approaches, such as combining axenic culture work with ultrastructural and molecular analysis. A variety of morphological characters have been used to define genera, including shape of the vegetative cell, structure and position of the chloroplast, number of nuclei per cell, presence or absence of pyrenoids, and presence or absence of flagellated cells (Starr, 1955). In theory this makes sense, because a certain combination of characters should describe a genus. For example, a walled planospore with flagella of equal length, which is the reproductive cell of an organism with a cupshaped chloroplast, was called Chlorococcum. Alternatively, a naked planospore from a cell with a netlike plastid was called Spongiochloris. However, despite being insightful, this does not always work in practice. As Trainor (1978) aptly pointed out, it could be difficult to identify plastid type if food reserves obscure it and/or flagellated stages might not always be observed, especially from live field material. So, do we make identification on a few live cells or fixed material? Before this question is answered, let us consider a few examples. Starr (1955) described a new monospecific genus, Neochloris, using a variety of morphological characters. Subsequently, 12 new species were added. Much later, Watanabe and Floyd (1989) studied nine species of Neochloris and divided them into three groups based on morphological characters. They investigated the ultrastructure of the motile cells by focusing on the flagellar apparatus (FA) and discovered that the basal bodies could be arranged three different ways: clock-

wise, (CW), counterclockwise (CCW), and directly opposite (DO). Using this stable FA character, the species did not group in the same way. Some species were retained with Neochloris (DO) and some were assigned to other genera [Chlorococcopsis (CW) and Parietochloris (CCW); Watanabe and Floyd, 1989]. Subsequently, Lewis et al. (1992) analyzed the same taxa for 18S rRNA and the results were concordant with the designation by Watanabe and Floyd (1989). However, examination of other characters in Neochloris and related taxa has created uncertainty (Deason et al., 1991; Kouwets, 1995). The classification of former species of Neochloris remains unresolved (Trainor and Morales, 1999). Lewis (1997) applied molecular techniques (18S rRNA) to Bracteococcus to determine its placement in the “larger picture of chlorophycean algae” because the motile cells have a FA of unusual orientation (CW). Lewis’ (1997) 18S sequence data confirmed that all nine species analyzed supported a clade with bootstrap support of 95% and were monophyletic, which demonstrated that the morphological characters used to distinguish the genus from other chlorococcalean algae were “reliable.” However, Bracteococcus has been interpreted to have CW basal body orientation and the 18S data pointed toward taxa with DO basal body orientation. Also, two Chlorococcum species did not form a clade, and the multinucleated zoosporic CW taxa Spongiochloris and Protosiphon “nested” with the uninucleated zoosporic CW taxa (Lewis, 1997). These data suggest that molecular characterization (e.g., 18S rRNA gene sequence alone) may not support the present classification based on morphological characters. A suite of stable characters (morphological, ultrastructural, biochemical, and molecular) may be required for the determination of some specific taxa. Trainor et al. (1976) proposed that Scenedesmus be divided into two groups—the obliquus or nonspiny type and the spiny type—based on morphological (LM and TEM/SEM) and reproductive characters. However, they took the conservative approach to taxonomy and postponed formal establishment of generic level taxa until more data were available. Kessler et al. (1997) reported that biochemical and physiological properties were not suitable for species differentiation of Scenedesmus, but they did identify two subgenera of Scenedesmus (Scenedesmus, nonspiny type and Desmodesmus, spiny type) using 16S rRNA gene sequence, DNA base composition, and DNA/DNA hybridization analysis. An et al. (1999) analyzed a more conserved region of DNA (ITS-2 rDNA sequence) in selected species of Scenedesmus and resolved a distinct separation between the subgenera, Scenedesmus and Desmodesmus, and proposed the recognition of both groups


as distinct genera. The ITS-2 gene sequence analysis was congruent with cell wall ultrastructure, but not with other morphological features (An et al., 1999). Thus, molecular data confirmed what had been originally proposed using morphological and reproductive characters. Taxa of the unicellular green alga Chlorella have been the organisms of choice for a variety of physiological and biochemical studies, including photosynthesis and nitrate reduction. However, the lack of obvious morphological characters combined with asexual reproduction only has created problems in delineating species. Numerous methods have been used, such as nutritional requirements, numerical classification strategies, combination of morphological and structural features, serological cross reactions, ultrastructure and chemical composition of the cell wall, pyrenoid ultrastructure, and a combination of biochemical and physiological characters (reviewed by Huss et al., 1999). The most convincing taxonomical scheme to date is based on a multimethod approach that compares 18S rRNA gene sequences, DNA base composition, and DNA hybridization values, combined with biochemical, physiological, and ultrastructure characters (Huss and Sogin, 1990; Huss et al., 1999). The results of Huss and co-workers showed that 19 Chlorella taxa were polyphyletic and dispersed over two classes of green algae, and they proposed that only four species be kept in the genus (glucosamine as a dominant cell wall component and presence of a double thylakoid bisecting the pyrenoid matrix), the other species being assigned to other genera (production of secondary carotenoids under nitrogen-deficient conditions (Huss et al., 1999). Molecular and TEM investigations of nonmotile greens have shown that many genera are polyphyletic. It is beyond the scope of this chapter to explore the systematics and evolution of green algae and phylogenetic relationships in detail, and the reader is advised to consult Mattox and Stewart (1984), Kantz et al. (1990), Wilcox et al. (1992), Floyd et al. (1993), Nozaki (1993), Friedl and Zeltner (1994), O’Kelly et al. (1994), Friedl (1995, 1997), Melkonian and Surek (1995), Nakayama et al. (1996), Booton et al. (1998), Hanagata (1998), Krienitz et al. (1999), Preisig (1999), and Sluiman and Guihal (1999). Thus, an extension of the classical approach to the taxonomy of nonmotile greens might use the following holistic protocol: Initial identification of field material (ideally live) is determined with the LM; cells are isolated into axenic culture, grown under different environmental conditions, and phenotypic plasticity, and biochemical and physiological changes are recorded; ultrastructural features are documented with confocal laser scanning microscope (CLFM), TEM, and/or SEM; and molecular analysis is conducted for

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the final determination of species (Trainor and Morales, 1999). However, in the not too distant future this entire procedure might be abbreviated by simply placing a drop of water from a pond, lake, or ocean on a surface containing fluorescent nucleic acid probes for identification of the major species present (Miller and Scholin, 1998). Nonmotile colonial or tetrasporalean algae often resemble the motile colonial algae. Bold and Wynne (1978) identified a problem with Gloeococcus “in which the flagellate cells move slowly within nonmotile colonies; this occurs also in the palmelloid phases of Chlamydomonas.” Further, they stated that “the decision where to classify a given organism may be difficult and is often subjective insofar as some tetrasporalean algae are concerned.” The holistic approach, as previously described, may give a definitive answer Despite the variable morphology, the inconsistent appearance of reproductive cells, and the similarity of morphological forms, it is possible to identify most genera of nonmotile coccoid and colonial green algae (live or fixed) with a classical taxonomic key and a LM. When making microscopic observations (preferably using phase contrast or differential interference contrast with a 40× or 60× dry objective or 100× oil objective), a series of questions should be asked to determine the genus, such as what is the cell shape, what is the chloroplast structure, are pyrenoids present or absent, are flagellated cells present or absent, are there any unusual features of the cell, are there distinguishing external wall structures, is a mucilaginous sheath present, is it free-living or attached, what is its habitat? With experience, researchers will gain the confidence to successfully identify many of the nonmotile green algae found in North America. TEM and SEM have been used for the identification of species, but it is beyond the scope of this chapter to discuss this approach. The reader is encouraged to consult PickettHeaps (1975), who has produced an extensive treatise on the cell biology and ultrastructure of green algae.

III. ECOLOGY AND DISTRIBUTION The nonmotile greens are ubiquitous and widely distributed in aquatic habitats throughout the North American continent. Aquatic habitats where these algae are found include temporary pools of water, waterfalls, ponds and lakes (e.g., Ankistrodesmus, Coelastrum, and Scenedesmus), rivers (e.g., Actinastrum and Dictyosphaerium), slow flowing streams (e.g., Hydrodictyon), marshes, and estuaries. Nonmotile greens are primarily a component of the plankton community (e.g., Desmodesmus, Golenkinia, Pediastrum, and

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Scenedesmus). They also may be found growing attached to rocks in lakes (e.g., Apatococcus and Desmococcus), on leaf surfaces or on other algae (e.g., Apiocyctis and Characium), and/or on the sediment surface. In the temperate zone, they are most abundant in freshwater ecosystems during the summer, when light and temperature are near their seasonal maximum and nutrients become limiting (e.g., N and P). Reynolds (1984) characterized freshwater plankton from temperate latitudes using trophic status and showed that green algae may be important in phytoplankton dynamics from spring through to autumn. HappeyWood (1988) reviewed the ecology of freshwater planktonic green algae. In high altitude lakes, plankton may be detectable in the spring under considerable cover of ice and snow, and small coccoid greens (e.g., Chlorella and Coccomyxa) are often the most common components. During the summer, in oligotrophic lakes, the smaller sized fractions of unicellular green algae appear as the dominant organisms (e.g., Sphaerocystis and Gloeocystis). In contrast, in well-mixed shallow eutrophic lakes, larger green algae (e.g., Coelastrum, Dictyosphaerium, and Pediastrum) may dominate during the summer. With respect to the seasonal successional process in oligotrophic to mesotrophic lakes, nonmotile greens appear to be restricted to a relatively short growth period defined by a narrow range of environmental conditions within which to successfully compete with a mixed assemblage of phytoplankton (Happey-Wood, 1988). This restricted period occurs during the early stratification of the lake when light is not limiting, and turbulence and turbidity (e.g., suspended solids or shading by other algae) are minimal. Once stratification is stabilized, the nonmotile greens begin to sink and decline in the water column, and sedimentation increases. Thus, their survival in the euphotic zone is dependent on the residence time together with the available nutrients in the epilimnion (Happey-Wood, 1988). Nonmotile greens have numerous adaptations to reduce sinking rates, including a mucilaginous sheath, wall ornamentation (e.g., bristles and spines), number of cells in a colony, and a shape other than spherical (Reynolds, 1984). Some nonmotile green algae (e.g., Chlorococcum submarinum and Chlorella) are capable of growing in saline conditions (e.g., inland saline lakes, estuaries, and marine coastal habitats; Shubert, 1998). Subaerial habitats also support many nonmotile green taxa, which may be easily observed growing on tree bark, wood fences, stone walls, gravestones, plaster walls, and so forth; in fact, any surface that holds some moisture. In extreme cases, this may result in damage to the surface over time (e.g., frescoes). Also, nonmotile green algae grow symbiotically with fungi in

lichens (e.g., Trebouxia, a phycobiont of many lichens, e.g., Cladonia), and with plants and animals (e.g., Chlorella in Hydra viridis, Paramecium bursaria, and Spongilla lacustris). Soil is a common habitat for nonmotile green algae (e.g., Apatococcus, Chlorococcum, Chlorosarcinopsis, and Tetracystis), which are widely distributed in a variety of soil types and microclimates (Starks et al., 1981). Chlorococcum is a common taxon found in soils. Archibald and Bold (1970) differentiated species of Chlorococcum by isolating clones into liquid culture medium (BBM) and scoring their response to various physiological/biochemical attributes (e.g., sensitivity to antibiotics, production of extracellular proteases, and amylase). Although this approach is scientifically sound, it can be problematical and appears to be little used by contemporary phycologists. Some algae grow on the surface of moist, bare soil (e.g., Protosiphon). As the soil dries, Protosiphon becomes orange–red (Bold and Wynne, 1978).

IV. COLLECTION AND PREPARATION FOR IDENTIFICATION Collecting planktonic nonmotile coccoid and colonial green algae is rather easy and straightforward. Whereas they are suspended in the water column, a plankton net with a mesh size of 20 µm is adequate for most taxa (nannoplankton and net plankton). A bottle sample can be taken, which is less concentrated and may require time to settle, but does allow quantification. Mosses, and stems and leaves of aquatic plants can be collected or hand squeezed for analysis of epiphytes. Algae on damp rock surfaces, walls, or bark can be removed with a penknife or other sharp instrument. Water from the habitat or distilled water should be added to the specimen vial. Soil samples can be collected with a trowel and stored in sterile plastic bags. Algae generally survive well after collection if the container is clear glass, illuminated, and kept cool (15–20°C) in an environmental growth chamber or on a window ledge with a northern exposure. Alternatively, culturing field material may yield a range of morphologies and/or reproductive structures for some genera (e.g., Chlorococcum, Desmodesmus, and Pediastrum) that may aid in identification of species. Stein (1973) described isolation and culture methods and media for growing algae. In addition, Prescott (1978) and Dillard (1999) may be consulted for additional information on the collection and preservation of algae. Wet mounts of fresh samples for microscopic analysis give the best results when a number 1 coverslip and at least 400× magnification are used. Resolution


can be enhanced with phase contrast or differential interference contrast. A 60× dry objective is preferable, because superior detail of the cells can be observed without the use of oil. If greater resolution is required (which is often the case with Chlorella and other extremely small green unicells), a 100× oil immersion objective may be used. For enhanced resolution, it is recommended that an additional drop of oil be placed on the condenser lens and the condenser carefully raised until the oil makes contact with the bottom of the slide. For health and safety reasons, the microscope oil should be labeled as PCB-free. If live material is not observed within 2–3 days after collection, it is recommended that the sample be fixed. The most common fixative is Lugol’s solution [2 g KI + 1 g resublimed I +300 mL distilled water (DW); it can be acidified with 10 mL glacial acetic acid or made neutral or slightly basic with 1 g of sodium acetate; it should be stored in a dark bottle at room temperature]. Lugol’s is very good for preserving small algae and flagellates. Lugol’s will stain starch dark blue–black, which is a simple test to separate the Chlorophyta from the Chrysophyta (which do not have true starch), because some look morphologically similar to the green algae. Add just enough drops of Lugol’s to give a pale straw color to the sample and store sam-

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ples in a cool dark place. M3 fixative is superior to Lugol’s and is excellent for long preservation. M3 fixative is made by adding 1 g I2 + 0.5 g KI + 5 mL glacial acetic acid +25 mL formalin +100 mL DW (R. Meyer, personal communication). Another simple fixative is 2–4% neutralized formalin. If samples are to be observed with epifluorescence, TEM, and/or SEM, EM grade glutaraldehyde (2.5–5% per volume) is the best choice. A cautionary note, glutaraldehyde is a strong fixative and human exposure should be minimized (e.g., wet mounts and light microscopy), because it can fix eye tissue. Another good fixative is FAA (a ratio of 10:7:2:1 parts of 95% ethanol: DW: formalin: acetic acid).

V. KEY AND DESCRIPTIONS OF GENERA A. Key This dichotomous taxonomic key has been constructed to be “user friendly” and is based on morphological characters seen with the LM. For some taxa, especially the spherical unicells, identification from field material may be difficult. Isolation and culturing of the organism may be required to observe motile cells.

1a.

Cells forming macroscopic growths……………………………………………………...........................................................................2

1b.

Cells solitary and/or not forming macroscopic growths………………………………..........................................................................8

2a.

Free-floating…………………………………………………………………………..................................................................................3

2b.

Usually attached………………………………………………………………………................................................................................4

3a.

Cells elongate, forming a netlike reticulum (Fig. 15B)…………………...........................................................................Hydrodictyon

3b.

Cells spherical, often with several long, fine pseudoflagella, arranged in groups of two or four; fragments of old parental walls may be visible (Fig. 24C)………......................................................................................................................................Schizochlamydella

4a.

Colony leaflike or a gelatinous sac or matrix…………………………………………............................................................................5

4b.

Colony not as above, but with cells embedded in stratified mucilage………………............................................................................7

5a.

Colony leaflike, cells in packets of two at the surface, lacking pseudoflagella (Fig. 21E)……………….......................……Phyllogloea

5b.

Colony a gelatinous sac or matrix…………………...................................………........................................................................……6

6a.

Cells arranged in packets of four (occasionally pairs) arranged around the edge of the colony, each cell with two pseudoflagella (Fig. 26F)…..............................................................................................................................................................………Tetraspora

6b.

Cells not in packets of four, cells feebly motile within gelatinous matrix (Fig. 13C)......................................................…Gloeococcus

7a.

Cells spherical, embedded in short, dichotomously branching, stratified gelatinous stalks, chloroplasts stellate (Fig. 15A) ….....................................................................................................................................................................................…Hormotila

7b.

Cells ellipsoidal, embedded in extensive, anastomosing, stratified gelatinous strands, chloroplasts cup-shaped (Fig. 14A) …...........................................................................................................................................................................……Gloeodendron

8a.

Cells growing within plants (endophytic) or animals (endozooic; including egg masses)………………........................................……9

8b.

Cells not growing endophytically or endozooically…………………………………............................................................................13

9a.

Cells growing in animals or egg masses…………………………………………........................................……...................................10

9b.

Cells growing among surficial cells of plants…………………......................................................................……........………………11

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10a.

Small spherical cells with a cup- or platelike chloroplast in protozoa or invertebrates (Fig. 5B)…......................................…Chlorella

10b.

Cells spherical to ellipsoidal with an axial, irregularly lobed chloroplast, in amphibian egg masses (often giving these a green color; Fig. 19D)….....................................................................................................................................................................……Oophilia

11a.

Largely colorless, threadlike growth on and within Sphagnum, chloroplasts concentrated in expanded ends of cells (Fig. 21D) ………………………….......................................................................................................................................................Phyllobium

11b.

Growth not threadlike, cells rounded…………………………………………………...........................................................................12

12a.

Cells irregularly oval; chloroplast cuplike in young cells, massive and diffuse in older cells, with thick lamellate walls, growing between host cells of Lemna and mosses (Fig. 5D)…………………………………………………..................................Chlorochytrium

12b.

Cells globular or flask-shaped, with rhizoidal lobes and extensions; parasitic within Ambrosia; protoplast reddish orange (Fig. 24A) …………..................................................................................................................................................................…Rhodochytrium

13a.

Cells or microscopic colonies attached to a substratum…………………....................................................................................……14

13b.

Cells or microscopic colonies not attached to a substratum………………................................................................................…….30

14a.

Colonies or aggregations of cells…………………………….........................................................................................………………15

14b.

Cells attached as individuals, without a communal point of attachment…………….........................................................................20

15a.

Cells attached at end of branching mucilaginous stalks, forming treelike colonies; sometimes attached to small crustacea or insect larvae (Fig. 5A)…........................................................................................................................................................…Chlorangiella

15b.

Cells not on branching stalks, occurring in an amorphous matrix, mucilaginous vesicle, or forming disklike thallus on the substratum………………….................................................................................................................................................………16

16a.

Colony mucilaginous, often amorphous or bulbous………………….........................................................................................……17

16b.

Colony not mucilaginous, closely appressed to substratum…………….....................................................................................……19

17a.

Colony pear-shaped or bulbous vesicle containing cells in pairs or fours with pseudoflagella projecting well beyond the colony envelope (Fig. 2C)………............................................................................................................................................................Apiocystis

17b.

Colony amorphous, cells lacking pseudoflagella……………………..............................................................................................…18

18a.

Cells embedded in concentrically stratified mucilage; may contain hematochrome (red pigment); often terrestrial (Fig. 20A) ............................................................................................................................................................................................…Palmella

18b.

Cells not embedded in stratified mucilage; aquatic (Fig. 20C)……................................................................................…Palmellopsis

19a.

Cells forming a pseudoparenchymatous small circular disk, bearing long erect pseudoflagella; chloroplast cup-shaped (Fig. 4C)……………......................................................................................................................................................…Chaetopeltis

19b.

Cells growing as a single layer of cells or pseudoparenchymatous disk of filaments; pseudoflagella absent; chloroplast parietal (Fig. 22D)....…………....................................................................................................................................................…Protoderma

20a.

Cells saccate, one end forming a rhizoid-like extension, the other end bulbous; chloroplast reticulate (Fig. 22E)……...…Protosiphon

20b.

Cells not as above, attached by well-defined stalk or over large part of cell surface..........................................................................21

21a.

Cells with a well-defined stalklike extension for attachment…………………………........................................................................22

21b.

Cells flattened onto substratum or attached by a broad face or pad…………………........................................................................25

22a.

Cells elongate–oval to fusiform, attached by a broad disk formed external to the cell wall (Fig. 4E)........................…Characiochloris

22b.

Attachment by stalk or pad that is part of the cell wall……......................................................................................................……23

23a.

Cells small, with a very fine tapering attachment; epiplanktonic; cup-shaped chloroplast with a single pyrenoid (Fig. 26A) ……………............................................................................................................................................…….....……Stylosphaeridium

23b.

Cells with a flattened pad where they attach…………………………………………...........................................................................24

24a.

Cells spherical, elliptical, or elongate with one to many disk-shaped chloroplasts lacking pyrenoids (Fig. 4F)…..............Characiopsis

24b.

Cells straight, slightly crescent-shaped, or sigmoid, with one to several parietal chloroplasts with pyrenoids; may be epiplanktonic (Fig. 4G)…......................................................................................................................................................................…Characium

25a.

Cells more or less spherical attached by a broad base or broad mucilage pad………........................................................................26

25b.

Cells ellipsoidal to flattened, or in a flattened casing………………………………….........................................................................27

26a.

Cells with anterior end down, attached by wide mucilage pad and enclosed in brown sheath; chloroplast in upper part of cell (Fig. 17A)…………..................................................................................................................................................……Malleochloris


261

26b.

Cells on broad attaching base; stalk almost lacking, thick brown sheath around, and much larger than, protoplast (Fig. 6A) …………….......................................................................................................................................................………Chlorophysema

27a.

Cells in angular casing or with angular outline……………...................................………………………….......................................28

27b.

Cells rounded (not angled) in face view…………………...................................……………………………........................................29

28a.

Cell with a cup-shaped chloroplast enclosed in flattened quadrangular casing with pores at corners through which pseudoflagella emerge (Fig. 22C)………....................................................................................................................................................Porochloris

28b.

Cells with octagonal outline and bipartite wall (Fig. 19A)……………..............................................................................Octagoniella

29a.

Cells lenticular, attached to submerged leaves of higher plants; cell wall thin against host, but thicker on free face (Fig. 10B) ……………………………………..........................................................................................................................................Ectogeron

29b.

Cells ellipsoidal, occurring in smooth, rounded casings with brownish encrustation around edge, on filamentous algae (Fig. 5C) …….................................................................................................................................................................................…Chloremys

30a.

Cells with spines or protuberances…………………………………………………...............................................................................31

30b.

Cells lacking spines or protuberances……………………………………………...................................…...........................................52

31a.

Cells solitary…………………………………………………………………………................................................................................40

31b.

Cells aggregated or forming colonies or coenobia……………………………………..........................................................................32

32a.

Cells regularly arranged, in pairs, fours, or multiples of four………………………...........................................................................33

32b.

Cells irregularly arranged colonies, sometimes in mucilage or cells attached to each other by strands……...................................…37

33a.

Cells arranged in pairs…………………………………………………………………............................................................................34

33b.

Cells usually fours or multiples of four, or if pairs, with few ( 600 described species; van Landingham, 1978), with some of the most difficult species to identify. Although hundreds of species of the genus Nitzschia have been described thus far, many appear to be synonyms, as new names are attributed to previously described taxa. This genus has received the attention of several taxonomists (LangeBertalot and Krammer, 1993; Krammer and LangeBertalot, 1988), but remains a difficult genus to work with at the species level. Genera belonging to the order Rhopalodiales, Epithemia and Rhopalodia, are asymmetric to the apical axis in valve view and possess a raphe that occupies a canal connecting with the interior of the cell via round or oval holes (portulae) (Round et al., 1990). In addition, the raphe of some species of Rhopalodia is also slightly elevated from the valve surface into a keel 669

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(Round et al., 1990). Cells are usually relatively large and heavily silicified. Most, and perhaps all, species in this order contain endocellular symbiotic cyanobacteria (blue green algae) (Drum and Pankratz, 1965; Geitler, 1977). Diatom cells containing endosymbiotic cyanobacteria have been shown to be capable of fixing nitrogen (Floener and Bothe, 1980). DeYoe et al. (1992) suggested that the endosymbionts have been modified into nitrogen-fixing “organelles” and their intracellular density is partially a function of the diatom’s nitrogen needs.

II. DIVERSITY AND MORPHOLOGY Diatom orders and genera considered in this chapter are listed in Table I. Diatoms in the orders Bacillariales, Surirellales, and Rhopalodiales share common pigments (chlorophylls and carotenoids) (Stoermer and Julius, 2002) with other diatoms (Bacillariophyta). Most taxa in this group are benthic in habitat and may inhabit dimly lighted microhabitats in deep lakes or turbid rivers and frequently maintain increased quantities of accessory pigments in such situations. Deep-living members of the Surirellaceae in particular are often deep chocolate brown in color (Round et al., 1990).

TABLE I Keeled or Canal-Bearing Freshwater Diatoms of North America Class

Order

Family

Genus

Bacillariophycidae Bacillariales Bacillariaceae Bacillaria Gmelin Cylindrotheca Rabenhorst Cymbellonitzschia Hustedt Denticula Ehrenberg Hantzschia Grunow Nitzschia Hassall Tryblionella W. Smith Surirellales Entomoneidaceae Entomoneis Ehrenberg Surirellaceae Campylodiscus Ehrenberg ex Kützing Cymatopleura W. Smith Stenopterobia Brébisson ex Van Heurck Surirella Turpin Rhopalodiales Rhopalodiaceae Epithemia Brébisson Rhopalodia O. Müller

Taxa within these three orders vary greatly in size. For example, the smallest members of the genus Nitzschia may be as small as 5 µm in length (Nitzschia inconspicua) whereas some species may be over two orders of magnitude larger (Nitzschia scalaris). Some species of Surirella and Cymatopleura (Surirellaceae) are among the most massive of freshwater diatoms. For example, Cymatopleura solea may constitute more than 50 times the biovolume of some co-occurring species (Lowe and Pan, 1996). It may be important to bear these size differences in mind when analyzing data on algal community structure, because a taxon comprising just a few percent of the community cell numbers may comprise a majority of the community biomass. All diatoms within these three orders possess a raphe and most species are highly motile. Genera within Bacillariales (Bacillaria, Cylindrotheca, Cymbellonitzschia, Denticula, Hantzschia, Nitzschia, and Tryblionella) have a raphe on a keel that is elevated above the surface of the valve face (Fig. 1). Heavy siliceous ribs (fibulae) are regularly spaced along the keeled raphe where it opens into the cell interior (Round et al., 1990). These fibulae vary in length among species and have been referred to as keel punctae or carinal punctae in older literature (Fig. 2) (Patrick and Reimer, 1966; Anonymous, 1975). The external raphe opening may be continuous from one pole of the diatom to the opposite pole or the raphe may be interrupted in the center of the valve. Genera within Surirellaceae (Surirellales), which include Campylodiscus, Cymatopleura, Stenopterobia, and Surirella, are generally heavily silicified and all have raphe systems that occupy both margins of each valve (Fig. 3). Thus, the raphe runs around the entire perimeter of each valve rendering them highly motile with substratum contact in almost any position. Fibulae associated with the raphes of species in the Surirellaceae are usually much broader than those in members of the Bacillariales. Entomoneis, in the family Entomoneidaceae (Surirellales) has a keeled raphe that is in two arches that are elevated a great distance from the valve surface, with each arch beginning at either end of the valve and terminating near the center of the valve. In addition, the entire frustule is often twisted about the apical axis, resulting in a relatively complex morphology and assuring its contact with substrata in the epipelic community (see Sec. V, Fig. 22). Much splitting and revision of taxonomic groups within the Bacillariales and Surirellales has occurred, particularly at the generic level (Round et al., 1990). Some of the proposed changes are a result of re-erecting older names that had been synonymized (e.g., Tryblionella), whereas other genera have been split from the

19. Keeled and Canalled Raphid Diatoms

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FIGURE 1 Scanning electron micrograph of a cross section of a frustule of Nitzschia sp. Note the keel (K) projecting from the margin of the frustule.

heavily silicified and robust and although they may be highly motile they often adnate on aquatic vegetation (epiphytic) (Marks and Lowe, 1993; Lowe, 1996).

III. ECOLOGY AND DISTRIBUTION

FIGURE 2 Keel puncta of Nitzschia amphibia, 2000×.

large genus Nitzschia based on details revealed largely by electron microscopy or by consideration of cytoplasmic characters (Psammodictyon, a marine genus). Genera within Rhopalodiales (Epithemia and Rhopalodia) have the raphe located in a canal that opens into the interior of the cell via round or oval pores (portulae). Epithemia and Rhopalodia also have internal fibulae, which usually cross the entire valve and appear as heavy lines when viewed under the light microscope (Fig. 4). Rhopalodia may have its canalled raphe slightly elevated into a keel (Round et al., 1990). Cells of diatoms belonging to the Rhopalodiales are usually

Species in the orders Bacillariales, Surirellales and Rhopalodiales are found worldwide in a variety of freshwater systems, including lakes, rivers, and wetlands (Stevenson et al., 1996). The Bacillariales and Surirellales also have estuarine and marine littoral species. The microhabitats of genera in the Bacillariales and Surirellales are primarily the epipelon and endopelon where the presence of the raphe in an elevated keel allows cells to move more efficiently over and through unconsolidated sediment. Substantial populations of Nitzschia can be found several hundred micrometers beneath the sediment (Fig. 5). In undisturbed sediments, populations of Nitzschia can accumulate to great numbers (Fig. 6), resulting in a golden brown color on the sediment surface. Many species of the Surirellales can also be found at considerable depth associated with lake sediments, especially species of Campylodiscus, Surirella, Entomoneis, Tryblionella, and Cymatopleura. Some of the benthic species of Surirella and Nitzschia are large enough to support

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FIGURE 3 Scanning electron micrograph of Cymatopleura solea with raphe in a wing around the entire valve margin.

FIGURE 4 Light micrograph of Epithemia turgida illustrating the fibulae (F) characteristic of genera in the Rhopalodiales. Scale the bar = 10 µm.

other epiphytic diatoms such as Synedra parasitica and Amphora perpusilla (Belanger et al., 1985). Several species of Bacillariales, Surirellales, and Rhopalodiales also occupy microhabitats other than those in the periphyton. Some species in the genus Nitzschia, particularly N. acicularis and N. holsatica, are lightly silicified and may be planktonic (Hustedt, 1942). Other species of keeled diatoms may be encountered in the seston of lakes and rivers (resuspended) but are rarely truly planktonic and are more properly tychoplanktonic, such as Cymatopleura solea and Entomoneis ornata. One genus, Hantzschia, contains a species (H. amphioxys), that is universally present in soils around the world (Round et al., 1990) and has been reported as part of the soil flora on every continent. Stenopterobia, a genus in the Surirellales, is almost entirely restricted to waters of low pH. Although a relatively small genus, its presence in diatom assemblages is an indication of acidic environments (Round et al., 1990). Stenopterobia sigmatella, a narrow sigmoid species is common in bogs and fens in North America (Stokes and Yung, 1986).

Many species of the genus Nitzschia are recognized as indicators of organic enrichment or pollution of the water in which they are found (Lowe, 1974). Cholnoky (1968) has attributed this pollution-tolerant behavior to the fact that many Nitzschia species are nitrogen heterotrophs and capitalize on organic nitrogen molecules in the water. Thus, they are found in largest numbers where organic nitrogen is abundant, areas of organically polluted water, and in this sense might be thought of as “pollution dependent” rather than “pollution tolerant.” Nitzschia palea is one of the most common and pollution-dependent species in this genus (Palmer, 1969). Tuchman (1996) reported that N. palea was able to utilize 21 different organic substrates. Members of the Rhopalodiales, Epithemia and Rhopalodia, contain endosymbiotic cyanobacteria first described by Geitler (1977) as “Spharoidkörper.” These were shown to be coccoid cyanobacteria (Drum and Pankratz, 1965) that are capable of nitrogen fixation within the diatom host (Floener and Bothe, 1980). Fairchild and Lowe (1984) and DeYoe et al., (1992) observed that these endosymbiotic cyanobacteria


FIGURE 5 Scanning electron micrograph of freeze-fractured stream sediments with a colony of Nitzschia filiformis in a mucilaginous tube several 100 µm below the sediment surface. Scale bar = 10 µm.

FIGURE 6 Scanning electron micrograph of freeze-fractured stream sediments with a colony of Nitzschia spp. in a layer on the sediment surface. Scale bar = 100 µm.

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confer a competitive advantage on species of Epithemia and Rhopalodia in microhabitats with inadequate nitrogen supplies and it has been suggested that these highly modified endosymbionts may be on the evolutionary pathway toward a nitrogen-fixing organelle in this family of diatoms (DeYoe et al., 1992). Species in the genera Epithemia and Rhopalodia are uncommon among the diatoms in their ability to fix nitrogen with endosymbiotic blue green algae (cyanobacteria). Epithemia and Rhopalodia are most common in alkaline water, while occupying microhabitats that are relatively poor in quantities of fixed nitrogen (NO3, NH4) (Fairchild et al., 1985). Such microhabitats are often in nitrogen-poor lakes and streams (Fairchild and Lowe, 1984; Bahls and Weber, 1988; Peterson and Grimm, 1992), usually on submerged plants (macrophyton) that provide colonizable surface area for diatoms and other microalgae (epiphyton) (Power, 1990) (Fig. 7). These host plants may be increasing PO4 availability to epiphytes (Burkholder et al., 1990), thus decreasing the local N:P ratio and conferring a competitive advantage on nitrogen-fixing epiphytes, such as heterocyst-bearing blue green algae and diatoms belonging to the Rhopalodiales (DeYoe et al., 1992). DeYoe et al. (1992) have shown that the number of endosymbionts per cell is a function of nitrogen availability in the microenvironment of the diatom cell.

IV. COLLECTION AND PREPARATION FOR IDENTIFICATION The reader is referred to Chapter 15 of this book for collection and preparation methodology useful for all diatoms. The keeled diatoms, however, warrant some special attention due to their predominance in the epipelon. If care is taken in the collection of epipelic algae, the investigator can be rewarded with collections rich in diatoms with little sediment or detritus. The tools of choice for collecting are an epipelic periphyton sampler (turkey baster) or micropipette with a rubber bulb for sucking up specimens. The epipelic periphyton sampler is a relatively coarse tool given the size of diatoms but if the sampler is deftly operated at the sediment surface and not thrust deeply into the sediment rich collections can be obtained. The micropipette is a better tool for fine-scale sampling where one is interested in different microhabitats and perhaps different clones of epipelic diatoms on the sediment. Careful observation in the field often reveals subtle patterns of algal distribution through slight changes in color, texture, and oxygen bubble distribution. Another habitat that should not be ignored is the endopelon. This community is best collected with a coring device and should be studied with scanning electron microscopy to observe patterns of distribution (Fig. 5) (Greenwood et al., 1999).

FIGURE 7 Scanning electron micrograph of Epithemia sp. (E) epiphytic on the filamentous green alga Dichotomosiphon, 1000×.


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V. KEYS AND DESCRIPTIONS OF GENERA See Figures 8–37.

A. Key to Orders 1a.

Two keels with raphes on each valve, located at the valve margin (Fig. 3)……………………………………………………...Surirellales

1b.

One raphe per valve, located either in a keel or in a canal…………………………………………………………...…………….….........2

2a.

Cells with raphe in a keel, most taxa symmetric to the apical axis, endosymbiotic cyanobacteria lacking…………………Bacillariales

2b.

Cells with raphe in a canal, always asymmetric to the apical axis and bearing endosymbiotic cyanobacteria…………..Rhopalodiales

B. Key to North American Genera of Bacillariales 1a.

Cells gregarious in large raftlike colonies; colonies dynamic with cells sliding back and forth—first stretching out in an elongated linear colony, then sliding back into a tabular colony of cells touching along the entire cell length (Figs. 9, 10)……….........Bacillaria

1b.

Cells usually singular; if colonial, colony not dynamic as above……………………………………………………………….…………...2

2a.

Cells weakly or strongly lunate, asymmetric to the apical axis………………………………………………………………..…………….3

2b.

Cells linear, sygmoid, or some other shape…………………………………………………………………………………………………….4

3a.

Cells elongate and slightly lunate; dorsal margin slightly convex, ventral margin slightly concave; raphe on concave margin on both valves (Fig. 8)…………………………………………………………………………………………….…………………………...Hantzschia

3b.

Cell strongly lunate with a strongly convex dorsal margin and a flat ventral margin; raphe on same margin of each valve (Fig. 12) ……………………………………………………………………………........................................................................Cymbellonitzschia

4a.

The two valves of the frustule twisted around each other, resulting in the raphe systems wrapping around the cell in a spiral pattern; in epipelic habitats, moving in a screwlike fashion through sediments (Fig. 11)…………………………………………....Cylindrotheca

4b.

Frustule not twisted……………………………………………………………………………………………...……………………………...5

5a.

Cells with fibulae extending entirely across valve face appearing like thick internal costa; raphe submarginal (Fig. 13) …………………………………………………………..............................................................................................................Denticula

5b.

Fibulae projecting, but not entirely, across valve face…………………………………………………………………………..……………6

6a.

Cells normally narrow, linear and straight or sygmoid, one plastid in each end of the cell; striae not interrupted by sterna (Figs. 15–21) …………………………………………………………………………………………………………..................…...Nitzschia

6b.

Cells usually robust and relatively broad; may be elliptical, linear, or panduriform; valve surface with an undulation along the apical axis; one plastid in each end of the cell; usually epipelic (Figs. 14, 34)………………………………………………………...Tryblionella

C. Key to North American Genera of Rhopalodiales 1a.

Cells strongly lunate but not canoe-shaped; raphe at distal ends ventral becoming more dorsal toward the proximal ends often forming a peak in the center of the valve (Fig. 28)………………………………………………………………………………....Epithemia

1b.

Valves of frustule lunate but frustule shaped like a canoe; thus, cells normally seen in girdle view; raphe normally along the dorsal margin of the valve (Figs. 27, 37)………………………………………………………………………………………………..…Rhopalodia

D. Key to North American Genera of Surirellales 1a.

Frustule with one raphe on each valve; cells with a high narrow keel that occupies an arch on each end of each valve; frustules often twisted about the apical axis, resulting in a cell that appears bilobate (Fig. 22)……………………………….…………….. Entomoneis

1b.

Frustule with two raphes on each valve; raphes marginal and cells usually relatively large and robust; growing most often on sediments…………………………………………………………………………………………………………........………………………...2

2a.

Valves saddle-shaped (Figs. 29, 30)………………………………………………………………………………….…………Campylodiscus

2b.

Valves may be twisted, but not saddle-shaped………………………………………………………………………………………………..3

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3a.

Valves with undulations or waves along valve face; frustule usually robust and heavily silicified (Figs. 3, 31, 32)……...Cymatopleura

3b.

Valves lacking undulations along valve face……………………………………………………...............................................................4

4a.

Valves S-shaped or straight and narrow; specimens usually restricted to acidic habitats (Fig. 33)………………………...Stenopterobia

4b.

Valves robust, broad, and heavily silicified; may be twisted, isopolar, or heteropolar, but not S-shaped (Figs. 23, 26, 34) ...............................................................................................………………………………………..........................................Surirella

E. Descriptions of Genera: Bacillariales Bacillaria Gmelin (Figs. 9, 10) Cells of Bacillaria are elongate and occur in unique colonies that dynamically change their morphology. The cells are held together along their margins by interlocking ridges and grooves (Fig. 9). Cells in the colony slide back and forth from a side-by-side position to a pole-to-pole position in an oscillating fashion. Individual frustules of Bacillaria possess a keeled raphe positioned near the center of each cell (Fig. 10). There are usually two plastids in each cell positioned toward the poles. Bacillaria paradoxa is a common North American species and is normally found in slightly brackish water and water high in dissolved solids. Colonies are normally epipelic but often become tychoplanktonic. Cylindrotheca Rabenhorst (Fig. 11) Frustules of Cylindrotheca are relatively long and narrow with attenuated apices. Each frustule is twisted about the apical axis so that the valve and girdle areas are spiraled around the length of the cell. The raphe is fibulate and marginal on each of the twisted valves. Cells have two to several platelike or disk-shaped plastids. Cylindrotheca is a relatively small genus (four species) found mostly in brackish water or marine habitats. The most widely distributed freshwater species is C. gracilis; it occurs primarily in the epipelon of streams of high conductance (Christensen and Reimer, 1968). Cylindrotheca can usually be identified under the microscope in living material by its screwlike movement across the substratum. The frustules are lightly silicified and are susceptible to destruction during acid or peroxide cleaning of samples. Cymbellonitzschia Hustedt (Fig. 12) Cells of Cymbellonitzschia are usually solitary or may form short chains in benthic areas of still water. Each cell has two small platelike plastids positioned on opposite poles of the cell. The valves are asymmetrical to the apical axis and thus have dorsal and ventral margins (Fig. 12). The fibulate raphe may be on the ventral margin of each valve (similar to Hantzschia), such as in the most common species (Cymbello-

nitzschia diluviana) but may be on the dorsal margin in other species. At first glance this taxon might be confused with Cymbella or Amphora due to its characteristic shape, but can quickly be identified as Cymbellonitzschia by the presence of the fibulate raphe. Denticula Ehrenberg (Fig. 13) Cells of Denticula are relatively small and usually solitary, but they may occur in short chains. Each cell contains two plastids on either side of the transapical plane near the center of the cell. Valves are symmetrical to both the apical and transapical axes and are linear to lanceolate with a fibulate raphe on each valve that is slightly eccentric on the valve surface (Fig. 13). Raphes on opposite valves display nitzschioid symmetry. Denticula is a relatively small genus that is uncommon, but occurs most abundantly in benthic hardwater habitats. This genus had formerly been included in the Epithemiaceae (Patrick and Reimer, 1975). More recently, Denticula has been suggested to be more closely related to Nitzschia and its allies by virtue of the “nitzschioid symmetry” of its raphe systems (Round et al., 1990). In addition, Denticula species in North America have not been recorded with cyanobacterial endosymbionts that appear universally in the Rhopalodiaceae, although Geitler (1977) indicated European species of the genus possess endosymbionts. Hantzschia Grunow (Fig. 8) Cells of Hantzschia are curved and thus are asymmetric to the apical axis in valve view (Fig. 8). The raphe occupies a keel on the concave (ventral) margin of each valve and its presence is best detected in the light microscope by the appearance of the associated fibulae. The convex (dorsal) margin of each valve lacks a raphe. Plastids may be rounded or lobed, are usually two in number, and are typically ventral. This is a relatively small genus but one species, Hantzschia amphioxys, is present in soil flora worldwide. This species is also a common component of atmospheric algal particulates. Nitzschia Hassall (Figs. 15–20) Cells of Nitzschia are usually long, straight, and narrow but may be ovoid or even slightly sygmoid.


FIGURE 8 Hantzschia amphioxys, valve view. Scale bar = 10 µm. FIGURE 9 Colony of Bacillaria paradoxa. Scale bar = 10 µm. FIGURE 10 Bacillaria paradoxa, valve view. Scale bar = 10 µm. FIGURE 11 Cylindrotheca gracilis, valve view. Scale bar = 10 µm. FIGURE 12 Cymbellonitzschia diluviana, valve view. Scale bar = 10 µm. FIGURE 13 Denticula tenuis, valve view. Scale bar = 10 µm. FIGURE 14 Tryblionella sp., valve view. Scale bar = 10 µm. FIGURE 15 Nitzschia dissipata, valve view. Scale bar = 10 µm. FIGURE 16 Nitzschia acicularis, valve view. Scale bar = 10 µm. FIGURE 17 Nitzschia amphibia, valve view. Scale bar = 10 µm. FIGURE 18 Nitzschia palea, valve view. Scale bar = 10 µm. FIGURE 19 Nitzschia denticula, valve view. Scale bar = 10 µm. FIGURE 20 Nitzschia sinuata v. tabellaria, valve view. Scale bar = 10 µm. FIGURE 21 Nitzschia angustata, valve view. Scale bar = 10 µm.

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FIGURE 22 Entomoneis ornata, valve view. Scale bar = 10 µm. FIGURE 23 Surirella tenera, valve view. Scale bar = 10 µm. FIGURE 24 Surirella linearis v. constricta, valve view. Scale bar = 10 µm. FIGURE 25 Surirella angustata, valve view. Scale bar = 10 µm. FIGURE 26 Surirella ovata, valve view. Scale bar = 10 µm. FIGURE 27 Rhopalodia gibba, valve view. Scale bar = 10 µm. FIGURE 28 Epithemia sp., valve view. Scale bar = 10 µm.


FIGURE 29 Campylodiscus noricus, valve view of saddle-shaped valve from the side. Scale bar = 10 µm. FIGURE 30 Campylodiscus noricus, valve view of saddle-shaped valve from the top. Scale bar = 10 µm. FIGURE 31 Cymatopleura elliptica, valve view. Scale bar = 10 µm. FIGURE 32 Cymatopleura solea, valve view. Scale bar = 10 µm. FIGURE 33 Stenopterobia sigmatella, valve view. Scale bar = 10 µm.

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FIGURE 34 Scanning electron micrograph of Tryblionella (foreground)

FIGURE 35 Scanning electron micrograph of Epithemia frustule

and Surirella (background). Note the longitudinal undulation of Tryblionella and the wing along the margin of Surirella with raphe in a wing around the entire valve margin.

in girdle view illustrating the slightly wedge-shaped nature of the cell.

FIGURE 36 Scanning electron micrograph of an Epithemia valve.

FIGURE 37 Scanning electron micrograph of Rhopalodia frustule illustrating the wedge-shaped nature of the cell, 2500×.


They usually occur singly but may form stellate colonies or live in mucilage tubes (Fig. 6). Cells usually contain two plastids that are toward each pole of the cell. The raphe system in Nitzschia is fibulate and is normally on or near the margin of the valve surface. The raphe is on opposite margins of the two valves of a frustule (nitzschioid symmetry), in contrast to the raphe position of Hantzschia. Nitzschia is a relatively large genus with hundreds of freshwater and marine species. Most species are epipelic in microhabitat but Nitzschia also contains planktonic, epilithic, and epiphytic species. Nitzschia contains many pollution-tolerant species (Lowe, 1974) that have been used as indicators of deteriorated water quality (Whitton et al., 1991; Whitton and Rott, 1996). Tryblionella W. Smith (Figs. 14, 21, 34) Frustules of Tryblionella are symmetric to both the apical and transapical axes in valve view (Fig. 14). Each valve possesses a marginal raphe elevated in a keel. Raphes of the two valves are on opposite margins of the frustule. Each valve is undulated about the apical axis (Round et al., 1990) (Fig. 34). Each cell contains two plastids located on opposite ends of the cell. Tryblionella is common but seldom abundant in the epipelon of a variety of hard-water habitats and can be distinguished from Nitzschia by its apically undulating valves.

F. Descriptions of Genera: Rhopalodiales Epithemia Brébisson (Figs. 28, 35, 36) Cells of Epithemia are lunate in valve view with a dorsal and ventral margin and usually occur singly (Fig. 28). The two valves are often oriented at a slight angle to each other resulting in wedge-shaped frustule (Fig. 35). A single large lobed plastid is located near the ventral margin of the cell. The raphe on each valve is ventral toward the ends of the valve and arcs toward the middle of the valve where it is more dorsal (Fig. 36). The raphe opens internally into a canal, which is connected to the cell interior by circular or subcircular holes or portulae. In addition, frustules are normally heavily silicified and the valves have thickened internal costae that appear as heavy lines in the light microscope. All species examined thus far contain endosymbiotic cyanobacteria-like cells that function in nitrogen fixation (Floener and Bothe, 1980). Epithemia is to be found in benthic hard-water habitats reaching maximum abundance in microhabitats where phosphorus is relatively more available (low N/P microhabitats) such as the surface of submerged aquatic plants.

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Rhopalodia O. Müller (Fig. 27) Cells of Rhopalodia are solitary and are dorsoventral (lunate) in valve view like Epithemia (Fig. 28). However, the cell is normally seen in girdle view because the entire cell is strongly wedge-shaped similar to a canoe (Fig. 37). This is a result of the cingulum being much wider on one side of the cell than the other. Cells have a single ventral platelike plastid similar to Epithemia. The raphe is normally along the dorsal margin of the valve and may be difficult to see in valve view. It opens into a canal similar to the structure displayed by Epithemia. Transapical costae are thickened internally appearing as fibulae. Endosymbiotic cyanophytes are present in all species of this genus studied and function in nitrogen fixation (Floener and Bothe, 1980). This genus is found in hard-water nitrogen-poor benthic habitats like those characteristic of Epithemia.

G. Descriptions of Genera: Surirellales Entomoneidaceae Entomoneis Ehrenberg (Fig. 22) Cells of Entomoneis are solitary and highly motile. The frustule is twisted about the apical axis (Fig. 22) and thus may lie in a variety of positions on the microscope slide. Frustules often appear in girdle view as hourglass-shaped or panduriform. They would appear slightly sigmoid in valve view but are rarely seen in this view. Each valve has a raphe in a keel that arches above the valve surface. Species of Entomoneis have one or two large platelike plastids. This genus is rarely abundant, but broadly distributed in epipelic habitats in water of high conductance. Entomoneis may occasionally become entrained in the water column as tychoplankton. Approximately six species have been reported from North America (Patrick and Reimer, 1975).

Surirellaceae Campylodiscus Ehrenberg ex Kützing (Figs. 29, 30) Cells of Campylodiscus are relatively large, solitary, and saddle-shaped (Fig. 29) and thus appear subcircular or crescent-shaped under the microscope, depending on their orientation. Each valve posseses two raphes that lie on the valve margins and are elevated on a wing supported by radially arranged ribs. Multiseriate striae alternate with the ribs. The two valves of the frustule have their apical axes at right angles to each other (Round et al., 1990). The single large plastid is divided into two plates appressed against each of the valve surfaces. It is useful to examine specimens of this large saddle-shaped diatom without a coverglass on the slide because the three-dimensional cells are easily broken under the weight of a cover glass.

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There is just one freshwater species in North America, Campylodiscus noricus, normally restricted to epipelic habitats in lentic ecosystems. Cymatopleura W. Smith (Figs. 3, 31, 32) This benthic diatom genus is characterized by having relatively large solitary cells with a raphe elevated on a wing along each margin of each valve. The valve surfaces display regular transapical undulations (Fig. 3). Valves are isopolar or heteropolar and may be elliptical, linear, or panduriform in shape (Round et al., 1990) and some species are slightly twisted about the apical axis (Hustedt, 1930). Each cell has a single plastid that is divided into two large plates appressed against each valve. This genus is quite common in epipelic habitats of lakes, rivers, and wetlands. The most common North American species is Cymatopleura solea, which is easily identified by its characteristic size and shape (Fig. 32). Stenopterobia Brébisson ex Van Heurck (Fig. 33) Cells of Stenopterobia are relatively long and narrow with a keel along both margins of each valve that is elevated from the valve surface (Fig. 33). Each cell has a single plastid divided into two plates lying against opposite valves. The cells may be either straight or sygmoid in outline. Few species have been reported from North America. Of these, Stenopterobia sigmatella is probably the most common (Stokes and Yung, 1986). This genus is closely related to Surirella, from which it can be differentiated by its narrow linear to sygmoid shape and its near restriction to acidic habitats. Species of Stenopterobia are normally benthic in acidic habitats. Surirella Turpin (Fig. 23–26, 34) Cells of Surirella are solitary and may be either isopolar or heteropolar and are slightly wedge-shaped in girdle view. Some species are twisted about the apical axis (Lowe, 1974; Krammer and Lange-Bertalot, 1988). Each cell contains one large platelike plastid divided into two halves with each half flattened against opposite valves of the frustule. Each valve has two raphes that are located on both margins of the valve. Many species are relatively large, heavily silicified, and may be ornamented with siliceous spines and protuberances. All species are benthic and are found most often on epipelic habitats but may also occupy the epilithon and epiphyton.

VI. GUIDE TO LITERATURE FOR SPECIES IDENTIFICATION The following is a list of key references for species identification. An attempt has been made to list refer-

ences from North America. However, many of the references listed here are from references outside North America. This is an unfortunate outcome resulting from our incomplete knowledge of the North American diatom flora. Bacillaria—Dodd (1987), Hustedt (1930), Krammer and Lange-Bertalot (1988) Campylodiscus—Hustedt (1930), Krammer and Lange-Bertalot (1988) Cylindrotheca—Christiansen and Reimer (1968) Cymatopleura—Hustedt (1930), Krammer and Lange-Bertalot (1988) Cymbellonitzschia—Hustedt (1930), Krammer and Lange-Bertalot (1988) Denticula—Johansen et al. (1990), Patrick and Reimer (1975) Entomoneis—Patrick and Reimer (1975) Epithemia—Patrick and Reimer (1975) Hantzschia—Mann (1977, 1980a, b 1981), Hustedt (1930), Krammer and Lange-Bertalot (1988) Nitzschia—Archibald (1970), Reimer, (1954), Mann (1986), Lange-Bertalot (1976, 1980), Krammer and Lange-Bertalot (1988), Lobban and Mann (1987) Rhopalodia—Patrick and Reimer (1975) Stenopterobia—Hustedt (1930), Krammer and Lange-Bertalot (1988) Surirella—Hustedt (1930), Krammer and LangeBertalot (1988) Tryblionella—Krammer and Lange-Bertalot (1988)

ACKNOWLEDGMENTS I thank the following individuals for assistance with the figures: Todd Clason, Jennifer Greenwood, David Johnson, Katherine Jones, Gina LaLiberte, and Susan Makosky.

LITERATURE CITED Anonymous. 1975. Proposals for a standardization of diatom terminology and diagnoses. Beihefte zur Nova Hedwigia, 53:323–354. Archibald, R. E. M. 1970. Key to the genus Nitzschia. Cont. News Lett. Limnological Society of South Africa 19:37–55. Bahls, L., Weber, C. I. 1988. Ecology and distribution in Montana of Epithemia sorex Kütz., a common nitrogen fixing diatom. Proceedings of the Montana Academy of Sciences 48:15–20. Belanger, S. E., Lowe, R. L., Rosen, B. H. 1985. Epiphytism of Synedra parasitica on Surirella robusta: Observations of populations and associations in a Virginia pond. Transactions of the American Microscopical Society 104:378–386. Burkholder, J. M. 1996. Interactions of benthic algae with their substrata, in: Stevenson, R. J., Bothwell, M. L., Lowe, R. L., Eds., Algal ecology. Academic Press, San Diego, pp. 253–297.

19. Keeled and Canalled Raphid Diatoms Burkholder, J. M., Wetzel, R. G., Klomparens, K. L. 1990. Direct comparison of phosphate uptake by adnate and loosely attached microalgae within an intact biofilm matrix. Applied and Environmental Microbiology 56:2882–2890. Cholnoky, B. J. 1968. Die Ökologie der Diatomeen in Binnengewasser. Cramer, Lehre, 699 p. Christensen, C. L., Reimer, C. W. 1968. Notes on the diatom Cylindrotheca gracilis (Breb. ex Kütz) Grun: Its ecology and distribution. Journal of the Iowa Academy of Science 75:36–41. DeYoe, H. R., Lowe, R. L., Marks, J. C. 1992. The effect of nitrogen and phosphorus on the endosymbiont load of Rhopalodia gibba and Epithemia turgida (Bacillariophyceae). Journal of Phycology 28:773–777. Dodd, J. J. 1987. Diatoms (The illustrated flora of Illinois). Southern Illinois Univ. Press, Carbondale, 477 p. Drum, R. W., Pankratz, S. 1965. Fine structure of an unusual cytoplasmic inclusion in the diatom genus Rhopalodia. Protoplasma 60:141–9. Fairchild, G. W., Lowe, R. L. 1984. Artificial substrates which release nutrients: Effects upon periphyton and invertebrate succession. Hydrobiologia 184:29–37. Fairchild, G. W., Lowe, R.L., Richardson, W. B. 1985. Nutrient-diffusing substrates as an in situ bioassay using periphyton: Algal growth responses to combinations of N and P. Ecology 66: 465–472. Floener, L., Bothe, H. 1980. Nitrogen fixation in Rhopalodia gibba, a diatom containing blue-greenish inclusions symbiotically, in: Schwemmler, W., Shenk, H. E. A., Eds., Endocytobiology: Endosymbiosis and cell biology, a synthesis of recent research, Vol. 1. de Gruyter, Berlin, pp. 541–552. Geitler, L. 1977. Zur Entwicklungsgeschichte der Epithemiaceen Epithemia, Rhopalodia and Denticula (Diatomophyceae) und ihre vermutlich symbiotischen Spharoidkörper. Plant Systematics and Evolution 128:259–275. Goldsborough, L. G., Robinson, G. G. C. 1996. Pattern in wetlands, in: Stevenson, R. J., Bothwell, M. L., Lowe, R. L., Eds., Algal ecology. Academic Press, San Diego, pp. 77–117. Greenwood, J., Clason, T., Lowe, R. L., Belanger, S. E. 1999. Examination of endopelic and epilithic algal community structure employing scanning electron microscopy. Freshwater Biology 41:821–828. Hustedt, F. 1930. Dr. L. Rabenhorsts Kryptogamen-Flora von Deutschland, Österreichs und der Schweiz, Vol. VII. Die Kieselalgen Deutschlands, Österreichs und der Schweiz unter Berücksichtigung der übrigen Länder Europas sowie der angrenzenden Meeresgebiete. Autorisierter Neudruck (1962), Johnson Reprint Corporation, New York. Hustedt, F. 1942. Das Phytoplankton des Süsswassers. Teil 2, Hälfte 2, Die Binnengewässer. Schweizerbart’sche Verlagsbuchhandlung, Stuttgart, 549 p. Johansen, J., Cognata, S. L., Kociolek, J. P. 1990. Examination of type material of Denticula rainierensis Sovereign. Memoirs of the California Academy of Sciences 17:211–219. Krammer, K., Lange-Bertalot, H. 1988. Süsswasserflora von Mitteleuropa. Bacillariophyceae, Part 2. Teil: Bacillariaceae, Epithemiaceae, Surirellaceae. Fischer, Stuttgart, 596 p. Lange-Bertalot, H. 1976. Eine Revision zur Taxonomie der Nitzschiae lanceolatae Grunow. Beihefte zur Nova Hedwigia 28:253–307. Lange-Bertalot, H. 1980. New species, combinations and synonyms in the genus Nitzschia. Bacillaria 3:41–77. Lange-Bertalot, H., Krammer, K. 1993. Observations on Simonsenia and some small species of Denticula and Nitzschia. Beihefte zur Nova Hedwigia 106:93–99. Lobban, C. S., Mann, D. G. 1987. The systematics of the tube-

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dwelling diatom Nitzschia martiana and Nitzschia section Spathulatae. Canadian Journal of Botany 65:2396–2402. Lowe, R. L. 1974. Environmental requirements and pollution tolerance of freshwater diatoms. EPA-670/4-74-007, 340 p. Lowe, R. L. 1996. Periphyton patterns in lakes, in: Stevenson, R. J., Bothwell, M. L., Lowe, R. L., Eds., Benthic algal ecology in freshwater ecosystems. Academic Press, San Diego, pp. 57–76. Lowe, R. L., Pan, Y. 1996. Use of benthic algae in water quality monitoring, in: Stevenson, R. J., Bothwell, M. L., Lowe, R. L., Eds., Benthic algal ecology in freshwater ecosystems. Academic Press, San Diego, pp. 705–739. Mann, D. G. 1977. The diatom genus Hantzschia Grünow, an appraisal. Beihefte zur Nova Hedwigia 54:323–354. Mann, D. G. 1980a. Hantzschia fenestrata Hust., Hantzschia or Nitzschia. British Phycological Journal 15:249–260. Mann, D. G. 1980b. Studies on the diatom genus Hantzschia, II. H. distinctepunctata. Beihefte zur Nova Hedwigia 33:341–352. Mann, D. G. 1981. Studies on the diatom genus Hantzschia, 3. Interspecific variation in H. virgata. Annals of Botany 47:377–395. Mann, D. G. 1986. Nitzschia subgenus Nitzschia (notes for a monograph of the Bacillariaceae. 2), in: Ricard, M., Ed., Proceedings of the 8th International Diatom Symposium. Koeltz, Koenigstein, pp. 215–226. Marks, J. C., Lowe, R. L. 1993. Interactive effects of nutrient availability and light levels on the periphyton composition of a large oligotrophic lake. Canadian Journal of Fisheries and Aquatic Sciences 50:1270–1278. Palmer, C. M. 1969. A composite rating of algae tolerating organic pollution. Journal of Phycology 5:78–82. Patrick, R., Reimer, C. W. 1966. The diatoms of the United States exclusive of Alaska and Hawaii. Monograph 13. Academy of Natural Sciences of Philadelphia, 672 p. Patrick, R., Reimer, C. W. 1975. The diatoms of the United States exclusive of Alaska and Hawaii. Monograph 13. Academy of Natural Sciences of Philadelphia, 672 p. Peterson, C. G., Grimm, N. B. 1992. Temporal variation in enrichment effects during periphyton succession in a nitrogen-limited stream ecosystem. Journal of the North American Benthological Society 11:20–36. Power, M. E. 1990. Effects of fish in river food webs. Science 250:811–814. Reimer, C. W. 1954. Re-evaluation of the diatom species Nitzschia frustulum (Kütz.) Grun. Butler University Botanical Studies 11:178–191. Round, F. E. 1981. The ecology of algae. Cambridge Univ. Press, New York. Round, F. E., Crawford, R. M., Mann, D. G. 1990. The diatoms. Biology and morphology of the genera. Cambridge Univ. Press, Cambridge, UK, 747 p. Stevenson, R. J., Bothwell, M. L. Lowe, R. L. Eds., Benthic algal ecology in freshwater ecosystems. Academic Press, San Diego, 753 p. Stoermer, E. F., Julius, M. L. 2002. Centric diatoms, in: Wehr, J. D., Sheath, R. G., Eds., Freshwater algae of North America. Academic Press, San Diego, pp. 559–594. Stokes, P. M., Yung, Y. K. 1986. Phytoplankton in selected LaCloche (Ontario) lakes, pH 4.2–7.0, with special reference to algae as indicators of chemical characteristics, in: Smol, J. P., Battarbee, R. W., Davis, R. B., Merilainen, J., Eds., Diatoms and lake acidity. Vol. 4. Junk, pp. Dordrecht, 4:57–72. Tuchman, N. C. 1996. The role of heterotrophy in algae, in: Stevenson, R. J., Bothwell, M. L., Lowe, R. L. Eds., Benthic algal ecology in freshwater ecosystems. Academic Press, San Diego, pp. 299–319.

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Van Landingham, J.L. 1978. catalogue of the fossil and Recent genera and species of diatoms and their synonyms. Part VI. Neidium through Rhoicosphenia. Cramer, Weinheim, Germany, pp. 2964–3605. Whitton, B. A., Rott, E. 1996. Use of algae for monitoring rivers, II.

Institut für Botanik, AG Hydrobotanik, Universität Innsbruck, Austria, 196 p. Whitton, B. A., Rott, E., Friedrich, G. 1991. Use of algae for monitoring rivers. Institut für Botanik, AG Hydrobotanik, Universität Innsbruck, Austria, 193 p.

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DINOFLAGELLATES Susan Carty Department of Biology Heidelberg College Tiffin, Ohio 44883

I. Introduction II. Morphology and Diversity A. Morphology B. Life Cycle C. Classification III. Ecology and Distribution A. Dinoflagellate Blooms B. Tropic States C. Specificity of Habitat D. Geographical Distribution IV. Collection and Preparation for Identification

I. INTRODUCTION Dinoflagellates (Division or Phylum Pyrrhophyta) are a group of primarily unicellular organisms united by a suite of unique characteristics, including flagellar insertion, pigmentation, organelles, and features of the nucleus, that distinguishes them from other groups. The name dinoflagellate comes from dinos (Greek), “whirling,” which describes their distinctive swimming pattern, and flagellum (Latin), “a whip.” Pyrrhophyta comes from the Greek pyrrh “flame colored,” “reddish.” Freshwater dinoflagellates, including one of the most recognizible unicells of all the plankton, Ceratium hirundinella, can be dominant members of the summer phytoplankton and may be responsible for taste and odor problems in drinking water. Currently there are about 250–300 species of freshwater dinoflagellates known worldwide, and about 150 have been reported from North America. Dinoflagellates, however, are best known to the Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

A. Collection B. Fixation C. Preparation for Identification V. Key and Descriptions of Genera A. Key B. Descriptions of Genera VI. Guide to Literature for Species Identification A. Compendia B. Local Floras Literature Cited

public as the source of marine red tides leading to various types of human illness caused by their toxins: paralytic shellfish poisoning (PSP), neurotoxic shellfish poisoning, diarrhetic shellfish poisoning, and ciguatera (Hallegraeff et al., 1995; Burkholder, 1998). These red tide species are marine or estuarine species, as is the ambush predator, the “cell from hell,” Pfiesteria piscicida (Burkholder et al., 1992; Steidinger et al., 1996a). While Pfiesteria is known for its fish-killing and neurological effects (Levin et al., 1997), its remarkable multistage life cycle has caught the attention of scientists from many disciplines who now ponder the placement of this extraordinary group of protists in the evolution of eukaryotes. Freshwater dinoflagellate taxa exhibit much of the variability found within their marine counterparts, including autotrophy, heterotrophy, and fish parasitism—apparently lacking only toxin production. It is the aim of this chapter to provide an introduction to the freshwater dinoflagellates, and most of the text will refer to the commonly encountered taxa. 685

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TABLE I Unique Features of Dinoflagellates Nuclear Permanently condensed chromosomes Dinomitosis (nuclear membrane intact, spindle external, chromosomes attached to nuclear envelope, nucleoli remain) Lack histones and nucleosomes Unusual base: 5-hydroxymethyl uracil Cellular Arrangement of flagella in cingulum and sulcus Pusule Trichocysts Chemical Dinosterol Peridinin Toxins

The characteristics that unite the dinoflagellates and make them unique (Table I) also make determining their phylogenetic affinities difficult (Table II). Dinoflagellates have a nucleus with permanently condensed chromosomes and unique bases, which lacks histones and nucleosomes (Soyer-Gobillard, 1996), although stages in some complex life histories may have a more typical eukaryotic nucleus (Steidinger et al., 1996a; Buckland-Nicks et al., 1997). The terms mesokaryotic (Dodge, 1965, 1966) and dinokaryotic have been used for this unique nucleus. Many autotrophic dinoflagellates have the unique carotenoid pigment peridinin associated with a unique peridinin-chlorophyll protein (Larkum, 1996) and an atypical photosynthesis with an unusual form of ribulose 1,5 bisphosphate carboxylase (RuBisCo), an enzyme critical to the initiation of the Calvin cycle (reviewed in Palmer, 1995, 1996). The distinctive whirling swimming pattern while being propelled forward is due to the presence of a tinsel flagellum in a transverse groove (cingulum) and a

whiplash flagellum in a longitudinal groove (sulcus), again, a unique organization (Fig. 1A). Internally distinctive organelles such as trichocysts (ejectile) and the pusule (nutrient uptake) are found. Plantlike features of dinoflagellates include cellulose walls in some taxa and the synthesis of starch. Distantly related photosynthetic protists may include the Euglenophyta (some nuclear similarities) and Cryptophyta. The animal-like features of some dinoflagellates (e.g., heterotrophy, trichocysts, eyespots), together with some strong evidence that dinoflagellates have come by photosynthesis through symbiosis (Tomas and Cox, 1973; Wilcox and Wedemayer, 1984; Fields and Rhodes, 1991; Chesnick et al., 1996), lead some researchers to look to the protozoa for nearest relatives. Many of the characters that make dinoflagellates unique are visible in the light microscope. Swimming cells show the pattern of whirling while being propelled forward and, if photosynthetic, are a (yellow-) golden (-brown) color. When cells are not moving, the cingulum is usually visible as a cinched-in waist, and the nucleus can be seen. The nucleus confirms the identification; it is large, filling 25–35% of the cell, and often centrally located, and the permanently condensed chromosomes appear as a fingerprint pattern of swirls. There is a broad range of cell size: small cells (the aptly named Peridinium inconspicuum) may be 10 µm wide by 12 µm long, and large Ceratium may reach 400 µm in length. This chapter deals with freshwater dinoflagellates, but there is also an extensive literature on marine dinoflagellates. In addition to their role as red-tide organisms, marine dinoflagellates are important member of oceanic phytoplankton, and as symbionts with reefbuilding corals (e.g., zooxanthellae such as Symbio-

TABLE II Comparison of Dinoflagellate Features to Other Taxa Dinoflagellate feature

Most like

Reference

(N base) hydroxymethyluracil 17S rRNA gene No histones Arch-shaped nuclear fibrils Chromosomes attached to nuclear membrane RuBisCo 18S rRNA gene Trichocysts Condensed chromosomes Chlorophyll c Store starch, cellulose walls 2Fe 2S ferredoxin 17S rRNA gene

Bacteriophage Herzog et al. (1984) Archaebacteria Herzog and Maroteaux (1986) Prokaryotes Bacteria Herzog et al. (1984) Bacteria Herzog et al. (1984) Bacteria Palmer (1996) Apicomplexa Wright and Lynn (1997a,b) Paramecium Hausmann (1978) Euglenoids Chromophytes Green algae/plants Chlorophytes Yoshikawa et al. (1997) Plants Herzog and Maroteaux (1986)

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FIGURE 1 Typical thecate motile dinoflagellate cell. (A). Ventral view. (B). Dorsal view.

dinium; Battey, 1992; Blank, 1992) they are vital to maintaining coral reef systems; bleaching occurs when dinoflagellates abandon their coral host. Dinoflagellates form symbiotic relationships with other organisms, including some jellyfish (Trench and Thinh, 1995). Some dinoflagellates are bioluminescent and have been widely studied to understand the circadian rhythm of the flashes (Knaust et al., 1998) and their function (Mensinger and Case, 1992). Another extensive literature exists on fossil dinoflagellate cysts. Dinoflagellates have a long (from the Silurian; Sarjeant, 1978), widespread, and extensive fossil record. Initially cysts were important stratigraphic markers used by oil companies (Lentin and Williams, 1989) and are increasingly used to understand global environmental conditions (MacRae et al., 1996). Work linking cysts to motile cells has clarified the identity of many fossil cysts (Wall and Dale, 1968). Currently, geologists are at the forefront of much of the work on dinoflagellate systematics (Fensome et al., 1993) and evolution (MacRae et al., 1996; Fensome et al., 1996). Books devoted to the biology of dinoflagellates include Spector (1984), Taylor (1987), (though many examples and topics relate to marine taxa), and Evitt (1985) (emphasis on cysts, but excellent diagrams and explanations of thecal morphology).

II. MORPHOLOGY AND DIVERSITY A. Morphology Freshwater dinoflagellates occur as single cells either in the plankton or attached to substrates such as fish, algal filaments, etc. Historically they have been described as thecate (armored with cellulose plates) or naked (lacking plates). In either case, a typical motile cell has the following characteristic features (Fig. 1A, B). A transverse groove or girdle, the cingulum, encircles the cell and divides it into an epitheca (anterior portion) and hypotheca (posterior portion), or an epicone and hypocone in taxa without plates. The cingulum houses a ribbon-like tinsel flagellum that is responsible for the whirling motion during swimming. The cingulum usually divides the cell into two approximately equal halves, but may be found higher, dividing cells into 1/3 epitheca, 2/3 hypotheca (Amphidinium Fig. 2C) or lower, dividing the cell into 2/3 epitheca, 1/3 hypotheca (Katodinium Fig. 2G). The ends of the cingulum may face each other or may be displaced to various degrees (very offset in Gyrodinium (Fig. 2F) and Gonyaulax (Fig. 3B). Displacement is measured in girdle widths and is usually left-handed (descending). A longitudinal groove, the sulcus, defines the ventral face

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FIGURE 2 Line drawings of unarmored dinoflagellates. No line scale, refer to text for cell size. (A) Actiniscus pentasterias v. arcticus. Arrows indicate rudimentary pentasters. (B) Pseudoactiniscus apentasterias. (C) Amphidinium klebsii. (D) Bernardinium bernardinense. (E) Gymnodinium acidotum. (F) Gyrodinium pusillum. (G) Katodinium spiroidinoides. (H) Cystodinedria inermis. (I) Cystodinium bataviense. (J) Hypnodinium sphaericum. (K) Dinococcus as Raciborskia bicornis. (L) Stylodinium globosum. (M) Tetradinium javanicum. (N) Oodinium limneticum. (O) Haidadinium ichthyophilum. (P) Dinastridium sexangulare. (Q) Phytodinium simplex. (R) Hemidinium as cyst form Gloeodinium montanum. (S) Rufusiella insignis. (T) Dinamœbidium coloradense, gymnodinioid cell (a), amoeboid cell (b), cysts on sand grain (c). p = “papilli-forming pseudopodia”, d = was not labeled, possibly a phagocytosed diatom. A and B are from Bursa (1969) with permission. C and D are from Thompson (1950) with permission. F is from Thompson (1947) with permission. I, J, L, M, R are from Thompson (1949) with permission. K is from Prescott (1951) with permission. N is from Jacobs (1946) with permission. O is from Buckland-Nicks et al. (1997) with permission. S is from Smith (1950) with permission. T is from Bursa (1970) and is reproduced from Arctic and Alpine Research with permission of the Regents of the University of Colorado.

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FIGURE 3 Line drawings of armored dinoflagellates. Where there is no line scale, refer to the text for cell size. (A) Exuviella compressa. (B) Gonyaulax spinifera. a, ventral; b, epitheca (s is a sulcal plate); c, hypotheca. (C) Thompsodinium intermedium. a, ventral (note trichocyst pores on plates); b, dorsal; c, epitheca; d, hypotheca. (D) Peridinium willei. a, ventral; b, dorsal; c, epitheca. (E) Peridiniopsis quadridens ventral. (F) Durinskia baltica. a, ventral; b, dorsal; c, epitheca. (G) Ceratium hirundinella, ventral H. Lophodinium polylophum. ventral. (I) Woloszynskia reticulata. a, dorsal; b, ventral; c, cyst. A, I are from Thompson (1950) with permission. C, E are from Carty (1989) with permission.

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of the cell, extends some distance into the hypotheca, may extend into the epitheca, and houses the whiplash flagellum, which propels the cell forward (Leadbeater and Dodge, 1967). Most dinoflagellates have a motile stage with a cingulum-sulcal arrangement at some time in their life cycle. The motile stage is frequently the assimilative stage during which nutrients are absorbed (photosynthetic or heterotrophic) but may be reproductive and/or dispersive. Nonmotile assimilative forms include the free-floating forms (Cystodinium (Fig. 2I), Hypnodinium (Fig. 2J), those with stalks attached to a substrate (Tetradinium (Fig. 2M), Stylodinium (Fig. 2L), and those attached directly to the substrate (Cystodinedria (Fig. 2H), Oodonium (Fig. 2N)). Nonmotile cysts may occur in the plankton or in sediments and may be more distinctive than the motile stage. Dinoflagellates are eukaryotic cells with membrane-bound organelles. The nucleus is usually large, variously shaped (round, C-shaped, curved), variously located (centrally, in epitheca or hypotheca), and with large, permanently condensed chromosomes. Mitochondria are tubular. Some species have a red stigma or eyespot in the sulcal region (Dodge, 1969), which may be part of a chloroplast or may be free (Dodge, 1969). Unique organelles include trichocysts and the pusule. Trichocysts are ejectile organelles whose construction and chemical composition are somewhat similar to those of the spindle trichocysts of Paramecium (Hausmann, 1978). The pusule is a tubular or vesicular organelle formed by invagination of the plasma membrane and is covered by additional membrane (Klut et al., 1987) located in the sulcus near the base of the flagella. Dodge (1972) identified seven types of pusule. Several functions have been hypothesized for it, including fluid intake for nutrients, a buoyancy device, waste expulsion, and osmoregulation (summarized in Spector, 1984). Experimental evidence of the uptake of molecular markers (Klut et al., 1987) would support nutrient uptake (Kofoid, 1909; Kofoid and Swezy, 1921). Most photosynthetic dinoflagellates have discoid or lobed chloroplasts located in a peripheral position. Chloroplasts are typically surrounded by three membranes, lack chloroplast endoplasmic reticulum (CER), and have thylakoid membranes in stacks of three and some type of pyrenoid (Dodge, 1975). Photosynthetic pigments are chlorophylls a and c2, with the carotenoid peridinin contributing the usually golden color, although cells may appear yellowish or almost brown (as in Peridinium gatunense). Other carotenoids include diadinoxanthin, dinoxanthin, and beta carotene (Jeffrey et al., 1975). Dinoflagellate photosynthetic systems are unique in having a water-soluble Chl

a-peridinin system and a membrane-bound Chl a-Chl c2-peridinin system (summarized in Iglesias-Prieto, 1996). Evidence of other chlorophylls (c1), carotenoids (fucoxanthin), phycoerythrin, and alloxanthin in some dinoflagellates suggests previous engulfment of diatoms, prymnesiophytes, and cryptomonads (MeyerHarms and Pollehne, 1998). Kleptoplastidy, the usage of prey chloroplasts, has been demonstrated for several dinoflagellates (Fields and Rhodes, 1991; Lewitus et al., 1999). Heterotrophic cells may lack chloroplasts but have a pink protoplasm (Gymnodinium helveticum, Entzia acuta), food vacuoles, and brightly colored (red, orange, yellow) accumulation bodies. Accumulation bodies may also be found in photosynthetic species. The storage material in most dinoflagellates is starch localized outside the chloroplast, although red oil droplets may be seen in the cytoplasm, especially near the end of the growing season. Dinoflagellates have a multilayered cell covering, the amphiesma or theca. The outermost layer is a unit membrane (considered by many the plasma membrane), beneath which are located vesicles that may contain plate material (Loeblich, 1969; Dodge and Crawford, 1970), then a membrane bounding the cell contents. Some dinoflagellates have a pellicle beneath the thecal layer (Morrill and Loeblich, 1981). Thecate dinoflagellates have cellulose plates in the vesicles beneath the outer membrane. Plate boundaries may have ridges. The taxonomy of thecate forms is based on the number and arrangement of plates. Kofoid (1909) proposed the system that is now widely used. Plates are arranged in concentric rings in relation to the cingulum, with prime (′) designations indicating which ring, and numbered from the most sulcal in a counterclockwise manner (Fig. 4A–D). Apical plates (′) are followed by precingular (′′), postcingular (′′′), and antapical (′′′′). Plates between apical and precingular are anterior intercalary (a), and plates between postcingular and antapical, not in contact with the sulcus, are posterior intercalary (p). Cingular and sulcal plates have also been designated (C for cingular, T for a transition plate between cingulum and sulcus, and S for sulcal: Sa, Sd, Ss, Sm, Sp, Spa) (Fig. 4E) (Balech, 1974, 1980). The plate pattern of Peridinium cinctum is 4′, 3a, 7′′, 5C, 5′′′, 2′′′′. Some species seem more prone to plate shifting, fusion, and splitting than others (Lefèvre 1932). There have been other systems devised to designate plates, notably that of Eaton (1980), who organized plates to reflect hypothesized evolutionary changes. Apical pores occur on some thecate species. The pore may be surrounded by a pore plate (Po), pore canal plates, and other small plates (Dodge and Hermes, 1981; Toriumi and Dodge, 1993). The pres-

20. Dinoflagellates

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FIGURE 4 Typical thecate motile cell (Peridinium gatunense). Plates are numbered according to the system of Kofoid (1909). (A) Ventral view. (B) Dorsal view. (C) Apical view. (D) Antapical view. (E) Sulcal plates, generalized diagram to show relative positions of plates. T, transition plate between cingular and sulcal,; Sa, sulcal anterior; Sp, sulcal posterior; Ss, sulcal sinister (left); Sd, sulcal dexter (right); Sm, sulcal medial; Spa, sulcal plate anterior to Sp.

ence of an apical pore alters the outline of the cell from smooth in those without a pore, to slight ridges, to a distinct chimney. Apical slits occur in Lophodinium and some species of Woloszynskia and may not be evident with the light microscope unless the cell ecdyses (sheds the theca). Some species have thick cellulose plates, some thinner, and some too thin to be resolved without electron microscopy. Plates have trichocyst pores and may have various types of ornamentation. Older cells show striated bands (Fig. 5J) where plates expand during growth, forming distinct overlap patterns. Cells with thicker plates may have plate extensions, either spines or lists (winglike flanges), along plate margins or on plate interiors (lists on Peridinium willei, Figs. 3D and 5I; Sphaerodinium fimbriatum, Figs. 6C and 7A; spines on Peridiniopsis quadridens, Fig. 3E). Plates may be involved in forming cell extensions called horns (Ceratium; Fig. 3G and 8D) or lobes (Peridinium cinctum f. tuberosum). Cell shape varies from spherical to oval/ovate, with alternative shapes appreciated by the taxonomist (i.e., Ceratium). There are degrees of dorsoventral compression from none (Peridinium gatunense) to extreme con-

cavity (Peridiniopsis polonicum). Lateral compression is rare in freshwater dinoflagellates (Amphidiniopsis being the exception). Apical–antapical depression is uncommon (some in Peridinium gatunense). Morphological variability within a species is slight, except for Ceratium hirundinella (Pearsall, 1929; Hutchinson, 1967). Athecate cells may be “gymnodinioid” or not. Gymnodinioid genera (Amphidinium, Bernardinium, Gymnodinium, Gyrodinium, Katodinium) are motile with a cingulum and sulcus. Their taxonomy is based on cell shape, location and completeness of cingulum, penetration of the sulcus into the epicone and hypocone, presence and color of chloroplasts, location of nucleus, color of cytoplasm, size, eyespot, and presence of accumulation bodies. Athecate cells that are nonmotile (nongymnodinioid) may have typical dinoflagellate coloring (Cystodinium, Hypnodinium, Tetradinium). Nonmotile genera are free floating or are attached by long (Stylodinium; Fig. 2L), short (Tetradinium, Fig. 2M), or rhizoidal (Oodinium, Fig. 2N) stalks or attachment discs (Dinococcus; Fig. 2K) or directly (Cystodinedria; Fig. 2H) to various substrata.

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FIGURE 5 Light (LM) and SEM micrographs of armored dinoflagellates, location included (county, state). Line scale on SEM = 10 µm, line scale on LM = 20 µm. (A) Gonyaulax spinifera, LM, note offset cingulum, apical pore, hypothecal spines (arrowhead) (Suffolk, NY). (B) Thompsodinium intermedium, LM; ventral view, empty cell (Suffolk, NY). (C) Thompsodinium intermedium, LM, note hypothecal fringe (arrowhead) (Suffolk, NY). (D) Thompsodinium intermedium, SEM, ventral view (Brazos, TX). (E) Thompsodinium intermedium, SEM, dorsal view; note large postcingular plates (Brazos, TX). (F) Thompsodinium intermedium, SEM, epithecal plate pattern; note apical pore, star pattern of apical and apical intercalary plates (Brazos, TX). (G) Peridinium gatunense, SEM, ventral view; note reticulate ornamentation (Burleson,TX). (H) Peridinium gatunense, LM, three cells; note round cross sections, “lumpy” appearence of cell. (I.) Peridinium willei, LM; note apical flange (arrowhead), posterior flanges (Seneca, OH). (J) Peridiniopsis polonicum, SEM, dorsal view; note two apical intercalary plates, striated growth bands (arrowhead), single posterior spine (Hamilton, OH). (K) Peridiniopsis polonicum LM, ventral view; note large 1’ plate, single posterior spine (Brazos, TX). (L) Peridiniopsis quadridens, LM, empty cell (Huron, OH). (M) Peridiniopsis penardiforme, LM (Huron, OH). (N) Durinskia baltica. (O) Durinskia Daltica, SEM, ventral view (Sandusky, OH).

20. Dinoflagellates

FIGURE 6 Line drawings of armored dinoflagellates. Where there is no line scale, refer to the text for cell size. (A) Hemidinium nasutum. side. (B) Glenodiniopsis steinii. a, ventral; b, dorsal; c, epitheca. (C) Sphaerodinium fimbriatum. a, ventral; b, epitheca. (D) Entzia acuta. a, ventral; b, epitheca. (E) Kansodinium ambiguum; a, ventral; b, hypotheca; c, epitheca. (F) Dinosphaera palustris. a, ventral; b. epitheca. (G) Amphidiniopsis sibbaldii. a, ventral; b, dorsal; c, epitheca. B is from Highfill and Pfiester (1992b) with permission. Cb is from Carty (1986) Fa is from Prescott (1951) with permission. Fb is from Kofoid and Michner (1912) G is from Nicholls (1998) with permission.

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FIGURE 7 Light (LM) and SEM micrographs of armored dinoflagellates, location included (county, state). Line scale on SEM = 10 µm, line scale on LM = 20 µm. (A) Sphaerodinium fimbriatum SEM, side view. Note frimbriae on plates (Williams, OH). (B) Sphaerodinium fimbriatum, LM, empty cell (Williams, OH). (C) Kansodinium ambiguum, LM (Hill, TX). (D) Kansodinium ambiguum, SEM, epitheca. Note apical pore, three apical plates (Hill, TX).

B. Life Cycle

TABLE III Sexually Reproducing Freshwater Dinoflagellates

1. Life Cycles of Motile Taxa

Taxon

Reference

Ceratium furcoides Ceratium cornutum Cystodinium bataviense Durinskia baltica Glenodiniopsis steinii Gloeodinium montanum Gymnodinium paradoxum Peridiniopsis cunningtonii Peridiniopsis lubieniensiforme Peridiniopsis penardii Peridinium bipes Peridinium cinctum Peridinium gatunense Peridinium inconspicuum Peridinium limbatum Peridinium volzii Peridinium willei Woloszynskia apiculata Woloszynskia pseudopalustre

Hickel (1988) Stosch (1965) Pfiester and Lynch (1980) Chesnick and Cox (1987, 1989) Highfill and Pfiester (1992a) Kelley and Pfiester (1990) Stosch (1972) Sako et al. (1985) Diwald (1938) Sako et al. (1987) Park and Hayashi (1993) Pfiester (1975) Pfiester (1977) Pfiester et al. (1984) Pfiester and Skvarla (1980) Pfiester and Skvarla (1979) Pfiester (1976) Stosch (1973) Stosch (1973)

Dinoflagellate life cycles include a growing, dividing, assimilative stage and a resting (cyst) stage (Fig. 9). About half of the genera have an assimilative stage that is motile, with a distinct cingulum, and are planktonic. Cells are typically haploid and divide mitotically to produce other assimilative cells or gametes. Cells undergoing mitosis may remain motile or form temporary nonmotile cells. Gymnodinium undergoes binary fission (Pfiester and Anderson, 1987). Some taxa ecdyse from the parental cell before division, and some may divide within the parental theca (Fig. 8G), which is then shed (Pfiester and Anderson, 1987). Ceratium donates part of the parental theca to each daughter cell (Fig. 8A). The sexual life cycles of some dinoflagellates have been determined (Table III). Assimilative, haploid cells undergo mitosis to produce cells that function as gametes. In culture, producton of gametes has been induced by nitrogen deficiency in some taxa (Pfiester, 1975, 1976, 1977; Chapman and Pfiester, 1995). Gametes may be the same size and shape as the parental cell (hologametes) or smaller. Gametes may be the same (isogamy) or different (anisogamy) sizes, clones may be monoecious and able to produce zygotes, or dioecious and require different clones to produce zygotes (Pfiester and Skvarla, 1979). Gametes fuse in the sulcal region, and, in Peridinium cinctum, the nuclei meet in a fertilization tube between the gametes (Pfiester, 1984). The resulting planozygote may have two trailing flagella and remain motile for a period of growth. Subsequently, it becomes a nonmotile

hypnozygote and settles to the sediment. Encystment occurs in north temperate areas during the autumn to overwinter. In subtropical Lake Kinneret, Israel, cysts are instead formed to oversummer extreme conditions (Pollingher et al., 1993). There is a period of dormancy following cyst formation before excystment can occur. The dormancy period may require cold and dark (Peridiniopsis cunningtonii, Sako et al., 1985; Peridinium bipes, Park and Hayashi, 1992) followed by light (Peridinium bipes, Park and Hayashi, 1992). Studies of excystment include long-term observations of Ceratium hirundinella in Esthwaite Water,

20. Dinoflagellates

FIGURE 8 Light (LM) and SEM micrographs of armored dinoflagellates, location included (county, state). Line scale on SEM = 10 µm, line scale on LM = 20 µm except as noted (LMs of Ceratium). (A) Ceratium hirundinella SEM. Note that reticulate ornamentation is heavier on older, lower (left in micrograph) sections and lighter on upper (regenerating) section; trichocyst pores are evident in upper section (Washington, TX). (B) Ceratium hirundinella f. hirundinella LM, line scale = 50 µm (Seneca, OH). (C) Ceratium brachyceros, LM, line scale = 50 µm (Huron, OH). (D) Ceratium hirundinella f piburgense, LM, line scale = 50 µm (Huron, OH). (E) Lophodinium polylophum SEM ventral view; note ridges (Brazos, TX). (F) Lophodinium polylophum, SEM, dorsal view (Brazos, TX). (G) Lophodinium polylophum, LM, dividing cell (Brazos, TX). (H) Entzia acuta, LM. Note posterior sulcal extension (arrowhead), apical pore (Stark, OH). (I) Woloszynskia reticulata SEM. Note heavy hypothecal sutures. (J) Woloszynskia SEM. Note thin, polygonal plates, (Fulton, OH). K. Hemidinium nasutum LM, (Brazos, TX). L. Hemidinium nasutum, LM. Note slashed appearance of cingulum (Brazos, TX). E, F, G are from Carty and Cox (1985) with permission.

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FIGURE 9 Generalized life cycle of a motile, photosynthetic, thecate dinoflagellate.

English Lake District, where a rise in temperature from 3°C to 5°C at the end of cold winter temperatures corresponded to a large increase in the number of vegetative cells in the plankton and empty cysts in the sediments (Heaney et al., 1983). In Japan, cysts collected in April from bottom sediments at 10–20°C showed maximum excystment in the lab between 20°C and 25°C, with none at 5°C (Kawabata and Banba, 1993). Work with other taxa found the optimum temperature for excystment of Peridiniopsis cunningtonii to be 22°C (Sako et al., 1985) and 15–25°C for Peridinium bipes (Park and Hayashi, 1992). The seemingly conflicting reports concerning the effects of temperature and light on excystment may indicate that beyond a required dormancy period, excystment is controlled by an internal biological clock (Perez et al., 1998; Rengefors and Anderson, 1998). Excystment ceases in temperate climates in midspring, possibly because of anoxia of sediments (Heaney et al., 1983).

2. Life Cycles of Nonmotile Taxa The order Phytodiniales (Dinococcales) includes cells that are nonmotile in the assimilative form and frequently include a parasitic or amoeboid stage. This group has complex, heteromorphic life histories, including parasitic or photosynthetic assimilative

stages, gymnodinioid or amoeboid swarmers (gametes?), and cysts. In the past, different generic names have been assigned to different stages of the same life cycle, such as Hemidinium (photosynthetic, motile swarmer) and Gloeodinium (cyst), Cystodinedria (cyst?), and Vampyrella (motile cell). It is also possible that different genus designations have been assigned to slight differences in morphology of the same entity, such as Cystodinium, Hypnodinium, and Dinococcus. Examples of complex life cycles include Stylodinium, which is an attached, round to oval cell on a stalk that produces amoebae that parasitize the filamentous chlorophyte Oedogonium. The amoebae then swell into the Stylodinium shape, which may release amoebae or gymnodinioid cells. Some gymnodinioid cells, with yellow-brown chromatophores and stigmas, behave like gametes (Pfiester and Popovsky, 1979). Cystodinedria inermis attached to Oedogonium was observed to release amoebae that later fed on filaments of Spirogyra, after which they became immobile, rounded up, and eventually took on the brownish appearance typical of Cystodinedria (Pfiester and Popovsky, 1979). Cystodinium bataviense reproduces both motile gymnodinioid zoospores and parasitic amoeboid stages (Pfiester and Lynch, 1980).

20. Dinoflagellates

The fish parasite Haidadinium also has a complex life history. The photosynthetic vegetative cyst, with dinokaryon, induces hyperplasia in stickleback fish, although a trophont (feeding) stage is rarely observed (Buckland-Nicks and Reimchen, 1995). This cyst stage repeatedly divides, producing dinospores, rhizopodial amoebae, or lobose amoebae. Motile dinospores contain chloroplasts and a dinokaryon and may be the infective stage. Rhizopodial amoebae are heterotrophic, ingest bacteria, and eventually produce yellow resting cysts. Lobose amoebae, with a eukaryotic nucleus, may generate spheroid amoebae (Buckland-Nicks and Reimchen, 1995). Many of the stages contain symbiotic bacteria. This organism is placed in the Phytodiniales, as its nutrition does not depend on fish, and it has features in common with other members of the order. Another fish parasite, Oodinium, has a parasitic trophont stage that feeds on the fish. A gymnodinioid swarmer is the infective form, settling onto the fish, extending a feeding tube, and then increasing greatly in size. This is followed by encystment and multiple divisions of the cyst, eventually producing the swarmer stage (Jacobs, 1946).

C. Classification A classification scheme is sought that reflects the phylogenetic relationship of dinoflagellates to other protists and relationships within the dinoflagellates. Dinoflagellates appear to be monophyletic, sharing a large suite of characters unique to themselves. This uniqueness is the root of the difficulty in determining evolutionary directions. Dinoflagellates lack an outgroup for comparison, and attempts have been made to organize the distinctive morphological groups without clear agreement on what consititutes ancestral characteristics; these include a plate-increase model and a plate-decrease model. In the former, few-plated taxa (Prorocentrum) become increasingly fragmented, leading through Peridinium-like taxa to many plated forms (Woloszynskia) and ultimately gymnodinioids (Loeblich, 1976; Taylor, 1980). The plate-decrease model places gymnodinioids in the ancestral position and ends with few plated Prorocentrum (Eaton, 1980; Dodge, 1983). There is also a plate fragmentation model that attempts to reconcile the variation in living cells with the fossil record (Bujak and Williams 1981). Molecular sequencing data are beginning to resolve some relationships and confirm monophyly (Saunders et al., 1997; Taylor, 1999). This monophyletic group has been historically claimed and named under both the Code of Botanical Nomenclature and Code of Zoological Nomenclature. Fensome et al., (1993) use

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botanical nomenclature, but call the division Dinophyta rather than Pyrrhophyta (used by phycologists) or a division name based on a genus. The present account recognizes dinoflagellates as the division Pyrrhophyta. Classification within the dinoflagellates (Tables IV and V) is complicated by species with complex life cycles. Since previous classification was based in part on morphology, there has been synonymizing as different parts of one species life cycle are united (Pfiester and Highfill, 1993). This chapter includes “cyst” names because (a) the cell may the assimilative form; (b) the cyst form may not yet be correlated with

TABLE IV Classification of Freshwater Dinoflagellatesa Kingdom: Protista Division: Pyrrhophyta Class: Dinophyceae Order Blastodiniales Family: Oodiniaceae Genus: Oodinium Order Dinamoebales Family Dinamoebaceae Genus: Dinamoebidium Order Gymnodiniales Family: Actiniscaceae Genera: Actiniscus, Pseudoactiniscus Family: Gymnodiniaceae Genera: Amphidinium, Bernardinium, Gymnodinium, Gyrodinium, Katodinium Order: Peridiniales Family: Gonyaulacaceae Genera: Gonyaulax, Thompsodinium Family: Peridiniaceae Genera: Peridinium, Peridiniopsis, Durinskia Family: Ceratiaceae Genus: Ceratium Family: Lophodiniaceae Genera: Lophodinium, Woloszynskia Family: Hemidiniaceae Genus: Hemidinium Family: Glenodiniopsidaceae Genera: Glenodiniopsis, Sphaerodinium Family: Dinosphaeraceae Genera: Dinosphaera, Entzia, Kansodinium Family Thecadiniaceae Genus: Amphidiniopsis Order: Phytodiniales (Dinococcales) Family: Phytodiniaceae Genera: Cystodinedria, Cystodinium, Dinastridium, Dinococcus, Haidadinium, Hypnodinium, Phytodinium, Rufusiella, Stylodinium, Tetradinium Order Prorocentrales Family Prorocentraceae Genus: Exuviaella a

Modified from Loeblich (1982), with new genera included and marine taxa excluded. Only genera reported from North America are included.

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TABLE V Number of Species Reported from North American Literaturea Genus (#sp + forms)

U.S.

Oodinium (1) Dinamœbidium (1) Actiniscus (2) Pseudoactiniscus (1) Amphidinium (6) Bernardinium (1) Gymnodinium (29) Gyrodinium (1) Katodinium (7) Gonyaulax (3) Thompsodinium (1) Peridinium (31+9) Peridiniopsis (17) Durinskia (1) Ceratium (5+8) Lophodinium (1) Woloszynskia (5) Hemidinium (2) Glenodiniopsis (1) Sphaerodinium (3) Dinosphaera (1) Entzia (1) Kansodinium (1) Amphidiniopsis (1) Cystodinedria (1) Cystodinium (6) Dinastridium (1) Dinococcus (2) Haidadinium (1) Hypnodinium (1) Phytodinium (1) Rufusiella (1) Stylodinium (3) Tetradinium (4) Exuviaella (1)

1 1

a

6 1 23 1 5 2 1 22 13 1 9 1 4 2 1 3

Canada

Mexico

Carib.+ C.A.

1 2 1 2 1 18

1

3

23 11 1 8 2 1 1 1 1

7 2 5 1

1 6 4 1

1 1 1

1 1 1 1 6 1 2 1 1 1 3 3 1

4

1 1

1 1

From Buckland-Nicks et al. (1997), Bursa (1969), Bursa (1970), Carty (1986), Carty (1993), Duthie and Socha (1976), Forest (1954) , Haberyan et al. (1995), Meyer and Brook (1969), Nicholls (1998), Ortega (1984), Popovsky (1970), Poulin et al. (1995), Stein and Borden (1979), Taft and Taft (1971), Thompson (1947), Thompson (1949), Thompson (1950), Whitford and Schumacher (1984).

an assimilative stage, or one cyst form may be correlated with more than one assimilative genus (as with the cyst genera Hypnodinium and Dinastridium); (c) many times only one form may be reported (either Hemidinium or Gloeodinium); and (d) the name of the motile cell rather than the cyst form may be used, according to the taxonomic convention. Linking cysts to motile cells from the marine environment allowed a better understanding of the taxonomic affinities of fossil cysts, many of which looked similar to modern cysts. The cyst genus Spiniferites was linked to the motile cell Gonyaulax, and it is proposed that even

though Spiniferites had precedent, the name of the motile cell would be used. By this convention, all species of Cystodinium would become species of Gymnodinium. Some freshwater dinoflagellate genera have a nonmotile assimilative stage, are capable of reproducing similar cells via autospores, and have a brief, motile (gametic?) stage (e.g., Cystodinium). It makes more sense that the assimilative form have precedence. Another complication for classification has been the division of species into thecate (with cellulose plates) or naked (lacking plates). While heavily thecate taxa are obvious in the light microscope, taxa with thin theca or apparently no theca are problematic. Steidinger et al., (1996b) have demonstrated that gymnodinioid (naked) cells may show evidence of plates when examined with the scanning electron microscope (SEM). Confounding the problems of life cycle and cell covering is the unknown degree of variability inherent in the arrangement of thecal plates. Taxa may be synonymized as they are recognized as nutritional or developmental variants, or speciated if differences are found to be stable. Some generic distinctions are unclear, such as those between Peridinium and Peridiniopsis. Peridinium consists of at least two genera, those with larger cells and three intercalary plates (Peridinium sensu stricto according to Boltovskoy in Bujak and Davies, 1983) and those with smaller cells and two intercalary plates (the Umbonatum group). The species within the genus Peridinium need careful evaluation. Popovsky and Pfiester (1990) synonymized many species in Peridinium, which makes keying them easier, but this is less accurate than consulting a reference that includes all species. Peridiniopsis has clearly defined species, but it is questionable that they belong in the same genus. The variety of plate patterns (Table VI), coupled with currrent understanding of plate variation due to plate shifting, loss, and fusion, should enable us to determine which are basal patterns and which are derived.

TABLE VI Apical Plate Patterns of Peridiniopsis Apical plates

Apical intercalary

Precingular

Example

3 3 4 4 4 4 5 5 5

1 1 0 0 1 1 0 0 1

6 7 6 7 7 6 6 7 7

P. borgei P. lindemannii P. penardii P. elpatiewskyi P. lubiensiforme P. cunningtonii P. cunningtonii P. thompsonii P. quadridens

20. Dinoflagellates

Separation of species into new genera can give us better clues to the evolution of the thecate taxa. Defining and distinguishing a species has always been a difficult task. A practical consideration should be if the taxon is morphologically distinctive enough to separate it from similar taxa, and whether this separation adds to our understanding of their ecological roles and evolutionary position. For example, forms of Ceratium hirundinella, hypothesized to be seasonal variations, may later be recognized as individual species based on distinctive cysts and stable morphology.

III. ECOLOGY AND DISTRIBUTION A. Dinoflagellate Blooms Generally, freshwater dinoflagellates are minor members of the summer phytoplankton maxima.

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However, a few species are capable of blooms. Most freshwater blooms are benign, almost unialgal assemblages (Tables VII and VIII). Harmful algal blooms (HABs) and red tides are generally not found in freshwater, although better definitions are needed (Smayda, 1997). The best known freshwater bloom former is Ceratium hirundinella, which has been studied in many parts of the United States, Canada, Europe, Africa, and Japan (Table VIII). Densities of dinoflagellates in a body of water are related to both bottom-up (factors promoting growth) and top-down (factors causing loss) processes. Factors promoting growth include inoculum, inorganic nutrients, vitamins, light, oxygen, temperature, pH, and lake morphometry (depth, stratification, ratio of epilimnion to hypolimnion) (Pollingher, 1987). Factors causing loss include predation, disease, life history characteristics (cyst production), and outflow.

TABLE VII Freshwater Dinoflagellate Blooms Organism

Location

Intensity

Peridinium bipes

Japan Japan Taiwan Israel Israel Finland OH, USA Canada England GA, USA USA Japan Japan OK, USA

Max 192 cells/ml 99% 95.7% biomass 95% biomass 1672 cells/ml “Virtual monoculture” 88% composition, 690 cells/ml 70% >95% sample 88%, 1868 cells/L 1.5 × 105 cells/L 4 × 104 cells/ml

Peridinium gatunense Peridinium inconspicuum Peridinium limbatum Peridinium lomnickii Peridinium pusillum Peridinium willei Peridiniopsis cunningtonii Peridiniopsis penardii Woloszynskia reticulata

“Bloom condition”

Comment

Reference

Reservoir Water odoriferous and brown

Park and Hayashi (1993) Yoshikawa et al. (1997) Wu and Chou (1998) Lindström (1991) Pollingher and Hickel (1991) Holopainen (1992)

Acidic lake, July–Aug Koryak (1978) Acid lake Eutrophic lake July, dystrophic lake Shallow, sheltered areas Reservoir, summer >17°C winter–spring August

Yan and Stokes (1978) Cranwell et al. (1985) Stoneburner and Smock (1980) Stewart and Blinn (1976) Sako et al. (1984) Sako et al. (1987) Pfiester et al. (1980)

TABLE VIII Ceratium hirundinella Blooms Location

Intensity

Comment

Utah L., USA (1) Eau Galle Res. WI, USA (2) Kam, NWT, Canada (4) Properus, NWT, Canada (4) Prelude, NWT, Canada (4) Grace, NWT, Canada (4) Long, NWT, Canada (4) Madeline, NWT, Canada (4) Ishitigawa Res. Japan (3) Esthwaite Water, GB (5) L. Balaton, Hungary (6) Goczalkowice Res. Poland (7) L. Sempach, Switz (8)

89–100% total standing crop 36–94% total biomass 55 × 103 cells/m3 50 × 103 cells/m3 48 × 103 cells/m3 92 × 103 cells/m3 26 × 103 cells/m3 100 × 103 cells/m3 1300 cells/ml 103 cells/ml 4.85 × 104 cells/ml 4.3 × 105 cells/dm3 380 cells/ml

Shallow, TDS 795–1650, high silt, PO4, NO3, pH 8.5+ Eutrophic, moderately alkaline, shallow Eutrophic, Aug, pH 6.9–7.9 Oligotrophic, Aug, DO > 80%, pH 7–7.4, low TP, low NO3 Oligotrophic, Aug, DO > 80%, pH 7–7.4, low TP, low NO3 Mesotrophic, June/Sept, pH 6.8–7.7 Mesotrophic, Aug/mSept, pH 6.8–7.7 Mesotrophic, Aug, pH 6.8–7.7

Shallow

(1) Whiting et al. (1978); (2) James et al. (1992); (3) Kawabata and Kagawa (1988); (4) Moore (1981); (5) Heaney and Talling (1980); (6) Padisák (1985); (7) Bucka and Zurek (1992); (8) Pollingher et al. (1993).

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Bottom-up factors include cysts that may eventually serve as inoculum and which form during a previous bloom (see also Section II.B on the Life Cycle). Ceratium hirundinella has a high percentage of conversion between assimilative cells and cysts but low cyst viability (1–6%) (Pollingher et al., 1993). Peridinium willei has a high rate of survival of cysts (50–81%), though fewer cysts are produced from the planktonic population (Pollingher et al., 1993). Peridinium gatunense also has a low (1%) proportion of the population forming cysts, but this is enough to produce blooms in Lake Kinneret (Pollingher, 1987). The actual timing, intensity, and duration of a bloom involve factors such as turbulence, which resuspends the cysts (Pollingher and Hickel, 1991), and water depth. Shallow lakes have greater emergence rates (Peridinium limbatum, Sanderson and Frost, 1996). Inorganic nutrients, particularly high levels of nitrates and phosphates, are often cited as factors necessary to trigger blooms (Whiting et al., 1978). Ceratium hirundinella was collected in higher numbers near a river inflow with elevated NO3 and PO4 than the reservoir (Kawabata and Kagawa, 1988). Uptake of PO4 by Ceratium hirundinella may be favored in June by downward migration at night (up to four meters) to the upper hypolimnion, but in July anoxia restricts migration and phosphate uptake, and a decrease in Ceratium hirundinella biomass is noted (James, et al., 1992). The mechanism explaining the effect of nitrates on blooms may be a low affinity for nitrate by nitrogen reductase, which allows the dinoflagellate to outcompete other algae (Witt et al., 1999, for Peridinium gatunense). Peridinium abundance may also correlate with phosphorus levels, but not with nitrogen (Wu and Chou, 1998). Other studies seem to indicate that above a minimal level nutrients are not limiting (Padisák, 1985; Sanderson and Frost, 1996). In addition to inorganic nutrients, many dinoflagellates seem to require vitamins such as B12 (Bruno and McLaughlin, 1977; Holt and Pfiester, 1981). Light is an important factor in the vertical distribution of dinoflagellates. Over 95% of cell counts were from subsurface waters (3–15 m) in Feitsui Reservoir, Taiwan (Wu and Chou, 1998). Heaney and Talling (1980) similarly found that C. hirundinella avoided surface waters of Esthwaite Water, U.K. In culture, higher light intensity yielded higher cell densities for this species (Bruno and McLaughlin, 1977), but in nature the compromise between surface light and nutrient supply at depth may explain why C. hirundinella can be situated at depths corresponding to about 10% of summer surface irradiance values (Heaney and Furnass, 1980), or even a deep chlorophyll maximum at 1–3% surface incident light (Gálvez et al., 1988; Echevarria and Rodriguez, 1994).

Most dinoflagellates are long day or warm temperature organisms with maximum growth during the summer. Ceratium hirundinella has been observed in plankton in March–April, with exponential growth during June and July, followed by a stationary period in July–September and a decline in October–November (Padisák, 1985). Growth occurs in subarctic lakes from June to September, with maximum abundance when water is between 4°C and 18°C, suggesting that temperature may not be a controlling factor. Blooms occur as days shorten and when there are strong thermoclines (Moore, 1981). In culture, an axenic strain of Ceratium hirundinella from Calder Lake, NY, grew at all tested pH values from 5.0 to 8.5, and grew best at pH 7.0–7.5 (Bruno and McLaughlin, 1977). This same strain also grew best at total dissolved solids ranging from 48 to 960 mg/L. (Bruno and McLaughlin, 1977), but in the field this species has been found blooming between 795 and 1650 mg/L (Utah Lake; Whiting et al., 1978). Interwoven in many of the resource factors is the influence of stratification. Dinoflagellates, being motile, can remain in the light, oxygenated, warmer surface waters and avoid sedimentation into the dark, anoxic hypolimnion (Heaney and Talling, 1980). Experiments with Ceratium in unmixed columns support these observations (Klemer and Barko, 1991). It has been suggested that in eutrophic lakes, dinoflagellates are absent only from system with high flushing rates or unstratified water columns (Klemer and Barko, 1991, citing Sommer et al., 1986). Top-down factors may be important for freshwater dinoflagellates, although little documentation exists, and some of that is conflicting. This is particularly true for Ceratium, which has been described as a “large, relatively ungrazed species” (Smayda, 1997). There even is some evidence that it reduces the number of zooplankters during a bloom (Bucka and Zurek, 1992). Another study in a fishless pond found that a crash in the population of Daphnia (filter feeding cladoceran) corresponded with a bloom of Ceratium hirundinella that displaced populations of smaller phytoplankton species that Daphnia fed upon. Lacking Daphnia as a food source, larger dipteran predators ate Ceratium (Xie et al., 1998). Other large dinoflagellates such as Peridinium limbatum may also be unaffected by grazing pressure, although effects may also depend on zooplankton size (Sanderson and Frost, 1996). Dinoflagellate losses can also occur through lake outflows, especially if wind direction has concentrated cells at the outflow end. Loss of 880 cells mL–1 × a discharge of 87 × 103 m3 d–1 produced a loss of 7.65 × 1013 cells d–1 (Heaney and Talling, 1980). Changing the

20. Dinoflagellates

site of withdrawal from the hypolimnion to the surface may also drastically reduce biomass (James et al., 1992). The number of dinoflagellates occurring in a body of water is also determined by biological factors such as motility and physical features of the lake. Motility allows for vertical migration related to a circadian rhythm. Ceratium hirundinella in particular may be localized at the surface during early hours and deeper by the afternoon (Heaney and Furnass, 1980; Padisák, 1985). In a deep, subarctic Canadian lake, densities of C. hirundinella were > 4 × 104 cells m–3 at 60 m and > 2 × 103 cells m–3 at 70-m depths (Moore, 1981). Ceratium also apparently position themselves between the foam rows caused by Langmuir circulation in larger lakes (Squires et al., 1979) and show variable horizontal distributions within reservoirs from inflow to dam (Padisak, 1985; Kawabata and Kagawa, 1988). High densities of Peridinium have also been found near the inflow (Wu and Chou, 1998).

B. Trophic States Dinoflagellates are highly variable in trophic status, including pigmented autotrophs, auxotrophs (require exogenous vitamins), mixotrophs (combine autotrophy with phagotrophy), and organotrophs (strict heterotrophs lacking chloroplasts) (Holt and Pfiester, 1981; Gaines and Elbrächter, 1987; Stoecker, 1998). Mixotrophic phagotrophy is exhibited in Ceratium hirundinella through production of a feeding veil (pallium), extracellular digestion, and a pseudopod that draws prey into the cell. Dinoflagellates utilizing a pallium begin the sequence with a pre-capture swimming pattern in which a tow filament rapidly connects with prey and a pseudopod is extended over the prey. Following digestion of the prey cell contents, the pallium is retracted into the theca (Jacobson and Anderson, 1986). Details of the feeding behavior of Peridiniopsis berolinensie indicate a chemosensory attraction to injured prey, attachment via a capture filament, extension of a feeding tube (form of peduncle), and suction of the prey contents (Calado and Moestrup, 1997). A feeding rate of 0.6 cells hr–1 has been measured with cryptophyte prey in the lab (Weisse and Kirchhoff, 1997). Organotrophy includes osmotrophy (absorbing dissolved organics), phagotrophy (particle ingestion, dinoflagellates ingest other dinoflagellates, diatoms, cyanobacteria, ciliates, metazoans), myzocytosis (cell contents sucked out via peduncle, seen in Katodinium fungiforme, Cystodinedria, and Stylodinium; Frey and Stoermer, 1980; Spero, 1985), and ectoparasitism of fish (Oodinium, Jacobs, 1946; Haidadinium, Buckland-Nicks et al., 1997).

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C. Specificity of Habitat Dinoflagellates occur most often in lentic habitats. Some are more common in large bodies of water, such as reservoirs and lakes (Ceratium hirundinella, Peridiniopsis polonicum), and others in ponds (Dinococcales found in duckweed ponds). Some species are more frequently encountered in soft water (acid to neutral), such as Peridinium limbatum, Gymnodinium caudatum, and Ceratium carolinianum), while others are more common in hard water (alkaline), such as Ceratium hirundinella. Some species may be more prevalent in eutrophic systems (Ceratium hirundinella) and others in brackish systems (Gonyaulax, Exuviaella). Unusual habitats may also harbor dinoflagellates. Rufusiella was reported “on the under surface of a dripping sandstone ledge” (Richards, 1962), and Dinamoebidium was attached to sand grains in an alpine stream (Bursa, 1970). Sand dwelling dinoflagellates are known from marine sands (Saunders and Dodge, 1984) and have been collected from a freshwater sandy beach (Nicholls, 1998). Fish parasites are well known, especially by aquarium hobbyists (Ling et al., 1993; Buckland-Nicks and Reimchen, 1995).

D. Geographical Distribution Too little is known about the geographic distribution of dinoflagellate species to be able to determine if patterns exist. However, of 11 armored species and 13 unarmored species first reported from North America (Table IX), only three (Peridinium gatunense, P. limbatum, Ceratium carolinianum) have been reported elsewhere. Some species, such as Ceratium hirundinella and Peridinium gatunense, are cosmopolitan, but the lack of reports for other species should not be considered evidence of rarity. Species may be common but unnoticed, and some species have patchy distributions. Three examples from my own experience may provide some insight. Lophodinium polylophum is probably a rare species. It was first collected in Paraguay (Daday, 1905) and has since been collected once from Mexico (Osorio-Tafall, 1942), and from one small pond in College Station, TX (Carty and Cox, 1985). It has not been reported in the literature since, although it is quite distinctive (Figs. 3H and 8E and F). A case of a common species absent from a state record is Peridiniopsis polonicum, another distinctive phytoplankter (Fig. 5J and K). It was not previously reported from Ohio (Taft and Taft, 1971) but has since been found in 24 countries (Carty, 1993; Carty and Fazio, 1997). It is generally widespread and commonly reported from North America and was probably absent

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TABLE IX Armored and Unarmored Freshwater Taxa First Identified in North America Name

Year

Locality

Ceratium carolinianum Peridinium limbatum Peridinium gatunense Peridinium wisconsinense Gymnodinium caudatum Oodinium limneticum Peridiniopsis thompsonii Gymnodinium marylandicum Tetradinium simplex Stylodinium longipes Rufusiella insignis Gymnodinium cruciatum Kansodinium ambiguum Thompsodinium intermedium Sphaerodinium fimbriatum Woloszynskia reticulata Woloszynskia cestocoetes Actiniscus canadensis Pseudoactiniscus apentasterias Dinamoebidium coloradense Katodinium auratum Amphidinium cryophilum Haidadinium ichthyophilum Amphidiniopsis sibbaldii

1850 1888 1925 1930 1944 1946 1947 1947 1949 1949 1949 1950 1950 1950 1950 1950 1950 1969 1969 1970 1970 1982 1997 1998

South Carolina New Jersey Panama Wisconsin Wisconsin Minnesota Kansas Maryland Michigan Maryland Kansas Kansas Kansas Kansas Kansas Kansas Kansas Northwest Territories Northwest Territories Colorado Colorado Wisconsin British Columbia Ontario

because of the limited geographical area covered by the authors. Thompsodinium intermedium was first reported by Thompson in 1950 from Kansas. Later it was found in Cuba (Popovsky, 1970), Texas (Carty, 1986, 1989), and one location in Ohio (Carty, 1993). I have since found large numbers in one deep kettle lake on Long Island, NY, and in large numbers in one tiny, brownwater pond on a peninsula in Belize, (Carty, unpublished observations). This may be a widespread species that is not recognized because of difficulties in identification.

IV. COLLECTION AND PREPARATION FOR IDENTIFICATION A. Collection Dinoflagellates are predominently planktonic and may be collected in whole-water samples or concentrated with a plankton net (10-µm mesh). It is advantageous to sample from different depths, in either discrete aliquots (e.g., Van Dorn sampler) or a tow, as the motile cells can travel throughout the water column and may not be at the surface. Squeezings from submerged vegetation (algal and macrophyte) can also be

collected (Thompson, 1947). Whole-water samples may be centifuged to concentrate organisms or placed in tapered-bottomed containers and the sediment examined. It is also useful to scrape fish. Any freshwater source may be examined, including reservoirs, lakes, ponds, marshes, stock tanks, permanent ditches, fish hatcheries, rivers, and creeks (see also Section III.C). While I have not sampled swimming pools, I have found dinoflagellates in waters treated with commercial shading-type chemicals. In reviewing 330 samples from Ohio, USA, collected in 1997, one-third (107) had no dinoflagellates, another third (114) had one species (frequently Ceratium hirundinella), and a third (107) had more than one, and up to nine, dinoflagellate species (Carty, unpublished observations). Dinoflagellates are in greatest abundance during the summer months, but autumn, spring, and winter collections should also be made. Some species seem more prevalent in cooler months, and some are exclusively cold-weather species. Thompson (1947) has collected cells from beneath ice.

B. Fixation Samples are usually fixed with Lugol’s iodine (to which glycerin is added), which is added until a tea color is reached. Lugol’s is the most widely used preservative, as it is known to maintain delicate flagella and its pH is not detrimental to cell coverings. Fixing a subsample of all collections while in the field will prevent further predation of algal cells by zooplankton and may highlight the presence of scarce dinoflagellates as iodine turns their starch black. After the live sample has been examined and found to contain useful material, the remainder of the sample may be preserved with Lugol’s or glutaraldehyde for later work with the SEM. Long-term storage of samples may be in Lugol’s, although the iodine bleaches with time and fungi can grow. Permanent slides may be prepared with syrup medium (Taft, 1978).

C. Preparation for Identitication Examination of living cells gives details of shape, coloring, size, and swimming pattern crucial for identification. Most naked dinoflagellates lose their shape at death and become spherical. Some species seem to congregate in the water outside of the coverslip, and this area should be examined. While most features can be seen with light microscopy, both phase and interference contrast optics improve the visualization of features. Positive identification of thecate dinoflagellates requires reconstruction of the plate pattern and/or

20. Dinoflagellates

recognition of certain defining characteristics. Many cells will retract from their outer wall, a useful feature for identifying the presence of a theca in those taxa with an obscure or thin theca such as Woloszynskia, especially in reaction to heat stress from the microscope light. Live samples may be left overnight, and the sedimented material may be examined the next day before too many cells have ecdysed. Both living cells and empty thecas may be rolled in the viewing field by gently blowing near the coverslip. Thecae may be dissociated and better seen after the addition of 5% sodium hypochlorite under the coverslip; slight to moderate pressure on the coverslip may be necessary for some species (Boltovskoy, 1975, 1976, 1989). Various stains may be used to aid in identification or to

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gain additional information about the cell. Examples of stains include Sudan stains for lipids, acetocarmine, feulgen, or propionic-lactic-orcein stain for the nucleus, and hydroiodic acid, trypan blue, or chlor-zinc iodine for cellulose (Pearsall, 1929; Sournia, 1978). Scanning electron microscopy (SEM) provides definitive information of plate shapes, numbers, and relationships and may be the easiest method for the reconstruction of sulcal plates. Samples preserved with 2% glutaraldehyde or other fixatives should be criticalpoint dried before sputter coating with gold-paladium. It may be necessary to remove the outer cell membrane to see thin plates (Steidinger et al., 1996b). Hexadimethyldisilazane (HMDS) and air drying may be satisfactory for heavily thecate species.

V. KEY AND DESCRIPTIONS OF GENERA A. Key The frequently encountered cell-stage in the life cycle of freshwater dinoflagellates. 1a.

Cell attached (or with attachment stalk and disc)...............................................................................................................................2

1b.

Cell free..............................................................................................................................................................................................9

2a.

No cingulum evident..........................................................................................................................................................................3

2b.

Attached to sand, moss, slight cingulum (Fig. 2T)......................................................................................................Dinamoebidium

3a.

Spherical cell on distinctive stalk, attached to filamentous algae (Figs. 2L and 10G).........................................................Stylodinium

3b.

Cell on short stalk, attachment disc or directly attached....................................................................................................................4

4a.

Cell contours rounded, sessile, attached to filamentous algae (Figs. 2 and 10D).............................................................Cystodinedria

4b.

Cell otherwise....................................................................................................................................................................................5

5a.

Cell ovoid, with short stalk, attached to fish......................................................................................................................................6

5b.

Cell with angles, drawn out into spines..............................................................................................................................................7

6a.

Cell ovoid, with short stalk (Fig. 2N)...................................................................................................................................Oodinium

6b.

Cell round with translucent, fenestrated matrix (Fig. 2O)................................................................................................Haidadinium

7a.

Oval cell with spines at both ends (Fig. 2K).......................................................................................................................Dinococcus

7b.

Angular cell........................................................................................................................................................................................8

8a.

Tetragonal cell with spines at corners (Figs. 2 and 10H)...................................................................................................Tetradinium

8b.

Irregularly polygonal cell, corners with blunt spines (Figs. 2 and 10I).............................................................................Dinastridium

9a.

No cingulum evident........................................................................................................................................................................10

9b.

Cingulum present.............................................................................................................................................................................15

10a.

Cell surrounded by sheath................................................................................................................................................................11

10b.

Thecate cell lacking sheath (Fig. 3A)...................................................................................................................................Exuviaella

11a.

Cells in thick mucilagenous sheaths.................................................................................................................................................12

11b.

Cells in firm, discrete envelope.........................................................................................................................................................13

12a.

Cell in several-layered envelope (like Gloeocystis; Figs. 2R and 10J).......................................................................Hemidinium (cyst)

12b.

Cell in many-layered, tubelike envelope (like Hormotilia) (Fig. 2S).......................................................................................Rufusiella

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13a.

Cell oval, brown, may have terminal spine(s) (Fig. 2J).....................................................................................................Cystodinium

13b.

Cell round .......................................................................................................................................................................................14

14a.

Cell with chloroplasts in roseate clusters, large vacuoles (Fig. 2J)...................................................................................Hypnodinium

14b.

Cell not as above (Figs. 2Q and 10F)...............................................................................................................................Phytodinium

15a.

Cingulum incomplete.......................................................................................................................................................................16

15b.

Cingulum encircles cell.....................................................................................................................................................................17

16a.

Cell photosynthetic (with chloroplasts; Fig. 6A)...............................................................................................................Hemidinium

16b.

Cell heterotrophic (lacking chloroplasts; Fig. 2D)...........................................................................................................Bernardinium

17a.

Cingulum not medial........................................................................................................................................................................18

17b.

Cingulum medial..............................................................................................................................................................................21

18a.

Cingulum divides cell into ≤1/3 epitheca, ≥2/3 hypotheca................................................................................................................19

18b.

Cingulum divides cell into 2/3 epitheca, 1/3 hypotheca (Fig. 2G).......................................................................................Katodinium

19a.

Cell thecate......................................................................................................................................................................................20

19b.

Cell nonthecate (Fig. 2C)................................................................................................................................................Amphidinium

20a.

Cell with two large plates, often near marine environment (Fig. 3A)....................................................................................Exuviaella

20b.

Cell with more plates, laterally compressed (Fig. 6G)..................................................................................................Amphidiniopsis

21a.

Cingulum ends offset more than 1.5 cingulum widths......................................................................................................................22

21b.

Cingulum ends offset none to less than 1.5 cingulum widths...........................................................................................................23

22a.

Cell athecate (Fig. 2F).......................................................................................................................................................Gyrodinium

22b.

Cell heavily thecate, often near marine environments (Fig. 3B)...........................................................................................Gonyaulax

23a.

Cell with distinctive plates................................................................................................................................................................24

23b.

Cell athecate, with very thin plates, or uncertain .............................................................................................................................31

24a.

Cell with one apical and 2–3 hypothecal horns (Figs. 3G and 8A–C)....................................................................................Ceratium

24b.

Cell without extended horns............................................................................................................................................................25

25a.

Cell heterotrophic, may have pink cytoplasm, 4′, 2a, 7′′, 5′′′, 1′′′′ (Figs. 6D and 8H).................................................................Entzia

25b.

Cell yellow-golden brown................................................................................................................................................................26

26a.

Cell has spine(s) on hypothecal plates..............................................................................................................................................27

26b.

Cell lacks spines, may have lists or fimbrae on plates.......................................................................................................................28

27a.

Plate tabulation 4′, 2–3a, 7′′, 5′′′, 2′′′′ (Figs. 4A–D)...............................................................................................Peridinium (in part)

27b.

Plate tabulation otherwise 3–5′, 0–1a, 6–7′′, 5′′′, 2′′′′ (Fig. 3E)...........................................................................Peridiniopsis (in part)

28a.

Cell with definite thick plate sutures only on hypotheca (Fig. 3I).....................................................................Woloszynskia reticulata

28b.

Sutures uniform on cell....................................................................................................................................................................29

29a.

Single, fringed list on 1′′′′ plate, 4′, 3a, 6′′, 5′′′, 2′′′′ (Fig. 3C).....................................................................................Thompsodinium

29b.

Plate tabulation otherwise................................................................................................................................................................30

30a.

Plate tabulation 4′, 2–3a, 7′′, 5′′′, 2′′′′ (Figs. 4A–D)...............................................................................................Peridinium (in part)

30b.

Plate tabulation otherwise, 3–5′, 0–1a, 6–7′′, 5′′′, 2′′′′ (Fig. 3E)..........................................................................Peridiniopsis (in part)

31a.

Athecate with cell membrane or pellicle...........................................................................................................................................32

31b.

Theca thin (can discern cell contents separate from outer covering), plate sutures not or barely visible............................................34

32a.

Cell from high Arctic lake, may have pentasters within....................................................................................................................33

32b.

Cell from more temperate climate, athecate (Figs. 2E and 10B).....................................................................................Gymnodinium

33a.

Cell with internal siliceous pentasters, rare (Fig. 2A)............................................................................................................Actiniscus

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33b.

Cell lacking pentasters, rare (Fig. 2B).........................................................................................................................Pseudoactiniscus

34a.

Cell with many, thin, polygonal plates..............................................................................................................................................35

34b.

Cell with definite plate pattern.........................................................................................................................................................36

35a.

Cell spindle shaped, with vertical ridges (Figs. 3H and 8E–G)........................................................................................Lophodinium

35b.

Cell round (Figs. 3I and 8I–J)..........................................................................................................................................Woloszynskia

36a.

Three apical plates, one antapical plate............................................................................................................................................37

36b.

Four apical plates, two antapical plates............................................................................................................................................38

37a.

Five precingular plates, apical pore, 3′, 1a, 5′′, 5′′′, 1′′′′ (Fig. 6E).....................................................................................Kansodinium

37b.

Six precingular plates, no apical pore, 3′, 1a, 6′′, 5′′′, 1′′′′ (Fig. 6F)..................................................................................Dinosphaera

38a.

Two anterior intercalary plates, 4′, 2a, 6′′, 5′′′, 2′′′′ (Fig. 3F)................................................................................................Durinskia

38b.

Four anterior intercalary plates........................................................................................................................................................39

39a.

Seven precingular plates, 4′, 4a, 7′′, 6′′′, 2′′′′ (Figs. 6C and 7A–B)................................................................................Sphaerodinium

39b.

Eight precingular plates, 4′, 4a, 8′′, 6–8′′′, 2′′′′ (Fig. 6B)................................................................................................Glenodiniopsis

FIGURE 10 Light micrographs of unarmored dinoflagellates, location included (county, state). Line scale = 20 µm. (A) Bernardinium bernardinense (Brazos, TX). (B) Gymnodinium fuscum (Burleson, TX). (C) Katodinium spiroidinoides (Marion, OH). (D) Cystodinedria inermis (Williams, OH). (E) Cystodinium bataviense (Montgomery, TX). (F) Phytodinium simplex (Brazos, TX). (G) Stylodinium globosum (Kenedy, TX). (H) Tetradinium javanicum (Brazos, TX). (I) Dinastridium sexangulare (Brazos, TX). (J) Hemidinium cyst stage (Brazos, TX).

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B. Descriptions of Genera This section includes the genus name, authority, figure references, the number of species reported from North America, name of species if just one, a brief description, plate pattern, sizes (based on North American specimens where possible), and topical reference if applicable. Several included taxa are from single (worldwide) reports; they are included to call attention to how little is known about dinoflagellates and how much remains to be done. This listing follows the order of genera in Table IV.

Order Blastodiniales Family Oodiniaceae Oodinium Chatton O. limneticum. (Fig. 2N) Epizooic, life cycle includes parasitic, nonmotile cell in the assimilative stage, a cyst, and gymnodinioid swarmers that infect fish. The parasitic cell has light-green to olive chromoplasts, starch, no eyespot, and grows from about 28 µm to 60 µm. It is attached by tentacle-like rhizoids and is enclosed in a thin cellulose wall. Spherical cysts are about 71 µm in diameter. Cysts undergo several divisions, the final forming gymnodinioid swarmers about 15 µm long. Swarmers have a deep cingulum dividing the cell into a larger epicone and smaller hypocone, and a sulcus that extends to the antapex, no eyespot, and numerous yellow-green chromoplasts (Jacobs, 1946). Lom (1981) differentiated parasitic dinoflagellates based primarily on host and features of the trophont stage (especially the mode of attachment). O. limneticum was transferred to Piscinoodinium, a genus erected for freshwater ectoparasites (Lom, 1981). O. limneticum does not attach to hosts like the type species P. pillulare, so further study is required. In addition to the original report, there has been one report from Mexico.

Order Dinamoebales Family Dinamoebaceae Dinamoebidium Pascher (Fig. 2T) Athecate, gymnodinioid/amœboid cell with irregularly scalloped periphery, nonmotile, no eyespot, with golden chromatophores, cells 19.8–38.5 µm long, 13–23 µm wide, amoebae 28–46.5 µm. Found in cold, alpine stream attached to sand grains and moss. Dinoflagellate features include a large nucleus and an “equatorial girdlelike groove” in the gymnodinioid cells. Amoeboid cells ingest algae. There has been one report from CO (USA) (Bursa, 1970).

Order Gymnodiniales Family Actiniscaceae Kützing Actiniscus Ehrenberg (Fig. 2A)

Athecate with

internal siliceous starlike element (pentaster), best known from fossils and marine species, found in Great Bear Lake, Canada. There has been one report (Bursa, 1969). The two freshwater species were not validly described, but marine species are known; additional work needs to be done. While no measurements are given in the text, based on the magnification of the figure, cells are about 59 µm long × 72 µm in diameter. Pseudoactiniscus Bursa P. apentasterias (Fig. 2B) Similar to Actiniscus but lacking pentasters, found in Canada, one report (Bursa, 1969). The genus and species have not been validly described, and additional work needs to be done. While no measurements are given in the text, based on the magnification of the figure, cells are about 70 µm long × 69 µm diameter. Family Gymnodiniaceae (Bergh) Schütt Amphidinium Claparède et Lachmann (Fig. 2C) Athecate, cingulum divides cell into ≤1/3 epitheca, ≥2/3 hypotheca, cells 18.6–22 µm long, 10–14 µm wide, 7–10 µm thick. Four species; reported infrequently from the United States and Canada. Bernardinium Chodat (Figs. 2D and 10A) Like Hemidinium with incomplete cingulum, but nonphotosynthetic, athecate. Cytoplasm may contain red or orange inclusions (food vacuoles?); cells 13–16 µm wide × 20–21 µm long, with eyespot. There have been several reports from the United States. Gymnodinium Stein (Figs. 2E and 10B) Classic “naked” dinoflagellate, some photosynthetic with peridinin, two bluegreen, others heterotrophic. Cingulum bisects cell. Must be examined alive for species determination. Cells small (G. triceratium 16 µm × 13 µm) to large (G. fuscum 55–60 µm × 80–100 µm). Genus reported (29 species) throughout the United States and Canada. Gyrodinium Kofoid et Swezy (Fig. 2F) Cingulum offset 1.5 times, athecate. One report from MD (USA) (Thompson, 1947); cells dorsoventrally compressed, with eyespot, chromatophores present, 25–32 µm long, 18–20 µm wide, 14–15 µm thick, collected in January. Katodinium Fott (Figs. 2G and 10C) Athecate, cingulum divides cell into 2/3 epitheca, 1/3 hypotheca, many heterotrophic; most are small, 4–16 µm × 6–12 µm. Massartia is a synonym. Reported infrequently from the United States and Canada. Must be viewed alive and swimming for genus and species identification (Christen, 1961). Species (7 known) are determined by size, shape, +/– stigma, +/– plastids, sulcal features.

20. Dinoflagellates

Order Peridiniales Schütt Family Gonyaulacaceae Lindemann Gonyaulax Diesing (Figs. 3B and 5A) Heavily thecate, mainly found in brackish water, most species marine; 2–4′, 0–3a, 5–6′′, 1p, 5–6′′′, 1–2′′′′, cingulum very displaced, cells 28–62 µm × 29–58 µm, infrequently reported (3 species). Thompsodinium Bourrelly (Figs. 3C and 5B–F) Photosynthetic golden chloroplasts, with eyespot, fringed list near antapex; 2′′′, 3′′′, and 4′′′ plate, 4′, 3a, 6′′, C6, 5′′′, 2′′′′, 4 sulcal plates, the 2a plate is variable in shape. Cells 28–43 µm long × 26–40 µm in diameter with slight dorsoventral compression; cingulum without displacement. Collected in the United States, Cuba, Belize (see IIID). Family Peridiniaceae Ehrenberg Peridinium Ehrenberg (Figs. 4A–D, 3D, 5G–I) Thecal tabulation 4′, 2–3a, 7′′, 5′′′, 2′′′′; heavily thecate, plates ornamented, most photosynthetic, either with an apical pore (Poroperidinium Lefèvre) or without (Cleistoperidinium Lefèvre). While most species (ca. 31 described) are round to oval, some have lobes (P. cinctum f. tuberosum, P. willei, P. limbatum), and some have distinctive lists (P. willei). Most of the subgroups within Peridinium produce large, robust cells 60–65 µm long and wide, the Umbonatum group includes smaller cells 13–30 µm long × 11–27 µm in diameter. (Lefèvre, 1932). Widely reported (especially P. inconspicuum and P. willei) from Canada, Caribbean, Mexico, and the United States. Peridiniopsis Lemmermann (Figs. 3E, 5J–M) Thecal tabulation 3–5′,0–1a, 6–7′′, 5′′′, 2′′′′, slight to moderately thecate, some with plate ornamentation, some photosynthetic, some heterotrophic, 32–47 µm long × 21–39 µm in diameter. The wide range of plate patterns requires plate determination for identification (Table VI). Species in the genus Glenodinium with known plate patterns were transferred to this genus by Bourrelly (1968), although the genus Glenodinium persists in the literature. Widely reported (17 species) from Canada, Caribbean, Mexico, and the United States. Durinskia Carty et Cox (Figs. 3F, 5N–O) Thecal tabulation 4′, 2a, 6′′, 5′′′, 2′′′′, photosynthetic, with eyespot, cell round to slightly oval, 26–33 µm long × 26–32 µm in diameter, with an apical pore, thin theca, no ornamentation. Collected from freshwater and saline environments. Infrequently reported, found in TX and OH (USA), and, as Peridinium dybowski,

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reported from Canada and MN (USA). As a Peridinium balticum isolate from the Salton Sea, CA (USA), it has been intensely studied because of its binucleate status (Tomas and Cox, 1973; Chesnick and Cox, 1987, 1989). Family Ceratiaceae (Schütt) Lindemann Ceratium Schrank (Figs. 3G and 8A–D) Thecal tabulation 4′, 5′′, 5′′′, 2′′′′; only genus with 1–2 horns formed from postcingular plates, with an apical horn (apical plates) which may have an apical pore and antapical horn (anatapical plates), heavily thecate, plates ornamented, mixotrophic, pale yellow to golden, no eyespot. Stubby C. brachyceros may be 33–40 µm wide × 65–80 µm long; C. hirundinella may be over 400 µm long. Ceratium hirundinella (and many of its forms) are widely reported. Family Lophodiniaceae Lemmermann Lophodinium Lemmermann (Figs. 3H and 8E–G) Thin theca of many hexagonal plates arranged into vertical ridges, photosynthetic with numerous oval golden chloroplasts, eyespot in sulcus, 42–44 (–62 in dividing cells) µm long × 31–41(–54) µm in diameter (OsorioTafall (1942) reports cells 70–80 µm long × 63–67 µm in diameter). Apical slit bounded by carina (ridge) not continuous with sulcus. Rarely encountered, it may have a brief planktonic stage. Reported from Mexico and TX (USA) (Carty and Cox, 1985). Woloszynskia Thompson (Figs. 3I and 8I–J) Cells round with numerous polygonal thin plates which are difficult to see, photosynthetic, infrequently reported. Size: 20–52 µm long × 14–46 µm wide, may have an apical slit. Distinctive cysts may verify the presence of the genus in the plankton. Reported (five species) from OK, TX, KS, OH, MD (USA). Family Hemidiniaceae Bourrelly Hemidinium Stein (Figs. 6A and 8K–L; cyst Figs. 2R and 10J) Motile cell with thin plates (very difficult to see), incomplete cingulum gives cell a “slashed” appearance, photosynthetic with golden brown chloroplasts, 24–29 µm long × 11–20 µm in diameter. Gloeodinium, a genus of immobile round cells in a thick gelatinous matrix was found to be a stage in the life cycle of Hemidinium (Pfiester and Highfill, 1993). The cyst (Gloeodinium) stage has 2–4 nonmotile, round cells in mucilage. Cells have numerous brown chromatophores, are 18–28 µm in diameter; colony is 69–74 µm in diameter. Cysts are epiphytic or free in the plankton.

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Family Glenodiniopsidaceae Schiller Glenodiniopsis Woloszynska (Fig. 6B) Thecal tabulation 4′, 4a, 8′′, 6–8′′′, 2′′′′; no apical pore, photosynthetic, thin asymmetrically arranged plates may require SEM for verification; 26–50 µm long × 26–33 µm in diameter (Highfill and Pfiester 1992a,b). Three reports (OK, MN, BC). Sphaerodinium Woloszynska (Figs. 6C and 7A–B) Thecal tabulation 4′, 4a, 7′′, 6′′′, 2′′′′; photosynthetic with golden chloroplasts, with an eyespot. Plate extensions give a distinctive cell outline for S. fimbriatum. Size: 42–53 µm long × 32–46 µm in diameter, infrequently reported. Genus is considered a synonym of Glenodinium, based on the description of Glenodinium cinctum with a horseshoe-shaped eyespot, which was also found in Sphaerodinium polonicum (Loeblich, 1980). There were no plates originally figured for Glenodinium, but there is a known pattern for Sphaerodinium. It is useful to maintain Sphaerodinium until Glenodinium is defined. Family Dinosphaeraceae Lindemann Dinosphaera Kofoid et Michener (Fig. 6F) Thecal tabulation 3′, 1a, 6′′, 5′′′, 1′′′′; no apical pore, photosynthetic, cells 25–30 µm diameter, 27–34 µm long. Reported, as Glenodinium palustre, from soft water lakes and bogs in WI (USA) (Prescott, 1951) and as Gonyaulax palustris from IA , IL, MA, MN. Entzia Lebour (Figs. 6D and 8H) Thecal tabulation 4′, 2a, 7′′, 5′′′, 1′′′′; heavily thecate with lightly reticulate ornamentation, related to marine genus Diplopsalis, heterotrophic, with apical pore, may have pink cytoplasm, distinctive feature a prominent sulcal list that extends past the antapex. Cell may ecdyse a pink, motile, gymnodinioid/katodinioid cell. Size: 30–38 µm long × 26–38 µm in diameter. Reported from OH (USA). Kansodinium Carty et Cox (Figs. 6E and 7C–D) Thecate, 3′, 1a, 5′′, 5′′′, 1′′′′, apical pore surrounded by apical collar, photosynthetic, with eyespot, 32–42 µm long × 27–39 µm in diameter × 27–31 µm thick, rounded cell with thin plates which require empty cells or SEM for verification. Reported from KS, TX (USA) (Carty and Cox, 1986). Family Thecadiniaceae Amphidiniopsis Woloszynska (Fig. 6G) Thecate, laterally compressed, Po, 4′, 3a, 7′′, 5c, 4(?)s, 5′′′, 2′′′′, apical pore, lacking chloroplasts, cells brownish-gray. Cingulum divides cell into about 1/5 epitheca, 4/5 hypo-

theca, cells 33–45 µm long, 19–28 µm lateral width, 24–34 µm dorsoventral width. Collected from sand, Ontario (Nicholls, 1998).

Order Phytodiniales (Dinococcales) Family Phytodiniaceae Klebs Cystodinedria Pascher (Figs. 2H and 10D) Oval cell, athecate, 38–48 µm long × 22–32 µm in diameter, epiphytic and parasitic on filaments (like Oedogonium, Zygnema, Spirogyra), nonmotile in assimilative stage. Assimilative cell gives rise to amoebae that parasitize green algal filaments and then form the ovoid shape. Formation of gymnodiniod cells is uncertain (Pfiester and Popovsky, 1979). May not be a dinoflagellate at all, but a digestive cyst of the protozoan Vampyrella (M. Elbrächter, personal communication, based on Röpstorf et al., 1994), or Vampyrella may be a stage in the dinoflagellate life history (Pfiester and Popovsky, 1979). The golden-brown color of the cell may be from the parasitized cell. This may not be a valid genus but rather part of the life cycle of another organism (Popovsky, 1982). Reported from OH (USA). Cystodinium Klebs (Figs. 2I and 10E) Athecate, Gymnodinium-like cell inside casing, photosynthetic, nonmotile in assimilative stage, planktonic, 65–118 µm long × 52–66 µm in diameter. Most commonly sampled from small ponds and marshes (Lemna ponds), several reports (six species) from the United States and Canada. Includes a parasitic amoeboid stage (Pfiester and Lynch, 1980). Dinastridium Pascher (Figs. 2P and 10I) Irregularly-shaped cells, usually with six sides (five to seven species), each angle with a single or double short spine, discoid, parietal plastids. Reproduce via gymnodinioid zoopores or autospores (Bourrelly, 1970). Considered by Popovsky and Pfiester (1990) to be hypnospores of other taxa, but recognizable in this form, described as follows: “color and chromatophores are typical for most dinoflagellates”; cell 28–40 µm in diameter (Forest, 1954). From TN (USA). Dinococcus Fott (Fig. 2K) Athecate, epiphytic with short stalk with one spine at either end of the elliptical cell, nonmotile in assimilative stage. Collected in WI (USA) by Prescott (1951) at 11 m depth. Cells 25–35 µm long including spines, 9–12 µm in diameter. Raciborskia Woloszynska is a synonym; may not be a valid genus but rather part of the life cycle of Cystodinium (Pfiester and Lynch, 1980). Haidadinium Buckland-Nicks et al., (Fig. 2O) Ectoparasite on stickleback fish, with complex life

20. Dinoflagellates

history including vegetative cysts (typical dinoflagellate nucleus and chloroplast), lobose and rhizopodial amoeboid stages, and swarmer stage. There has been one report from British Columbia, from small lakes in Sphagnum bogs with low cation concentrations (Buckland-Nicks et al., 1997). Hypnodinium Klebs (Fig. 2J) Spherical planktonic cell, photosynthetic with chloroplasts arranged in roseate clusters, athecate, nonmotile in assimilative stage, 64–66 µm in diameter. There have been a few reports from MD, MN, NC, OH, Quebec. Phytodinium Klebs (Figs. 2Q and 10F) Athecate, nonmotile, round cells enclosed by thick wall, photosynthetic with oval chloroplasts, similar to Cystodinium, reproduce via autospores There has been one report (Meyer and Brook, 1969, from MN, USA) from a dystrophic pond. Rufusiella Loeblich (Fig. 2S) Athecate, cells embedded in an asymetrically layered mucilagenous envelope. Noted to produce gymnodinioid swarmers (Thompson in Smith, 1950), but later work found thecate Hemidinium-like motile cells. Cell single or two to four, protoplast brownish, may contain red oil globules. Cells 30–83 µm in diameter, 41–100 µm long including sheath. Cells collected from scrapings or grown on solid media have typical eccentrically layered sheath; cells grown in liquid culture are more “Gloeodinium”-like in appearance. Collected in KS (USA) as scrapings from wet areas (Richards, 1962). Stylodinium Klebs (Figs. 2L and 10G) Athecate, epiphytic, nonmotile in assimilative stage, may contain colored globules, round cell atop distinct stalk, cell 20 µm long × 17.5 µm in diameter; stalk 12.5 µm long. Parasitic on Oedogonium and Fragilaria, produces gymnodiniod zoospores or amoebae (Pfiester and Popovsky, 1979). Three species known; river habitats, few reports from BC, MD, MN, NY, OH. Tetradinium Klebs (Figs. 2M and 10H) Athecate, epiphytic, photosynthetic with many golden-brown discoid chloroplasts, nonmotile in assimilative stage. Name derived from tetragonal shape of cell; has two spines at each of the four corners. Cell about 20–73 µm across, stalk length variable, 13–23 µm. Reported (four species) from the United States and Canada, attached to filamentous algae (Thompson, 1949).

Order Prorocentrales Bourrelly Family Prorocentraceae Engler Exuviaella Cienkowski (Fig. 3A)

Thecate, usually

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a marine genus, two large heavy plates with smaller plates at the apex. One report from MD (USA), with two parietal chromatophores, a large posterior nucleus, no eyespot, 22–26 µm long, 15–18 µm in diameter, 11–12 µm thick (Thompson, 1950). Two species placed in Prorocentrum have been found in fresh waters of Tasmania, Australia (Croome and Tyler, 1987). Heavily thecate species with two large plates are placed in Prorocentrum or Exuviaella. Dodge and Bibby (1973) merged the two genera into Prorocentrum; McLachlan et al., (1997) reinstated Exuviaella. Grzebyk et al., (1998), using 18S rDNA sequences and morphological differences, allowed that there was sufficient heterogeneity to support separation of Prorocentrum species at the genus level.

VI. GUIDE TO LITERATURE FOR SPECIES IDENTIFICATION There are a few keys to species within genera; Lefèvre (1932) for Peridinium and Christen (1961) for Katodinium are two. Compendia should be consulted for the dinoflagellate species within each genus. Local floras include illustrations and may contain keys to species found in that area. Where appropriate I have included citations to pertinent literature under the genus description.

A. Compendia Lefèvre, M. 1932. Monographie des espèces d’eau douce du genre Peridinium. Archivio Botanico 2:1–208. Mém. No. 5. Need to check for current genus, as many species of Peridinium have been transferred. Good illustrations and comprehensive. Popovsky, J., Pfiester, L. A. 1990. Süßwasserflora von Mitteleuropa, Band 6: Dinophyceae (Dinoflagellida). Gustav Fischer Verlag, Jena, 272 pp. Many lumped species that are probably valid. Check other sources. Schiller, J. 1933/37. Dinoflagellatae (Peridineae), in: Kolkwitz, R., Ed., Rabenhorst’s KryptogamenFlora von Deutschland, Österreich und der Schweiz, 2. Aufl., 10(3):1–2. Starmach, K. 1974. Cryptophyceae, Dinophyceae, Raphidophyceae. Flora Slodkowodna Polski 4:1–520. Panstwowe Wydawnictwo Naukowe. Warszwa, Kraków. Need to check for current genera names, since some taxa have been transferred. Good illustrations and comprehensive.

B. Local Floras Forest, H. S. 1954. Handbook of algae with special reference to Tennessee and the southeastern United States. The University of Tennessee Press, Knoxville, TN. Prescott, G.W. 1951. Algae of the western Great Lakes area exclusive of desmids and diatoms. Cranbrook Institute of Science Bulletin 31:1–946. Taft, C. E., Taft, C. W. 1971. The algae of western Lake Erie. Bulletin of the Ohio Biological Survey 4:1–189.

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Tiffany, L. H., Britton, M. E. 1952. The algae of Illinois. University of Chicago Press, Chicago, 407 pp. Wailes, G. H. 1934. Freshwater dinoflagellates of North America. Museum and Art Notes, Vancouver City Museum 7, (Supp. II):1–10, 4 plates. Whitford, L. A., Schumacher G. J. 1984. A manual of fresh-water algae. Sparks Press, Raleigh, NC.

ACKNOWLEDGMENTS I would like to acknowledge my dinoflagellate mentors, Elenor Cox and Lois Pfiester, friends and family who have been encouraging, Thomas Bermudez for the original drawing 6Ca, Victor W. Fazio, III, for the original drawing 3Ba, the reviewers and editors of the manuscript, the EM center and Dan Schwab at Bowling Green State University (OH), and the librarians at Heidelberg College for cheerful, extensive processing of interlibrary loan requests. Special thanks to Vic Fazio and Dale Ritter for patient reading of the manuscript.

LITERATURE CITED Balech, E. 1974. El genero Protoperidinium Bergh, 1881 (Peridinium Ehrenberg, 1831, partim). Revista del Museo Argentino de Ciencias Naturales “Bernardino Rivadavia” (Buenos Aires) IV, No. 1. Balech, E. 1980. On thecal morphology of dinoflagellates with special emphasis on circular and sulcal plates. Anales del Instituto Ciencias del Mar y Limnologia, Universidad Nacional Autonoma de Mexico 7:57–68. Battey, J. F. 1992. Carbon metabolism in zooxanthellae-coelenterate symbioses, in: Reisser, W., Ed., Algae and symbioses. Biopress Limited, Bristol, England, pp. 153–187. Blank, R. J. 1992. Taxonomy of Symbiodinium—the microalgae most frequently found in symbiosis with marine invertebrates, in: Reisser, W., Ed., Algae and symbioses. Biopress Limited, Bristol, England, pp. 189–197. Boltovskoy, A. 1975. Estructura y estereoultraestructura tecal de dinoflagelados. II. Peridinium cinctum (Müller) Ehrenberg. Physis Seccion B Los Aguas Continentales y Sus Organismos 34:73–84. Boltovskoy, A. 1976. Estructura y estereoultraestructura tecal de dinoflagelados. III. Peridinium bipes Stein forma apoda, n.f. Physis Seccion B Los Aguas Continentales y Sus Organismos 35:147–155. Boltovskoy, A. 1989. Thecal morphology of the dinoflagellate Peridinium gutwinskii. Nova Hedwigia 49:369–380. Bourrelly, P. 1968. Notes sur les Péridiniens d’eau douce. Protistologica 4:5–16. Bourrelly, P. 1970. Les Algues d’Eau Douce. Initiation à la Systématique. Tome III, Les algues bleues et rouges: Les Eugléniens, Peridiniens et Cryptomonadines. Éditions N. Boubée & Cie, Paris, 512pp. Bruno, S. F., McLaughlin, J. J. A. 1977. The nutrition of the freshwater dinoflagellate Ceratium hirundinella. Journal of Protozoology 24:548–553. Bucka, H., Zurek, R. 1992. Trophic relations between phyto- and zooplankton in a field experiment in the aspect of the formation

and decline of water blooms. Acta Hydrobiologica 34:139–155. Buckland-Nicks, J., Reimchen, T. E. 1995. A novel association between an endemic stickleback and a parasitic dinoflagellate. 3. Details of the life cycle. Archiv für Protistenkunde 145:165–175. Buckland-Nicks, J., Reimchen, T. E., Garbary, D. J. 1997. Haidadinium ichthyophilum gen. nov. et sp.nov. (Phytodiniales, Dinophyceae), a freshwater ectoparasite on stickleback (Gasterosteus aculeatus) from the Queen Charlotte Islands, Canada. Canadian Journal of Botany 75:1936–1940. Bujak, J. P., Davies, E. H. 1983. Modern and fossil Peridiniineae. AASP Contribution Series Number 13. American Association of Stratigraphic Palynologists, Dallas, 203pp. Bujak, J. P., Williams, G. L. 1981. The evolution of dinoflagellates. Canadian Journal of Botany 59:2077–2087. Burkholder, J. M. 1998. Implications of harmful microalgae and heterotrophic dinoflagellates in management of sustainable marine fisheries. Ecological Applications 8(Suppl.): S37–S62. Burkholder, J. M., Noga, E. J., Hobbs, C. H., Glasgow, H. B. 1992. New “phantom” dinoflagellate is the causative agent of major estuarine fish kills. Nature 358:407–410. Bursa, A. S. 1969. Actiniscus canadensis n. sp., A. pentasterias Ehrenberg v. arcticus n. var., Pseudoactiniscus apentasterias n. gen., n. sp., marine relicts in Canadian arctic lakes. Journal of Protozoology 16:411–418. Bursa, A. S. 1970. Dinamoebidium coloradense spec. nov. and Katodinium auratum spec. nov. in Como Creek, Boulder County, Colorado. Arctic and Alpine Research 2:145–151. Calado, A. J., Moestrup, Ø. 1997. Feeding in Peridiniopsis berolinensis (Dinophyceae): New observations on tube feeding by an omnivorous, heterotrophic dinoflagellate. Phycologia 36:47–59. Carty, S. 1986. The taxonomy and systematics of freshwater armored dinoflagellates. Ph.D. dissertation, Texas A&M University, 286pp. Carty, S. 1989. Thompsodinium and two species of Peridiniopsis (Dinophyceae): taxonomic notes based on scanning electron micrographs. Transactions of the American Microscopical Society 108:64–73. Carty, S. 1993. Contribution to the dinoflagellate flora of Ohio. Ohio Journal of Science 93:140–6. Carty, S., Cox, E. R. 1985. Observations on Lophodinium polylophum (Dinophyceae). Journal of Phycology 21:396–401. Carty, S., Cox, E. R. 1986. Kansodinium gen. nov. and Durinskia gen. nov.: Two genera of freshwater dinoflagellates (Pyrrhophyta). Phycologia 25:197–204. Carty, S., Fazio, V. W., III. 1997. //aves.net/algaeweb/ppolncum.htm. Chapman, A. D., Pfiester, L. A. 1995. The effects of temperature, irradiaance, and nitrogen on the encystment and growth of the freshwater dinoflagellates Peridinium cinctum and P. willei in culture (Dinophyceae). Journal of Phycology 31:355–359. Chesnick, J. M., Cox, E. R. 1987. Synchronized sexuality of an algal symbiont and its dinoflagellate host, Peridinium balticum (Levander) Lemmermann. BioSystems 21:69–78. Chesnick, J. M., Cox, E. R. 1989. Fertilization and zygote development in the binucleate dinoflagellate Peridinium balticum (Pyrrhophyta). American Journal of Botany 76:1060–1072. Chesnick, J. M., Morden, C. W., Schmieg, A. M. 1996. Identity of the endosymbiont of Peridinium foliaceum (Pyrrhophyta): Analysis of the rbcLS operon. Journal of Phycology 32:850–857. Christen, Von H. R. 1961. Über die Gattung Katodinium Fott (= Massartia Conrad). Schweizerische Zeitschrit für Hydrologie 23:309–341. Cranwell, P. A., Robinson, N., Eglinton, G. 1985. Esterified lipids of the freshwater dinoflagellate Peridinium lomnickii. Lipids 20:645–651.

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21

CRYPTOMONADS Paul Kugrens

Brec L. Clay*

Department of Biology Colorado State University Fort Collins, Colorado 80523

UTEX Culture Collection University of Texas Austin, Texas 78713

I. Introduction II. Unique Features of Cryptomonads A. External Cell Architecture B. Periplast Structure C. Flagella and Flagellar Apparatus D. Ejectisomes E. Ejectisome Digestion Vesicles F. Mitochondria and Chloroplasts G. Nucleomorphs H. Reproduction I. Nucleus and Mitosis J. Contractile Vavuoles K. Starch III. Origin of Cryptomonads IV. Ecology A. Abiotic Factors B. Biological Factors C. Cryptomonad Endosymbiotins and Pathogens E. Types of Nutrition—Carbon Sources V. Collection, Preparation for Isolation, and Culturing

I. INTRODUCTION Cryptomonads, cryptoprotists, or cryptophytes, as these algae are commonly called, are unicellular, biflagellate protists. They are variously classified as belonging to the phylum (division) Cryptophyta, class Cryptophyceae, order Cryptomonadales, or phylum Cryptista sensu Cavalier-Smith (1986). Cryptomonads are important primary producers in freshwater and marine habitats (Gillott, 1990; Klaveness, 1988a, b), and many are cosmopolitan in their distribution, although they appear to be more common in cooler water. Many

VI. Classification, Key, and Descriptions A. Introduction B. General Features Useful in Determining Genera and Species C. Classification of the Phylum Cryptophyta D. Key E. Freshwater Cryptomonad Genera and Species F. Guide to Literature for Species Identification VII. Availability of Cryptomonads VIII. Family Kathablepharidaceae A. Ecology B. Cell Structure C. Classification of Kathablepharidaceae D. Isolation and Culturing Techniques for Kathablepharids Literature Cited

are collected in phytoplankton samples, but their cells are extremely delicate and rupture when fixatives are added or when temperatures are elevated. As a consequence, their numbers generally are low in preserved samples and therefore are assumed to be a small and obscure taxonomic group of protists. To the contrary, they often assume dominant phytoplankton status in temperate lakes and reservoirs, where they may dominate the under-ice, early spring and late fall populations. In fact, the variations in cell structure discovered with specialized electron microscopic techniques (Hill, 1991a, b; Hill and Wetherbee, 1986, 1988, 1989;

* Present address: CH Diagnostic and Consulting Service, Loveland, Colorado 80538. Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

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Kugrens and Lee, 1986, 1991; Kugrens et al., 1986, 1987; Lee and Kugrens, 1986; Clay and Kugrens, 1999a–c; Clay et al., 1999) strongly indicate that there are numerous unrecognized freshwater genera and species (Andersen, 1992). While cryptomonads represent a well-circumscribed group of algae, there is another group of colorless flagellates that historically has been included in the Cryptophyta. This group, the katablepharids, includes the genera Leucocryptos and Kathablepharis, which are common in both freshwater and marine habitats. Except for the presence of ejectisomes and placement of flagella, their other cellular features do not support the placement of this group within the cryptomonads (Lee and Kugrens, 1991b; Lee et al., 1991; Vørs, 1992a, b; Clay and Kugrens, 1999a, b). Currently their systematic/phylogenetic position remains undetermined, but, for convenience, we have retained this group in the Cryptophyta, realizing that they probably are not cryptomonads. For clarity in presentation, the discussion and description of cryptomonad characteristics are presented in the first portion of this chapter (Figs. 1–16), and a separate description of kathablepharids (Figs. 17 and 18) follows the cryptomonad description.

II. UNIQUE FEATURES OF CRYPTOMONADS A. External Cell Architecture The external cell architecture is influenced primarily by a furrow/gullet complex (Figs. 1, 7A–C, 8A, 11A–C, 12A–C, 16A, C) which tends to impart an asymmetrical shape to the cells. In addition, cell shapes may be oval, compressed, lunulate, caudate, acute, elongate, sigmoid, or otherwise contorted. All cells possess an anterior, outwardly facing depression called a vestibulum, from which the flagella originate on the right side. In freshwater crytpomonads the contractile vacuole is located in the anterior end of the cell and usually discharges through a predetermined site in the dorsal portion of the vestibulum (Kugrens et al., 1986). In some genera, such as Campylomonas, a vestibular ligule, which is a small, flat extension of the cell, covers the discharge site of the contractile vacuole (Hill, 1991c; Kugrens and Lee, 1991; Kugrens et al., 1986). A gullet, some type of furrow, or a combination of a furrow-gullet (Fig. 7A–C) is one of the primary diagnostic features for the genera. A furrow is a ventral groove, of variable length, that begins in the vestibular region of the cell and extends posteriorly, terminating somewhere in the anterior half of the cell (Hill and Wetherbee, 1986, 1988, 1989; Klaveness, 1985;

Kugrens et al., 1986; Munawar and Bistricki, 1979). A tubular invagination called the gullet may extend posteriorly from the vestibulum or from the end of the furrow (Munawar and Bistricki, 1979; Hill and Wetherbee, 1986, 1988, 1989; Kugrens et al., 1986). Several types of furrows have been described in cryptomonads (Munawar and Bistricki, 1979; Klaveness, 1985; Kugrens et al., 1986) and these variations have become major features in cryptomonad systematics (Hill and Wetherbee, 1986, 1988, 1989; Hill, 1990, 1991b; Clay et al., 1999). There are at least five variations in the furrow/gullet complex (Kugrens et al., 1986). Furthermore, a gullet may have evolved from the fusion of a furrow (Kugrens and Lee, 1991; Clay et al., 1999). Consequently, many genera have a combination of a furrow and gullet, and these may represent intermediate stages in the evolution of a gullet. Cryptomonads may have only a gullet (Figs. 11A, C, 12A–C, 13A, B), a simple furrow only (Fig. 16A, B, 8A), a simple furrow and gullet, or a complex furrow structure with or without a gullet (Fig. 16A). The gullet and furrow are lined with large ejectisomes, and ejectisome discharge in this area would not destroy cell organelles by this action. Specialized furrow plates may be associated with each type of furrow, thereby providing additional variations to the furrow structure. Furrow plates are of two types, scalariform (Fig. 1) and fibrillar. A scalariform furrow plate is in the form of a ladder that has sides connected by lateral, crystalline “rungs.” Fibrillar furrow plates are made up of microfibrils that are oriented parallel to each other and occur as a thin plate along one side of the furrow. Surprisingly little is known about the actual cell architecture in cryptomonads. Prior to Hill’s studies, as well as our own, we relied on light microscopic descriptions for this aspect (Skuja, 1948; HuberPestalozzi, 1950; Butcher, 1967; Bourelly, 1970) because most electron microscopic preparatory procedures tend to distort the cell shapes. Parducz’s (1967) fixation preserves cell shape well; however, freeze drying is the best method for examining the external features and determining whether a gullet and/or furrow is/are present. In fact, the morphology of cryptomonad cells has been drastically revised as a result of studies using cryofixation techniques. The existence of tubular gullets, furrows, and furrow/gullet combinations and their variations has proved to be a significant delineator of genera and should serve to enhance light microscopic identifications. Unfortunately, facilities that utilize electron microscopic cryotechniques are fewer, and it is becoming increasingly difficult to find a facility that still employs these techniques.

21. Cryptomonads

Ventral Flagellum Dorsal Flagellum Contractile Vacuole Tubular Hair Rhizostyle Periplastidial Space

Furrow/Gullet

Furrow Plate

Large Ejectisome Starch Inner Periplast Plate Nucleomorph Chloroplast

Chloroplast Endoplasmic Reticulum Small Ejectisome

FIGURE 1 Diagram of a generalized cryptomonad cell, showing the cellular details described in the text.

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External Face

Internal Face

Fiber Scale

Integral Membrane Protein Ejectisome

Cell Membrane

Periplast Plates of Internal Periplast Component (IPC)

A

External Face

Internal Face Fiber

Scale

Integral Membrane Protein Periplast Sheet of Internal Periplast Component (IPC)

Cell Membrane

Ejectisome

Pore

B FIGURE 2 Diagrams of two major periplast types and their association with the plasma membrane and ejectisomes. (A) Periplast structure in cells where the inner periplast component is composed of plates. The plates are attached to the plasma membrane by transmembrane particles. (B) Periplast structure in cells having an inner periplast component consisting of a single sheet. The sheet is not associated with the plasma membrane.

21. Cryptomonads

Vestibulum Contractile Vacuole Dorsal Flagellum Ventral Flagellum

A Furrow Fold Chloroplast Stoma Pyrenoid Ejectisome Nucleus

Side View

C

Ventral View

D

B

Oval Periplast Plate

E

F

FIGURE 3 Light micrographic illustrations of Goniomonas and Cryptomonas spp. (A) Goniomonas truncata, showing the flattened nature of cells, the dorsal nucleus, and the dorsally inserted flagella. Vertical striations shown in the diagrams are visible with a light microscope. (B) Cryptomonas ovata from various views. (C) Cryptomonas obovata. (D) Cryptomonas erosa. (E) Cryptomonas ovolinii. Scale bars = 10 µm.

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FIGURE 4 Light micrographs of Goniomonas and Cryptomonas species. (A) Goniomonas truncata cell, showing the location of the nucleus (N) and ejectisomes (E). (B) G. truncata cell, showing surface striations (arrows). (C) Cryptomonas obovata cells, with a pyrenoid visible in some cells. (D) Cryptomonas ovata cells, with two pyrenoids (arrows) in each cell. Flagella (F) and ejectisomes (E) are also evident. (E) Cryptomonas ovata cell, showing ejectisomes (E) around furrow, with flagella and a nucleus (N) also visible. (F) Cryptomonas phaseolus cell. (G) Cryptomonas tetrapyrenoidosa, showing four pyrenoids (arrows). (H) Cryptomonas erosa with the furrow flanked by ejectisomes (E), a portion of a flagellum (F), and the location of the vestibulum. Scale bars = 10 µm.

21. Cryptomonads

Vestibulum Vestibular Ligule Contractile Vacuole

Ventral View

Flagellum Ejectisome Furrow Gullet

Side view

Nucleus Pyrenoid

A

Inner Periplast Sheet

B

Side view

Ventral View

F

C

D

E

FIGURE 5 Diagrams of genera belonging to the family Campylomonadaceae. (A) Campylomonas reflexa. (B) Cryptomonas (Campylomonas) rostratiformis. (C) Cryptomonas (Campylomonas marssonii. (D) Cryptomonas (Campylomonas) platyuris. (E) Chilomonas paramecium. (F) Chilomonas acuta. Scale bars = 10 µm.

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FIGURE 6 Light micrographs (differential interference contrast) of several species from the Campylomonadaceae and Pyrenomonadaceae. (A) Campylomonas rostratiformis cell, with its characteristic rhinote anterior and numerous ejectisomes (E) lining the furrow. (B) Somewhat flattened cell of Cryptomonas (Campylomonas) rostratiformis, showing multiple pyrenoids (arrows). (C) Cryptomonas (Campylomonas) platyuris in ventral view, showing the broad shape of the cell in this view and the ejectisomes (E) lining the furrow. The cell is filled with considerable starch. (D) Chilomonas paramecium cell, with starch filling most of the cell. (E) Pyrenomonas ovalis cell with ejectisomes (E). (F) Pyrenomonas ovalis with the characteristic prominent pyrenoid (P). (G) Higher magnification of a P. ovalis cell, showing periplast plates (arrows). (H) Storeatula rhinosa cell with a prominent pyrenoid (P). (I) S. rhinosa cell as viewed obliquely, showing the asymmetrical cell shape and a prominent pyrenoid (P). (J) Storeatula sp. cell with an extensive ejectisome region and a prominent pyrenoid (P). Portions of the flagella are visible and are adhering to the cell. Scale bars = 10 µm.

21. Cryptomonads

FIGURE 7 Scanning electron micrographs of Cryptomonas and Campylomonas species. (A) Cell of Cryptomonas tetrapyrenoidosa with long (LF) and short (SF) flagella inserted on the right side of the vestibulum (V). A long furrow extends from the vestibulum, and a stoma (S) is present. (B) Cell of Campylomonas rostratiformis with long (LF) and short (SF) flagella inserted on the right side of the vestibulum. A vestibular ligule (vl) is seen attached to the dorsal wall of the vestibulum. A slightly curved, oblique furrow (F) runs for approximately one-third of the cell length. Note the rostrate anterior of the cell. (C) Oblique view of a cell of Campylomonas reflexa, showing the long (LF) and short (SF) flagella inserted on the right side of the vestibulum. A vestibular ligule (vl) attaches to the dorsal wall of the vestibulum. A furrow (F) extends posteriorly for a third of the cell length. Note the reflexed tail (arrow). (D) Lateral view of C. reflexa, showing the reflexed shape of the cell. A and B are from Kugrens et al. (1986), with permission.

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FIGURE 8 Scanning electron micrographs Campylomonas platyuris. (A) Ventral view with long (LF) and short (SF) flagella inserted in the vestibulum (V). A vestibular ligule (vl) occurs on the dorsal side of the vestibulum. An oblique furrow (F) runs for almost one-half of the cell length from the vestibulum. (B) Dorsal view of C. platyuris. Note the slightly reflexed cell shape. A is from Kugrens et al. (1986) with permission.

Dorsal Flagellum Contractile Vacuole

Ventral Flagellum Vestibulum Furrow Gullet Ejectisome

Pyrenoid

Chloroplast

Inner Periplast Sheet

Inner Periplast Plates

A

B

FIGURE 9 Light microscopic illustrations of red-colored cryptomonad species. (A) Pyrenomonas ovalis. (B) Storeatula rhinosa. Scale bars = 10 µm.

21. Cryptomonads

FIGURE 10 Scanning electron micrographs of red-colored cryptomonad species. (A) Ventral view of Rhodomonas ovalis, showing long (LF) and short (SF) flagella inserted on the right side of the vestibulum (V). A short furrow (F) and vestibulum are shown near the anterior, ventral surface. Note the absence of distinct plates at the cell posterior (arrow). (B) Lateral view of R. ovalis. (C) Oblique view of a cell of Storeatula rhinosa, showing long (LF) and short (SF) flagella inserted on the right side of the vestibulum (V). (D) Lateral view of S. rhinosa, showing the narrower elongate shape of the cell. C and D from Kugrens et al. (1999), with permission.

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Flagellum Pyrenoid Starch

Contractile Vacuole Vestibulum Ejectisome Gullet Nucleus

Hexagonal Periplast Plates

Chloroplast

A

C

B

D Starch

Vestibulum

Nucleus Ventral Views of Cells

E

F

FIGURE 11 Light microscopic illustrations of blue-green cryptomonad species. (A) Komma caudata. (B) Chroomonas oblonga. (C) Chroomonas coerulea. (D) Chroomonas nordstedtii. (E) Chroomonas pochmanni. (F) Hemiselmis amylosa. Scale bars = 10 µm. Scanning electron micrographs of blue-green cryptomonad species. (A) Cell of Chroomonas coerulea, showing a short flagellum (SF) and a long flagellum (LF) inserted subapically in the right side of the vestibulum (V). (B) Cell of Chroomonas oblonga, showing a short flagellum (SF) and a long flagellum (LF) inserted subapically in the right side of the vestibulum (V). (C) Ventral view of a cell of Chroomonas sp., showing a short flagellum (SF) and a long flagellum (LF) inserted subapically in the right side of the vestibulum (V). Scale bars = 10 µm.

21. Cryptomonads

FIGURE 12 Light micrographs of blue-green cryptomonads species. (A) Komma caudata cell showing the typical shape. (B) Komma caudata cell with a dorsal pyrenoid (P) and a nucleolus located in the nucleus (N). (C) Chroomonas pochmanni cell, displaying the typical shape and a pyrenoid (P) and ejectisomes (E). D) Chroomonas coerulea cells with some stigmas (S) and pyrenoids (P) visible. (E) Slightly flattened C. coerulea cells, showing the stigma (S) associated with the pyrenoid (P). (F) Chroomonas nordstedtii cell displaying the typical cell shape. A pyrenoid (P) and starch grains are present in the cell. (G) Hemiselmis amylosa illustrating the typical bean-shaped cell in lateral view. A flagellum (F) and pyrenoid (P) are evident. (H) Hemislemis amylosa cell showing two flagella arising from a slight depression near the middle of the cell. (I) Slightly larger Hemiselmis amylosa cell with a pyrenoid (P). Scale bars = 10 µm, unless otherwise indicated.

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FIGURE 13 Scanning electron micrographs of blue-green cryptomonad. (A) Ventral view of a cell of Komma caudata, showing a short flagellum (SF) and a long flagellum (LF) inserted subapically in the right side of the vestibulum (V). Note the acuminate tail. (B) Lateral view of Komma caudata. Plate elevations are visible. C. Ventral view of a cell of K. pochmanni, showing a short flagellum (SF) and a long flagellum (LF) inserted subapically in the right side of the vestibulum (V). (D) Lateral view of K. pochmanni. C is from Kugrens and Lee (1991), with permission.

21. Cryptomonads

FIGURE 14 Scanning electron micrographs of blue-green cryptomonads. (A) Cell of Chroomonas coerulea, showing a short flagellum (SF) and a long flagellum (LF) inserted subapically in the right side of the vestibulum (V). Rectangular surface plates are visible. (B) Cell of Chroomonas oblonga, showing rectangular surface palted, and the short (SF) and long (LF) flagellar insertion. (C) Lateral view of Hemiselmis amylosa cell with long flagella (LF) and short flagella (SF) inserted in the vestibulum (V), approximately one-third of the distance down from the cell apex. (D) Ventral view of P. amylosa. showing the location of the vestibulum (V). A and B after Kugrens et al. (1986), and C and D are from Clay and Kugrens (1999b), with permission.

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FIGURE 15 Scanning electron micrographs of Plagioselmis and Cryptomonas ovata. (A) Cell of Plagioselmis nanoplanctica, showing the long (LF) and short (SF) flagella inserted on the right side of the vestibulum (V). A furrow (F) extends from the vestibulum for approximately half the length of the cell. Note the midventral band (mvb) on the cell posterior. (B) Slightly lateral view of P. nanoplanctica to show the cell shape. (C) Cell of Cryptomonas ovata with long (LF) and short (SF) flagella inserted on the right side of the vestibulum (V). A long furrow extends from the vestibulum. Note the stoma (S). (D) Dorsal view of Cryptomonas ovata.

21. Cryptomonads

Contractile Vacuole Furrow Pyrenoid Hexagonal Periplast Plate

A

B Anterior Flagellum Cell Covering Cytostomal Ring Pharyngeal Microtubule Food Vacuole Digested Chloroplast Mitochondrion Ejectisomes Vesicle Nucleus Nucleolus Posterior Flagellum

C

FIGURE 16 Light microscopic illustrations of Plagioselmis and Kathablepharis species. (A) Diagram of Plagioselmis nanoplanctica. (B) Kathablepharis ovalis, showing its general features. (C) Diagram of a Kathablepharis phoenikoston cell, as interpreted from electron microscopic data.

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FIGURE 17 Light micrographs of Plagioselmis and Kathablepharis. (A) Lateral view of Plagioselmis nanoplanctica, showing the typical comma shape of the cell. The pyrenoid is dorsal (arrow). (B) Ventral view of Plagioselmis nanoplanctica, showing that this cell is broad in this view. The pyrenoid is on the dorsal side of the cell (arrow). (C) Kathablepharis ovalis with ingested food in an enlarged food vacuole. A row of ejectisomes (E) extends posteriorly from the site of flagellar insertion. The nucleus (N) is in the posterior of the cell. (D) Kathablepharis phoenikoston cell with the nucleus in the posterior. Globular contents probably represent ingested food. Scale bars = 10 µm.

B. Periplast Structure (Fig. 2) Periplasts are special cell coverings found only in cryptomonads, although the same term is applied to euglenoids, which have a different arrangement. In cryptomonads periplasts consist of inner and surface components (Hibberd et al., 1971; Hill and Wetherbee, 1986, 1988, 1989; Kugrens et al., 1987; Kugrens and Lee, 1991; Wetherbee et al., 1986, 1987; Hill, 1990, 1991b; Clay and Kugrens, 1999a–c) with the plasma membrane located between the two components. Both components are variable in their structure, depending on the genus. The inner periplast component (IPC) represents the protein component beneath the plasma membrane and is of two general types. One type consists of multiple plates of various shapes (Hibberd et al., 1971; Hill and Wetherbee, 1986, 1988, 1989; Kugrens and Lee, 1986; Hill, 1990, 1991b), and a second type consists of a single sheet (Grim and Staehelin, 1984; Kugrens and Lee, 1986; Hill, 1991b). The plates of the inner periplast component in the first type are connected to the cell membrane by intramembrane particles or proteins (Brett and Wetherbee, 1986; Hill and Wetherbee, 1986, 1988, 1989; Kugrens and Lee, 1986, 1991; Wetherbee et al., 1986, 1987; Clay et al., 1999). The arrangement

of these IMP domains conforms to the plate shapes (Brett and Wetherbee, 1986; Kugrens and Lee, 1986, 1991; Wetherbee et al., 1987; Clay et al., 1999). Variations in the shapes of these plates are major features in establishing genera. Multiple-plated periplasts may have hexagonal, square, oval/round, rectangular, or irregularly shaped plates making up the inner and/or surface components. In genera with a periplast sheet, which is a continuous internal protein cover, there are numerous closely spaced pores through which the ejectisome membranes penetrate to dock with the cell membrane (Grim and Staehelin, 1984; Kugrens et al., 1994). The periplast sheet itself is not connected to the plasma membrane. The surface periplast component (SPC) does not have a sheetlike variant, but consists of plates, heptagonal scales, mucilage, or a combination of any of these. It appears that both types of inner periplast components, and some of the surface plates, are composed of protein (Gantt, 1971; Faust, 1974). Since plate shapes are used for generic designations, it is critical that the plate shapes be determined accurately. Oakley and Santore (1982), Gantt (1971), and Faust (1974) suggested that periplast shapes may undergo conformational changes. However, plates do not undergo shape changes unless they are subjected

21. Cryptomonads

FIGURE 18 Scanning electron micrographs of colorless cryptomonad and kathablepharid species. (A) Cell of Goniomonas truncata with both flagella (F) inserted on the dorsal side of the vestibulum (V). Note some unilateral spikes (sp) remaining on the left flagellum. A ventral furrow (F) continues from the vestibulum and features a persistent opening termed the stoma (S). A second tubular invagination termed the infundibulum (I) is present on the left side of the cell. (B) Cell of Chilomonas paramecium showing two subapically inserted flagella (F) on the right side of the vestibulum (V). Note the vestibular ligule (vl) attached to the dorsal side of the vestibulum. A short furrow (F) extends from the vestibulum. (C) Cell of Kathablepharis ovalis, showing two subapically inserted flagella (F) arising from a flagellar mound (fm). A cytostome (C) is located between the flagellar mound and the cell apex. (D) Cell of Kathablepharis phoenikoston, showing an anteriorly directed flagellum (AF) and a trailing flagellum (TF), both inserted subapically. A and B are from Kugrens and Lee (1991), C is from Lee and Kugrens (1991), and D is from Clay and Kugrens (1999a), with permission.

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to drastic treatments, such as desiccation, fixation, or excessive centrifugation. Any technique other than quick freezing may create artifacts, particularly in those periplasts having circular or oval plates, the shapes of which may be modified by pressures from adjacent plates. For example, in instances where the plates are approximately the same size, each plate is surrounded by six others (1 by 6 arrangement). When the plates are forced against each other, a hexagonal pattern may result (Kugrens et al., 1987). Some studies attempted to use either light (Novarino, 1993a,b) or scanning electron microscopy (Santore, 1977; Novarino, 1991a,b; Novarino and Lucas, 1993; Novarino et al., 1994) to determine plate shapes, but with a few exceptions (Munawar and Bistricki, 1979; Klaveness, 1985; Kugrens et al., 1986; Hill, 1990), scanning electron microscopy is inadequate for studying subsurface components. If possible, quickfreezing freeze-fracture procedures should be used to study periplast plate shapes. With this technique cells are quick-frozen (“slammed”) without pretreatment or fixation (Boyne, 1979; Chandler, 1984; Phillips and Boyne, 1984). Consequently, the periplast shapes in cryptomonads and the intimate association that exists between the plates and the plasma membrane can be examined accurately.

C. Flagella and Flagellar Apparatus (Fig. 1) With the exception of Goniomonas, where the flagella are inserted on the dorsal side of the vestibulum, the flagella of cryptomonad cells are inserted subapically on the right side of the cell. The two flagella are subequal in length and consist of a dorsal and ventral insertion. Bipartite tubular hairs occur on at least one of the flagella (Kugrens et al., 1987; Clay et al., 1999), and there appear to be at least five variations in the arrangement of tubular and nontubular hairs on the flagella (Kugrens et al., 1987). Most of these features can be seen only with EM. The most common arrangement of hairs on the flagella is one in which the longer or dorsal flagellum bears two laterally opposed rows of tubular hairs and the shorter or ventral flagellum bears a single row of hairs. Tubular hairs on the dorsal flagellum have one solid extension called a terminal filament, whereas the tubular hairs of the ventral flagellum have two unequal terminal filaments. In addition, flagella may bear heptagonal scales (Pennick, 1981; Lee and Kugrens, 1986). Instead of tubular hairs, Goniomonas has a unilateral row of curved spikes on one of its flagella and fine, nontubular hairs on both flagella (Kugrens et al., 1987; Kugrens and Lee, 1991). The flagellar transition region is unique and con-

sists of a doublet system of septa in all cryptomonads (Grain et al., 1988; Kugrens and Lee, 1991). A rhizostyle is an integral component of the flagellar apparatus in most cryptomonads, and it consists of microtubules that originate near the basal bodies and then extend posteriorly into the cell. One type of rhizostyle, found in Chilomonas (Roberts et al., 1981; Kugrens and Lee, 1991), Hanusia phi (Gillot and Gibbs, 1983; Gillott, 1990; Deane et al., 1998) Teleaulax (Hill, 1991b), Storeatula (Hill, 1991c), Geminigera (Hill, 1991c) and Proteomonas (Hill and Wetherbee, 1986), passes close to the nucleus and terminates near the posterior end of the cell. Each microtubule has a wing-like extension (lamella), and the lengths of these lamellae may vary with the respective microtubule (Gillot and Gibbs, 1983). A second type of rhizostyle, reported for Cryptomonas ovata (Roberts, 1984; Hill, 1990) and Cryptomonas theta (= Guillardia theta) (Gillot and Gibbs, 1983), lacks wings on the microtubules. Usually, this rhizostyle terminates anterior to the nucleus (Roberts, 1984).

D. Ejectisomes (Fig. 2) The cryptomonad ejectisomes (formerly called trichocysts) were the first to be described (Anderson, 1962), but different types have been discovered in other organisms, and these are noted later. Ejectisomes are the extrusive organelles of all cryptomonads, and they appear to be identical in all genera that have been investigated (Kugrens et al., 1994). Cryptomonad cells contain two sizes of ejectisomes (Schuster, 1970; Kugrens et al., 1994; Clay et al., 1999). Large ejectisomes are located near the gullet/furrow complex, whereas small ejectisomes occur elsewhere in the peripheral cytoplasm. Both types of ejectisomes consist of two unequal sized components that are joined together and enclosed by a membrane (Kugrens et al., 1994). Each of these components has a tightly wound, tapered ribbon. The widest part of the tape is toward the outside of the ribbon. The smaller ribbon generally faces toward the outside of the larger ribbon. The tape has a crystalline substructure (Morrall and Greenwood, 1980; Grim and Staehelin, 1984; Kugrens et al., 1994). The ejectisomes discharge when the organism is irritated. Discharged ejectisomes form a long tube, with the short portion oriented at a slight angle to the long tube. The tube is formed because of a spiral rolling of the ribbon (Kugrens et al., 1994). The ejectisome ribbons are formed in Golgiderived vesicles, where they are tightly wound. Initially, the tape consists of a few turns, but the number increases as material is added. As the ejectisomes mature and enlarge, they are transported to the periph-

21. Cryptomonads

ery of the cell. The ejectisome membranes contain intramembrane particles (IMPs) arranged in a rosette configuration (Grim and Staehelin, 1984), and these attach the ejectisome membrane to the cell membrane in a process called docking. In those species with a single periplast sheet, such as Chilomonas, there are numerous pores within the sheet. The ejectisomes are anchored to the sides of these pores (Grim and Staehelin, 1984). Some Pyramimonas species (chlorophytes) and the colorless Kathablepharis (Lee and Kugrens, 1991; Kugrens et al., 1994; Clay and Kugrens, 1999a,b) and Leucocryptos (Vørs, 1992a, b) also possess ejectisomes; however, they differ in structure from cryptomonad ejectisomes. In all three genera, only the large ribbon makes up the ejectisome. After release all form a tube, just as in cryptomonads, but the small component is lacking. Discharge of ejectisomes is forceful and rapid, propelling the organism in the direction opposite the discharge. Thus the ejectisome could function as an escape mechanism to avoid predators, or it could be a defense mechanism to inflict damage on a potential predator (Kugrens et al., 1994).

E. Ejectisome Digestion Vesicles Ultrastructural evidence indicates that some vesicles in cryptomonads are specialized for ejectisome autolysis (Kugrens et al., 1994). These vesicles originate from the fusion of several ejectisome chambers and continue to enlarge by the fusion of additional ejectisome membranes. Individual ejectisomes disaggregate within these vesicles. In older vesicles, components of expanded ejectisomes make up most of the contents. In later stages most of the tubular, expanded components of ejectisomes are no longer recognizable, and the contents appear fibrillar or granular. The vesicle sizes are larger in cells from older cultures, and there may be several vesicles per cell. Golgi vesicles have been observed to fuse with the existing vesicles, perhaps to add lytic enzymes. The vesicles apparently represent specific repositories for defective or surplus ejectisomes; thus they represent another unique component of cryptomonad cells. These may be the refractive vesicles that are frequently seen with the light microscope and formerly may have been referred to as the Corps de Maupas (Lucas, 1970b).

F. Mitochondria and Chloroplasts (Fig. 1) A single reticulate mitochondrion with flattened cristae (Santore and Greenwood, 1977; Roberts et al., 1981; Kugrens and Lee, 1991) apparently occurs in cells of all cryptomonads.

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Chloroplasts may be olive green, brownish, bluegreen, or red, depending on the pigments present. Pigments consist of chlorophylls a and c2, alpha and beta carotene, alloxanthin, diadinoxanthin, and several forms of blue and red phycobiliproteins called Cr-phycocyanin and Cr-phycoerythrin, to differentiate them from cyanobacterial and rhodophyte phycobiliproteins (Glazer and Appel, 1977; Hill and Rowan, 1989). With the exception of a marine endosymbiont (Hibberd, 1977), only one or two chloroplasts occur in the cells of pigmented genera (Santore, 1984, 1987; Hill, 1991a,b,c). Either Cr-phycocyanin or Cr-phycoerythrin (Hill and Rowan, 1989) is located in the intrathylakoidal lumens of the photosynthetic lamellae (Gantt et al., 1971; Faust and Gantt, 1973; Gantt, 1979, 1980; Ludwig and Gibbs, 1989). Chloroplasts are surrounded by a double membrane called the periplastidial envelope, periplastidial compartment, periplastidal complex, or chloroplast endoplasmic reticulum (CER), which originates from an evagination of the outer membrane of the nuclear envelope and surrounds the chloroplast, starch granules, and a reduced nucleus known as the nucleomorph (Gillott and Gibbs, 1980; Santore, 1982c; Ludwig and Gibbs, 1985a). Starch granules are formed within the periplastidial compartment, not in the chloroplast, and they generally are associated with a pyrenoid if present. The number of thylakoids penetrating the pyrenoid has been suggested as a possible taxonomic character (Santore, 1984). Thylakoids in the chloroplasts usually are arranged in pairs (Gantt et al., 1971; Dwarte and Vesk, 1982, 1983; Santore, 1984), sometimes in groups of three (Klaveness, 1981; Hill, 1991b), or in stacks of variable number (Hill, 1991b). Chilomonas has a reduced chloroplast which lacks pigments and is called a leucoplast (Sespenwol, 1973; Heywood, 1988; Kugrens and Lee, 1991). Goniomonas lacks plastids and a nucleomorph, consequently it also lacks the periplastidial compartment.

G. Nucleomorphs (Fig. 1) Nucleomorphs are highly reduced endosymbiont nuclei located in the periplastidial compartment (Gillott and Gibbs, 1980; McKerracher and Gibbs, 1982; Morrall and Greenwood, 1982; Santore, 1982c, 1984, 1987; Ludwig and Gibbs, 1985a; Kugrens and Lee, 1989, 1991). They represent a vestigial nucleus, which remains from an ancestral endosymbiont, possibly a red alga (Douglas et al., 1991; McFadden et al., 1997). The nucleomorph is small, it is limited by a double membrane and it contains DNA (Ludwig and Gibbs, 1985a; Douglas et al., 1991; McFadden, 1993; McFadden et al., 1997). In addition, the nucleomorph

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contains a fibrillo-granular region and dense bodies. Its location within the compartment may have systematic applications (Santore, 1984; Hill and Wetherbee, 1989), specifically in Rhodomonas and Storeatula, where the nucleomorph is located in the pyrenoidal bridge (Hill and Wetherbee, 1989; Novarino, 1991a,b).

H. Reproduction Reproduction usually occurs by mitotic divisions, although sexual cycles have been documented for Proteomonas (Hill and Wetherbee, 1986) and Chroomonas (Kugrens and Lee, 1988). Resistant spore production is rare, but some species produce cysts or palmelloid stages to withstand adverse conditions (Santore, 1978). It has been suggested that palmelloid cells, surrounded by extensive mucilage, might be an adaptation to deter grazing (Klaveness, 1988a).

I. Nucleus and Mitosis The nucleus is located in the cell posterior. In interphase it contains dispersed chromatin and a prominent nucleolus. The outer membrane of the nuclear envelope expands to form the chloroplast endoplasmic reticulum around the chloroplast, nucleomorph, and starch. The Golgi apparatus, the gullet/furrow, and contractile vacuole in freshwater species are situated anterior to the nucleus. Mitosis has been studied in Chroomonas salina (Oakley and Dodge, 1973, 1976; Meyer and Pienaar, 1981, 1984b), Cryptomonas sp. (Oakley and Bisalputra, 1977; Oakley and Heath, 1978), Cryptomonas theta (McKerracher and Gibbs, 1982), and Chroomonas africana (Meyer and Pienaar, 1981, 1984b). Generally, microtubules proliferate near the flagellar bases at the onset of mitosis. The nuclear envelope disaggregates as the microtubules move away from the basal bodies to form a spindle. At metaphase the chromosomes appear as a solid mass. Chromosomal microtubules terminate in the chromosomal mass (Oakley and Dodge, 1973, 1976; McKerracher and Gibbs, 1982), but kinetochores, which are special sites of microtubular attachment to chromosomes, have not been observed. At anaphase the solid chromosomal mass splits and the two masses move to the poles. A cytokinetic ring is formed at metaphase. The ring constricts to cleave the cells into two daughter cells. Cytokinesis and cell separation follow a pole reversal when daughter cells are formed (Perasso et al., 1993).

J. Contractile Vavuoles (Fig. 1) Most freshwater cryptomonads possess a pulsating vacuole, known as a contractile vacuole, that functions

in osmoregulation. The contractile vacuole expels excess water and waste metabolites from the cell (Patterson, 1981). Since freshwater cryptomonads exist in a hypo-osmotic medium, there is a net influx of water into the cell, and the contractile vacuole is the organelle that actively expels water from the cell to prevent a rupture of the cell, since a cell wall is lacking. Contractile vacuoles generally are located in the anterior of the cell.

K. Starch (Figs. 1, 3, 6C, D, 9) The principal storage product is starch, and it is found as granules in the periplastidial space and not in the chloroplast. If a pyrenoid is present some starch accumulates around the pyrenoid in large plates. Cryptomonad starch is an α-1-4-glucan composed of 30% amylose and amylopectin and is similar to starch found in green algae and dinoflagellates (Antia et al., 1979), it stains purple with iodine.

III. ORIGIN OF CRYPTOMONADS Cryptomonads consist of both colorless and pigmented cells, and one of the colorless forms lacks any vestiges of plastids. With the exception of Goniomonas, cryptomonads have one of the most complex cells known, consisting of four genomes—the host genome, the mitochondrial genome, the chloroplast genome, and the endosymbiont nuclear genome. These associations apparently originated from three distinct symbiotic events: two prokaryote–eukaryote events and a eukaryote–eukaryote endosymbiotic association (McKerracher and Gibbs, 1982; Ludwig and Gibbs, 1985a,b, 1989; Gillott, 1990; Douglas et al., 1991; McFadden et al., 1994, 1997). Therefore, except for Goniomonas (McFadden et al., 1994), cryptomonads consist of a eukaryotic host cell, two prokaryotic endosymbionts (the mitochondrion and chloroplast), and a eukaryotic endosymbiont. The endosymbiotic events leading to a pigmented cryptomonad cell presumably are due to a secondary endosymbiosis whereby a colorless phagocytic host cell ingested a red algal cell (McFadden, 1993; McFadden et al., 1997). Once the red algal cell was ingested, it became surrounded by two concentric membranes, one of which is continuous with the outer membrane of the host nuclear envelope. Therefore, the inner membrane probably represents the plasma membrane of the red algal cell, whereas the outermost of these membranes may have originated from the food vacuole membrane fusing with the outer membrane of the nuclear envelope, which bore 80S ribosomes, thereby creating one

21. Cryptomonads

continuous outer membrane. This outer membrane subsequently became studded with 80S ribosomes. Presumably, this occurred by lateral diffusion, through the lipid bilayer, of pre-existing ribosome receptors (i.e., ribophorins) derived from the outer nuclear membrane, resulting in an outer membrane covered with 80S ribosomes. These 80S ribosomes occurring on the outer membrane are distinct from the 80S ribosomes found in the cytoplasm of the red algal endosymbiont. Over time, numerous genes of the endosymbiont were translocated to the host nucleus. This event necessitated the evolution of a complex protein import mechanism whereby chloroplast gene products transcribed on host cytosolic ribosomes could be targeted through the two topogenically unique membranes of the chloroplast endoplasmic reticulum, back into the chloroplast. Two putatively novel N-terminal signal sequences would serve as the import mechanism. Together, these complex events resulted in the chloroplast and the endosymbiont cytoplasm being surrounded by two membranes. This complex, including the smooth inner membrane, is termed the chloroplast endoplasmic reticulum (CER), and the former endosymbiont cytoplasm, with its contents that surround the chloroplast(s), is termed the periplastidial space or compartment. The endosymbiont subsequently transferred many of its genes to the host nucleus. These evolutionary events created a situation in which the endosymbiont cell became fully dependent on the host for its survival. Molecular evidence indicates that the nucleomorph is of red algal origin, and the presence of starch in the periplastidial cytoplasm is visual evidence of this origin. Moreover, cryptomonad chloroplasts contain phycobiliproteins similar to those found in red algae (Glazer and Appell, 1977) and possess the type I purple form of Rubisco, which, among eukaryotes, occurs only in red algae and cryptomonads (Martin et al., 1992). The nucleomorph contains three linear chromosomes bearing multiple ribosomal RNA genes (McFadden et al., 1997). Finally, it is now widely accepted that extant photosynthetic cryptomonads evolved monophyletically from the permanent fusion of a phagocytic protozoan host and a red algal unicell which originally possessed both phycoerythrin and phycocyanin phycobiliproteins. Differential loss of phycoerythrin or phycocyanin produced photosynthetic cryptomonads that are either blue-green or red to reddish brown in coloration. The most comprehensive molecular data regarding cryptomonad phylogeny (Marin et al., 1998) indicate that a switch in coloration occurred from a red to a blue-green cryptomonad, and perhaps back to a red cryptomonad again. Such an event is biochemically feasible (Wemmer et al., 1993; Glazer and Wedemayer,

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1995), and in this context, it is proposed that the photosynthetic ancestor lost phycocyanin and allophycocyanin. Thus, phycoerythrin was the ancestral accessory pigment in these cryptomonads. and bluegreen cryptomonads were derived from phycoerythrincontaining types. Blue-green cryptomonads actually possess true phycoerythrins, but they appear blue-green because linear phycoerythrobilin chromophores have been replaced with phycocyanobilin chromophores (Apt et al., 1995; Glazer and Wedemeyer, 1995). Recent cryptomonad molecular phylogenies (Marin et al., 1998; Clay and Kugrens, 1999; Clay et al., 1999) are largely consistent with the most recently proposed cryptomonad classification system (Clay et al., 1999). In this scheme the division Cryptophyta consists of two classes with the following three orders: the Goniomonadales, which contains a single family; the Cryptomonadales, which contains two families; and the Pyrenomonadales, which contains five families. Six of the eight families appear to be natural groups (i.e., monophyletic) and are supported by molecular phylogenies. The remaining two families represent well-circumscribed taxa, but each also contains a monotypic clade. Based on early sequence data, the host nuclearencoded rRNA appeared to be related to the protozoan Acanthameba and the green algal lineage. However, considerably more sequence data have accumulated in recent years, and this proposed relationship is no longer tenable. At present, the large-scale phylogenetic affinities of the host component of cryptomonads continues to be one of the major conundrums in evolutionary protistology. Ultrastructurally, they appear to be related to the “stramenopiles” sensu Patterson (Heterokonta sensu Cavalier-Smith, 1986) by virtue of possessing tubular hairs and chloroplasts not free in the cytosol, but rather delimited by the two extra membranes of the chloroplast endoplasmic reticulum. Cryptomonads, however, have flat cristae, which demonstrates their affinities with other platycristate eukaryotes. Sequence analyses have not provided evidence of the nature of the host that engulfed the endosymbionts. It has been suggested that Glaucophytes are the sister group to the cryptomonads (Cavalier-Smith et al., 1996), and this is consistent with the analyses by Bhattacharya et al. (1995). It should also be realized that in all published trees to date, cryptomonad host affinities are contravening, and none are considered reliable. In contrast, the origin of the cryptomonad nucleomorph is well understood and is corroborated by several lines of evidence. In particular, many independent molecular analyses support the hypothesis that nucleomorph encoded ribosomal rRNA genes are related to red algae (Douglas et al., 1991; Cavalier-Smith et al., 1996; McFadden et al., 1997).

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IV. ECOLOGY Cryptomonads are ubiquitous and have been reported from nearly all types of water throughout the world, including arctic, temperate, and tropical oceans; streams, lakes, and reservoirs; and environments of variable salinity (Klaveness, 1988a,b). However, it is the lakes in temperate regions of the world where cryptomonads display their largest diversity, and where they are found under a wide variety of conditions (Taylor et al., 1979). Several have even exploited intracellular environments, serving as functional endosymbiotic chloroplasts for some ciliates (Hibberd, 1977; Klaveness, 1988a) and dinoflagellates (Lewitus et al., 1999). However, the number of cryptomonad species generally is underestimated in phytoplankton inventories, largely because of the preservation of samples with destructive fixatives such as formaldehyde (Klaveness, 1988a).

A. Abiotic Factors In lentic, estuarine, and marine habitats, cryptomonads are permanent residents of the phytoplankton community. In fact, when other populations are diminishing, cryptomonads increase in numbers (Rott, 1983; Klaveness, 1988a, b). In many lakes there appears to be a population peak during autumn destratification when the epilimnion and hypolimnion mix (Pollingher, 1981). In an extensive survey conducted by Taylor et al., (1979) of lakes in the eastern and southeastern United States, cryptomonads seemed to prefer colder waters, an observation that we have also made in the Rocky Mountain region. In small temperate lakes, they display a variety of seasonal strategies, including stable stratified populations, diel vertical migration, and formation of resting stages during some parts of the year. In most lakes, cryptomonads exhibit maximal population densities far below the surface (Reynolds, 1980, 1984; Rott, 1983). Optimal depths have been reported from 15 to 25 m, with the deepest occurring in late spring and early summer and the shallowest in late autumn and early winter. This pattern occurs primarily in more productive, buffered lakes with low turbulence and, therefore, reduced but presumably adequate light. In low buffered, eutrophic lakes, the decrease in pH due to photosynthesis favors productivity of cyanobacteria and a decrease in cryptomonads. Several cryptomonad species are able to thrive under low light conditions through the phenomenon of chromatic acclimation. Moreover, some are adept at surviving prolonged periods of darkness. One particular strain was shown to survive a dark period for more

than 24 weeks, whereas two species, Hemiselmis virescens and Rhodomonas lens, survived for only a 4-week maximum. Survival during winter under lake ice in near-dark conditions has been shown, in part, to be an effect of low temperatures resulting in low respiration, a change in lipid composition (Henderson and Mackinlay, 1989), and low grazing pressures (Morgan and Kalff, 1979). These phenomena, coupled with chromatic acclimation to efficiently harvest available light, appear to be sufficient for winter survival. Day length and its relation to water depth have been shown to be important factors shaping phytoplankton communities (Arvola et al., 1991). Because of the attenuating effects of water, daybreak occurs later and nighttime occurs earlier with increasing depth. At extremes, low light intensity translates into low productivity, and extreme light intensity has proved adverse to some phytoplanktonic algae. Consequently, day length acts as a selective force that favors those planktonic algae most capable of responding to the day length–light intensity variable. Cryptomonads are capable of sensing light intensity and avoid the extremes through diel, vertical migration behavior (Watanabe et al., 1976; Watanabe and Furuya, 1982a, b; Arvola et al., 1991). Variation in pH among lakes causes differences to appear in cryptomonad populations and even among strains of the same species isolated from different sources (Pringsheim, 1968; Klaveness, 1988a). However, it is often difficult to determine what is ultimately responsible for these differences, since other environmental factors are coupled with pH under natural conditions. Nevertheless, laboratory studies indicate that strains of a given species isolated from different geographic regions exhibit growth at restricted pH ranges that correlate with those from the locality where they were isolated (Pringsheim, 1968). Another laboratory study conducted on a strain of Rhodomonas lacustris showed good growth between pH 6 and pH 8.5, although pH 10 was tolerated in the light cycle in unbuffered media, but growth was reduced (Klaveness, 1977). As with other environmental parameters, cryptomonads occur over a wide range of pH, but generally the range of pH tolerance is narrow. Based on our own observations, cryptomonads in Colorado and Wyoming favor alkaline conditions and are most prominent in lakes with pH values above 7.5. It has been proposed that algae with a chloroplast endoplasmic reticulum, such as the cryptomonads, might have an advantage in high-pH environments (Lee and Kugrens, 1998, 2000). Dissolved inorganic carbon (DIC) is present mainly as bicarbonate in waters that have a high pH. The carbon-fixing enzyme Rubisco can only utilize DIC in the form of CO2. Therefore, if the

21. Cryptomonads

space within the chloroplast endoplasmic reticulum is acidic, then it could be a reservoir of DIC in the form of CO2 that algae without chloroplast endoplasmic reticulum would not have. The availability of CO2 in this space would impart a competitive advantage to these algae in waters high in pH and low in CO2. A putative cryptomonad, Cyanomonas, has been reported to cause massive catfish deaths in Texas ponds (Pfiester and Holt, 1978). Neither the organism nor the toxin was isolated; therefore, the identity of Cyanomonas remains in doubt. Furthermore, the light micrographs presented in this publication lacked the resolution needed to determine whether it indeed was a Cyanomonas or even a member of the cryptomonads. Rhodomonas sp. also has been implicated in exotoxin production (Stemberger and Gilbert, 1985), although it was not demonstrated that the observed inhibition of rotifer growth was specifically due to Rhodomonas. Bacteria in the cultures may have been the causative agents.

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plast remains functional for a period of time (Schnepf et al., 1989; Lewitus et al., 1999). Kleptoplastidy of cryptomonad chloroplasts occurs in ciliates (Stoecker and Silver, 1990) and dinoflagellates (Skovgaard, 1988; Schnepf et al., 1989; Putt, 1990), and these chloroplasts photosynthesize and produce starch, which may be an available carbon source for the host (Putt, 1990; Schnepf and Elbrächter, 1992), or it may fulfill metabolic requirements under limited food availability (Lewitus et al., 1999). Kleptoplastidity is particularly important in the survival of the ichthyotoxic dinoflagellate Pfiesteria piscicida, where cryptomonad chloroplasts are selectively ingested by nontoxic zoospores. Cryptomonad chloroplasts are retained for approximately 9 days, and they remain functional during that time, as determined by the uptake of C14-bicarbonate (Lewitus et al., 1999). This retention of chloroplasts promotes the survival of this intermediate stage in the life cycle of Pfiesteria.

B. Biological Factors Cryptomonads are optimal or near-optimal food organisms for zooplankton (Guillard, 1975; Klaveness, 1984; Stemberger and Gilbert, 1985; Sarnelle, 1993). Additionally, they are routinely ingested by various colorless dinoflagellates, ciliates, and Kathablepharis sp. (Stemberger and Gilbert, 1985; Clay and Kugrens, 1999a,b; Lewitus et al., 1999). Observations of Daphnia hyalina and Diaptomus gracilis showed that Rhodomonas sp. was present most of the time (Ferguson et al., 1982). Moreover, various studies suggest that when given a choice among flagellate food items, rotifers appear to select cryptomonads preferentially (Stemberger and Gilbert, 1985), and the presence of cryptomonads enhances the reproduction of planktonic rotifers (Edmondson, 1965). Pejler (1977) observed that various rotifers prefer Rhodomonas to Chrysochromulina. Given that many zooplankton selectively choose cryptomonads as prey, grazing probably plays a significant role in regulating planktonic cryptomonad population dynamics (Sarnelle, 1993). Cryptomonad populations generally reach a maximum following periods of moderate turbulence, when they are disseminated throughout the water column and mixed with higher nutrient waters (Reynolds, 1984), and grazing is reduced. When turbulence decreases and nutrients are depleted, grazing once again reduces numbers, thereby regulating the populations. Perhaps one of the most interesting aspects of cryptomonad ecology is the phenomenon known, as kleptoplastidy, which is a process in which the chloroplast of an ingested photoautotroph is retained, and the chloro-

C. Cryptomonad Endosymbiotins and Pathogens Cryptomonad cells are susceptible to prokaryotic or eukaryotic infections; however, pathogenicity in the cryptomonads has not been the object of detailed studies. Although mixotrophy is rare in cryptomonads (see Section IV.D), bacteria can enter the cell and become endosymbionts, as demonstrated by Schnepf and Melkonian (1990). These bacteria apparently do not have any adverse effects on cells and might represent a mutualistic association. In addition, these bacteria harbor bacteriophages (viruses which infect the bacteria). However, other bacteria and viruses (Pienaar, 1976) may adversely affect cryptomonads (Klaveness, 1982). Klaveness (1982) has shown that Cualobacter can attach to cells externally, causing malformations of the cells. Canter (1968) reported that certain cryptomonads are vulnerable to chytrid parasites. Specifically, Rhizophydium fugax has been observed on species of Cryptomonas that are resting in palmelloid colonies. The chytrid may initially be attracted to the polysaccharide mucilage that envelopes the cells of the palmelloid colony. Upon reaching a cell, the chytrid apparently situates itself in the furrow, and the rhizoidal system invades the cell through this depression. Chytrids have also been reported parasitizing Chilomonas striata (Caljon, 1983). In addition to chytrid parasites, intracellular parasites of undetermined taxonomic status have been observed in Cryptomonas (= Campylomonas) rostratiformis (Ettl and Moestrup, 1980).

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D. Types of Nutrition—Carbon Sources Cryptomonads are photoautotrophic, heterotrophic, or mixotrophic. Photoautotrophs synthesize organic molecules from CO2 by photosynthesis. Heterotrophic cryptomonads require some organic materials for their carbon source, via osmotrophy or phagotrophy. Osmotrophs utilize dissolved organic matter, whereas phagotrophs ingest particulate matter, including other organisms. Both types are restricted to colorless cryptomonads and kathablepharids. For instance, Chilomonas is colorless; thus, it is unable to manufacture its own basic organic molecules. It is strictly osmotrophic and obtains organic compounds by incorporating dissolved organic molecules into its cell for metabolism (unpublished observations). It does not ingest particulate materials, the type of nutrition that this alga was assumed to have. In fact, the cell covering is a major obstacle to phagotrophy. Goniomonas, on the other hand, is phagotrophic and routinely ingests bacteria (Mignot, 1965), presumably through a specialized structure in the cell known as the infundibulum (Mignot, 1965; Kugrens and Lee, 1991). Whether it is also osmotrophic is unknown. The vast majority of cryptomonads, however, are strictly photoautotrophic and do not require dissolved organic matter (DOM) in their metabolism. Several studies indicated that dissolved organic matter does not enhance the growth of photosynthetic cryptomonads (Lewitus and Caron, 1991; Arvola and Tulonen, 1998). Reported increased growth when DOM was added probably was due to bacterial respiration, where the bacteria oxidized the organic molecules and released CO2. Cryptomonad growth increased because of increased CO2 for photosynthesis (Arvola and Tulonen, 1998) and not through a utilization of dissolved organic matter. Mixotrophy is an ecologically important type of nutrition in many flagellates (Boraas et al., 1988) where a photosynthetic organism can also ingest particulate matter, primarily other cells, both prokaryotes and eukaryotes. This type of nutrition is common in chrysophytes, but it also has been reported in a few cryptomonads (Tranvik et al., 1989; Kugrens and Lee, 1991). For example, Cryptomonas was studied with respect to bacterial ingestion (Tranvik et al., 1989), but electron microscopic examinations were not conducted. One species of Chroomonas may be mixotrophic, as determined by ultrastructural studies (Kugrens and Lee, 1991). This study revealed a specialized bacterial incorporation vesicle and bacteria in various stages of digestion; it is the only genus of cryptomonad in which mixotrophy has been documented with electron microscopy. As was the case with phagotrophy in Chilo-

monas, mixotrophy in other cryptomonads actually is precluded because of the presence of the periplast and furrow plates, which would impede phagocytosis.

V. COLLECTION, PREPARATION FOR ISOLATION, AND CULTURING Cryptomonads can be collected with phytoplankton nets from lakes or other bodies of standing water, or by grab samples. Attached cryptomonads in mucilage can be scraped off various substrata with a putty knife and placed in a collecting bottle with water, where they become motile. Samples must be kept cold during transport. Fixation of cells is impractical since cells either rupture or distort drastically. Lugol’s fixative, however, provides the least distortion, but because iodine is a component of the fixative, the cells often appear purple because of stained starch, and cells are not their original color. For proper identification living cells must be examined with a microscope. Photomicrography is difficult since usually the cells are actively swimming, and a flash attachment for photomicrography is helpful. Phase contrast or differential interference contrast (DIC) is also helpful in identifying features of cells. Before isolations are attempted, field samples should be enriched with growth media to establish populations that are capable of growing in a given medium. Then individual cells are isolated in the medium with confidence that they will grow. Isolations of cryptomonads are most successful with the serial dilution pipetting technique (Hoshaw and Rosowski, 1973), with the use of either a dissecting microscope or inverted microscope, depending on the dexterity of the individual. All freshwater cryptomonads studied thus far grow profusely in sterilized lake water with added Bold’s Basal Medium (Nichols, 1973) or Alga-Gro (Carolina Biological Supply Company) concentrate at 40 mL per L of lake water. Cultures should be grown in media with a pH of 7.8 or higher and maintained at approximately 18°C (the optimum temperature range is 16°–20°C) in 16:8 h light:dark regimes. In addition, a sterilized wheat seed must be added to colorless cultures such as Chilomonas and Goniomonas. Kathablepharis can only be maintained in mineral medium that also contains its food organism. For instance, K. ovalis requires Chrysochromulina parva, and K. phoenikoston requires Chroomonas.

VI. CLASSIFICATION, KEY, AND DESCRIPTIONS A. Introduction By virtue of their unicellularity and small size,

21. Cryptomonads

proper cryptomonad identification generally cannot be accomplished by light microscopy. Therefore, current and future classification schemes must rely on electron microscopic techniques to reveal many of the unique features of cryptomonad cells (Brett and Wetherbee, 1986; Dodge, 1969; Dwarte and Vesk, 1983; Faust, 1974; Grim and Staehelin, 1984; Hibberd et al., 1971; Gantt, 1971, 1980; Greenwood and Griffiths, 1971; Hill, 1991a, b; Hill and Wetherbee, 1986, 1988, 1989; Klaveness, 1985; Kugrens and Lee, 1987, 1991; Kugrens et al., 1986, 1987; Lucas, 1970a, b, 1982; Munawar and Bistricki, 1979; Santore, 1977, 1982a, b, 1983, 1984, 1987; Sespenwol, 1973; Wetherbee et al., 1986). The pertinent features were described in detail earlier in this chapter. Furthermore, with the use of information from the structures discussed it has been possible to delineate 18 genera, 11 of which occur in freshwater. Cryptomonas, Campylomonas, and Komma are strictly freshwater genera. It is significant that the number of genera has increased since Santore’s 1984 and 1987 review articles, in which he recognized only five genera. Since that time, an expansion and revision of genera has occurred, as well as the elimination of the genus Rhodomonas (Erata and Chihara, 1989; Novarino, 1991; Novarino and Lucas, 1993), which was erected by Santore (1984, 1987), although there were compelling arguments to retain the genus (Hill and Wetherbee, 1989; Hill, 1991a). More recent ultrastructural investigations also point out the need for reexamining the structural characters that were proposed by Santore (1984, 1987) and Novarino (1991, 1994) as generic characters. More importantly, a correct interpretation of the actual structures must be made since the key characters, such as cell architecture and periplast, easily produce artifacts. Therefore, specialized techniques for scanning electron microscopy and freeze-fracture must be employed to observe the unaltered variations in these features (Munawar and Bistricki, 1979; Grim and Staehelin, 1984; Klaveness, 1985; Brett and Wetherbee, 1986; Wetherbee et al., 1986, 1987; Hill and Wetherbee, 1986, 1988, 1989; Hill 1990, 1991b; Kugrens et al., 1986, 1987; Wetherbee et al., 1986; Kugrens and Lee, 1987, 1991). For information on earlier classification schemes based on light microscopy, the publications by Bourrelly (1970), Huber-Pestalozzi (1950), and Skuja (1939, 1948) should be consulted for freshwater cryptomonads, whereas the extensive treatise by Butcher (1967) is the major reference for marine cryptomonads. Since cryptomonads and kathablepharids are combined in this chapter, the unifying characteristics of this grouping would be the presence of ejectisomes and the

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subapical insertion of flagella. The following section, however, involves only those that are within the cryptomonads with periplasts. Kathablepharids are described separately at the end of the chapter.

B. General Features Useful in Determining Genera and Species Initially, strains can be separated artificially into two broad groups based on the presence or absence of pigments. Then a second separation can be based upon the presence of either phycocyanin (blue) or phycoerythrin (red) as the major accessory pigment (Hill and Rowans, 1989). The blue-green and non-blue-green colored genera are separated easily into distinct groups when live cells are observed. The third separation could be based on life histories. For instance, the marine Proteomonas apparently is the only genus studied that displays an alternation of generations. The remainder reproduce asexually or are haplobionts (Kugrens and Lee, 1988). The fourth and most specific method for separating genera is based on a combination of characters involving the furrow/gullet complex (Figs. 1, 7, 8) and the type of periplast component (Fig. 9), primarily the inner component. Plate types, sizes, shapes, and plate arrangements vary among genera. The first periplast separation is based on laminate (single sheet) plate forms vs. multiple plated forms. Genera in each group could further be separated even when pigmentation is lacking. For instance, Chilomonas has a single inner periplast sheet (Grim and Staehelin, 1984; Kugrens and Lee, 1991), and Goniomonas has rectangular plates (Kugrens and Lee, 1991). In the multiple-plated forms, plate shapes, the size of plates, and their arrangement could further delineate genera, in conjunction with the furrow/gullet type and cell shape. The following characteristics are most useful in delineating species (based mainly on SEM features); a brief discussion of these features follows.

1. Flagellar Hair Arrangement and Scale Morphology The arrangement of tubular and/or nontubular hairs, scales, and other structures on flagellar surfaces could circumscribe species. Variations were reported by Kugrens et al. (1986) and Lee and Kugrens (1986).

2. Variations in the Structure of the Flagellar Apparatus The reconstruction of the entire flagellar apparatus for each genus, particularly, the complexity of the rhizostyle, can be examined for comparative purposes in delineating species, as well as for possible future phylogenetic considerations.

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3. The Presence and Structural Variations of the Furrow Plate There are currently two types associated with cryptomonad cells. These appear to correlate with the type of periplast for a given species. Other variations are expected to be found and could serve as an additional character when morphological cladistic analyses are conducted.

4. The Number and Location of the Nucleomorph(s) in the Periplastidal Compartment Variations have been summarized by Santore (1982c); and these variations were used in delineating Pyrenomonas/Rhodomonas and Storeatula (Hill and Wetherbee, 1989; Kugrens et al., 1999), and Teleaulax, Geminigera, and Campylomonas (Hill, 1991b). The genera that have a nucleomorph embedded within a pyrenoid that bridges two lobes of a chloroplast constitute the family Pyrenomonadaceae (Clay et al., 1999). In fact, these genera form a phylogenetic clade based on ssu rDNA sequence data (Marin et al., 1998; Clay et al., 1999).

5. Presence and Location of Eyespots Eyespots have been confirmed ultrastructurally only for Chroomonas spp. (Santore, 1987; Hill, 1991a) Many cryptomonads have orange bodies that do not represent eyespots.

6. Types of Thylakoid Arrangements within the Chloroplasts Several arrangements have been reported, with doublet thylakoids being the most common arrangement. However, caution must be exercised since environmental conditions may influence this feature (Klaveness, 1981).

7. Type of Scales Comprising the Outer Periplast Component Only one type has been found to date, but the sample size has been limited and other types are expected. Only heptagonal scales have been reported (Pennick, 1981; Hill, 1990, 1991b; Lee and Kugrens, 1986).

8. The Number, Location, and Types of Pyrenoids Pyrenoids may be absent, or there may be one, two, or several pyrenoids per chloroplast (HuberPestalozzi, 1950). Thylakoids may or may not penetrate the pyrenoid, and this character may be a diagnostic feature for species determination (Clay and Kugrens, 1999; Clay et al., 1999).

9. The Number of Chloroplasts per Cell This variation may be species specific or may even be useful in determining genera (Hill, 1991b).

C. Classification of the Phylum Cryptophyta The following classification scheme for cryptomonads conforms to the rules and regulations set forth by the International Code of Botanical Nomenclature (ICBN). This scheme is based on the most current and reliable ultrastructural and phylogenetic information and was proposed by Clay et al. (1999a). Phycobiliprotein pigment types are important features in this classification scheme.

Phylum Cryptophyta (syn. Cryptista) Cavalier-Smith (1986) Plastidial complex with nucleomorphs may be present or absent; chloroplasts (when present) contain chlorophylls a and c2, and phycobiliproteins are located in the lumen of the thylakoids; bipartite tubular hairs on flagella occur in members possessing the plastidial complex; cell covering comprises inner and superficial periplast components (IPC and SPC); ejectisomes are present. Two classes are recognized: Class Goniomonadophyceae Cavalier-Smith (1993) Plastids and nucleomorphs are absent. Bipartite tubular hairs on flagella are lacking. Spikes occur on one flagellum. Cells possess an infundibulum. One order: Order Goniomonadales (Goniomonadida) Novarino and Lucas (1993) Diagnosis identical to the class. Family Goniomonadaceae Hill (1991a) Synonymous with Cyathomonadaceae Pringsheim (1944). Characters as for order. Goniomonas Stein. Class Cryptophyceae Plastidial complex with nucleomorphs present. Chloroplasts possess either the phycobiliprotein Crphycoerythrin or Cr-phycocyanin in the intrathylakoidal space. Leucoplast is present in some. Bipartite tubular hairs appear on at least one flagellum. Two orders: Order Cryptomonadales Not equivalent to Cryptomonadales sensu Novarino and Lucas (1993). Chloroplasts possess the phycobiliprotein Cr-phycoerythrin 566 (PE III). Leucoplast is present in some. Two families:

21. Cryptomonads

Family Cryptomonadaceae Furrow and gullet complex with a stoma present. The IPC comprises multiple plates. Nucleomorphs are positioned between the pyrenoid and nucleus. Possess a short rhizostyle without wings (lamellae). A fibrous furrow plate is present. One genus: Cryptomonas Ehrenberg Family Campylomonadaceae Furrow and gullet present; IPC composed of a sheet; nucleomorphs positioned between pyrenoid and nucleus or similar position if a leucoplast is present; possesses a long, keeled flagellar rhizostyle with wings (lamellae); scalariform furrow plate present; vestibular ligule present. Two genera: Campylomonas Hill Chilomonas Ehrenberg Order Pyrenomonadales Not equivalent to Pyrenomonadales sensu Novarino et Lucas (1993, 1995). Chloroplasts possess the phycobiliprotein Cr-phycoerythrin 545 (PE I) or Cr-phycoerythrin 555 (PE II), never Cr-phycoerythrin 566 (PE 566); or possess Crphycocyanin. Four families: Family Pyrenomonadaceae Novarino et Lucas (1993) Chloroplasts possess Cr-phycoerythrin 545 (PE I); nucleomorphs positioned within pyrenoid. Two genera: Pyrenomonas Santore, Synonym: Rhodomonas Karsten Storeatula Hill Rhinomonas Hill et Wetherbee

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Family Geminigeraceae Clay, Kugrens et Lee Chloroplasts possess Cr-phycoerythrin 545 (PE I); IPC comprises a sheet or a sheet and multiple plates if diplomorphic; nucleomorphs never positioned in the pyrenoid; possesses a long, keeled rhizostyle with wings (lamellae); scalariform furrow plate present. Five genera, and all are marine: Geminigera Hill Teleaulax Hill Hanusia Deane, Hill, Brett, et McFadden Guillardia Hill et Wetherbee Proteomonas Hill et Wetherbee Family Chroomonadaceae Clay, Kugrens, et Lee (1999) Chloroplasts possess Cr-phycocyanin 630 (PC III), 645 (PC IV), or Cr-phycocyanin 569; rhizostyle absent. Three genera: Chroomonas Hansgirg Falcomonas Hill Komma Hill Family Hemiselmidaceae Butcher (1967) Chloroplasts possess Cr-phycocyanin 615 (PC II) or Cr-phycoerythrin 555 (PE II), never possess the other three types of phycocyanins or other two types of phycoerythrins; gullet only; nucleomorphs positioned anterior to pyrenoid; rhizostyle absent; thylakoids penetrate pyrenoid; flagella inserted laterally. One genus: Hemiselmis Parke

D. Key 1a.

Cells colorless.....................................................................................................................................................................................2

1b.

Cells pigmented..................................................................................................................................................................................4

2a.

Cells with leucoplast..........................................................................................................................................................Chilomonas

2b.

Cells without leucoplasts and chloroplast endoplasmic reticulum......................................................................................................3

3a.

Cells with a furrow/gullet complex..................................................................................................................................Goniomonas

3b.

Cells lacking a furrow/gullet but have a distinct feeding apparatus...............................................................................Kathablepharis

4a.

Cells blue-green in color, because of presence of phycocyanin............................................................................................................5

4b.

Cells olive, brown, or red in color because of presence of phycoerythrin............................................................................................7

5a.

Flagella inserted approximately one-third of the cell length behind anterior.......................................................................Hemiselmis

5b.

Subapically inserted flagella, near anterior end...................................................................................................................................6

6a.

Cells with hexagonal inner and surface periplast plates...........................................................................................................Komma

6b.

Cells with inner and surface rectangular periplast plates..................................................................................................Chroomonas

7a.

Cells with multiplated periplast..........................................................................................................................................................8

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7b.

Single sheetlike inner periplast component, cells may appear somewhat contorted...........................................................................10

8a.

Cells with square inner periplast plates with beveled corners..........................................................................................Pyrenomonas

8b.

Cells with periplast plates with hexagonal or oval shapes..................................................................................................................9

9a.

Inner periplast plates oval; cells with a complex furrow..................................................................................................Cryptomonas

9b.

Inner periplast plates hexagonal; cells with a simple furrow..............................................................................................Plagioselmis

10a.

Furrow absent, gullet only ...................................................................................................................................................Storeatula

10b.

Simple furrow and gullet present.................................................................................................................................Campylomonas

E. Freshwater Cryptomonad Genera and Species The following descriptions provide characteristics of genera and some species currently recognized. Scanning electron micrographs (SEMs) and light micrographs and diagrams are provided so that cellular features can be interpreted properly. Furthermore, SEM is becoming more common in phytoplankton identification, and the SEMs provided should serve as baseline information for identification. In addition to descriptions for genera, some of the most commonly found species for each genus are described and depicted.

tangular plates that are not offset. Goniomonas has freshwater and marine representatives. Refer to Mignot (1965), Schuster (1968), Hill (1991a), Kugrens and Lee (1991), and Kugrens (1998) for additional information. Goniomonas truncata (Figs. 6B, 7A) Cells 5–12 µm long, 3–5 µm wide, and 4–10 µm deep. This species generally possesses the features listed for the genus. Order Cryptomonadales Family Cryptomonadaceae

Class Cryptophyceae Order Goniomonadales Family Goniomonadaceae Goniomonas Stein (Syn. Cyathomonas (Figs. 3A, 4A, B, 19A) This genus represents a cell that is related to the ancestral type and is the least complex cryptomonad. It lacks any vestige of a plastid and nucleomorph, thus a periplastidial compartment is lacking. However, Goniomonas definitely is a cryptomonad based on the presence of ejectisomes, the structure of the flagellar transition region, and the presence of a periplast. Large ejectisomes are arranged in a ring around the anterior of the cell (Mignot, 1965; Schuster, 1968; Kugrens and Lee, 1991; Kugrens, 1998), and small ejectisomes occur at the corners of the periplast plates, just beneath the periplast, but these are not evident with the light microscope. Cells are laterally compressed, colorless, and phagocytic on bacteria. The nucleus is situated in the dorsal portion of the cell. Flagella are inserted on the dorsal side of the vestibulum, with one flagellum bearing recurved spines, while the other flagellum has fine fibrillar hairs (Kugrens and Lee, 1991). The vestibulum connects to a ventral furrow, which connects to a furrow with posterior stoma; a gullet is absent. An opening, the infundibulum, is located on the left side of the cell and presumably is the ingestion site for particulates. The periplast has inner and outer rec-

Cryptomonas Ehrenberg (Figs. 3B–E, 4C–H, 6A–C, 7, 8, 15C, D) Cells often form palmelloid colonies with cells embedded in extensive mucilage. Motile cells possess two flagella that originate from the right side of the vestibulum, and flagella have the most common arrangement of tubular hairs. Cells have a complex type of vestibular-furrow-gullet complex, with the furrow consisting of furrow ridges, furrow folds, and a persistent oval opening called the stoma that is located at the posterior of the furrow. The furrow appears to have the ability to open and close. The periplast consists of an inner component of round to ovalshaped plates and a surface component of a thin layer of fibrils. The periplastidial compartment contains two chloroplasts with two pyrenoids not traversed by thylakoids, and two nucleomorphs, each located between the nucleus and the pyrenoids. Chloroplasts possess Crphycoerythrin with maximum absorption at 566 nm. This genus is ubiquitous in temperate lakes, reservoirs, and streams and occurs only in freshwater habitats. Light microscopic observations generally are unreliable with respect to species characteristics. For instance, the type species, C. ovata, was described as lacking pyrenoids, but pyrenoids were found (Roberts, 1984; Hill, 1991c; Fig. 4D,E). Based on this character, it would meet the characteristics for C. pyrenoidosa Harvey. For further information on the structure of this genus refer to Santore (1977, 1984), Munawar and

21. Cryptomonads

Bistricki (1979), Roberts (1984), Brett et al. (1986), Kugrens et al. (1986), Kugrens and Lee (1987), and Hill (1991b). Cryptomonas ovata (Figs. 3B, 4D, E, 15C, D) Cells are ellipsoid to oval, and they may be slightly curved. Cells measure 20–80 µm in length, 6–20 µm in width, and 5–18 µm in depth and appear somewhat flattened. The furrow is complex, and cells have a short gullet. There are two chloroplasts per cell, each with a pyrenoid, that are olive green to dark brown in color. Cryptomonas obovata (Fig. 3C) Cells are 24–46 µm long and 13–24 µm in diameter and slightly curved. The vestibulum is below the cell apex, imparting a lobed appearance to the cell apex. Cells have two olive or brown chloroplasts without pyrenoids. Usually many starch grains are present in the cells. Cryptomonas phaseolus (Figs. 3D, 4F) This is the smallest Cryptomonas species, measuring 8–13 µm in length and 5–8 µm in diameter. It is ellipsoid in lateral view and oval in cross section, with rounded ends. The anterior end has a rounded protrusion anterior to the site of flagellar insertion. The posterior of the cell is slightly narrower than the anterior end. Each cell has two brownish chloroplasts without pyrenoids. Cryptomonas tetrapyrenoidosa (Fig. 4G) Cells measure 20–60 µm in length, 10–27 µm in width, and 5–17 µm in depth. There are two chloroplasts per cell, each with two pyrenoids; thus there are four total pyrenoids. The periplast type has not been investigated. Cryptomonas erosa (Figs. 3E, 4H) Cells are oval or slightly elliptical, flat and slightly contorted, ranging in size from 13 to 45 µm in length, and from 6 to 26 µm in width. There are two chloroplasts per cell which are olive, and pyrenoids are absent. The periplast type has not been investigated. Cryptomonas ozolini Skuja (Fig. 3F) Cells are slightly egg shaped and compressed. The anterior end is the widest portion of the cell. Cells measure 17–29 µm in length, 9–13 µm in width, and 6–9 µm in depth. Cells contain two olive-green chloroplasts, each with a pyrenoid. The ultrastructure of this species indicates that it should be a new genus (unpublished observations).

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Family Campylomonadaceae Campylomonas Hill (Figs. 5A–D, 6A–C, 7B–D, 8B) Members of this genus were formerly placed in the genus Cryptomonas because of a lack of ultrastructural evidence that demonstrated that the two genera are different (Hill, 1991b). A basic distinction at the light microscopic level is that Campylomonas spp. are slightly contorted, sigmoid-shaped cells with a characteristic recurved posterior, which imparts a sigmoid shape to the cells in lateral view. At the electron microscopic level, the main distinguishing features are the presence of a periplast sheet, a vestibular ligule, and a simple furrow with a gullet of variable length extending posteriorly from the furrow. A surface periplast component may be lacking, or it may consist of fibrillar material or heptagonal scales. Cells contain two chloroplasts, which may or may not have pyrenoids. If pyrenoids are present, each chloroplast has its own pyrenoid and it is not traversed by thylakoids. Two nucleomorphs are located either near and posterior to each pyrenoid when present, or close to the nucleus. Chloroplasts possess Cr-phycoerythrin with a maximum absorption at 566 nm, but not in quantities that impart a reddish coloration. The genus is strictly freshwater. For additional information refer to Munawar and Bistricki (1979), Klaveness (1985), Kugrens et al. (1986), Kugrens and Lee (1987), Hill (1991b). Note that in all publications, except for Campylomonas reflexa Hill (1991b), Campylomonas is still incorrectly identified as Cryptomonas. Campylomonas reflexa (Figs. 5A, 7C, D) This is the type species, and it has the characteristics described for the genus. Two pyrenoids are present. Nucleomorphs are located posterior to the pyrenoids and anterior to the nucleus. Cells are highly variable in size and can range from 15 to 60 µm in length and from 10 to 30 µm in width. Campylomonas rostratiformis (= Cryptomonas rostratiformis) (Figs. 5B, 7B). This species is the largest cryptomonad, ranging in size from 45 to 80 µm in length, 16 to 40 µm in width, and from 14 to 24 µm in depth. It has not been officially transferred to Campylomonas, although it has been shown that it does not belong in Cryptomonas (Kugrens and Lee, 1986; Kugrens et al., 1987). Cells are slightly recurved at the posterior, and they have a rostrate anterior. The furrow is curved slightly toward the left, and the vestibular ligule is pointed and attached to the left side of the vestibulum. The cells have two chloroplasts, each with numerous pyrenoids.

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Starch is often present in large amounts throughout the chloroplast, obscuring the pyrenoids. Campylomonas platyuris (Figs. 5D, 6C, 8) As was the case with C. rostratiformis, this species also has not been transferred officially to Campylomonas, even though it possesses the periplast and furrow/gullet complex of Campylomonas (Kugrens and Lee, 1986; Kugrens et al., 1987). Cells range in size from 30 to 55 µm in length, from 15 to 28 µm in width, and from 9 to 16 µm in depth, and are characterized by a flattened posterior portion or tail when viewed from the side. There are two chloroplasts per cell, but pyrenoids are lacking. Considerable starch usually is present in the cells. Campylomonas marssoni (Fig. 5C) Cells range in size from 16 to 38 µm in length and from 8 to 14 µm in width. Cells are somewhat fusiform and slightly sigmoid in shape, with a pointed posterior end. Each cell contains two chloroplasts without pyrenoids, which differs from C. reflexa. Chilomonas Ehrenberg (Figs. 5E, 6D, 18B) Three freshwater species have been described. The cells are colorless, but not phagocytic. The furrow/gullet complex in C. paramecium consists of a vestibulum, a short furrow, and a long tubular gullet (Kugrens and Lee, 1991). A vestibular ligule covers the area of contractile vacuole discharge. Both flagella have a unilateral row of tubular hairs (Kugrens and Lee, 1991). The inner periplast component consists of an inner sheet with numerous ejectisome pores, and the sheet is not connected to the plasma membrane (Grim and Staehelin, 1984; Kugrens et al., 1986). The surface periplast component consists primarily of fibrils. Ejectisomes are attached to the pore edges in the periplast sheet. The periplastidial compartment contains leucoplasts that lack thylakoids, numerous large starch grains, and two nucleomorphs located in the periplastidial compartment anterior to the nucleus. Refer to Anderson (1962), Schuster (1970), Sespenwol (1973), Roberts et al. (1981), Grim and Staehelin (1984), Kugrens et al. (1986), Kugrens and Lee (1987, 1991), Heywood (1988), and Kugrens (1998) for additional information. Chilomonas paramecium (Figs. 5E, 6D, 19B) Cells are 20–40 µm long and 10–20 µm in diameter, with a rhinote anterior and a blunt, reflexed posterior, imparting a sigmoid shape to the cells. Two leucoplasts and nucleomorphs are located in the

periplastidial compartment. Evolutionarily this species appears to be derived from Campylomonas (Clay et al., 1999). Chilomonas acuta (Fig. 5F) This species has been described from freshwater, but it has not been cultured. It is possible that this genus might be Leucocryptos acuta in Bourrelly (1970). Its ultrastructural features are unknown. This species was observed in an enrichment culture from a reservoir near Severance, Colorado, but isolation was unsuccessful. Order Pyrenomonadales Family Pyrenomonadaceae Pyrenomonas Santore (= Rhodomonas Karsten) (Figs. 6E–G, 9B, 10A, B) Pyrenomonas is the most accepted name, but some authors continue to use Rhodomonas; the generic names are synonyms. Cells may be red, brown or golden brown, in coloration. However, in freshwater collections we have encountered only red-colored forms. Cells have a short furrow and a deep gullet. The periplast consists of inner more or less square plates with beveled corners. The plates taper slightly toward the posterior. The surface periplast component consists of intertwining fibrils. The periplastidial compartment usually has a single, bilobed chloroplast with a pyrenoid situated between the two lobes of the chloroplast. Thylakoids do not traverse the pyrenoid, and a nucleomorph is located in an invagination of the pyrenoid. Chloroplasts contain a preponderance of phycoerythrin. For additional information refer to Santore (1984), Erata and Chihara (1989), Hill and Wetherbee (1989), Novarino (1993), and Kugrens et al. (1999). Pyrenomonas ovalis (Figs. 6E–G, 9B, 10A, B) Cells are oval to ellipsoid, 14–15.5 µm long and 7–8 µm wide and have a single red chloroplast with two lobes. The pyrenoid is attached to both lobes, forming a bridge between the two lobes, making the chloroplast appear H-shaped. The nucleomorph is embedded within the pyrenoid. Cells have a short furrow and an anterior tubular, deep gullet. Currently this is the only species described from freshwater, and it was collected from Great Western reservoir near Broomfield, Colorado (Kugrens et al., 1999). Storeatula Hill (Figs. 6H–J, 9A, 10C, D) Cells are ellipsoid with a slightly rhinote anterior. A furrow is lacking, and the tubular gullet extends to

21. Cryptomonads

approximately the middle of the cell and is lined with several rows of ejectisomes. The periplast has an inner sheet and an outer component of coarse fibrils. The periplastidial compartment contains a single, bilobed chloroplast, with a pyrenoid that connects the two lobes of the chloroplast. A nucleomorph is located in an anterior groove or depression in the pyrenoid. Chloroplasts contain the biliprotein Cr-phycoerythrin. Only one freshwater species has been identified thus far. For additional information refer to Hill (1991b) and Kugrens et al. (1999). Storeatula rhinosa (Figs. 6H–J, 9A, 10C, D) Cells are 16–20 µm long, 7–8 µm wide, and 8–10 µm deep and ellipsoid with a slightly pointed anterior; a furrow is lacking, with a tubular gullet; single chloroplast with pyrenoid. This species has been collected from Hanratty’s Ditch near Beulah, Colorado, and Sheldon Lake and North Shields Pond in Fort Collins, Colorado. Family Chroomonadceae Komma Hill (Figs. 11A, 12A, B, 13A, B) Based on one isolate, this genus originally was described as being comma-shaped or acuminate with a rounded anterior end, tapering to a pointed or acutely rounded posterior and the absence of a furrow. A tubular gullet extends posteriorly from the base of the vestibulum. The periplast consists of relatively small internal and surface hexagonal plates, with the surface plates being crystalline in composition, and, occasionally, rosulate, heptagonal scales lie on the surface of these external plates. The periplastidial compartment contains a single blue-green chloroplast with a central pyrenoid lacking traversing thylakoids and projects from the chloroplast. The chloroplast occupies a dorsocentral position in the cell and contains C-phycocyanin with a maximum absorption at 645 nm. The nucleomorph is situated at the level of the pyrenoid. Refer to Hill (1990) for more specific descriptions. Other bluegreen cryptomonads also have hexagonal plates (Kugrens and Lee, 1991), but they are not comma shaped. The genus is strictly freshwater, and only one species has been described. Komma caudata (Figs. 11A, 12A, B, 13A, B) Any blue-green acuminate cryptomonads probably represent Komma. Cells are comma shaped or acuminate with an acute posterior end and a rounded anterior end. Cells measure 8–12 µm in length and 4–6 µm in width. Cells contain a single, dorsal, blue-green chloroplast with a single pyrenoid projecting from the center

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of the chloroplast. Cells lack a furrow and possess a gullet only. Chroomonas Hansgirg (Figs. 11B–E, 12C–F, 13C, D, 14B) Cells are subobovate and chloroplasts are bluegreen in color. Cells lack a furrow, but a tubular gullet extends posteriorly from the vestibulum. Inner and outer components of the periplast consist of offset rectangular plates (Hill, 1991a), with the anterior of the plate edges raised, because of rows of intramembrane particles in the cell membrane attaching the plates tightly at the posterior end of each plate. Scales or fibrils may be present, in addition to the surface plates in some species. The periplastidial compartment contains one or two chloroplasts that may have a pyrenoid. The nucleomorph usually is located near the pyrenoid. Chloroplasts contain Cr-phycocyanin, imparting a blue-green color to the cells. A stigma may be present in chloroplasts of some species. Refer to Dodge (1969), Gantt (1971), Antia et al. (1973), Meyer and Pienaar (1984), Kugrens et al. (1986), Kugrens and Lee (1987), and Hill (1990) for additional features and descriptions of Chroomonas spp. Chroomonas oblonga (Fig. 11B) Cells are ellipsoid, measuring 15 µm in length and 6 µm in width. Two chloroplasts are present per cell, each with a pyrenoid. A stigma may be present. The periplast comprises small, rectangular plates in the inner and surface components. Isolated from Fossil Creek south of Fort Collins, Colorado. Chroomonas coerulea (Figs. 11C, 12D, E, 14A) Cells are ellipsoid and sometimes slightly concave dorsiventrally, measuring 8–12 µm in length and 4–6 µm in width. The cell posterior is rounded. Inner and surface periplast components consist of small rectangular periplast plates. A single blue-green chloroplast is present, with a single pyrenoid and a prominent stigma, which is associated laterally with the pyrenoid. Only a vestibulum and gullet are present. Flagella are shorter than the cell length. Cells often form extensive mucilage in which groups of cells remain embedded and attached to a substrate. Ubiquitous in Rocky Mountain lakes, reservoirs, and streams. Chroomonas pochmanni (Figs. 11E, 12C, 13C, D) Cells are barrel shaped to ovoid, 10–18 µm long and 8–13 µm wide, and blue-green in color. A single chloroplast and a massive pyrenoid are present in the cell. This species may be mixotrophic (Kugrens and

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Lee, 1990, 1991). Cells have a gullet but lack a furrow. One massive blue-green chloroplast with a large pyrenoid is present per cell. A large, prominent contractile vacuole is located in the anterior of the cell. In addition a bacterial ingestion vacuole also may be visible with a light microscope. Numerous large starch grains usually are present in the cell. This species does not conform to the characters for Chroomonas or any other blue-green cryptomonad and therefore will need to be a new genus. Chroomonas nordstedtii (Figs. 11D, 12F) Cells are slightly elongated and egg-shaped, with the posterior larger in diameter than the anterior, with the anterior end obliquely truncate. Cells are slightly curved on the dorsal side and range in size from 10 to 30 µm in length and 7 to 15 µm in diameter, which is larger than described by Huber-Pestalozzi (1950). A blue-green parietal chloroplast with a prominent pyrenoid and a small stigma fill the posterior threefourths of the cell. Common in Wyoming lakes. Family Hemiselmidaceae Hemiselmis Parke (Figs. 11F, 12H–I, 14C, D) Cells are rounded anteriorly and posteriorly, slightly flattened dorso-ventrally, and appear somewhat bean-shaped in the lateral view, becausae of the vestibulum and gullet located approximately one-third of the cell length from the anterior. A furrow is absent. The surface periplast component comprises large hexagonal plates, and the inner component probably consists of a periplast sheet. The periplastidial compartment contains a single dorsal, boat-shaped chloroplast with a centrally situated, stalked pyrenoid that is traversed by a single thylakoid. The nucleomorph is located anterior to the pyrenoid. Chloroplasts are blue-green in color because of the presence of Cr-phycocyanin. This genus is ubiquitous in Colorado and Wyoming lakes, specifically Lake John and Cowdrey Lake in Colorado and Diamond and Twin Buttes Lakes in southern Wyoming. Currently there is only one freshwater species described, and it is the smallest cryptomonad described from fresh water. Hemiselmis amylifera (Figs. 11F, 12H–I, 14C, D) Cells generally are suspended throughout the water column and usually do not swim unless disturbed. Cells range in size from 4 to 5.5 µm in length and from 2.5 to 3 µm in width and are 3 µm in depth. They are slightly compressed laterally, and they are ovate or bean-shaped in the lateral view. The vestibulum and gullet are oval and shallow, located one-third the dis-

tance from the anterior. Cells have few ejectisomes and a single parietal chloroplast with a prominent dorsal pyrenoid in the anterior portion of the cell. Cells are blue-green in color. Cryptomonads of Uncertain Taxonomic Status Cyanomonas Oltmanns (Figures Not Available) It must be pointed out that the existence of this genus is in doubt. Cells contain several blue-green chloroplasts. It has never been cultured and might represent a species of Chroomonas, in which large starch grains have been mistaken for chloroplasts (Hill, 1990). The cell shape is similar to that of some Chroomonas spp. Since this genus has not been investigated with the electron microscope, and it has not been cultured, its status as a legitimate genus remains in doubt. There have been suggestions that numerous large starch grains that have a blue-green refraction might have been mistaken for chloroplasts. Nevertheless, a description is provided. One species, C. americana, has been described, and there has been one collection reported in North America, that from the Charles River near Boston, Massachusetts. Plagioselmis Butcher (Figs. 15A, B, 16A, 17A, B) This genus originally was described as being from the marine environment by Butcher (1967), with P. prolonga representing the type species, until Novarino et al. (1994) transferred the freshwater Rhodomonas minuta into the genus Plagioselmis, as P. nanoplanctica. Cells are pink or red in color because of the presence of phycoerythrin, and they are commashaped, with an acute tail that lacks plates. There is a ventral furrow (= sulcus, a deep furrow rather than a gullet). The periplast consists of an internal component (IPC) of hexagonal plates, whereas rosette scales make up the surface periplast component (SPC). The acute tail has a continuous periplast sheet rather than plates. Thylakoids usually occur in groups of three (Klaveness, 1981). Novarino et al. (1994) described several of these features from three isolates of Plagioselmis, but one major characteristic differed among the isolates. This was used to delineate taxa at the species level. For instance, they noted that some strains of P. prolonga and P. nanoplanctica lacked a furrow (= sulcus). Cells have a depression on the ventral side, but they attributed these depressions to folds that were induced by cell shrinkage. If the absence of a furrow indeed is correct, then either the characters for Plagioselmis must be expanded or the nonfurrow strains should be described as a new genus; an investigation is needed to determine whether the folds are artifacts or whether the collapsed folds were obscure a furrow. However, it

21. Cryptomonads

is clear from the figures provided in this chapter that Plagioselmis has a furrow. P. nanoplanctica (Fig. 15A, B, 16A, 17A, B) This is the only recognized freshwater species. Cells are comma-shaped with an acute posterior end, and chloroplasts are red or pink colored. Cells range in size from 12 to 18 µm in length and from 8 to 10 µm in diameter at the widest portion of the cell. This is the only recognized freshwater species at this time.

F. Guide to Literature for Species Identification 1. Campylomonas—Hill (1991c), Kugrens et al. (1986) 2. Chilomonas—Hill (1991), Kugrens and Lee (1991) 3. Chroomonas—Hill (1991a) 4. Cryptomonas—Kugrens et al. (1986) 5. Goniomonas—Hill (1991), Kugrens and Lee (1991) 6. Hemislemis—Clay and Kugrens (1999b) 7. Kathablepharis—Lee and Kugrens (1991), Lee et al. (1991), Clay and Kugrens (1999a, c) 8. Komma—Hill (1991a) 9. Plagioselmis—Novarino et al. (1994) 10. Rhodomonas/Pyrenomonas—Kugrens et al. (1999) 11. Storeatula—Kugrens et al. (1999)

VII. AVAILABILITY OF CRYPTOMONADS Cryptomonad cultures are available from a variety of sources. At Colorado State University 172 strains of cryptomonads are maintained, and most are not duplicate strains. In addition, Dr. Michael Melkonian’s laboratory at the University of Cologne and Dr. Dag Klaveness’s laboratory at the University of Oslo, Norway, have numerous cryptomonads in culture. Each of these culture collections contains over 50 isolates. Other culture collections with a considerable number of cryptomonad cultures are the University of Texas Culture Collection, the Japanese NIES Collection, the Culture Collection of Algae and Protozoa, and Provasoli-Guilliard Culture Collection of Marine Phytoplankton (CCMP) at the Bigelow Laboratories, Boothbay Harbor, Maine.

VIII. FAMILY KATHABLEPHARIDACEAE The flagellates belonging to this family include the genera Kathablepharis and Leucocryptos, but the dis-

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tinction between the two is rather tenuous (Vørs, 1992a, b). Higher taxonomic affiliations are not possible at this time, and the family is considered incertae sedis (Clay and Kugrens, 1999). As mentioned earlier, these probably should be placed in their own phylum, but currently they are still classified as cryptomonads. These organisms represent one of the most enigmatic groups of protists. Some of their cellular structures are unusual and are not found in any other groups, yet other cellular structures transcend several groups. Their classification is uncertain, but traditionally the katablepharids have been placed in the cryptomonads because of the presence of ejectisomes and the site of flagellar insertion. These features were based on earlier light microscopic observations, but electron microscopic examination has revealed distinct differences. For instance, the ejectisomes are not similar to either the cryptomonad ejectisomes or the Pyramimonas parkeii ejectisomes. Kathablepharis also has a complex feeding apparatus similar to that of suctorian ciliates or the apical complex found in apicomplexans (Kugrens et al., 1994), based on the similarity of the conoid rings and associated microtubules in the feeding apparatus, resembling an apical complex. In his treatment of the kingdom Protozoa, CavalierSmith (1993) places Kathablepharis in the phylum Opalozoa, a position somewhat removed from that of the rest of the cryptomonads, because it differs in many ultrastructural aspects, including the presence of tubular mitochondrial cristae and the purported absence of the subsidiary scroll that is present in the ejectisomes of all cryptomonads. In addition, a periplast is absent in kathablepharids, and they possess the distinct feeding apparatus described earlier (Lee and Kugrens, 1991b).

A. Ecology Based on collecting data from the Rocky Mountain region, kathablepharids appear to be tolerant of a variety of temperatures, pH, salinities, and nutrient conditions, a fact also noted by Vørs (1992) in her autecological studies. The lack of more global information regarding these flagellates probably is due to the fact that they may be overlooked in plankton samples or misidentified, especially in fixed material. Kathablepharids are voracious predators, attacking their prey in groups, and the size of their groups ranges up to several hundred cells. They feed on both bacteria and various eukaryotes, but each freshwater species that has been studied in detail appears to prefer a specific food organism (Lee and Kugrens, 1991; Lee et al., 1991). For instance, Kathablepharis ovalis feeds on the chrysophyte Chrysochromulina parva, and K. phoenikoston prefers to feed on Chroomonas.

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B. Cell Structure (Fig. 8C) 1. Cell Covering A distinctive cell covering surrounds the cell, including the flagella (Fig. 8C). The cell covering is composed of outer and inner components. The outer component of the cell covering occurs in distinct rows encircling the body of the cell. The inner component of the cell covering occurs between the outer compartment of the wall and the plasma membrane. The inner component is composed of randomly arranged fibrils and covers the whole cell. The outer compartment of the cell covering is absent over the area of the cytostome, the area where the flagella are inserted into the cell and the area posterior to the cytostome, where the rows of ejectisomes occur under the plasma membrane. The outer cell covering appears to be made up of rows of hexagonal subunits, perhaps scales (Lee and Kugrens, 1991), which overlap each other in the rows. The rows of subunits are oriented approximately 45° to the cell surface. The subunits and thus the rows are approximately 25 nm wide.

2. Flagella Flagella are inserted subapically, and their orientation is variable among species. The flagella appear thick when viewed with the light microscope because of a scale covering.

3. Feeding Apparatus All kathablepharids have a distinctive, complex feeding apparatus that is similar to that of suctorian ciliates or apicomplexans (Kugrens et al., 1994; Lee et al., 1991). Depending on the species, the feeding apparatus consists of a stack of 2 to 10 rings that are located just posterior to the depression at the anterior end known as the stoma or mouth. Microtubular bundles are attached to these rings, and the microtubules extend toward the posterior of the cell. Small vesicles occur inside along the cytopharyngeal microtubules. The successive rings increase in diameter when they are farther from the stoma, forming a cone or “conoid” of rings. The microtubules collectively are known as the cytopharyngeal skeleton (Vørs, 1992), and there are two of these circular arrays. One array is associated with the conoid rings, and the others occur just beneath the cell cover and probably function as a cytoskeleton. This second array of microtubules is known as the pellicular skeleton (Vørs, 1992).

4. Nucleus and Mitosis The interphase nucleus is typical of a eukaryotic nucleus, with chromatin attached to the inner membrane of the nuclear envelope. A single nucleolus

occurs in the nucleus. When cells divide, the nucleolus disperses, the nuclear envelope detaches from the chromatin and is converted into rough endoplasmic reticulum, and the chromatin condenses into a single disc-shaped mass where individual chromosomes cannot be discerned. Microtubules penetrate the chromosome mass, but kinetochores were not observed. Spindle microtubules end in a number of minipoles in the cytoplasm. The chromosome mass separates at anaphase, and each mass migrates to the poles. Then the nuclear envelope attaches to the chromatin and the nucleolus reappears. Cytokinesis is longitudinal, forming two cell products.

5. Mitochondria Mitochondria have tubular cristae, and they usually are found between the outer and inner arrays of microtubules of the pellicular and cytopharyngeal skeletons. Serial sections of the cell have not been made; thus there is the possibility that only one mitochondrion occurs per cell.

6. Ejectisomes Ejectisomes consist of only one component rather than two as described for cryptomonads. Large ejectisomes occur in the area of the cell posterior to, and to the right of, the flagella. The large ejectisomes occur in one or two rows oriented parallel to the long axis of the cell. Smaller ejectisomes occur under the plasma membrane in the medial and posterior areas of the cell. Both large and small ejectisomes consist of a single ribbon wound into a spiral that is contained within a membrane. On discharge into the surrounding medium, the ejectisomes consist of a long, straight ribbon. After discharge of the ejectisome, the edges of the ribbon roll inward, creating a tubular structure. Near the tip of the discharged ejectisome, the ribbon tapers rapidly to a spatula-like point.

7. Food Vacuoles Food vacuoles are located in the posterior portion of the cell, and food particles are transferred from the cytosome to these food vacuoles. Both bacteria and chloroplasts from the food organisms are commonly found in these food vacuoles.

8. Alveolate-like Structures Alveoli are flattened vesicles that occur beneath the plasma membrane and are characteristic of dinoflagellates and ciliates. Kathablepharids have similar vesicles that are derived from endoplasmic reticulum. These differ from dinoflagellates, however, because they have ribosomes attached to them.

21. Cryptomonads

C. Classification of Kathablepharidaceae The colorless flagellate Kathablepharis Skuja comprises eight freshwater species based on light microscopic studies. Two of the most common freshwater representatives, K. ovalis and K. phoenikoston, are described in this chapter. Kathablepharis Skuja (Figs. 16B, C, 17C, D, 18C, D) Cells of Kathablepharis vary in size and morphology, but all have two flagella that are subapically inserted on the ventral side, a continuous covering of fused scales outside of the plasma membrane, and ejectisomes. Cells may be oval to cylindrical in shape. Chloroplasts are absent, although when chloroplasts are present in food vacuoles, there could be the impression that cells possess chloroplasts. Two species of freshwater Kathablepharis are discussed. Kathablepharis ovalis (Figs. 16B, 17C. 18C) K. ovalis is a common flagellate in freshwater habitats in the Rocky Mountain region, occurring in a variety of lentic and lotic habitats. It is small and colorless and often contains one to several cells of Chrysochromulina parva, making it appear as though it contains chloroplasts. Cells range in size from 8 to 15 µm in length, but the size is dependent on the number of ingested cells. Cells have two subapical flagella, and the flagella emerge laterally from a subapical mound. The anterior flagellum is approximately 15 µm long, and the posterior flagellum is approximately 12 µm long. The flagella are encased in the same cell covering that encloses the entire cell. The cells have a large central nucleus. One to several large food vacuoles occupy the posterior portion of the cell. Two arrays of microtubules, one inside the other, begin at the anterior end of the cell and continue into the posterior region of the cell. A Golgi apparatus is just anterior to the nucleus and inside the inner array of microtubules. Six large ejectisomes occur in two rows posterior to and just to the right of the flagella, while smaller ejectisomes occur under the plasma membrane in the posterior and medial portion of the cell. At the light microscopic level, cells of K. ovalis are ovate to subovate, with both flagella directed anteriorly during swimming. At the ultrastructural level, the feeding apparatus in K. ovalis has two cytopharyngeal rings. K. ovalis has been collected from ponds in the Department of Energy’s Rocky Flats Nuclear Weapons Plant, Jefferson County; in Horsetooth Reservoir and North Shields Pond, Larimer County, and in South Delaney Buttes Lake and Lake John, Jackson County, Colorado, USA. This organism, however, is easily overlooked because it is small and colorless and has no

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striking features when examined in the light microscope. In addition to dispersed cells, swarms of Kathablepharis are common, consisting of aggregations of 20–100 cells that attack the prey cell. Kathablepharis phoenikoston Skuja (Figs. 16C, 17D, 18D) Cells of K. phoenikoston are cylindrical and have one anteriorly directed flagellum and one trailing flagellum when swimming. K. phoenikoston possesses 9 to 10 conoid-like rings that are associated with the feeding apparatus. The cell covering and other features are similar to those described for K. ovalis.

D. Isolation and Culturing Techniques for Kathablepharids Isolation involves the same techniques as described for cryptomonads. In addition to the usual mineral media, these organisms require the presence of their specific food organism in the culture since they do not survive on bacteria alone. The mineral medium promotes growth of the photosynthetic prey organism so that kathablepharids can actively feed and survive by ingesting the selected alga.

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Cryptophyceae. II. The eyespot. British Phycological Journal 17:113–119. Ludwig, M., Gibbs, S. P. 1985a. DNA is present in the nucleomorph of cryptomonads: Further evidence that the chloroplast evolved from a eukaryotic endosymbiont. Protoplasma 127:9–20. Ludwig, M., Gibbs, S. P. 1985b. DNA is present in the nucleomorph of cryptomonads: Further evidence that the chloroplast evolved from a eukaryotic endosymbiont. Protoplasma 127:9–20. Ludwig, M., Gibbs, S. P. 1989. Localization of phycoerythrin at the lumenal surface of the thylakoid membrane in Rhodomonas lens. Journal of Cell Biology 108:875–884. Lund, J. W. G. 1962. A rarely recorded but very common British alga, Rhodomonas minuta Skuja. British Phycological Bulletin 2:133–139. Marin, B., Klingberg, M., Melkonian, M. 1998. Phylogenetic relationships among the Cryptophyta: Analysis of nuclear-encoded SSU rRNA sequences support the monophyly of extant plastidcontaining lineages. Protistology 149:265–276. Martin W., Sommerville C. C., Loiseaux-de Goer S. 1992. Molecular phylogenies of plastid origins and algal evolution. Journal of Molecular Evolution 35: 385–404. McFadden, G. I. 1993. Second-hand chloroplasts: Evolution of cryptomonad algae. Advances in Botanical Research 19:189–230. McFadden, G. I., Gilson, P. R., Douglas, S. E., Cavalier-Smith, T., Hofmann, C. J. B., Maier, U-G. 1997. Bonsai genomics: Sequencing the smallest eukaryotic genomes. Trends in Genetics 13:46–49. McFadden, G. I., Gilson, P. R., Hill, D. R. A. 1994. Goniomonas: rRNA sequences indicate that this phagotrophic flagellate is a close relative of the host component of cryptomonads. European Journal of Phycology 29:29–32. McKerracher, L., Gibbs, S.P. 1982. Cell and nucleomorph division in the alga Cryptomonas. Canadian Journal of Botany 60:2440–2452. Meyer, S. R., Pienaar, R. N. 1981. The ultrastructure of mitosis and cytokinesis in a new species of Chroomonas (Cryptophyceae). Electron Microscopy Society of Southern Africa Proceedings 11:163–164. Meyer, S. R., Pienaar, R. N. 1984a. The microanatomy of Chroomonas africana sp. nov. (Cryptophyceae). South African Journal of Botany 3:306–319. Meyer, S. R., Pienaar, R. N. 1984b. Mitosis and cytokinesis in Chroomonas africana Meyer, Pienaar (Cryptophyceae). South African Journal of Botany 3:320 –330. Mignot, J. P. 1965. Etude ultrastructurale de Cyathomonas truncata From. (Flagelle Cryptomonadine). Journal de Microscopie (Paris) 4:239–252. Morgan, K., Kalff, J. 1975. The winter dark survival of an algal flagellate Cryptomonas erosa (Skuja). Internationale Vereinigung für Theoretische und Angewandte Limnologie Verhandlungen 19:2735–2740. Morrall, S., Greenwood, A. D. 1980. A comparison of the periodic substructure of the trichocysts of the Cryptophyceae and Prasinophyceae. Biosystems 12:71–83. Morrall, S., Greenwood, A. D. 1982. Ultrastructure of nucleomorph division in species of Cryptophyceae and its evolutionary implications. Journal of Cell Science 54:311–328. Munawar, M., Bistricki T. 1979. Scanning electron microscopy of some nanoplankton cryptomonads. Scanning Electron Microscopy 3:247–252. Nichols, H. W. 1973. Growth media—freshwater, in: Stein, J. R., Ed., Handbook of Phycological Methods. Culture Methods and Growth Measurements. Cambridge University Press, New York, pp. 7–24 Novarino, G. 1991a. Observations on Rhinomonas reticulata comb.

nov. and R. reticulata var. eleniana var. nov. (Cryptophyceae), with comments on the genera Pyrenomonas and Rhodomonas. Nordic Journal of Botany 11:2453–2252. Novarino, G. 1991b. Observations on some new and interesting Cryptophyceae. Nordic Journal of Botany 11:599–611. Novarino, G. 1993a. A comparison of some morphological characters in Chroomonas ligulata sp. nov. and C. placoides sp. nov. (Cryptophyceae). Nordic Journal of Botany 13:583–589. Novarino, G. 1993b. Possible detection of the periplast areas and the nucleomorph of cryptomonads by light microscopy: Some early observations by Künstler, Skuja and Hoolande. Quekett Journal of Microscopie (Paris) 37:45–51. Novarino, G., Lucas, I. A. N. 1993. Some proposals for a new classification system of the Cryptophyceae. Botanical Journal of the Linnean Society, London 111:3–21. Novarino, G., Lucas, I. A. N., Morrall, S. 1994. Observations on the genus Plagioselmis (Cryptophyceae). Cryptogamie, Algologie 15:87–96. Oakley, B. R., Bisalputra, T. 1977. Mitosis and cell division in Cryptomonas (Cryptophyceae). Canadian Journal of Botany 55:2789–2800. Oakley, B. R., Dodge, J. D. 1973. Mitosis in the Cryptophyceae. Nature 244:521–522. Oakley, B. R., Dodge, J. D. 1976. The ultrastructure of mitosis in Chroomonas salina (Cryptophyceae). Protoplasma 88:241–254. Oakley, B. R., Heath, I. B. 1978. The arrangements of microtubules in serially sectioned spindles of the alga Cryptomonas. Journal of Cell Science 31:53–70. Oakley, B. R., Santore, U. J. 1982. Cryptophyceae: Introduction and bibliography, in: Rosowski, J. R., Parker, B. C., Eds., Selected papers of phycology. Allen Press, Lawrence, KS, pp. 682–686. Parducz, B. 1967. Ciliary movement and coordination in ciliates. International Review of Cytology 21:91–128. Patterson, D. J. 1981. The behaviour of contractile vacuole complexes of cryptophycean flagellates. British Phycological Journal 16:429–439. Pennick, D. L. 1981. Flagellar scales in Hemiselmis brunnescens Butcher and H. virescens Droop (Cryptophyceae). Archiv für Protistenkunde 124:267–270. Pejler, B. 1977. Experience with rotifer cultures based on Rhodomonas. Archiv für Hydrobiologie Ergebnisse der Limnologie 8:264–266. Perasso, L., Brett, S. J., Wetherbee, R. 1993. Pole reversal and the development of cell asymmetry during division in cryptomonad flagellates. Protoplasma 174: 19–24. Pfiester, L. A., Holt, J. R. 1978. A freshwater “red tide” in Texas. Southwestern Naturalist 23:103–110. Phillips, D., Boyne, A. F. 1984. Liquid nitrogen-based quick freezing: Experiences with bounce-free delivery of cholenergic nerve terminals to a metal surface. Journal of Electron Microscopy Technique 1:9–29. Pienaar. R. N. 1976. Virus-like particles in three species of phytoplankton from San Juan Island, Washington. Phycologia 15:185–190. Pollinger, U. 1981. The structure and dynamics of the phytoplankton assemblages in lake Kinneret, Israel. Journal of Plankton Research 3:93–105. Pringsheim, E. G. 1968. Zur kenntnis der Cryptomonaden des Süsswassers. Nova Hedwigia 16:367–401. Putt, M. 1990. Metabolism of photosynthate in the chloroplastretaining ciliate Loboea strobila. Marine Ecology Progress Series 60:271–282. Reynolds, C. S. 1980. Phytoplankton assemblages and their periodicity in stratifying lake systems. Holarctic Ecology 3:141–159.

21. Cryptomonads Reynolds, C. S. 1982. Phytoplankton periodicity: Its motivation, mechanisms and manipulation. Freshwater Biological Association Annual Report 3:141–159. Reynolds, C. S. 1984. Phytoplankton periodicity: The interactions of form, function and environmental variability. Freshwater Biology 14:111–142. Roberts, K. R. 1984. Structure and significance of the cryptomonad flagellar apparatus. I. Cryptomonas ovata (Cryptophyta). Journal of Phycology 20:159–167. Roberts, K. R., Stewart, K. D., Mattox, K. R. 1981. The flagellar apparatus of Chilomonas paramecium (Cryptophyceae) and its comparison with certain zooflagellates. Journal of Phycology 17:159–167. Rott, E. 1983. Sind die Veränderung im Phytoplanktonbild dem Pilburger Sees Auswirkungen der Tiefenwasserableitung? Archiv für Hydrobiologie Supplement band 67:29–80. Santore, U. J. 1977. Scanning electron microscopy and comparative micromorphology of the periplast of Hemiselmis rufescens, Chroomonas sp., Chroomonas salina and members of the genus Cryptomonas (Cryptophyceae). British Phycological Journal 12:255–270. Santore, U. J. 1978. Light- and electron-microscopic observations of the palmelloid phase in members of the genus Cryptomonas (Cryptophyceae). Archiv für Protistenkunde 120:420–435. Santore, U. J. 1982a. Comparative ultrastructure of two members of the Cryptophyceae assigned to the genus Chroomonas—with comments on their taxonomy. Archiv für Protistenkunde 125:5–29. Santore, U. J. 1982b. The ultrastructure of Hemiselmis brunnescens and Hemiselmis virescens with additional observations on Hemiselmis rufescens and comments about the Hemiselmidaceae as a natural group of the Cryptophyceae. British Phycological Journal 17:81–89. Santore, U. J. 1982c. The distribution of the nucleomorph in the Cryptophyceae. Cell Biology International Reports 6:1055–1063. Santore, U. J. 1983. Flagellar and body scales in the Cryptophyceae. British Phycological Journal 18:239–248. Santore, U. J. 1984. Some aspects of taxonomy in the Cryptophyceae. New Phytologist 98:627–646. Santore, U. J. 1987. A cytological survey of the genus Chroomonas— with comments on the taxonomy of this natural group of the Cryptophyceae. Archiv für Protistenkunde 134:83–114. Santore, U. J., Greenwood, A. D. 1977. The mitochondrial complex in Cryptophyceae. Archives of Microbiology 112:207–218. Sarnelle, O. 1993. Herbivore effects on phytoplankton succession in a eutrophic lake. Ecological Monographs 63:129–149. Schnepf, E., Elbrächter, M. 1992. Nutritional strategies in dinoflagellates: a review with emphasis on cell biological aspects. European Journal of Protistology 28:3–24. Schnepf, E., Melkonian, M. 1990. Bacteriophage-like particles in endocytic bacteria of Cryptomonas (Cryptophyceae). Phycologia 29:338–343. Schnepf, E., Winter, S., Mollenhauer, D. 1989. Gymnodinium aeruginosum (Dinophyta): A blue-green dinoflagellate with a vestigial, anucleate, cryptophycean endosymbiont. Plant Systematics and Evolution 164:75–91. Schuster, F. L. 1968. The gullet and trichocysts of Cyathomonas truncata. Experimental Cell Research 49:277–284. Schuster, F. L. 1970. The trichocysts of Chilomonas paramecium. Journal of Protozoology 17:521–526. Sespenwol, S. 1973. Leucoplast of the cryptomonad Chilomonas paramecium; evidence for the presence of a true plastid in a colorless flagellate. Experimental Cell Research 76:395–409.

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22

BROWN ALGAE John D. Wehr Louis Calder Center—Biological Station and Department of Biological Science Fordham University, Armonk, New York 10504

I. Introduction II. Diversity and Morphology A. Diversity and Classification B. Morphology and Reproduction III. Ecology and Distribution A. Ecological Factors B. Geographical Distribution

I. INTRODUCTION Brown algae, the Phaeophyceae (or Fucophyceae; Christensen, 1978), are a class (or division, Phaeophyta; Papenfuss, 1951) of algae consisting mainly of complex, macroscopic seaweeds whose brown color comes from a carotenoid pigment, fucoxanthin, and in some species, various phaeophycean tannins. Of perhaps 2000 species (in 265 genera) of brown algae (Van den Hoek et al., 1995), less than 1% are known from freshwater habitats, although some marine species may colonize brackish waters (Wilce, 1966; Dop, 1979; West and Kraft, 1996). Various authors cite between 3 and 7 genera, and up to 12 species of freshwater brown algae worldwide (see Sect. 21.II.A). Members of the group have many features in common with chrysophytes, synurophytes (Chrysophyta), and diatoms (Bacillariophyta), including chloroplast structure (thylakoids in stacks of three, girdle lamella, chloroplast endoplasmic reticulum), heterokont motile stage (unequal flagella), major pigments (chlorophylls a, c1, and c2, β-carotene, violaxanthin, diatoxanthin, and large amounts of fucoxanthin), as well as the storage reserve laminarin (Craigie, 1974; Goodwin, 1974; Pueschel and Stein, 1983; Lee, 1989). However, no Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

IV. Methods for Collection and Identification V. Key and Descriptions of Genera A. Key B. Descriptions of Genera VI. Guide to Literature for Species Identification Literature Cited

members of the Phaeophyceae are unicellular or colonial in the vegetative phase—the predominant morphology in other golden-brown groups. Brown algae have cell walls composed of cellulose, which is often supplemented with the mucopolysaccharide alginic acid. In seaweeds, this material is produced in sufficient quantities in some species to be harvested for commercial purposes, but in freshwater species, alginates appear to be less prevalent. Further descriptions of the group can be found in reviews by Papenfuss (1951), Van den Hoek et al. (1995), and Graham and Wilcox (2000). Freshwater species of brown algae have been known for more than 100 years [Pleurocladia was described by Braun (1855), Heribaudiella by Gomont (1896); Bourrelly (1981)], but today most are still known only from scattered locations. Freshwater phaeophytes occur in a variety of streams and rivers, as well as in the littoral zone of lakes, but their biology has largely remained obscure for most phycologists and freshwater ecologists. This is unfortunate, because some species, most notably Heribaudiella fluviatilis, can at times be one of the dominant species of benthic algae in smaller rivers (Kann, 1978a). One reason for their obscurity may be that most species form crusts or 757

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brown colonies that may be mistaken for members of other algal groups or lichens. The most recent monograph of freshwater algae in North America (Smith, 1950) listed only one species (H. fluviatilis), which was recorded from only one location [considered doubtful by Smith (1950), but see Sect. V.B], and even its identity has been questioned (Pueschel and Stein, 1983). Since that time, other genera and many more locations have been identified on this continent, suggesting that at least some species are less rare than previously thought, although their distribution is still far from well known. Whitford (1977) proposed that few species of freshwater algae may actually be rare, but instead are simply under-reported. Several species long thought to be uncommon have turned out to be fairly cosmopolitan, such as several species of freshwater red algae (Sheath and Hambrook, 1990). For these reasons, Whitford challenged biologists to describe the habitats of newly discovered algae more carefully. Because of the paucity of information on freshwater Phaeophyta, the present chapter describes all known taxa, although at present only five species (in four genera) have thus far been confirmed from sites in North America.

II. DIVERSITY AND MORPHOLOGY A. Diversity and Classification Freshwater brown algae are undoubtedly the least diverse of all groups of freshwater algae. Although some species can at times form substantial populations, no habitats are known that have several species of freshwater brown algae within a single location. Kann (1993) has observed filamentous Pleurocladia and encrusting Heribaudiella occurring together on stones in the littoral zone of Lake Erken, Sweden, and KuselFetzmann (1996) noted that the Pleurocladia may grow as an epiphyte on Heribaudiella in some Austrian streams. More than 60 years ago, Israelsson (1938) demonstrated that different species of freshwater brown algae exhibit different geographic patterns, which appear to be the result of different ecological requirements. Given the small number of species overall, it is not surprising that their local diversity is low. Accounts of the number of genera and species of phaeophytes from freshwater vary among authors, largely due to lack of study. Their classification is also unsettled, mainly because of uncertainty regarding the reliability of certain morphological features (e.g., branching pattern, colony shape, presence of hairs) as taxonomic attributes, the possible synonymy of several taxa, and whether historical collections were described and identified accurately (Waern, 1952; Müller and Geller, 1978; Dop, 1979; Bourrelly, 1981; Pueschel and

Stein, 1983; Wehr and Stein, 1985). Further work with field populations and cultured material to better describe their reproduction and genetic relationships will undoubtedly reveal new groupings and perhaps new (or fewer) species within the group. The present account recognizes six freshwater genera and seven species worldwide within the division (Table I). Of these, five species have thus far been reported from sites in North America. Following the general schemes of Bold and Wynne (1985) and Van den Hoek et al. (1995), all fresh water phaeophytes are classified as members of the Ectocarpales (five genera) or Sphacelariales (one genus, two species).

B. Morphology and Reproduction The morphologies of all freshwater phaeophyte species are based upon a relatively simple filamentous structure and do not form parenchymatous (tissue-like) thalli, characteristic of more complex brown seaweeds. Their size range is also substantially smaller than those that colonize marine habitats. Crustose forms, although visually conspicuous, may only be 10–30 cells tall (1–2 mm), and form colonies of perhaps 0.2–50 cm2 in area (Wehr and Stein, 1985; Kusel-Fetzmann, 1996). Filamentous forms can form macroscopic tufts 2–10 mm in size. Several colonies may coalesce to form larger expanses on rocks, but these dimensions are in great contrast to species of intertidal brown algae, which reach sizes of several meters, or subtidal kelp forests that may be as tall as 20–60 m (Bold and Wynne, 1985). Among freshwater forms, three basic morphologies are seen (Fig. 1): (1) most consist of uniseriate (single axis), branched filaments (members of the Ectocarpales), which develop to form either (Fig. 1A) spreading or cushion-like tufts [Bodanella, Ectocarpus, Pleurocladia, Porterinema (syn. = Pseudobodanella)]; (2) others consists of prostrate filaments, which produce an upright series of densely packed, vertical filaments, forming a crustose morphology (Heribaudiella; Fig. 1B); and (3) in Sphacelaria cells are arranged in multiseriate (multiaxial), branched filaments, and also form spreading cushions on submerged substrata (Fig. 1C). Thalli in most species seem capable of forming hyaline, multicellular filaments or hairs (Waern, 1952; Wilce, 1966; Dop, 1979; Schloesser and Blum, 1980; Yoshizaki et al., 1984; Kusel-Fetzmann, 1996; Wujek et al., 1996). It seems doubtful that these hairs can be used as diagnostic features, as studies suggest they may be produced in response to reduced Cl– or long photoperiods (Dop, 1979), or P-limitation (Fig. 2A). Such patterns are similar to those observed in filamentous species of green algae (Gibson and Whitton, 1987)

22. Brown Algae

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TABLE I Species of Brown Algae Reported from Freshwater Environments, with Morphology (UF = Uniseriate Filaments; CR = Crustose, MF = Multiseriate Filaments), Habitats, and Localities Taxon

Morphology

Habitat

Localities

Ectocarpales Bodanella lauterborni

UF

Lake

Ectocarpus siliculosusb

UF

Stream, estuary

Pleurocladia lacustris

UF

Stream, lake

Heribaudiella fluviatilisc

CR

Stream, lake

Porterinema fluviatile

UF

Lake

North America: unknown Other: Lake Constance,a Europe North America: unknown Other: Hopkins River, Australia North America: Green River (UT, CO), Devon Island ( NWT) Other: Austria, Germany, Poland, Scandinavia, England North America: at least 30 sites Other: many locations in Europe, also Japan, China North America: unknownd Other: Germany, Netherlands, United Kingdom

Sphacelariales Sphacelaria fluviatilis

MF

Stream, lake

MF

Lake

S. lacustris

North America: Gull Lake, MI Other: China North America: Lake Michigan Other: unknown

a

Known locally as Bodensee. Ectocarpus confervoides has been collected from the River Werra, Germany, polluted by potassium mines (Geißler, 1983). c Previously reported as Lithoderma arvernensis, L. fluviatile, and L. fontanum; L. zonatum (Jao, 1941) is retained by some authors. d Freshwater and euryhaline (= Pseudobodanella peterfii) reported from North America from marine and estuarine sites only.

b

and cyanobacteria (Sinclair and Whitton, 1977). A few species (e.g., Pleurocladia) become encrusted with CaCO3, which may cause the thallus to appear pale brown or gray macroscopically; microscopically carbonates may even cloak filaments in a crystalline tube (Kirkby et al., 1972; Kusel-Fetzmann, 1996). All known freshwater species have a diplohaplontic life history (both diploid and haploid vegetative phases) and most are isomorphic (diploid and haploid stages identical or very similar). For details on the alternation of generations in members of this division, see Papenfuss (1951) and Van den Hoek et al. (1995). Cellular features of freshwater brown algae are much like that of the division (Schloesser, 1977; Pueschel and Stein, 1983; West and Kraft, 1996); differences among genera are discussed later (Sect. V.B). Cells contain one to several golden-brown chloroplasts, which may be discoid, ribbon-like, or irregular-shaped, and usually parietal; pyrenoids are present in some species. The ultrastructure of freshwater species investigated (Heribaudiella, Sphacelaria) suggest typical phaeophyte features: thylakoids in triplets, chloroplast envelope consisting of four membranes, and plasmodesmata traversing crosswalls (Schloesser, 1977; Schloesser and Blum, 1980; Timpano, 1980; Pueschel and Stein, 1983). Most species possess numerous refractive bodies, including physodes, darkly pigmented bodies that may store phaeophycean tannins (fucosan), other

polyphenolics, and terpenes (Chadefaud, 1950; Graham and Wilcox, 2000; for cytological methods, see Sect. IV). When thalli are exposed to the air these tannins become oxidized and darken (Lee, 1989), giving dried or exposed colonies a dark brown or black color. Reproductive structures are distinctive for this group, but life cycles are incompletely known among the freshwater species. In general, species within the Ectocarpales and Sphacelariales produce two types of terminal sporangia: unilocular (large, single chamber) and plurilocular (multichambered), although in some species, only one of the two structures has been observed (Papenfuss, 1951; Hamel, 1931–1939; Bourrelly, 1981). Unilocular sporangia are typically produced on sporophytes (diploid), appear as large, ovate or clavate structures (Figs. 1A, C, 3F), and are the usual sites of meiosis. Initially unilocular sporangia (often arising from elongated filaments; Svedelius, 1930; Kumano and Hirose, 1959) contain several brown chloroplasts, which later condense. Following meiosis the sporangium produces (usually eight) biflagellate zoospores (or zooids), which are thought to serve as gametes (Kumano and Hirose, 1959; Müller and Geller, 1978). Plurilocular sporangia (Fig. 2F) are produced on either gametophyte (1n) or sporophyte (2n) plants, which divide repeatedly from erect narrow threadlike cells (e.g., Ectocarpus, Heribaudiella) or

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FIGURE 1 Three general morphologies exhibited by freshwater brown algae: (A) uniseriate, branched filaments forming cushion-like tufts (e.g., Pleurocladia); (B) branched prostrate filaments giving rise to tightly packed, vertical filaments, forming crustose thalli (e.g., Heribaudiella); (C) multiseriate branched filaments (quasicorticated), forming spreading cushions (Sphacelaria).

terminal branches (e.g., Bodanella) to form multicellular structures that produce asexual zoospores or zoospores that later settle and germinate to produce new filaments. In Heribaudiella and Porterinema, biflagellate zoospores are pear-shaped with two laterally inserted flagella, and possess a single parietal chloroplast and an apical stigma (Kumano and Hirose, 1959; Dop, 1979). Dispersal of freshwater phaeophytes is likely favored by zoospores released from unilocular sporangia. After release they attach to available substrata and

form germination tubes that later form filaments that develop the typical prostrate or disclike basal system (Yoshizaki et al., 1984). More recent success in isolating some freshwater phaeophytes into pure culture [Bodanella; Müller and Geller (1978); Pleurocladia; Kusel-Fetzmann and Schagerl (1992) and KuselFetzmann (1996); and Ectocarpus, West and Kraft (1996)] offers promise in addressing questions of relatedness among genera and species, dominant reproductive or ploidy phases, morphological plasticity, and mechanisms of their reproduction and spread. For

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FIGURE 2 Freshwater phaeophytes, Pleurocladia, Bodanella, Ectocarpus (scale bars = 10 µm): (A) and (B) differences in hair formation by Pleurocladia lacustris in response to P-rich (A, +22 µM NaH2PO4) and P-limited (B, no P added) conditions in culture; hairs (B, upper) are terminal, hyaline, multicellular filaments 100–300 µm long; (C) and (D) Bodanella lauterborni, showing detail of cells and branching pattern (C) and general morphology (D); (E) and (F) Ectocarpus siliculosus, showing sparse branching pattern, ribbon-like chloroplasts (E, photo by Jason Sonneman, with permission), and plurilocular sporangium (arrow) (F, photo by Jason Sonneman, with permission).

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FIGURE 3 Freshwater phaeophyte, Heribaudiella (scale bars = 10 µm, except where indicated): (A) and (B) macroscopic appearance of individual colonies (A, McKenzie River, Oregon) and coalescing colonies (B, Bonaparte River, British Columbia) on rocks; (C) prostrate filaments; (D) columns of vertical filaments following removal from rocks and pressure applied to the coverslip; (E) detail of cells and chloroplasts in vertical system; (F) unilocular sporangium.

22. Brown Algae

example, it is possible that in some freshwater species, portions of the typical phaeophyte life cycle may not occur. Further work on the reproductive ecology of freshwater brown algae based on field populations (as shown for the red alga Batrachospermum; Hambrook and Sheath, 1991), in conjunction with purified cultures, is clearly needed for members of this group.

III. ECOLOGY AND DISTRIBUTION A. Ecological Factors Due to the relatively sparse literature on freshwater brown algae, knowledge of the ecological factors governing their abundance and distribution is correspondingly limited. In some accounts, precise or even approximate locations of populations are difficult to discern (see methods, Sect. IV). However, despite these gaps in our knowledge, some generalizations can be made. All freshwater species are benthic and most are epilithic in habit, particularly Heribaudiella fluviatilis, which almost exclusively colonizes stones in streams or lakes. With this encrusting species, there appears to be a preference for more resistant rocks, such as basalt, quartz, schist, and gneiss (Allorge and Manguin, 1941; Wehr and Stein, 1985), although Kusel-Fetzmann (1996) also reported this alga colonizing bricks in one Austrian stream. Other taxa, such as Pleurocladia and Porterinema fluviatile, are fairly nonspecific with regard to substratum. Some are epiphytes on larger algae (e.g., Cladophora, Rhizoclonium) and macrophytes (Phragmites, Typha) or may colonize artificial substrata (Israelsson, 1938; Waern, 1952; Kirkby et al., 1972; Dop, 1979). Pleurocladia lacustris has been found attached to stones and boulders in the Green River (Utah–Colorado; Ekenstam et al., 1996) and in several streams in southern Austria (Kusel-Fetzmann, 1996). It may also be attached to reeds in the littoral zone of lakes, such as in Lake Wigry, Poland (Szymanska and Zakrys, 1990), Brasside Ponds, United Kingdom (Kirkby et al., 1972), and several Swedish lakes (Israelsson, 1938). Pleurocladia was also observed on glass slides that were placed in Lake Erken, Sweden (Kann, 1993). Porterinema fluviatile similarly colonizes reed stems, as well as submerged glass slides in eutrophic lakes (Dop and Vroman, 1976). With further study, the ecological breadth of this alga may prove to be quite broad, as earlier studies have reported it from brackish sites (0–8‰); and growing as an endophyte in Enteromorpha and Cladophora (Waern, 1952). As mentioned earlier, some of those with encrusting or entangled growth forms may also be complexed with CaCO3, particularly Sphacelaria spp. and P. lacustris (Israelsson, 1938; Waern, 1952; Kirkby

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et al., 1972; Schloesser and Blum, 1980). Although several species are found in both lakes and rivers, certain species, especially Heribaudiella fluviatilis, Pleurocladia lacustris, and Sphacelaria fluviatilis, appear to be best developed in flowing waters (Jao, 1943; Kann, 1978a; Wehr and Stein, 1985; Kusel-Fetzmann, 1996). Among those species that occur in running waters, most reports mention that freshwater brown algae occur in rocky, clear-water streams and are largely absent from turbid or muddy habitats (Budde, 1927; Fritsch, 1929; Allorge and Manguin, 1941; Jao, 1941; Chadefaud, 1950; Holmes and Whitton, 1975; Kann, 1966, 1978a, b; Starmach, 1977; Wehr and Stein, 1985; Yoshizaki and Iura, 1991; Kusel-Fetzmann, 1996). The most complete (although still fragmentary) ecological data for any freshwater phaeophyte is for Heribaudiella fluviatilis. In summarizing sites throughout Europe at the time, Israelsson (1938) noted that the species colonized a wide range of streams and lakes spanning oligotrophic to eutrophic conditions. More recent data suggest that this species most often occurs in stony streams, with moderately alkaline water (most ≥ pH 7.0), but fairly broad Ca, P, and N concentrations (Table II). Kann (1966, 1978a, b) reported that Heribaudiella is typical of “calcium-poor, summer warm streams,” although her data indicate that the species is rarely observed in either extremely softwater or hardwater systems, or at very low nutrient levels. Nearly all populations occupy clear-water habitats, although a few may tolerate moderate levels of humic materials (Kann, 1978a; Wehr and Stein, 1985), but not low pH, humus-rich waters (Israelsson, 1938). Although this alga is most commonly a lotic species, it has also been reported from rocky-shore habitats in some European lakes (Kann, 1945, 1993). Many studies from Europe and Japan (e.g., Budde, 1927; Fritsch, 1929; Geitler, 1932; Allorge and Manguin, 1941; Yoneda, 1949; Holmes and Whitton, 1975, 1977a; Kusel-Fetzmann, 1996) report that Heribaudiella often co-occurs with the encrusting red alga Hildenbrandia (particularly in shaded reaches), although this pattern is far from consistent. Holmes and Whitton (1977a, b, c) report both species in the rivers Tees, Swale, and Wear (United Kingdom), but each alga has also been recorded from sites where the other is apparently absent. Jao (1944) described Heribaudiella (as Lithoderma zonatum; see Sect. V.B.) as an indicator species for rapidly flowing, stony streams in China, along with the encrusting cyanobacterium Schizothrix (also Homoeothrix, Lemanea, Bangia, and several diatom species), but without Hildenbrandia. Kann (1978a, b) suggests that Heribaudiella and Hildenbrandia often occur separately because the latter tends to colonize more Ca-rich streams. Surveys of North American

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TABLE II Summary of Selected Ecological Conditions of Streams and Rivers from Three Regions in Which Heribaudiella fluviatilis Has Been Collected [N/A = Data Not Available; Based on Kann (1978a), Holmes and Whitton (1977a, b, 1981), Wehr and Stein (1985), Kusel-Fetzmann (1996), and Wehr (unpublished data)] Variable

Northern UK

Austria

North America

Light Substratum Geology Width (m) Depth (cm) Current velocity (cm s–1) Temperature (°C) Conductance (µS cm–1)a pH PO4–P (µg P L–1) NO3–N (µg N L–1) NH4–N (µg N L–1) Ca (µg L–1)

Open to partial shade Boulders, cobbles Basalt, sandstone 5–25 N/A Up to 1700 3–20 150–600 7.5–8.0 20–1300 200–1050 20–300 20–50

Shaded Boulders, cobbles, pebbles, bricks Quartz, basalt, marble 1–10 10–100 N/A 5–21 110–640 7.4–8.3 17–62 300–3600 20–280 30–45

Open to partial shade Boulders, cobbles Basalt, quartz, granite 1–60 10–100 200–1600 5–25 50–450 7.0–8.7 < 2.5–25 2–200 < 2–100 10–70

a

Corrected to 25°C.

streams thus far have encountered Heribaudiella only without Hildenbrandia (Wehr and Stein, 1985; Wehr, unpublished), but with other macroalgal species (e.g., Nostoc parmelioides, N. verrucosum, Cladophora glomerata). Kann (1945, 1978a) has also pointed out that several encrusting forms (e.g., Chamaesiphon, Gongrosira, Heribaudiella, Homoeothrix) may co-occur in turbulent streams, simply because they are well adapted to these habitats. Taken together, these community-level data suggest that so-called associations of algae described in earlier publications (e.g., Geitler, 1932; Waern, 1938; Luther, 1954) may not be consistent in different parts of the world. Some forms are equally common in lakes and rivers, and Pleurocladia lacustris is the most widely reported freshwater species from lentic systems (although it is also present in streams). This species colonizes a wide range of substrata, including stones, wood, many aquatic plant species, and artificial substrata, yet in freshwaters, it occurs in a fairly narrow range of chemical conditions. Early distribution data for Europe suggest that Pleurocladia prefers nutrientrich, strongly calcareous waters (Israelsson, 1938). Subsequent studies support this suggestion; nearly all reports of freshwater population sites from Europe and North America mention eutrophic conditions and the presence of CaCO3 precipitates associated with older colonies of Pleurocladia (Waern, 1952; Kirkby et al., 1972; Szymanska and Zakrys, 1990; Kann, 1993; Ekenstam et al., 1996; Kusel-Fetzmann, 1996; D. Ekenstam, pers. comm.). Limited chemical data report specific conductance levels > 600 µS cm–1 (Kirkby et al.,

1972; Kusel-Fetzmann, 1996) and its absence in nearby sites with conductance ⬇ 450 µS cm–1 (Kusel-Fetzmann, 1996). Algal species that often co-occur with Pleurocladia, including Gloeotrichia pisum, Rivularia spp., and Chaetophora incrassata, are also typical of nutrient-rich, hard waters (Kann, 1993). Pleurocladia has been observed in Hell Kettles, a series of very hardwater ponds in northern England (B. A. Whitton, pers. comm.), also colonized abundantly by Chara hispida and known for their especially rich marsh flora (Wheeler and Whitton, 1971). Pleurocladia may also occur in some sites with Heribaudiella, especially those with higher dissolved calcium (Kann, 1993; KuselFetzmann, 1996; D. Ekenstam, pers. comm.). Occurrences of P. lacustris in some brackish or intermittently marine habitats (Waern, 1952), including arctic sites in North America (Wilce, 1966), suggest that the alga may also be a euryhaline species. However, Waern (1952) noted that, despite a large number of freshwater Pleurocladia populations in northern Europe, no nearby marine (or even brackish) populations have been identified (for further discussion of disjunct distributions, see Sect. III.B). Incomplete knowledge of the ecological factors affecting Pleurocladia lacustris is undoubtedly due to a paucity of collections. KuselFetzmann (1996) pointed out that the alga was first discovered in Austria after observing “curious pale hairs” when examining crusts of Heribaudiella; it was an epiphyte on the other alga and produced small cushion-like colonies on rocks. Kirkby et al. (1972) comment that Pleurocladia was observed only on rotting (not recently dead) Typha leaves in the littoral

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zone of a eutrophic pond in England, whereas Waern (1952) observed that Pleurocladia may also grow as an endophyte in aquatic plants and macroalgae, which may lead to its under-reporting. Future studies require careful attention to the observations made by earlier researchers, as well as more complete collection of ecological data at these sites. The light requirements of freshwater phaeophytes have not been studied in detail, although scattered reports suggest that Heribaudiella may more often colonize shaded reaches of streams (Kusel-Fetzmann, 1996). Surveys of other rivers in North America and the United Kingdom have encountered this alga in habitats spanning a broad range of light environments, from small shaded streams (< 1 m wide) to wide rivers (> 50 m) with little or no shade (Holmes and Whitton, 1977a, b, c; Wehr and Stein, 1985). In contrast, Bodanella lauterborni has been collected only from deep epilithic habitats (> 15 m) on limestone rocks in lakes (Geitler, 1928; Müller and Geller, 1978; Kann, 1982). Similarly, the only known populations of Sphacelaria lacustris have been collected from deep (5–15 m), poorly illuminated areas of the sublittoral region of western Lake Michigan, and not in shallower habitats (Schloesser and Blum, 1980). Culture studies with this alga found that optimum growth and reproduction (gemmae-like propagules and unilocular sporangia) was achieved under reduced (screened) light levels (reported as 1000 lux) and short-day (8L : 16D) conditions (Schloesser, 1977; Schloesser and Blum, 1980). Very little is known of the competitive abilities or community importance of brown algae in freshwater habitats. Many populations of Heribaudiella fluviatilis colonize and may completely encrust certain rocks, with few other species present (Holmes and Whitton, 1975; Wehr, unpublished data). Heribaudiella may overgrow crusts of Hildenbrandia rivularis in streams where they co-occur (Fritsch, 1929; Geitler, 1932; Kusel-Fetzmann, 1996). In running waters, Sphacelaria fluviatilis apparently also grows in nearly monospecific stands (Jao, 1944). Thalli of Heribaudiella are usually free of epiphytes, but may occasionally serve as a host for diatoms, small cyanobacterial species (Chamaesiphon incrustans, Homoeothrix varians), chantransia stages of red algae, and Pleurocladia lacustris (Svedelius, 1930; Pueschel and Stein, 1983; Kusel-Fetzmann, 1996). The influence of herbivores on any freshwater species is unknown, although studies of marine species suggest that the large quantities of polyphenolics produced by many phaeophytes (> 2% of dry mass) may inhibit herbivore activity (Targett and Arnold, 1998). Field and lab herbivory experiments are clearly needed for freshwater species.

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B. Geographical Distribution Despite several hundred documented populations of freshwater phaeophytes recognized world wide, knowledge of their distribution and biogeography is fragmentary. The paucity of information on these algae makes each discovery of a new locality still worthy of publication, and renews speculation as to their origins (e.g., West, 1990; Kusel-Fetzmann, 1996; Wujek et al., 1996). Some may still be safely regarded as rare (or at least very poorly known), whereas others are world wide in their distribution. Many species distributions are regarded as disjunct, such as Sphacelaria fluviatilis, an epilithic species with only two known locations, a stream in south-central China and a small lake in Michigan (Jao, 1943; Thompson, 1975; Wujek et al., 1996). Sphacelaria lacustris is thus far known only from western Lake Michigan (Schloesser and Blum, 1980). Bodanella lauterborni is apparently known from only three locations, all in western Europe (Bourrelly, 1981). Heribaudiella fluviatilis may be the most widespread of all freshwater phaeophytes, occurring in many locations in Europe, western North America, Japan, and China (Wehr and Stein, 1985; KuselFetzmann, 1996), although there are currently no records from Africa, South America, Australia, or New Zealand, despite many phycological studies in these areas. More recent surveys of more than 250 river reaches for H. fluviatilis in North America have located 30 populations, all within western coniferous or boreal forests, and none have been located in any biomes east of the Mississippi River or south of Oregon (Wehr, unpublished). The discovery of Heribaudiella from a stream near Yellowknife, Northwest Territories (Sheath and Cole, 1992), extends its distribution more than 1000 km north (11°N latitude). No populations are known from Mexico or Central America, although given the diversity of freshwater habitats in these regions (see Chap. 2), it is reasonable to expect this alga in rocky streams from these regions as well. Comments more than 70 years ago by Budde (1927) and Fritsch (1929) that this species is easily missed are still true. Because more than 99% of all known species within the Phaeophyceae occupy marine habitats, questions often focus on the possible dispersal and adaptation of taxa from marine to freshwater habitats. A review of the published ecological information for all freshwater phaeophytes suggests little evidence of (at least recent) marine invasions by most species. One reason is that at least four of the seven recognized species from fresh waters have no counterparts in marine environments. Two exceptions, Ectocarpus siliculosus and Porterinema fluviatile, seem to be true euryhaline species that colo-

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nize a broad range of salinities, including fresh waters (Dop, 1979; West and Kraft, 1996). Pleurocladia lacustris has been almost entirely reported from freshwater locations distant from the ocean, but at least one North American location is intermittently saline, and morphological data strongly indicate that these populations are the same species (Wilce, 1966). Current biogeographic data, however, show no patterns that suggest a marine invasion. Analyses of Pleurocladia distribution patterns in northern Europe (Israelsson, 1938; Waern, 1952) indicate a complete lack of marine populations in a region where freshwater populations are most common. With additional freshwater populations in Austria, France, and the Ukraine, data suggest that the freshwater history of this species may be quite old, perhaps pre-glacial (Waern, 1952). The discovery of Pleurocladia lacustris in the Green River (Utah, Colorado) more than 1000 km from any marine water (Ekenstam et al., 1996), is in agreement with reports from European sites (Szymanska and Zakrys, 1990; Kusel-Fetzmann, 1996). Similarly, the North American distribution of Heribaudiella fluviatilis in the Northwest Territories, British Columbia, Washington, Montana, Oregon, and Utah shows a near absence of the alga from river sites near the coast and an abundance of populations in interior and upland regions (Wehr and Stein, 1985; West, 1990; D. Ekenstam, pers. comm.). In addition, none of the inland populations of Heribaudiella, Pleurocladia, or Sphacelaria in North America or Europe are reported to be influenced by elevated salinity. Finally, there are no studies that have yet identified marine species of Phaeophyta from inland saline lakes, which are scattered across most continents. An exception may be found in Ectocarpus siliculosus, which was discovered in a waterfall of the Hopkins River (Australia), roughly 40m above sea level, with a specific conductance of 3.0 mS cm–1; an isolate of this population has been shown to tolerate a wide range of salinities in culture (West and Kraft, 1996). As this is the first documented case of a population of Ectocarpus from any freshwater site, it is too early to speculate on the causes for its distribution. An intriguing but obscure species, Porterinema fluviatile, has been sampled from many brackish and freshwater sites in Europe (Waern, 1952), whereas Wilce et al. (1970) described a North American population from a freshwater site adjacent to a salt marsh in Massachusetts. It has since been sampled from freshwater sites in Netherlands and isolated into culture, using Wood’s Hole freshwater medium (Dop, 1979). The ecological and distributional history of this species complex clearly requires further attention. Molecular analyses (e.g., 18S rRNA and rbcL genes) in conjunction with biogeo-

graphic studies of most freshwater phaeophytes are sorely needed to elucidate distribution patterns and genetic relationships among these apparently disjunct populations.

IV. METHODS FOR COLLECTION AND IDENTIFICATION Because all known freshwater brown algae are benthic, methods used for sampling or removing various substrata are needed (e.g., Weitzel, 1979; Stevenson, 1996). Also, most species are macroalgal, that is, colonies or filaments which are recognizable (if not easily identified) with the naked eye. Nonetheless, freshwater phaeophytes are still cryptic and difficult to find. A survey of > 1000 stream segments in North America (Sheath and Cole, 1992) located only one additional population of Heribaudiella fluviatilis, from a stream in the Northwest Territories. In most investigations, new populations are discovered by researchers who have encountered the species nearby or elsewhere (Wehr and Stein, 1985; Kusel-Fetzmann, 1996). Because colonies or thalli of benthic algae may be inconspicuous in the field, Sheath (Chap. 5) recommends the use of a plastic view box, which provides a clear view through calm water and allows the investigator to distinguish different growth forms, pigmentation, and microhabitats much more easily. The present author has found this device reduces search time in both streams and shallow littoral areas of lakes. If quantitative samples are needed, transects (along tape measures sampled at regular intervals) or quadrats may be required. With macroalgae some authors may use visual estimates of cover for visually distinct physiognomies (e.g., Holmes and Whitton, 1977a, c; Wehr and Stein, 1985; Sheath and Cole, 1992) and later assign species names based on microscopic examination. This chapter thus provides descriptions of genera (Sect. V.B.) based on macroscopic and microscopic appearances, which may be helpful in field sampling. Our understanding of the distribution and ecology of freshwater brown algae requires many more thorough surveys. With the development of inexpensive global positioning systems (GPS), relatively precise geographic information (latitude, longitude, altitude) should be relatively easy to collect in all future studies. Whenever possible, ecological data, particularly type of substratum and size, current velocity, irradiance, temperature, conductance, turbidity, and water chemistry (especially pH, N, P, Ca), should be measured in all collections and surveys of freshwater phaeophytes. In addition, simple relative scales of abundance or cover estimates (Wehr and Stein, 1985; Sheath and Cole, 1992) will

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greatly aid future syntheses of the ecology of this group of algae. In rivers, surveys have also been conducted by wading specific lengths (10-m to 0.5-km reaches) and recording presence or absence, relative abundance, or percentage of cover estimates of each macroalga from that area (Holmes and Whitton, 1977a, b). A field microscope (e.g., Swift Instruments, San Jose, CA) is helpful in distinguishing colonies or crusts that appear similar in the field (see also Holmes and Whitton, 1975). Methods for removal of material from substrata depend on the growth form and type of substratum. Encrusting or firmly attached epilithic species (mainly Heribaudiella, Sphacelaria) are best sampled by removing entire rocks when possible, then scraping material (using a razor blade) into vials for later identification, whereas smaller stones may be transported intact. It is important to note the macroscopic appearance (shape, margin, size, thickness) and color of the colonies in the field during sampling, as material may change even during brief storage times. Most epiphytic forms (e.g., Pleurocladia) in fresh water are usually firmly attached to plant hosts, permitting removal of plants or plant fragments with dip nets or collected using SCUBA. However, more delicate or gelatinous forms may be best sampled using forceps or a modification of the half-bottle sampler (Douglas, 1958), which can isolate water plus algal material and permit removal of thalli without loss. Kann (1976, 1978a) has also detected some species of brown algae (Pleurocladia) from lakes by observing settled zoospores on artificial (glass, plastic) substrates, although their selectivity for or against freshwater phaeophytes is unknown. Material is best examined live and soon after collection for recognition of pigmentation, chloroplast form, and presence of hairs. Algal samples can be kept alive for several days if stored cool (5–10°C) and wet or moist. Some authors (Waern, 1952; Kusel-Fetzmann, 1996; West and Kraft, 1996) have reported that filamentous forms retain their normal growth form in sample water for several weeks or months, and some species (as discussed earlier) can be brought into culture using standard media (Müller and Geller, 1978; Schloesser and Blum, 1980; Kusel-Fetzmann and

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Schagerl, 1992). Identification of some species frequently requires reproductive structures (Sect. V). Their position, number, and arrangement are diagnostic, but there is some doubt whether shape of sporangia may be used (Waern, 1952; West and Kraft, 1996). Motile stages are readily produced with field material or in culture if maintained under moderate (10–20°C) temperatures (Kumano and Hirose, 1959; Müller and Geller, 1978; Wehr, unpublished). Long photoperiods (16 : 8) apparently favor formation of plurilocular sporangia (and zooids) in Porterinema (Dop, 1979), whereas short-day conditions (8 : 16) may induce the production of unilocular sporangia in Sphacelaria lacustris (Schloesser and Blum, 1980). If samples are to be stored for long periods, excised specimens are best preserved using 2–3% glutaraldehyde or 2% paraformaldehyde, and stored cool and in the dark. Alternatively, encrusting species may be “preserved” on rocks (and suitable for herbaria) by simply air-drying the entire rock, although cells and plastids tend to become distorted upon drying. Isolates of Sphacelaria lacustris from Lake Michigan that were grown on agar plates, later were air dried to produce thin, dry specimens that were stored on herbarium sheets (Scholesser and Blum, 1980). Inspection of this material (holotype in U.S. National Herbarium, Algal Collection) more than 20 years later found that most of the morphological and cellular features were retained (Wehr, unpublished). When preparing thalli for light microscopy, specimens of crustose species may require vigorous chopping (with razor blade) and/or crushing (with cover slip) to separate densely packed filaments. The morphology of filamentous forms may also be more clearly revealed in squash preparations. Species that have become calcified with CaCO3 (especially Pleurocladia, Porterinema) may require treatment with dilute acid (1–2% HCl) before observation. Stains may be used to emphasize important structures, particularly cresyl blue (Chadefaud, 1950; Schloesser and Blum, 1980), which may be used to demonstrate physodes (which store phaeophycean tannins). Vanillin–HCl may also be used, which stains physodes red (Chadefaud, 1950: Lee, 1989).

V. KEY AND DESCRIPTIONS OF GENERA A. Key Taxa not reported from North America in fresh waters are marked with an asterisk. 1a.

Thalli small cushion-like tufts or expanses of spreading filaments (Fig. 1A, C)........…..……..…..……..…..………..…….....…..……..2

1b.

Thalli not cushion-like (Figs. 1B, 3C); creeping or crustose…………………..……..…..……...............................................................3

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John D. Wehr

2a.

Filaments uniseriate and multiseriate; with numerous disc-shaped chloroplasts (Figs. 1C, 4E, F)...........……………..….…Sphacelaria

2b.

Filaments uniseriate only; single (rarely two) chloroplasts; basal filaments curved or arching; erect system spreading (comblike) (Fig. 4A, B) ....…………………..……..…..……..…..………..……..…..……..…..………..……..…..……..…..…….............Pleurocladia

3a.

Thalli crustose, forming dark brown patches on stones; branched basal filaments with short, densely packed upright filaments (Figs. 1B, 3) ……….….…………………..……..…..……..…..………..……........…..………..……..…..……..…..…………Heribaudiella

3b.

Thalli not crustose, spreading or simple, variously branched.............………………..……..…..……..…..…................…………….….4

4a.

Filaments sparingly branched, vegetative cells narrow, cylindrical, chloroplasts several, ribbon-like; plurilocular sporangia (if present) narrow-elongate (Fig. 2E)…..……..…..……..…..………..……..…..……..…..………..……..…..……..…..……...........Ectocarpus*

4b.

Filaments frequently or irregularly branched, cells inflated or quadrate, chloroplasts few to several, plurilocular sporangia unknown or (if present) broad or inflated in shape…..……..…..……..…..…….……..……..…..……..…..………..……..…........……..…..……..5

5a.

Branched filaments with prostrate and erect forms ….…….….……………………..……..…..……..…..………..……..…..……....…...6

5b.

Branched filaments prostrate only, creeping along substrata; may have rhizoid-like branches, chloroplasts many Fig. 2C, D) ….…………………………………………………..……..…..……..…..……..........................…..……..…..……..…..…………Bodanella*

6a.

Basal filaments often curved or arching, erect filaments spreading; cells with single (rarely two) chloroplast, unilocular sporangia ovoid.....………………………..……..…..……..…..………..……..…..……..…..………..……..…..……..…..……...........….Pleurocladia

6b.

Basal and erect filaments irregularly arranged, with two parietal, lobed chloroplasts; plurilocular sporangia in crown-shaped clusters of four or more (Fig. 4C, D)…..……..…..……..…..………..……..…..……..…..………..……..…..……..…....……Porterinema

B. Descriptions of Genera Ectocarpales Bodanella Zimmermann (Fig. 2C, D) Thalli basal or creeping filaments form on rocky substrata, without erect filaments. Filaments are uniseriate, frequently but irregularly branched, composed of irregularly shaped cells; inflated, quadrate, angular, ovoid, or “wavy” in shape; vegetative cells are 10–16 µm wide, 10–25 µm long. General form may be confused with Sphacelaria lacustris, but the only latter genus possesses multiseriate axes; filaments of Bodanella are uniseriate. Terminal, short, narrow hairs (6–10 µm diameter), or basal rhizoid-like filaments may also be present. Parietal chloroplasts are small, numerous (10–15 per cell), and discoid. Unilocular sporangia are ovoid or globose; 15–20 µm wide by 25–30 µm long. Zoospores are pyriform (10–12 µm by 5–6 µm), with laterally inserted flagella. Plurilocular sporangia are unknown. A monotypic genus, Bodanella lauterborni was named for its original location, Bodensee (Lake Constance, Austria–Germany), where it colonizes limestone deep (15–35 m) in lakes (with Hildenbrandia and Cladophora). It is not known from North America; worldwide distribution consists of three European lacustrine populations, in Lake Constance (Zimmermann, 1928; Müller and Geller, 1978), Lunzer Untersee (Austria; Geitler, 1928), and Traunsee (Austria; Kann, 1982). Ectocarpus Lyngbye (Fig. 2E, F) The freshwater form is sparingly or irregularly

branched; thalli are mostly erect filaments consisting of cylindrical (isodiametric) cells 15–40 µm diameter, up to four times as long as broad. Narrow, hair-like filaments also are present (8–12 µm diameter). Chloroplasts are several (2 to 4?), ribbon-like, lobed, and parietal, with pyrenoids, arrangement variable (netlike, spiral, etc.). Plurilocular sporangia are terminal, narrowly ellipsoid, conical, or linear, with numerous divisions; length is 70–200 (500) µm by 15–35 µm diameter. Unilocular sporangia are unknown from freshwater populations, but well established in marine population (Lee, 1989). Ectocarpus is an ecologically and geographically widespread marine and estuarine genus (Müller, 1979), recently discovered (E. siliculosus Dillw.) in a freshwater waterfall of the Hopkins River (Australia) with other freshwater taxa (Mougeotia, Cladophora; West and Kraft, 1996). This is the only record of any freshwater phaeophyte in the southern hemisphere, according to Entwisle et al. (1997). The alga is unknown from freshwater sites in North America, but is common in coastal brackish sites on this continent. It was also observed (E. confervoides) with freshwater taxa (e.g. Anabaena, Cyclotella, Scenedesmus) in the River Werra (Germany) in sites polluted by potassium mine waste (Geißler 1983). Heribaudiella Gomont (Fig. 3A–F) Thalli are olive-brown to dark brown crusts on rocks in streams and lakes; colonies are 1–5 cm (up to ⬇ 20 cm) diameter with rounded or irregular outline (Fig. 3A), but with distinct margins (Chamaesiphon spp. also forms brown crusts, but margins are indis-

22. Brown Algae

FIGURE 4 Freshwater phaeophytes, Pleurocladia, Porterinema, Sphacelaria (scale bars = 10 µm, except where indicated): (A) and (B) Pleurocladia lacustris, detail of filament and chloroplasts, showing centrifugal or arched growth patterns, with unilocular sporangium (arrow) (A); and erect, branched filaments, showing one or two parietal chloroplasts per cell (B, photo by E. L. Kusel-Fetzmann, with permission); (C) and (D) Porterinema fluviatile, showing irregularly branched filaments with short erect filaments (C) and clusters of filaments with apical, unilocular sporangia (arrows); (E) and (F) Sphacelaria lacustris (holotype, from Lake Michigan) [(E) filament showing complex, multiply branched, multiseriate growth form, (ch) many small chloroplasts and (P) physodes (photo reproduced with permission of the Journal of Phycology from Schloesser and Blum (1980) 16:201–207, Fig. 5); (F) detail of multiseriate primary axis; note cell divisions in two planes (from herbarium specimen)].

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tinct smudges or flecks; Holmes and Whitton, 1975; Wehr and Stein, 1985). Multiple colonies may coalesce to cover entire rocks or boulders (Fig. 3B). When scraped off carefully, colonies appear as a series of vertical columns at low magnification (Fig. 3D). Filaments in basal system are repeatedly branched (Fig. 3C); erect filaments are sparingly and dichotomously branched. Heribaudiella forms an erect system of appressed vertical filaments that do not easily separate under pressure; cells are mostly quadrate, 8–15 µm diameter, 5–15 cells long. Chloroplasts are oval or discoid, numerous (4–10 per cell; Fig. 3E); physodes are present. Multicellular hyaline hairs may be present (up to 1 mm long). Unilocular sporangia are terminal, inflated, ovoid or clavate (Fig. 3F); 10–25 µm wide, 15–35 µm long [development described by Svedelius (1930), Fig. 4]. Biflagellate zoospores are pyriform or irregular shape (⬇ 6–8 µm). Plurilocular sporangia are obscure (reported as rare), produced terminally in narrow-celled columns four (rarely eight) cells tall; immature plurilocular sporangia are difficult to distinguish from smaller vegetative filaments. Svedelius (1930) united several taxa (e.g., Lithoderma fluviatile, L. fontanum) under this name. Lithoderma zonatum was described from fresh waters by Jao (1941), based on frequent plurilocular sporangia (reported occasionally in H. fluviatilis) and several layers or zones of erect filaments; this taxon was still reported by Bourrelly (1981). Jao (1941) was apparently unaware of Svedelius’ study, because this layered morphology is exactly shown in his earlier study (Svedelius, 1930, Fig. 13) and was suggested to perhaps represent annual layers. For these reasons, the Chinese species (L. zonatum Jao) should be regarded as synonymous with Heribaudiella fluviatilis (Aresch.) Sved. No freshwater taxa are now assigned to the genus Lithoderma. A monotypic genus, Heribaudiella fluviatilis is the most widely observed freshwater phaeophyte worldwide, reported from at least 30 locations in western North America, but no extant populations are known east of the Mississippi or south of Oregon. A very early record (Collins et al., 1898; as Lithoderma fluviatile) from Island Brook, Connecticut, was included in the Phycotheca Boreali-Americana, although its identity has been questioned due to its possible marine habitat (Smith, 1950) and appearance of some samples (Pueschel and Stein, 1983). Further examination by the present author of the 1888 material deposited in the New York Botanical Garden, University of Michigan, and U.S. National (Smithsonian) herbaria suggests this population may in fact be Heribaudiella fluviatilis, based on morphology (vertical series of erect filaments) and co-occurring diatoms (> 99% were freshwater species; Wehr, unpublished data). However, surveys of

Island Brook in 1998 and adjacent streams failed to find an extant population in the area (Wehr, unpublished). In North American streams it often co-occurs with Audouinella hermannii, Chamaesiphon spp., Nostoc parmelioides, N. verrucosum, and Cladophora glomerata; in Europe, Hildenbrandia rivularis may colonize the same rock. Pleurocladia A. Braun (Figs. 2A, B, 4A, B) Thalli are small, brown to pale brown or tan (depending on calcification) hemispherical tufts or cushions (up to 3 mm diameter by 100–300 µm tall) on rocks or attached to plants (angiosperms, mosses, macroalgae), or endophytic, present in both streams and lakes. Waern (1952) reports that colonies appear macroscopically like a brown-colored Gloeotrichia. In less calcareous regions, colonies may be gelatinous. At low magnification (100⫻), colonies appear as a dense network of radiating, branched filaments (Fig. 2B). Two filament systems are evident: (1) creeping or basal system, with (infrequently) branched filaments usually consisting of rounded or inflated cells 8–16 µm diameter (occasionally elongate), often exhibiting centrifugal or arched growth patterns (Fig. 4A), which give rise to (2) upright, irregularly (alternate or opposite) branched long filaments (Fig. 4B), which are usually narrower (6–12 µm), elongate (cells 12–35 µm long), and more nearly isodiametric. Vegetative cells contain one (rarely two) large golden-brown parietal chloroplast (with pyrenoids); darker granules (physodes?) and refractive (lipid?) bodies may be common. Unilocular sporangia are common, single, clavate or globose [15–30 µm in diameter by 25–60 (–80) µm long], and borne laterally or terminally (Fig. 1A). Plurilocular sporangia are uncommon in freshwater populations (but see Waern, 1952), linear-elongate, narrow. Long (100–300 µm) multicellular hairs (5–7 µm diameter) are common in field populations (environmentally induced; Fig. 2A, B), arising from upright filaments, giving the colony a fuzzy appearance when viewed macroscopically. Pleurocladia lacustris is currently known from a few freshwater sites in North America [Wyoming, Colorado (Green River), and Devon Island, Northwest Territories]; suitable freshwater habitats undoubtedly occur in other locations on this continent. Many more populations are known from Europe. The species has also been recorded from marine and brackish habitats. The relationship between P. lacustris and related species and genera are discussed by Waern (1952), Wilce (1966), and Bourrelly (1981). Porterinema Waern (Fig. 4C, D) Thalli are monostromatic, brown disc-shaped plates of loosely arranged filaments. The genus occurs

22. Brown Algae

as an epiphyte on or endophyte in other algae (e.g., Rhizoclonium, Enteromorpha) or macrophytes (e.g., Elodea); it may also colonize stones and artificial substrata (e.g., glass slides). Thalli are creeping, composed of irregularly branched filaments, with short (a few cells) erect filaments produced infrequently (Fig. 4C). Basal cells are barrel-shaped, or occasionally enlarged on proximal ends (6–12 µm diameter by 6–12 µm long); erect cells are few but more elongate (up to 40–50 µm long). Vegetative cells occur with one to three lobed, golden-brown, parietal chloroplasts. Terminal, multicellular hairs (3–8 µm diameter; up to 200 µm long) are common and may be sheathed at their base. Unilocular sporangia are rarely reported, on basal or erect filaments, pear- or club-shaped (15–30 µm wide; up to 80 mm long). Plurilocular sporangia (6–8 µm diameter) are common, intercalary (occasionally terminal), typically four-celled clusters (or “crowns”) on pedicels (short filaments) or sessile; sometimes they are produced in clusters of up to 32 sporangia. One species, Porterinema fluviatile, is distributed mainly among brackish sites in Europe and North America, but several truly freshwater sites are known in Europe (Waern, 1952; Dop, 1979), and one site in North America: a stream draining into a salt marsh near Ipswich, Massachusetts (Wilce et al., 1970). As such it should be regarded as part of the North American freshwater algal flora, but requires further study. The report of a new genus, Pseudobodanella peterfii in Europe (Gerloff, 1967), appears to be identical to Porterinema fluviatile, lacking only hairs, and is very likely synonymous (Bourrelly, 1981; D. M. Müller, pers. comm). Bourrelly (1981) and Dop (1979) suggest that past records of Apistonema pyrenigerum (previously classified in Chrysophyceae), such as from a small pond in the United Kingdom (Belcher, 1959), are also Porterinema. Other synonymous or related taxa (e.g., Apistonema expansum, Porterinema marina) have been similarly considered by Wilce et al. (1970) and Dop (1979).

Sphacelariales Sphacelaria Lyngbye (Fig. 4E, F) Freshwater thalli are small (1–2 mm) brown tufts or cushions on rocks in streams or lakes; they may be calcified. Vegetative growth is the result of basal (creeping) and erect filaments; rhizoidal cells form where basal filaments contact substrata. The genus is distinguished by axes variably multiaxial (biseriate or multiseriate) and uniaxial (uniseriate); branches are hemiblastic (primordial cells arising from upper position), resulting in apical growth pattern. Branching pattern is irregular (S. lacustris) or opposite (S. fluviatilis); it may become pseudoparenchymatous. Cells

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comprising the main axis are rectangular or inflated (12–25 µm diameter); they are broader prior to lateral cell division, cylindrical on erect filaments. Cells contain numerous (10–20), small (3–8 µm), peripheral, discshaped chloroplasts and physodes (especially meristematic regions); pyrenoids are lacking. Multicellular hairs (> 500 µm length) develop from basal and erect filaments in some plants. Unilocular sporangia are known in one species (S. lacustris; see Schloesser and Blum, 1980, Fig. 3). Plurilocular sporangia are unknown in fresh waters for either species. Clusters of vegetative, gemmae-like propagules common; sessile or borne on short (1- or 2-celled) branches. Two freshwater species are known: S. lacustris, reported on rocks in western Lake Michigan at depths of 5–15 m (Schloesser and Blum, 1980); S. fluviatilis, reported in rapidly flowing water in the Kialing River, China (Jao, 1943, 1944), and in the shallow (≤ 1 m) littoral zone of Gull Lake, Michigan (Thompson, 1975; Timpano, 1978, 1980; Wujek et al., 1996). The two species were separated largely on the basis of branching pattern (alternate or irregular in S. lacustris, opposite in S. fluviatilis), lateral cell divisions (infrequent and irregular in S. lacustris, two to four regular divisions in main axes of S. fluviatilis), and hairs (lacking in S. fluviatilis?) (Schloesser and Blum, 1980). Further studies are needed on the ecological requirements, geographic distribution, and genetic differences of the two species. No populations are yet known from Europe, Central America, or South America.

VI. GUIDE TO LITERATURE FOR SPECIES IDENTIFICATION All but one freshwater phaeophyte genus (Sphacelaria) are monotypic; however, the primary literature is still helpful for identification. A few general keys are useful for most freshwater species (Starmach, 1977; Bourrelly, 1981), but the reader should be aware that recent combinations (e.g., Pseudobodanella = Porterinema) are not included. Many of the most complete descriptions may be older literature, come from other continents, or are in German or Japanese. 1. Bodanella—Zimmermann (1928), Müller and Geller (1978) 2. Ectocarpus—West and Kraft (1996) 3. Heribaudiella—Holmes and Whitton (1975), Yoshizaki et al. (1984), Kusel-Fetzmann (1996) 4. Pleurocladia—Wilce (1966), Kirkby et al. (1972), Kusel-Fetzmann (1996) 5. Porterinema—Waern (1952), Dop (1979) 6. Sphacelaria—Jao (1941), Schloesser and Blum (1980), Wujek et al. (1996)

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ACKNOWLEDGMENTS I express my thanks to Dean Blinn, Janet Stein, and Brian Whitton, who helped instill in me an enthusiasm for stream algae. Advice, samples, records, and/or photographs from Robert Wilce, Dieter Müller, John West, Elsa Kusel-Fetzmann, Devon Ekenstam, Robert Sims, and Willem Prud’homme van Reine were especially helpful in the preparation of this manuscript. Loans of herbarium specimens from the National Herbarium (Smithsonian), University of Michigan, New York Botanical Garden, and Trinity College, Connecticut, were extremely useful. Thanks also to many people who have helped in my searches for freshwater phaeophytes, Deb Donaldson, Syd and Dick Cannings, Bob Sheath, Greg Flannery, and Alissa Perrone.

LITERATURE CITED Allorge, P., Manguin, E. 1941. Algues d’eau douce des Pyrénées basques. Bulletin de la Société Botanique de France 88:159–191. Belcher, J. H. 1959. Some uncommon Chlorophyceae from the Lee Valley. British Phycological Bulletin 1:73–74. Bold, H. C., Wyne, M. J. 1985. Introduction to the algae. 2nd ed. Prentice-Hall, Englewood Cliffs, NJ, 720 pp. Bourrelly, P. 1981. Les algues d’eau douce, Vol. II. Les algues jaunes et braunes, Chrysophycées, Phaeophycées, Xanthoophycées, et Diatomées, rev. ed. Soc. Nouvelle des Éditions Boubée, Paris. Braun, A. 1855. Decade XLV + XLVI, in: Rabenhorst, L., Ed. (1848–1860) Die Algen Sachsens, Respective Mittel-Europa’s. Dresden, Germany. Budde, H. 1927. Die Rot- und Braunalgen des Westfälischen Sauerlandes. Berischte der Deutschen Botanischen Gesellschaft 45:143–150. Chadefaud, M. 1950. Observations cytologiques sur la Phéophycée d’eau douce: Heribaudiella fluviatilis (Aresch.) Sved. Bulletin Société Botanique de France 97:198–199. Christensen, T. 1978. Annotations to a textbook of phycology. Botanisk Tidsskrift 73:65–70. Collins, F. S., Holden, I., Setchell, W. A. 1898. Phycotheca borealiAmericana, Vol. XI. Malden, MA, 536 p. Craigie, J. S. 1974. Storage products, in: Stewart, W. D. P., Ed., Algal physiology and biochemistry. Univ. California Press, Berkeley, pp. 206–235. Dop, A. J. 1979. Porterinema fluviatile (Porter) Waern (Phaeophyceae) in the Netherlands. Acta Botanica Neerlandica 28:449–458. Dop, A. J., Vroman, M. 1976. Observations on some interesting freshwater algae from the Netherlands. Acta Botanica Neerlandica 25:321–328 Douglas, B. 1958. The ecology of the attached diatoms and other algae in a small stony stream. Journal of Ecology 45:295–322. Ekenstam, D., Bozniak, E. G., Sommerfeld, M. R. 1996. Freshwater Pleurocladia (Phaeophyta) in North America. Journal of Phycology (Supplement) 32:15. Entwisle, T. J., Sonneman, J., Lewis, S. H. 1997. Freshwater algae in Australia. A guide to conspicuous genera. Sainty Associates, Potts Point, NSW, Australia. Fritsch, F. E. 1929. The encrusting algal communities of certain fastflowing streams. New Phytologist 28:165–196. Geibler, U. 1983. Die salzbelastete Flußstrecke der Werra—ein

Binnenlandstandort für Ectocarpus confervoides (Roth) Kjellman. Nova Hedwigia 37:193–217. Geitler, L. 1928. Über die Tiefenflora an Felsen im Lunzer Untersee. Archiv für Protistenkunde 62:96–104. Geitler, L. 1932. Notizen über Hildenbrandia rivularis und Heribaudiella fluviatilis. Archiv für Protistenkunde 76:581–588. Gerloff, J. 1967. Eine neue Phaeophyceae aus dem Süsswasser: Pseudobodanella peterfii nov. gen. et nov. spec. Revue Roumaine de Biologie—Botanique 12:27–35. Gibson, M. T., Whitton, B. A. 1987. Hairs, phosphatase activity and environmental chemistry in Stigeoclonium, Chaetophora and Draparnaldia (Chaetophorales). British Phycological Journal 22:11–22. Gomont, M. 1896. Contribution à la flore algologique de la HautAuvergne. Bulletin de la Société Botanique de France 43:373–393. Goodwin 1974. Carotenoids and biliproteins, in: Stewart, W. D. P., Ed., Algal physiology and biochemistry. Univ. California Press, Berkeley, pp. 176–205. Graham, L. E., Wilcox, L. W. 2000. Algae. Prentice-Hall, Upper Saddle River, NJ. Hambrook, J. A., Sheath, R. G. 1991. Reproductive ecology of the freshwater red alga Batrachospermum boryanum Sirodot in a temperate headwater stream. Hydrobiologia 218:233–246. Hamel, G. 1931–1939. Phaéophycées de France. Trait de Botanique 47. Paris, 432 p. + 10 pl. Holmes, N. T. H., Whitton, B. A. 1975. Notes on some macroscopic algae new or seldom recorded for Britain: Nostoc parmelioides, Heribaudiella fluviatilis, Cladophora aegagropila, Monostroma bullosum, Rhodoplax schinzii. Vasculum 60:47–55. Holmes, N. T. H., Whitton, B. A. 1977a. The macrophytic vegetation of the River Tees in 1975: Observed and predicted changes. Freshwater Biology 7:43–60. Holmes, N. T. H., Whitton, B. A. 1977b. The macrophytic vegetation of the River Swale, Yorkshire. Freshwater Biology 7:545–558. Holmes, N. T. H., Whitton, B. A. 1977c. Macrophytes of the River Wear: 1966–1976. Naturalist (Hull) 102:53–73. Holmes, N. T. H., Whitton, B. A. 1981. Phytobenthos of the River Tees and its tributaries. Freshwater Biology 11:139–163. Israelsson, G, 1938. Über die Süsswasserphaeophycéen Schwedens. Botanische Notiser 1938:113–128. Jao, C.-C. 1941. Studies on the freshwater algae of China. VII. Lithoderma zonatum, a new freshwater member of the Phaeophyceae. Sinensia 12:239–44. Jao, C.-C. 1943. Studies on the freshwater algae of China. XI. Sphacelaria fluviatilis, a new freshwater brown alga. Sinensia 14:151–154. Jao, C.-C. 1944. Studies on the freshwater algae of China. XII. The attached algal communities of the Kialing River. Sinensia 15:61–73. Kann, E. 1945. Zur Ökologie der Litoralalgen in ostholsteinischen Waldseen. Archiv für Hydrobiologie 41:14–42. Kann, E. 1966. Der Algenaufwuchs in einigen Bächen Österreichs. Verhandlungen—Internationale Vereinigung für Theoretische und Angewandte Limnologie 16:646–654. Kann, E. 1976. Algenaufwuchs unter natürlichen Bedingungen auf Kunststoffen. Chemie Kunststoffe Aktuell 2:63–71. Kann, E. 1978a. Systematik und Ökologie der Algen österreichischer Bergbäche. Archiv für Hydrobiologie (Supplement) 53:405–643. Kann, E. 1978b. Typification of Austrian streams concerning algae. Verhandlungen—Internationale Vereinigung für Theoretische und Angewandte Limnologie 20:1523–1526. Kann, E. 1982. Qualitative Veränderungen der litoralen Algenbiocönose österreichischer Seen (Lunzer Untersee, Traunsee, Attersee) im Laufe der letzen Jahrzehnte. Archiv für Hydrobiologie Supplement 62:440–490.

22. Brown Algae Kann, E. 1993. Der litorale Algenaufwuchs im See Erken und in seinem Abfluß (Uppland, Schweden). Algological Studies 69:91–112. Kirkby, S. M., Hibberd, D. J., Whitton, B. A. 1972. Pleurocladia lacustris A. Braun (Phaeophyta)—a new British record. Vasculum 57:51–56. Kumano, S., Hirose, H. 1959. On the swarmers and reproductive organs of a phaeophyceous fresh-water alga of Japan, Heribaudiella fluviatilis (Areschoug) Svedelius. Bulletin of the Japanese Society of Phycology 7:45–51 (in Japanese). Kusel-Fetzmann, E. L. 1996. New records of freshwater Phaeophyceae from lower Austria. Nova Hedwigia 62:79–89. Kusel-Fetzmann, E., Schagerl, M. 1992. Verzeichnis der Sammlung von Algen-Kulturen an der Abteilung für Hydrobotanik am Institut für Pflanzenphysiologie der Universität Wien. Phyton 32:209–234. Lee, R. E. 1989. Phycology, 2nd ed. Cambridge Univ. Press, Cambridge, UK. Luther, H. 1954. Über Krustenbewuchs an Steinen fliessender Gewässer, speziell in Südfinnland. Acta Botanica Fennica 55:1–66. Müller, D. G. 1979. Genetic affinity of Ectocarpus siliculosus (Dillw.) Lyngb. from the Mediterranean, North Atlantic and Australia. Phycologia 18:312–318. Müller, D. G., Geller, W. 1978. Einige Beobachtungen an Kulturen der Süsswasser-Braunalge Bodanella lauterborni Zimmermann. Nova Hedwigia 29:735–741. Papenfuss, G. F. 1951. Phaeophyta, in: Smith, G. M., Ed., Manual of phycology. Chronica Botanica, Waltham, MA, pp. 119–158. Pueschel, C. M., Stein, J. R. 1983. Ultrastructure of a freshwater brown alga from western Canada. Journal of Phycology 19:209–215. Schloesser, R. E. 1977. The identification of a new freshwater brown alga from the Lake Michigan sublittoral zone. M.Sc. thesis, University of Wisconsin, Milwaukee, 80 p. Schloesser, R. E., Blum, J. L. 1980. Sphacelaria lacustris sp. nov., a freshwater brown alga from Lake Michigan. Journal of Phycology 16:201–207. Sheath, R. G., Cole, K. M. 1992. Biogeography of stream macroalgae in North America. Journal of Phycology 28:448–460. Sheath, R. G., Hambrook, J. A. 1990. Freshwater ecology, in: Cole, K. M., Sheath, R. G., Eds., Biology of the red algae. Cambridge Univ. Press, Cambridge, UK, pp. 423–453. Sinclair, C., Whitton, B. A. 1977. Influence of nutrient deficiency on hair formation in the Rivulariaceae. British Phycological Journal 12:297–313. Smith, G. M. 1950. The fresh-water algae of the United States, 2nd ed. McGraw–Hill, New York. Starmach, K. 1977. Phaeophyta—brunatnice. Rhodophyta—krasnorosty. Flora s ⁄l odkowodna Polski, Vol. 14. Panstwowe Wydawnictwo Naukowe, Warsaw/Krakow. Stevenson, R. J., 1996. An introduction to algal ecology in freshwater benthic habitats, in: Stevenson, R. J., Bothwell, M. L., Lowe, R. L., Eds., Algal ecology: Freshwater benthic ecosystems. Academic Press, San Diego, pp. 3–30.

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Svedelius, N. 1930. Über die sogenannten Süsswasser-Lithodermen. Zeitschrift für Botanik 23:892–918. Szymanska, H., Zakrys, B. 1990. New phycological records from Poland. Archiv für Hydrobiologie Supplement 87:25–32. Targett, N. M., Arnold, T. M. 1998. Predicting the effects of brown algal phlorotannins on marine herbivores in tropical and temperate oceans. Journal of Phycology 34:195–205. Thompson, R. H. 1975. The freshwater brown alga Sphacelaria fluviatilis. Journal of Phycology (Supplement) 11:5. Timpano, P. 1978. A preliminary report of the fresh-water phaeophyte, Sphacelaria fluviatilis. Journal of Phycology (Supplement) 14:34. Timpano, P. 1980. The ultrastructure of the fresh-water phaeophyte, Sphacelaria fluviatilis. Journal of Phycology (Supplement) 16:44. Van den Hoek, C., Mann, D. G., Jahns, H. M. 1995. Algae. An introduction to phycology. Cambridge Univ. Press, Cambridge, U.K. Waern, M. 1938. Om Cladophora aegagropila, Nostoc pruniforme, och andra alger i Lilla Ullevifjärden, Mälaren. Botanische Noten 1938:129–142. Waern, M. 1952. Rocky-shore algae in the Öregund Archipelago. Acta Phytogeographica Suecica 30:1–298. Wehr, J. D., Stein, J. R. 1985. Studies on the biogeography and ecology of the freshwater phaeophycean alga Heribaudiella fluviatilis. Journal of Phycology 21:81–93. Weitzel, R. L. [Ed.] 1979. Methods and measurements of periphyton communities: A review. ASTM special technical publication 690. American Society for Testing and Materials, Philadelphia. West, J. A. 1990. Noteworthy collections. Washington. Heribaudiella fluviatilis (Areschoug) Svedelius. Madroño 37:144. West, J. A., Kraft, G. T. 1996. Ectocarpus siliculosus (Dillwyn) Lyngb. from Hopkins River Falls, Victoria—the first record of a freshwater brown alga in Australia. Muelleria 9:29–33. Wheeler, B. D., Whitton, B. A. 1971. Ecology of Hell Kettles. 1. Terrestrial and sub-aquatic vegetation. Vasculum 55:25–37. Whitford, L. A. 1977. Are there any rare freshwater algae? Journal of Phycology 13:73. Wilce, R. T. 1966. Pleurocladia lacustris in Arctic America. Journal of Phycology 2:57–66. Wilce, R.T., Webber, E. E., Sears, J. R. 1970. Petroderma and Porterinema in the New World. Marine Biology 5:119–135. Wujek, D. E., Thompson, R. H., Timpano, P. 1996. The occurrence of the freshwater brown alga Sphacelaria fluviatilis Jao from Michigan. Michigan Botanist 35:111–114. Yoneda, Y. 1949. Notes on the freshwater algae of Kikusui-sen, a rheocrene at Yoro-mura in Province Mino. Journal of Japanese Botany 24:169–175. Yoshizaki, M., Iura, K. 1991. Notes on Heribaudiella fluviatilis from Chiba Prefecture and Ibaraki Prefecture. Chiba Seibutu-si 40:37–39 (in Japanese). Yoshizaki, M., Miyaji, K,, Kasaki, H. 1984. A morphological study of Heribaudiella fluviatilis (Areschoug) Svedelius (Phaeophyceae) from Central Japan. Nankiseibutu 26:19–23 (in Japanese). Zimmermann, W. 1928. Über Algenbestände aus der Tiefenzone des Bodensees. Zur Ökologie und Soziologie der Tiefseepflanzen. Zeitschrift für Botanik 20:1–28 + 2 pl.


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USE OF ALGAE IN ENVIRONMENTAL ASSESSMENTS R. Jan Stevenson

John P. Smol

Department of Zoology Michigan State University East Lansing, Michigan 48824

Department of Biology Paleoecological Environmental Assessment and Research Laboratory (PEARL) Queen’s University Kingston, Ontario Canada K7L 3N6

I. Introduction II. Goals of Environmental Assessment with Algae III. Sampling and Assessing Algal Assemblages for Environmental Assessment A. Sampling Algae in Freshwater Habitats B. Attributes of Algal Assemblages for Environmental Assessment IV. Developing Metrics for Hazard Assessment A. Relating Goals to Ecological Attributes

I. INTRODUCTION Algae have long been used to assess environmental conditions in aquatic habitats throughout the world. During the early part of the twentieth century, algae were exploxed as indicators of organic pollution in European streams and rivers (Kolkwitz and Marsson, 1908). Between 20 and 50 years ago, use of algal indicators of environmental conditions flourished based on the environmental sensitivities and tolerances of individual taxa and species composition of assemblages (e.g., Butcher, 1947; Fjerdingstad, 1950; Zelinka and Marvan, 1961; Slàdecek, 1973; Lowe, 1974; LangeBertalot, 1979). Nutrient stimulation of algal growth made algae part of the problem in the eutrophication of lakes such that trophic status of lakes was also characterized by the amount of algae (Vollenweider, 1976; Carlson, 1977). In North America, Ruth Patrick and C. Mervin Palmer were pioneers in the developFreshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

V. VI. VII. VIII.

B. Testing Metrics C. Multimetric Indices D. Multivariate Statistics and Hazard Assessment Exopsure Assessment: What Are Environmental Conditions? Stressor–Response Relations Risk Characterization and Management Decisions Conclusions Literature Cited

ment of large monitoring programs to assess the ecological health of rivers and nuisance algal growths (Patrick, 1949; Patrick et al., 1954; Palmer, 1969). More recently, the sensitivity of many algal taxa to pH, combined with preservation of certain algal cell wall components (e.g., diatom frustules and chrysophyte scales) in sediments, has been employed to assess problems with acid deposition and to determine if rates of lake acidification have been enhanced by human contributions to acid deposition (Smol, 1995; Battarbee et al., 1999). Government agencies throughout the world now use algae to monitor and assess ecological conditions in many types of aquatic ecosystems (e.g., Weber, 1973; Dixit and Smol, 1991; Dixit et al., 1992, 1999; Bahls, 1993; Kentucky Division of Water, 1993; Whitton and Rott, 1996; Biggs et al., 1998; Kelly et al., 1998; Stevenson and Bahls, 1999). Thus, characterization of algal assemblages has been important in environmental assessment, both in indicating changes in 775

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environmental conditions that impair or threaten ecosystem health and in determining if algae themselves are causing problems. Algae are particularly valuable in environmental assessments. Algae are the base of most aquatic food chains, are important in biogeochemical cycling, and serve as habitat for many organisms in aquatic ecosystems (e.g., Minshall, 1978; Wetzel, 1983; Power, 1990; Carpenter and Kitchell, 1993; Vymazal, 1994; Bott, 1996; Lamberti, 1996; Mulholland, 1996; Wetzel, 1996). Thus, a natural balance of species and assemblage functions is important for ecosystem health (Angermeier and Karr, 1994). Increases in algal biomass and shifts in species composition can cause problems with many ecosystem services by causing taste and odor problems in water supplies (Sigworth, 1957; Palmer, 1962; Arruda and Fromm, 1989), toxic algal blooms (Bowling and Baker, 1996; Burkholder and Glasgow, 1997), and low dissolved oxygen levels (Lasenby, 1975). In many aquatic habitats, algae are the most diverse assemblage of organisms that can be easily sampled and readily identified to species (particularly diatoms and desmids). The great species-specific sensitivity of algae to environmental conditions and their high diversity in habitats provide the potential for very precise and accurate assessments of the physical, chemical, and biological conditions that may be causing problems. Moreover, algae and paleolimnological techniques can be used to infer historical conditions in lakes, wetlands, and even reservoirs and rivers (Fritz, 1990; Smol, 1992). Algae occur in all aquatic habitats, so they could be very valuable for comparison among ecosystems with the same group of organisms. From a logistical perspective, algae are relatively easy to sample, and analysis is relatively inexpensive compared with bioassessment with other groups of organisms. In addition, many characteristics of algal assemblages can be measured and used as multiple lines of evidence for whether ecological integrity has been altered and the causes of those alterations. Algal bioassessment complements physical and chemical data by providing corroborative evidence for environmental change. Both structural and functional characteristics of algae can be used to assess environmental conditions in aquatic habitats. Algal biomass (measured as chlorophyll a, cell numbers, and/or algal biovolume; Stevenson, 1996) can be used to indicate the presence of toxic pollutants as well as trophic status and nuisance algal growths (Carlson, 1977; Dodds et al., 1998). Taxonomic composition and diversity of algal assemblages are used to assess ecological health of habitats and to infer probable environmental causes of ecological impairment (e.g., Patrick et al., 1954; Smol,

1992; Stevenson and Pan, 1999). Ratios of chemicals in algal samples can be used to indicate algal health (phaeophytin:chlorophyll a) and nutrient limitation (N:P) (Weber, 1973; Hecky and Kilham, 1988; Biggs, 1995). Photosynthesis, respiration, and phosphatase activity are examples of algal metabolism that can be used to assess the amount of algae in habitats, physiological impairment, and phosphorus limitation (Blanck, 1985; Hill et al., 1997; Newman et al., 1994). In this chapter, the abundant and diverse methods of using algae to assess environmental conditions in all aquatic habitats are organized in a risk assessment framework (U.S. Environmental Protection Agency, 1992, 1996, 1998). Many reviews of algal methods for environmental assessment have been published in recent years (Stevenson and Lowe, 1986; Round, 1991; Coste et al., 1991; Smol, 1992; Whitton and Kelly, 1995; Rosen, 1995; Reid et al., 1995; Lowe and Pan, 1996; Stevenson, 1998; McCormick and Stevenson, 1998; Wehr and Descy, 1998; Kelly and Whitton, 1998; Kelly et al., 1998; Ibelings et al., 1998; Prygiel et al., 1999a; Stevenson and Pan, 1999; Stevenson and Bahls, 1999; see many chapters in Whitton et al., 1991; Whitton and Rott, 1996; Stoermer and Smol, 1999; Prygiel et al., 1999b). We take this abundance of recent reviews as an indication of the growing importance of algae in environmental assessment. In our chapter, we emphasize understanding the goals of environmental programs, developing and testing hypotheses that address program goals, and selecting the simplest and most direct methods for achieving program goals. We present the characteristics of algae that can be used in environmental assessments and then elaborate on how these characteristics can be related by using them in the five steps of ecological risk assessment. Although algae have been used for such assessments in habitats throughout the world, great potential exists for developing indices that more directly meet the needs of specific environmental assessment programs. Thus, in this chapter, we present the many approaches for developing algal methods for environmental assessment, and then we describe the application of algal methods for assessment.

II. GOALS OF ENVIRONMENTAL ASSESSMENT WITH ALGAE The goals of environmental assessment programs can be established by legislation, by government officials and policy decision makers, by scientists, or by the general public. In most cases, scientists play an important role in translating the official goals of an environmental program into hypotheses that can be tested and

23. Use of Algae in Environmental Assessments

in developing a practical study plan that can be implemented within the budget allocated for the project. The United States Environmental Protection Agency (U.S. EPA) risk assessment and risk management framework (U.S. EPA, 1992, 1996, 1998) (Fig. 1) is valuable for translating the many goals of ‘environmental problem solving’ into a series of testable hypotheses and for providing a sound scientific approach for solving problems. The overall goals of environmental assessment, with algae or other organisms, are to characterize the effects or potential effects of human activities and to implement management strategies that reduce the risk of ecological impairment and restore valued ecological conditions. In addition to the actual state of the ecosystem, factors such as economic, social, and legal issues may affect decisions about how to protect or restore valued ecological characteristics (Fig. 1). Because of the complexity of many environmental issues, clearly stated goals permit the development of testable hypotheses, sampling and statistical design, and choice of the best methods. The ecological risk assessment (ERA) framework helps to organize and relate the many issues associated with environmental problems and form these hypotheses. The ERA helps distinguish between the ecological conditions that the public wants to protect, the stressors that threaten those conditions, the human activities causing those stressors, and the many other factors involved in decisions on how to solve the problem. In general, most environmental assessments involve one or more steps in the ERA (Fig. 1). A full risk assessment involves five steps of ERA: problem formulation, hazard (response) assessment, exposure (stressor) assessment, evaluating the stressor–response relationships, and then characterizing the risk asso-

FIGURE 1 Elements of the risk assessment and risk management framework. Modified from U.S. Environmental Protection Agency (1996).

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ciated with each stressor and responses of interest. Problem formulation is the identification of the ecological attributes and stressors with which the public are most concerned. We may start from the perspective that a valued ecological attribute, such as water clarity or biotic integrity (sensu Karr and Dudley, 1981), is threatened or impaired and that we need to determine the cause of the problem. Are aesthetics or taste and odor impaired by nuisance growths of algae? Are toxic algal blooms occurring? Alternatively, we may be concerned about how a stressor, such as acid deposition or nutrients, could be affecting ecosystems. During problem formulation, identifying and distinguishing valued ecological attributes and stressors is very important, so that cause–effect relationships can be identified and targeted in an ERA. Our adaptation of the ERA for algal bioassessments has broad applications in the integration of the diversity of information that can be obtained in ecological assessments with algae and applying them to hazard assessment and exposure assessment. Hazard assessment is determining whether the ecological conditions in a habitat are impaired and is an assessment of the dependent variable (valued ecological attribute or an indicator of that attribute, a response variable) in the problem (Fig. 2). For example, have algae accumulated to nuisance levels or have sensitive species been lost from the habitat. Exposure assessment is an evaluation of the intensity, frequency, and duration of altered habitat conditions or contamination. For example, what is the pH, total phosphorus concentration, or organic load in the habitat, and how long does it last? Exposure assessment is a measurement of the stressor, which is the independent variable in the stressor– response relationship. The stressor–response relationship permits determination of the stressor that is likely to be most threatening or causing impairment of ecological conditions (Fig. 2). Stressor–response relationships may be found in previously published literature or in studies that accompany the ERA. Ecological risk associated with each stressor should then be characterized by comparing assessed response (hazard assessment) and stressor levels (exposure assessment) with the stressor–response relationship (Fig. 2). One or more stressors should be identified as being the likely stressors that most threaten impairment or cause impairment (see Stevenson, 2001, for more discussion). Most stressors should be too low to cause the observed impairment. However, at least one stressor, or an interaction of multiple stressors, should be high enough to cause the observed response. Algae can be used in ERA to determine whether a problem exists, to infer levels of specific stressors in a

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targeted habitat within the water body, and budget. A complete discussion of the most appropriate sampling methods and design is beyond the scope of this chapter, but we will review some of the important issues related to sampling for bioassessment. In general, the same basic approaches are useful for solving most problems in any water body or habitat. Algal indicators may, however, be more precise if they are refined with regional datasets for specific water body types, but this problem will be discussed later with development of indicators of exposure assessment (weighted averaging inference models). FIGURE 2 Response–stressor relationship between hypothetical ecological response and stressor with hazard assessment and exposure assessment indicated. Acceptable levels of the ecological response are indicated by the horizontal dotted line. The stressor level that produces that level of response is indicated by the vertical dashed line. The observed ecological response (indicated by horizontal solid arrow marked HA on the y axis) is below acceptable levels. Two stressors were measured or inferred based on algal indices (indicated by the vertical arrows marked EA (exposure assessment) on the x axis). One stressor (indicated by EA1) is too low to cause the observed response, whereas the other has a high certainty of causing the observed ecological condition.

habitat, and to characterize stressor–response relationships. Algal indices are used to assess both stressors as well as valued ecological attributes. Probably more than any other group of organisms, algae have been used to infer physical and chemical conditions (potential stressors) in a habitat through the determination of species composition of assemblages and species’ ecological preferences. Potential for inferring stressor conditions exists for other organisms, but algae are used much more often than fish, macroinvertebrates, or insects in terrestrial habitats to infer levels of pH, conductivity, trophic status, and sewage contamination. Therefore, distinguishing between whether algal indices are indicating valued ecological attributes or stressors is important for relating results of algal bioassessments to goals of the ERA. This opportunity to use algae to determine whether problems exist, potentially even forecast problems, and to diagnose causes of problems should be emphasized in algal bioassessments.

III. SAMPLING AND ASSESSING ALGAL ASSEMBLAGES FOR ENVIRONMENTAL ASSESSMENT A. Sampling Algae in Freshwater Habitats Sampling techniques and design may vary with the objectives of the assessment, probable factors affecting algae and anticipated problems, water body type,

1. Sampling Design Objectives of an environmental assessment should be defined as clearly as possible for the formulation of hypotheses and a sampling design (e.g., Green, 1979). Testable hypotheses should be formed so that results provide answers that address the objectives of the assessment and that have a defined error or uncertainty. Testing hypotheses requires a sampling design that includes replicate sampling or some form of assessment of error variation. Estimates of ecological health in small, local studies are usually based on replicate sampling at all sites and provide means and estimates of variation at each site to test the hypothesis that conditions at a tested site are significantly different from conditions at a reference site or from a criterion. However, replicate sampling at all sites in large surveys is often not affordable; then replicate sampling at a random subset of sites (often 10% or more) can be used to characterize variation in estimates of ecological health. The error variation associated with the random subset of sites in large surveys is a measure of the precision (standard deviation or standard error) of assessments at all sites. Before sampling, investigators should define the extent of each sample site so that it can be repeated at all locations. The open-water region of lakes usually defines the horizontal extent of plankton habitats sampled at a lake site, but the vertical extent (depth of sampling) may vary with goals of the project. In many river programs, for example, sampling one riffle has been considered sufficient to characterize the ecological health of a stream with benthic algae (Bahls, 1993), but in other programs the extent of the habitat is defined as a stream reach with a length that is 40 times the width of the stream (Klemm and Lazorchak, 1994; Stevenson and Bahls, 1999). Similarly, a representative portion of wetland should be chosen to sample (e.g., Stevenson et al., 1999). Most algal sampling strategies focus on a specific habitat within the water body (Wehr and Sheath, Chap. 2, this volume), such as plankton, algae on rocks


(epilithic) in riffles, or algae on plants (epiphytic). Alternatively, objectives of a project may call for characterizing the diversity of algae in a habitat and consequently sampling all suitable habitats within a water body (defined site) (Porter et al., 1993). The presumed advantage of sampling targeted habitats is that algal indicators are more sensitive and can more precisely detect changes in environmental conditions if interhabitat variability is reduced (Rosen, 1995). In a recent review, Kelly et al. (1998) make strong arguments for sampling rocks or other hard substrata, if they are present. In some cases, however, sampling the same targeted habitat in all water bodies of a project is impractical. Although finding plankton in lakes is usually not a problem, finding cobble riffles in all streams or open water in all wetlands can be a problem. Multihabitat sampling is one solution to the problem of habitat diversity among sites. One advantage to multihabitat sampling is a more complete assessment of all taxa at a site, which potentially is a better characterization of biodiversity and biointegrity than assemblages from targeted habitats. Another solution is to classify streams and wetlands by size and hydrogeomorphology (Vannote et al., 1980; Biggs and Close, 1989; Rosgen, 1994; Biggs, 1995; Brinson, 1993; Goldsborough and Robinson, 1996; Biggs et al., 1998), so they have similar habitats, and then to develop sampling strategies and indicators for specific hydrogeomorphic classes of streams and wetlands. Estimates of algal biomass, which are particularly important in characterizing trophic state, require quantitative sampling of habitats. Habitats are quantitatively sampled by measuring the volume of water collected or area of substratum sampled and accounting for the proportion of sample assayed. Algal attributes (e.g., biomass or productivity) can be expressed on a volume-specific or area-specific basis by correcting measurements for volume or area sampled and proportion of sample assayed (Wetzel and Likens, 1991; APHA, 1998). The main disadvantage of quantitative sampling is the time required and the practicality of precisely characterizing the area sampled. Measuring sample volume, in the field or in the laboratory, requires relatively little extra time. The benefits of quantitative sampling are also reduced when habitat conditions are spatially or temporally variable such that biomass is affected. Thus, quantitative sampling is particularly problematic in structurally diverse and hydrologically variable streams and wetlands. Qualitative algal sampling is recommended in habitats that have great spatial and temporal variation and when sufficient time is not available to measure the substratum area. One goal of qualitative sampling could be sampling all species at a site (Porter et al.,

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1993), which would call for sampling all habitats, water column, rocks, plants, and sediment in different physical settings (e.g., light, depth, current velocity, etc.). Another goal could be to determine the dominant algae at a site, which would require estimating the relative areas of different habitats and proportional sampling of those habitats. Variation in quantitative or qualitative estimates of algal attributes, due to spatial variation in habitat conditions, can be reduced by composite sampling. Reducing spatial variation calls for subsampling of many areas throughout the defined extent of the study site and putting all subsamples into a composite sample. Variation in estimates of algal attributes due to temporal variation in habitats is more difficult to reduce because it requires sampling throughout a study period and return visits to a site, which may not be practical. To reduce effects of temporal variation on quantitative attributes in highly variable ecosystems like streams, it is best to sample after an extended (1–2week) period of stable habitat conditions so that algal assemblages have reached peak or sustainable biomass and a relatively predictable state (Stevenson, 1990; Peterson and Stevenson, 1992; Biggs, 1996; Stevenson, 1996). Semiquantitative approaches for assessing algal biomass and percentage cover of different algal groups have been used to reduce field and lab time and to increase the spatial and temporal extent of algal assessments in streams. Secchi disc transparency is a semiquantitative approach for assessing plankton biomass in lakes (Brezonik, 1978; Davies-Colley and Vant, 1988; Wetzel and Likens, 1991). In streams, visual characterization of algal type, percentage cover, filament length, and periphyton mat thickness along multiple transects have been used in many situations (Holmes and Whitton, 1981; Sheath and Burkholder, 1985; Rout and Gaur, 1990; Stevenson and Bahls, 1999). These techniques can easily be employed in all sampling programs because they require little time and can provide biomass assessments over large areas. They may be excellent quantitative tools for volunteer programs because they require little taxonomic expertise.

2. Sampling Techniques a. Sampling Present-Day Assemblages Numerous algal habitats can be sampled within water bodies. Phytoplankton can be sampled at specific depths with Van Dorn, Kemmerer, or similar discrete-depth samplers (APHA, 1998). Depth-integrated samples can be collected with devices (e.g., peristaltic pumps) that allow water to slowly enter a sampling chamber or by compositing samples collected from specific depths.

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Phytoplankton biomass is almost always expressed per unit volume (e.g., Wetzel and Likens, 1991). Qualitative samples can be collected with plankton nets; however, we recommend collecting whole water samples with known volumes whenever possible so that small algae are not missed and samples can be assayed quantitatively. Algae in whole water samples can be concentrated by filtering or settling (e.g., Wetzel and Likens, 1991; APHA, 1998). Metaphyton are macroalgal and microalgal masses suspended in the water column and entangled among macrophytes or along shorelines, typically in slow or still water (Hillebrand, 1983; Goldsborough and Robinson, 1996). Quantitative sampling of metaphyton requires collecting algae from a vertical column through the assemblage. Coring tubes can be used to isolate and collect a column of metaphyton. Scissors are useful for cutting horizontal filaments that block the insertion of the tube through the metaphyton assemblage. The depth of the core should not extend to the substratum surface. The diameter of the core depends upon the spatial variability of the metaphyton, on necessary sample size, and on the ability to isolate the core of algae from surrounding metaphyton (Stevenson, personal observation). Metaphyton in the form of unconsolidated green clouds requires wider cores (ca. 10 cm) because filaments are difficult to isolate in narrow cores. Narrower cores (ca. 3 cm) can be used to sample consolidated microalgal mats. Metaphyton biomass should be expressed on an areal basis (e.g., m–2). Qualitative samples of metaphyton can be gathered with grabs, forceps, strainers, spoons, or cooking basters. Benthic algae are sampled by scraping hard or firm substrata, such as rocks, plants, and tree branches, usually after they have been removed from the water (Stevenson and Hashim, 1989; Aloi, 1990; Porter et al., 1993). Cores of algae should be collected on soft or unconsolidated substrata, such as sediments and sand (Stevenson and Stoermer, 1981; Stevenson and Hashim, 1989). Area of substrata sampled should be recorded to quantify samples. Some substrata, such as bedrock and logs, cannot be removed from the water. In those cases, vertical tubes can be used to isolate an area of substratum. After algae are scraped from the substratum in the tube, algae and water in the tube can be removed with a suction device. Artificial substrata are also used to assess benthic algal assemblages (Patrick et al., 19547; Tuchman and Stevenson, 1980; Aloi, 1990). They are typically uniform substrata (e.g., glass or acrylic slides, clay tiles, acrylic or wooden dowels) that can be used across many water body types (streams, rivers, wetlands, lakes). If placed in similar light and current environ-

ments in all habitats, differences in assemblages among sites should be highly sensitive to water chemistry. However, placement and sampling of artificial substrata require more than one trip to sample sites. Samples are often lost because they are subject to vandalism. Finally, assemblages on artificial subtrates may not reflect historical changes in habitats or changes in physical habitat structure as well as assemblages on natural substrata (see also Section II). Most large national and state programs have chosen to sample natural substrata (Porter et al., 1993; Klemm and Lazorchak, 1994), but artificial substrata can be valuable in smaller-scale programs where travel time is limited or in ecosystems with great habitat diversity. b. Sampling Historic Assemblages Sediment sampling in lakes, streams, rivers, and wetlands with deposited sediments can include an algal assemblage that has accumulated for months, years, or centuries, depending upon the depth and disturbance of sediments. A large variety of coring apparatuses are available to retrieve sediment cores (several are illustrated in Smol and Glew, 1992), with the choice of equipment largely dependent on the type of system being studied and the temporal resolution required. The resolution is also dependent on the type of sectioning techniques and equipment one uses. Close-interval sectioning equipment and techniques (e.g., Glew, 1988) are available that can provide lake managers with a high degree of temporal resolution. The overall paleolimnological approach is summarized in the accompanying schematic (Fig. 3). Once the study site is chosen, a sediment core is removed, usually from near the center of the lake. In general, the central, flat portion of a basin integrates indicators from across the lake, and so a more holistic record of past environmental change is archived. Once the core is retrieved and sectioned, the “depth-time” profile must be established. This requires dating a sufficient number of sediment layers to attain a reliable chronology. For most paleolimnological studies dealing with recent environmental assessments, 210 Pb dating is most often used (Oldfield and Appleby, 1984), as the half-life (22.26 years) of this naturally occurring isotope enables one to date, with reasonable certainty, approximately the last century or so of sediment accumulation. In some lake systems, close to annual (and sometimes subannual) resolution is possible. Analyzing a sediment core at close intervals (e.g., every cm or every 0.5 cm) is time consuming and may not be practical for some large-scale, regional environmental assessments. For these cases, paleolimnologists have sometimes used a “snapshot” approach, which


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FIGURE 3 Schematic diagram showing the major steps involved in a paleolimnological assessment. Modified from Dixit et al., (1992a).

attempts to estimate what conditions were like before anthropogenic impacts and how much degradation has occurred. This so-called top/bottom approach is a very simple but effective tool for obtaining regional assessments of environmental change. Paleolimnologists remove surface sediment cores as they would in a detailed paleoenvironmental assessment; but instead of sectioning and analyzing the entire core, they simply analyze, for example, diatom valves and/or chrysophyte scales in the top 1 cm of sediment (= present-day conditions) and from a sediment level known to have been deposited before anthropogenic impact (i.e., the bottom sediment section). This before-and-after approach has been effectively used to infer environmental change, such as acidification (Cumming et al., 1992a; Battarbee et al., 1999; Dixit et al., 1999) and eutrophication (e.g., Dixit and Smol, 1994; Hall and Smol, 1999) that has occurred on regional scales. In addition to providing some estimate of degradation, these paleoenvironmental data also provide important information on the natural background conditions of a system and therefore provide important mitigation targets for environmental remediation efforts (Smol,

1992). To determine the rates and trajectories of past changes, more detailed paleoenvironmental assessments are required. The next step is to recover any paleoenvironmental information archived in dated sediment cores. Our focus here is on the paleophycological data, but many other types of proxy data are available. For example, past changes in terrestrial vegetation can be inferred from the analyses of fossil pollen grains (the field of palynology); paleomagnetic measurements and other techniques can be used to estimate past erosion rates (e.g., Dearing et al., 1987); and isotope and geochemical analyses of metals and other contaminants (e.g., PCBs, DDT, etc.) can be analyzed from the sedimentary profiles (Autenrieth et al., 1991). Despite these other powerful approaches, the mainstay of many paleolimnological assessments is algal data. Virtually every algal group leaves some sort of morphological or chemical fossil in the sedimentary record, but the indicators that are most often used are diatom valves (Dixit et al., 1992), chrysophyte scales and cysts (Smol, 1995), and fossil pigments (Leavitt, 1996). As shown in examples given later in this chapter, these indicators can be used

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to reconstruct past limnological characteristics (such as pH, eutrophication variables, and salinity). Paleolimnological approaches have been subjected to a large amount of quality assurance and quality control considerations; if undertaken carefully and correctly, the paleolimnological approach is robust, reproducible, and powerful.

B. Attributes of Algal Assemblages for Environmental Assessment Many attributes of algal assemblages can be used to assess environmental conditions in a habitat or site (Table I). Structural attributes (e.g., species composition) and functional attributes (e.g., productivity) can be measured in the field or the laboratory. The diversity of these attributes and the pros and cons of their uses have been discussed in recent studies (Stevenson, 1996; Stoermer and Smol, 1999; Stevenson and Pan, 1999; Stevenson and Bahls, 1999). In this treatment, we will focus more on the value of these attributes in detecting effects of humans on ecological systems. For this review, we will use a set of terms recommended by Karr and Chu (1999) to clarify discussions of using biological data for bioassessment. An attribute

TABLE I Basic Attributes of Algal Assemblages That Can Be Measured and Potentially Be Used to Assess Environmental Conditions Structural attributes Biomass chl a Ash free dry mass Cell density Cell biovolume Taxonomic composition Species relative abundances Species relative biovolume Functional group biovolume Diversity Species richness Genus richness Evenness Chemical composition chl a:(Phaeophytin+Chl a) ratio chl a:ash free dry mass ratio P or N/ash free dry mass N:P ratio of algal assemblages Functional attributes Photosynthesis rates Respiration rates Net primary productivity Growth rates Nutrient uptake rates For a more detailed list with literature citations, see McCormick and Cairns (1994).

is any characteristic of an assemblage that can be measured, such as chlorophyll a (chl a), number of species, or net primary productivity. A metric is an attribute that responds to human disturbances of habitats. Some attributes, such as the number of diatom genera in a 200-valve count, may not reliably respond to human impacts. Karr and Chu (1999) also recommend the use of the term index for statistical and other mathematical summaries of many metrics and indices. Multimetric indices of biotic integrity, such as the diatom bioassessment index (Kentucky Division of Water, 1993), are examples of multiple metrics being averaged or summed to compose a single, summary index. Many of the water quality indices that are commonly used in Europe (e.g., pollution tolerance index (Lange-Bertlalot 1979), generic diatom index (Coste and Ayphassorho, 1991), and trophic diatom index (Kelly and Whitton, 1995)) should probably be characterized as metrics, according to the method of Karr and Chu (1999), but we do not recommend renaming these indicators and being encumbered by semantics. Algal assemblages can be characterized through the use of two basic kinds of attributes, structural and functional (Table I). Structural attributes are instantaneous characterizations of assemblages, such as biomass per unit area or volume of habitat, taxonomic and chemical characterizations of community composition, and diversity of community taxa (e.g., species richness). Functional attributes are measures or indicators of assemblage metabolism, such as photosynthetic rate (gross primary production), respiration rate, net primary productivity, nutrient cycling, phosphatase activity, and population growth rates. Functional assessments usually require more time in the field or multiple trips, so are used less in routine environmental assessments. However, they can be important for understanding impairment of algal and microbial activity.

1. Biomass Biomass of algae usually increases with resource availability and decreases with many stressors caused by humans (Vollenweider, 1976; Dodds et al., 1998). Removal of riparian (stream-side) canopies along streams and nutrient loading in all water bodies increase light, temperature, and nutrient availability, which can limit algal growth rates and biomass accrual (see reviews in Biggs, 1996; Hill, 1996; DeNicola, 1996; Borchardt, 1996). Sediments, toxic substances, and removal of benthic habitat can limit algal growth and accrual (Genter, 1996; Hoagland et al., 1996). Because biomass and the potential for nuisance algal growths vary temporally with season and weather (Whitton, 1970; Wong et al., 1978; Lembi et al., 1988), timing of sampling is important. In most habi-


tats, peak biomass occurs after periods of undisturbed habitat conditions (e.g., post-flood) when algal biomass has had an opportunity to accrue. Spring and summer blooms of phytoplankton are common in lakes with spring turnover and warm summer temperatures (Wetzel, 1983; Harper, 1992). Nuisance algal growths may occur, with filamentous benthic algal accrual during seasonal optima (Whitton, 1970; Biggs and Price, 1987; Dodds and Gudder, 1992; Lembi et al., Chap. 24, this volume), or in the water column during summer low flow, when water residence time is sufficient for algal accrual in the water column (Bowling and Baker, 1996). Biomass is an important attribute in environmental assessments because it is related to productivity and nuisance problems. Biomass of algal assemblages can be estimated with laboratory assays of chl a, dry mass, ash-free dry mass, algal cell density, biovolume, or chemical mass of samples. All of these measurements have pros and cons (see Stevenson, 1996, for a review), because none directly measure all constituents of algal biomass or only algal biomass. However, all are reasonable estimates of algal biomass in different situations. Chl a must be extracted from cells in organic solvents, such as acetone or methanol, and then assayed by spectrophotometry, fluorometry, or high-performance liquid chromatography (HPLC) (Lorenzen, 1967; Mantoura and Llewellyn, 1983; Wetzel and Likens, 1991; APHA, 1998; Van Heukelem et al., 1992; Millie et al., 1993). Spectrophotometric and flourometric chl a assays should be corrected for phaeophytin. Dry mass and ash-free dry mass are measured by drying and combusting samples (APHA, 1998). Cell density is measured after cells are counted microscopically (Lund et al., 1958; APHA, 1998; Stevenson and Bahls, 1999). Algal biovolume can be measured by distinguishing sizes of cells during microscopic counts, multiplying biovolume by cell size for all size categories, and finally summing biovolumes for all size categories in the sample (Stevenson et al., 1985; Wetzel and Likens, 1991; APHA, 1998; Hillebrand et al., 1999). Because of variation in vacuole size and cell walls (e.g., SickoGoad et al., 1977), cell surface area may be a valuable indicator of biomass because much cytoplasm is within 1–2 µm of the cell membrane. Biomass can also be estimated rapidly with field assays, such as secchi depth in the water column and percentage cover and thickness of algal assemblages on substrata (Wetzel and Likens, 1991). Secchi depth characterizes light attenuation in the water column; assessments of algal biomass by this method are confounded by suspended inorganic materials, dissolved substances, and other factors (Preisendorfer, 1986). The strengths of assessing benthic algal biomass from

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percentage cover and thickness of algal assemblages is that biomass throughout a stream reach can be readily characterized (Holmes and Whitton, 1981; Sheath and Burkholder, 1985; Stevenson and Bahls, 1999). Remote sensing of algal biomass also shows promise for assessing spatially and temporally variable growths (Cullen et al., 1997). These rapid assessment techniques may permit more thorough spatial and temporal assessments, which may improve the notoriously variable relationships between biomass and nutrient concentrations or loading.

2. Taxonomic Composition Taxonomic composition of algae is a powerful tool for assessing biotic integrity and diagnosing the direct and indirect causes of environmental problems (Stevenson, 1998). Differences in taxonomic composition of assemblages between an assessed site and a reference (desired) site can indicate impairment of biotic integrity and environmental conditions, if natural variation in assemblage composition is well documented (e.g., McCormick and O’Dell, 1996; McCormick and Stevenson, 1998). When natural seasonal or interhabitat variation in composition is not well known, changes in taxonomic composition can be related to human activities by comparing shifts in taxonomic composition to environmental change with autecological characteristics of species and relating inferred environmental changes to human activities (e.g., Kwandrans et al., 1998). Shifts in functional groups of algae (defined as different growth forms and divisions of algae; e.g., Pan et al., 2000) can also indicate an important change in food quality and in habitat structure for benthic invertebrates. For example, the food quality and accessibility of diatoms are usually greater than cyanobacteria and filamentous green algae for many herbivores (Porter, 1977; Lamberti, 1996). In addition, the habitat structure for benthic invertebrates differs greatly with changes from microalgae (e.g., diatoms) to macroalgae (e.g., Cladophora) (Holomuzki and Short, 1988; Power, 1990). Taxonomic composition of algal assemblages can provide a highly precise and accurate characterization of biotic integrity and environmental conditions (Stoermer and Smol, 1999). Taxonomic composition of assemblages develops over periods of time, ranging from weeks to years, and should reflect environmental changes during that period. Even though taxonomic composition varies spatially and temporally in a water body, autecological characterizations of environmental conditions based on taxonomic composition should consistently reflect the physical and chemical changes caused by humans. For example, if trophic status of a habitat is being assessed, only low-nutrient indicator

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taxa should occur in low-nutrient habitats, even though temperature and shading and stage of community development may change with time and local habitat structure. Species presence and success in assemblages are fundamentally constrained by environmental conditions and interactions (e.g., competition) with other species in the habitat (Stevenson, 1997). Thus, attributes of assemblages based on percentage taxonomic similarity of assemblages at a test site and a reference site (Raschke, 1993; Stevenson, 1984) and percentage sensitive species should be good metrics because typically they sensitively, precisely, and monotonically change along gradients of human disturbance. Taxonomic composition of algal assemblages usually requires microscopic assessments of samples, but some basic information about growth form and class can be obtained with rapid field assessments (e.g., Sheath and Burkholder, 1985; Stevenson and Bahls, 1999). The methods used for microscopic identification and counting of algae depend upon the objectives of data analysis and type of sample. A two-step process has been adopted by the national stream assessment programs in the United States. The first step is to count all algae and identify only nondiatom algae in a wet mount at either 400× (e.g., Palmer cell) or 1000×, if many small algae occur in samples. Algae can be counted in wet mounts at 1000× with an inverted microscope (Lund et al., 1958) or with a regular microscope by drying samples onto a coverglass, inverting the sample onto a microscope slide in 20 µL of water, and sealing the sample by ringing the coverglass with fingernail polish or varnish (Stevenson, unpublished method). The second step is to count diatoms after oxidizing organic material out of diatoms and mounting them in a highly refractive mounting medium (Stevenson and Bahls, 1999). This technique provides the most complete taxonomic assessment of an algal assemblage. Using counts of 300 algal cells, colonies, or filaments and about 500 diatom valves is a standard approach of some U.S. national programs (Porter et al., 1993; Pan et al., 1996) and usually provides relatively precise estimates of the relative abundances of the dominant taxa in a sample. Alternatively, counting rules have been defined so that cells of all algae are identified and counted until at least 10 cells (or natural counting units = cells, colonies, or filaments) of the 10 dominant taxa are counted (Stevenson, unpublished data). This type rule, rather than a fixed total number of cells, ensures precision in estimates of a specified number of taxa. Some assessment programs primarily use diatoms (Bahls, 1993; Kentucky Division of Water, 1993; Kelly et al., 1998; Kwandrans et al., 1998), because the number of species in diatom assemblages is usually sufficient to show a response.

Taxonomic composition can be recorded as presence/absence, percentage or proportional relative abundances, percentage or proportional relative biovolumes, or absolute densities and biovolumes of taxa (cells or µm3 cm–2 or mL–1). Although there is no published comparison of these forms of data, they represent scales of biological resolution and probably reflect a gradient from least sensitive to most sensitive and least variable to most variable. Presence/absence records of species should be based on observations of thousands of cells and conceptually should reflect longterm changes in habitat conditions if immigration and colonization of habitats are a selective barrier. Relative abundances and biovolumes of taxa probably reflect recent habitat conditions more than long-term conditions because of recent species responses to environment. Densities and biovolumes of taxa change daily, so absolute densities and biovolumes may be too sensitive to detect more long-term environmental changes. Relative abundance of cells is more commonly used than relative biovolumes (because of ease of use), but the latter is particularly valuable when cell sizes vary greatly among taxa within samples.

3. Diversity Richness and evenness of taxa abundances are two basic elements of diversity (Shannon, 1948; Simpson, 1949; Hurlbert, 1971) of biological assemblages. Richness and evenness are hypothesized to decrease with increasing human disturbance of habitats; however, evenness of species abundances may increase if toxic stresses retard the growth of dominant taxa more than rare taxa (e.g., Patrick, 1973). Two problems develop with use of diversity measures in environmental assessment: standard counting procedures may not accurately assess diversity (Patrick et al., 1954; Stevenson and Lowe, 1986), and diversity may not change monotonically across the gradient of human disturbance (Stevenson, 1984; Jüttner et al., 1996; Stevenson and Pan, 1999). Species diversity and evenness are highly correlated with standard 300–600 cell counts (Archibald, 1972). In these counts many species have usually not been identified, so richness is more a function of evenness than evenness is a function of richness (Patrick et al., 1954; Stevenson and Lowe, 1986). As shown by standard counting procedures, nonmonotonic (showing both positive and negative changes as the independent variable increases) responses of algal diversity to some environmental gradients seem to be related to maximum evenness of tolerant and sensitive taxa at midpoints along environmental optima, to fewer species being adapted to environmental extremes at both ends of environmental gradients, and to subsidy-stress perturbation gradients


(Odum et al., 1979). Despite these difficulties, species richness and evenness may respond monotonically (having only positive or negative changes, but not necessarily linear changes, as the independent variable increases), sensitively, and precisely to gradients of human disturbance in some settings and should be tested for use as metrics.

4. Chemical Composition The chemical composition of algal assemblages can be used to assess the trophic status of water bodies (e.g., Carlson, 1977), such as total phosphorus (TP) and nitrogen (TN) concentrations of water and periphyton (Dodds et al., 1998; Biggs, 1995). TN:TP ratios are widely used to infer which nutrient regulates algal growth (Healey and Hendzel, 1980; Hecky and Kilham, 1988; Biggs, 1995). In many of these assessments, most of the total P and N are particulate, and much of the particulate matter is algae. Thus, measurements of TP or TN per unit volume or area of habitat largely reflect the amount of algae in the habitat. Of course, the most widespread use of trophic assessments with TP and TN is phytoplankton in lakes (Carlson, 1977), but use has also been proposed for streams, rivers, and wetlands (Dodds et al., 1998; McCormick and Stevenson, 1998). TP and TN per unit biomass in benthic algae have also been positively correlated to benthic algal biomass in streams; however, negative density-dependent effects may reduce biomass-specific concentrations of benthic algal TP and TN and confound estimates of P and N availability to cells (Humphrey and Stevenson, 1992). Volume-specific, area-specific, and biomass-specific estimates of TP and TN do increase monotonically with most gradients of human disturbance and may be good metrics for trophic status in streams, rivers, and wetlands as well as lakes. Chemical assessments are also valuable for monitoring heavy metal contamination in rivers, lakes, and estuaries (Briand et al., 1978; Whitton et al., 1989; Say et al., 1990). Many algae accumulate heavy metals when exposed to them in natural environments (Whitton, 1984). While toxicity of heavy metals to algae is one reason for monitoring heavy metals in algae, other reasons include bioaccumulation and metal removal from waste streams and movement of heavy metals into the food web (Whitton and Shehata, 1982; Vymazal, 1984; Radwin et al., 1990).

5. Functional Attributes Metabolism of algal assemblages is highly sensitive to environmental conditions and is important to the assessment of ecosystem function and many ecosystem services. Estimates of photosynthesis (gross primary

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productivity), respiration, net primary productivity, nutrient uptake and cycling, and phosphatase activity are common functions measured in ecological studies (Bott et al., 1978; Healey and Hendzel, 1979; Wetzel and Likens, 1991; Marzolf et al., 1994; Hill et al., 1997; Whitton et al., 1998). These techniques are rarely incorporated into routine monitoring and survey work because they require more field time than typical water, phytoplankton, and periphyton sampling. However, they can be valuable additions to bioassessment projects. Biggs (1990) describes using algal growth rates to assess stream enrichment. Metabolism can be based on an area-specific, volume-specific, or biomassspecific basis. The most direct measurement of cellular performance is biomass-specific rates of metabolism; however area-specific and volume-specific measurements directly relate to community performance and ecosystem services. Caution in the accurate use of these attributes must be exercised. Area- and volume-specific measurements of productivity and respiration increase with biomass in the habitat, irrespective of human influence of biomass-specific rates. However, for periphyton, biomass-specific rates of productivity and nutrient uptake decrease substantially with increasing biomass in the habitat (e.g., Hill and Boston, 1989), presumably because of shading and impairment of nutrient mixing through the microbial matrix (Stevenson and Glover, 1993). Gross and net productivity and respiration can be measured in the field with light and dark chambers and changes in oxygen concentration (Bott et al., 1978; Wetzel and Likens, 1991). Alternatively, productivity can be estimated with changes in oxygen concentration in the water during the sampling period or at two locations in a stream, if diffusion of oxygen from the water column is properly accounted for (Kelly et al., 1974; Marzolf et al., 1994). Furthermore, algae secrete an enzyme called phosphatase when in low-P environments. The phosphatase enzyme cleaves PO4 from organic molecules and makes it biologically available. Phosphatase is measured with water samples in the laboratory (Healey and Hendzel, 1979).

6. Bioassay In this discussion we define bioassays as in-lab culture of organisms in waters from the study site. A valuable, field-based use of this technique is the Selenastrum bottle assay, in which known quantities of this highly culturable green alga are added to water from the study site and growth is monitored over a predefined period (Cain and Trainor, 1973; U.S. Environmental Protection Agency, 1971; Trainor and Shubert, 1973; Greene et al., 1976; Ghosh and Gaur, 1990; McCormick et al., 1996). Alternatively, plank-

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tonic or benthic assemblages from reference or test sites could be cultured in bioassays with waters from those habitats or different dilution levels of effluents entering those regions. Twist et al. (1997) introduced the novel approach of embedding test organisms in alginate and culturing them in situ. Nutrient-diffusing substrata, microcosms, and mesocosms are also valuable bioassay techniques in field settings (e.g., Cotê, 1983; Fairchild et al., 1985; Gensemer, 1991; Hoagland et al., 1993; Lamberti and Steinman, 1993; Thorp et al., 1996). Response of organisms to bioassays can provide another valuable line of evidence for identifying causes of environmental stress. Bioassay results with specific chemicals or effluents added can be used to confirm cause–effect relations between parameters for which only observational correlations can be obtained in field surveys (e.g., McCormick and O’Dell, 1996; Pan et al., 2000).

IV. DEVELOPING METRICS FOR HAZARD ASSESSMENT A. Relating Goals to Ecological Attributes Hazard assessments are the determination of the intensity, spatial extent, frequency, and duration of environmental problems or the threat of environmental problems in which ecological conditions do not meet designated use (U.S. EPA, 1996). Designated use describes the goals for environmental protection in U.S. management plans, such as preserving biotic integrity and biodiversity, maintaining fishable and swimmable conditions, minimizing taste and odor problems or risks to human health in water supplies, optimizing sustainable fisheries production, and protecting human health. Designated use emphasizes valued ecological attributes that the public wants to protect. Many algal attributes can be related to these designated uses. We recommend selecting as many metrics as possible to develop multiple lines of evidence to help assess ecological conditions, which can be altered in many ways. Thus, hazard assessment requires identifying the goals of environmental assessment, selecting algal metrics that represent qualities of ecosystems that are related to designated uses, and then measuring those algal metrics to determine if goals are being met. One of the most fundamental goals of environmental assessment is to determine if the natural balance of flora and fauna has been altered in a habitat. The concept of biotic integrity and natural balance of flora and fauna is a legislated goal in the United States’ Clean Water Act (Karr and Dudley, 1981; Adler, 1995), is fundamental to ecosystem protection and sustaining biodiversity (Angermeier and Karr, 1994), and is

broadly applied in U.S. monitoring programs in which indices of biotic integrity are used to identify ecological problems (Plafkin et al., 1989; Barbour et al., 1999; Karr and Chu, 1999). Biotic integrity or ecosystem health can be defined as the similarity between assemblages in an evaluated habitat and assemblages in a set of reference habitats (sensu Hughes, 1995). That assessment of similarity can be based on structural and functional characteristics. In most cases, assessment of biotic integrity has been based on changes in diversity, species composition, functional groups (such as proportions of diatoms versus green algae and cyanobacteria), and changes in ecological conditions inferred by species composition and species autecological characteristics (Karr, 1981; Smol, 1992; Kerans and Karr, 1994; Stevenson and Bahls, 1999). Alternatively, more specific assessments of algal nuisance can be the goals of projects. Such nuisances may cause reduced water clarity, hypolimnetic deoxygenation, taste and odor problems, habitat alteration, or toxic effects on other organisms, including humans (Carmichael, 1994, Chap. 24). In these cases, algal biomass or specific problem taxa may be important attributes for assessment. Attributes that respond to gradients of human disturbance are classified as metrics. A good metric for hazard assessment is unambiguous, sensitive, precise, reliable, and transferable among regions and perhaps water body types (Murtaugh, 1996; Karr and Chu, 1999). McCormick and Cairns (1994) list other ideal qualities of indicators that should also be considered, such as relevance, redundancy, and cost-effectiveness. An unambiguous attribute responds monotonically to increasing levels of human disturbance (Fig. 4a and b). An attribute that can be equal at low and high levels of human disturbance (Fig. 4c) is ambiguous and should not be used as a metric. A good attribute may respond nonlinearly to human disturbance, but most change substantially (i.e., be sensitive) over the range of human disturbance being assessed. Precision (low variability in repeated measures) of metrics is important for detecting responses. In addition, metrics will be much more effective if they respond to human disturbance at many times of year and if they respond in many regions. Two categories of attributes can be identified that characterize or indicate valued ecological attributes, and these categories are distinguished by the degree to which reference conditions need to be characterized. All attributes in the first category can be directly related to gradients of human disturbance by regression analysis to determine their use as metrics without rigorous identification and characterization of reference conditions and assemblages at reference sites. The first category includes attributes such as biomass,


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FIGURE 4 Examples of attribute responses that provide useful metrics. (A) Negative attribute responses along a gradient of human disturbance. (B) Positive responses along a gradient of human disturbance. (C) Ambiguous responses along a gradient of human disturbance. The different lines in each figure (A–C) represent different patterns that would fit the categories represented in the figures (negative, positive, and ambiguous, respectively).

species richness, Shannon (1948) diversity, relative abundances of specific nuisance taxa or functional groups of taxa, relative abundances of pollution-sensitive and pollution-tolerant taxa, and function of assemblages (Tables I and II; Stevenson and Bahls, 1999). A second category of metrics requires comparison of differences between reference and test sites as the

response variable to the gradient of human disturbance (Table II). Similarity of species composition between test sites and reference sites is one potential metric. Similarity of species composition between reference and test sites should decrease with increasing human disturbance at test sites. Many different formula can be used to calculate percentage similarity. One compares

TABLE II Examples of Metrics That Are Based on Taxonomic Composition of Assemblages and Autecological Characteristics of Species Class

Stressor

Representative references

SC

BI

Pollution in general (based on species)

Lange-Bertalot (1979), Descy (1979), Coste (1982), Bahls (1993)

SC

BI

Pollution in general (based on genera)

Rumeau and Coste (1988), Coste and Ayphassorho (1991)

SC

D

pH

Whitmore (1989)

SC

D

Trophic status

Whitmore (1989), Kelly and Whitton (1995)

SC

D

Organic wastes

Zelinka and Marvan (1961), Palmer (1969), Sládecˇ ek (1973, 1986), Watanabe et al. (1986)

SC

D

Salinity

Zeimann (1991)

WA

D

pH

Charles and Smol (1988), ter Braak and van Dam (1989), Sweets (1992), Cumming et al. (1992a, b)

WA

D

Salinity

Fritz (1990), Cumming and Smol (1992)

WA

D

TP

Anderson et al. (1993), Reavie et al. (1995), Pan et al. (1996), Pan and Stevenson (1996)

WA

D

Fish presence

Kingston et al. (1992)

Metrics are classified based on whether they are based on simple categorical autecological characterizations (SC) or accurate weighted-average (WA) autecological characterizations and on whether they could be used to infer biotic integrity (BI) of sites or are highly diagnostic (D) of stressors that may threaten or impair biotic integrity.

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relative abundances of species in two samples and bases similarity upon the sum of the lower relative abundances of each taxon in the two communities. The percentage community similarity (PSc; Whittaker, 1952) is a straightforward example, where PSC = Σi = 1,Smin(ai, bi) Here ai is the percentage of the ith species in sample a, and bi is the percentage of the same ith species in sample b. A second kind of similarity measurement is based on a distance measurement, which is a dissimilarity measurement rather than a similarity measurement, because the index increases with the greater dissimilarity (Pielou, 1984; Stevenson, 1984). Euclidean distance (ED) is a standard, where ED = √(Σi = 1,S(ai – bi)2) When using these dissimilarity indices, we recommend log-transforming relative abundances to reduce the importance and variability of common taxa. A third category of similarity indices only compares the observed taxa at an assessed site with the taxa that are expected at that site to determine the proportion of species that have been lost from assessed sites. This technique has been used widely with macroinvertebrates (Moss et al., 1999) and was recently employed with diatom genera (Chessman et al., 1999). This approach has the potential for increasing the precision of algal metrics by distinguishing species in three categories: species that should be there and are still there, species that have been lost, and species that have invaded. First, loss of species is an important impairment of biodiversity. Second, diagnosis of causes of impairment may be substantially enhanced by linking autecological information to whether species have resisted disturbance, not resisted disturbance, or invaded to exploit disturbance. In our application of the ERA framework, we emphasize the distinction between algal indicators that characterize designated use and ecological values in which the public are most interested and algal indicators that diagnose the stressors that may threaten or cause impairment of designated use. Some protocols recommend that metrics indicating the status of designated use be called “response” or “condition” indicators, and metrics that diagnose the physical, chemical, or biological factors that could be impairing designated use be called “stressor” or “causal” indicators (Paulsen et al., 1991; U.S. Environmental Protection Agency, 1998). This distinction of types of metrics emphasizes the diversity of information that can be obtained with algal assessments and how to apply that information in environmental problem solving. Therefore, algal indi-

cators that use environmental preferences of species, such as weighted average indices of pH or TP, will be described later under exposure assessment, rather that here under hazard assessment.

B. Testing Metrics Metrics can be tested with measurements of attributes at multiple sites with varying levels of human disturbance and either parametric or nonparametric statistical methods (see Sokal and Rohlf, 1998). Sites with different levels of human impact should be chosen and sampled to assess the ecological response to human disturbance. The level of human disturbance at sites can be characterized with multiple lines of evidence. When point sources of pollution occur, environmental gradients are relatively simple to establish with a reference condition upstream from the point source and a decreasing gradient of disturbance at increasing distances downstream from the point source. When non-point-source pollution is a concern, human disturbance can be estimated by land use type, intensity, and proximity to a habitat or by concentrations of contaminants. The multivariate nature of complex non-pointsource contamination can be simplified with the use of ordination techniques and axis scores as a ranking scale of human disturbance. Reference sites help define expected conditions in a habitat if it had not been affected by human activities (Hughes, 1995); these are typically the least impacted ecosystems in the region. Reference sites can be sites upstream from a point source of pollution in a stream, whereas test sites can be downstream. Alternatively, reference sites for a specific climatic and hydrogeomorphic class of habitats can be defined as a set of sites with lowest human disturbance or greatest riparian buffer within their watersheds. Reference sites may have the lowest level of a specific stressor in them, such as low phosphorus and other specific indicators of human disturbance. Algae are particularly useful in establishing reference conditions in lakes and wetlands, where sediments are continuously deposited because algal remains in sediments from times of low human disturbance can be used to infer historical conditions in those habitats. Paleolimnological approaches provide direct measurements of long-term environmental trends at a specific site, which increases certainty about how fast and the extent to which a system is deteriorating. To propose realistic mitigation procedures, paleolimnological reconstruction of past conditions can provide a realistic target for restoration. Long-term data can also show critical loads of pollutants or stressors that a system can handle before negative effects are manifested


(Smol, 1990, 1992, 1995; Anderson and Battarbee, 1994; and papers in Stoermer and Smol, 1999). Paleolimnological approaches are based on fairly straightforward principles. Under ideal conditions, sediments slowly accumulate at the bottom of lakes, without disruptions. Certainly, in some cases, problems may occur (e.g., excessive bioturbation), but these problems can usually be recognized and assessed. Over time, therefore, the history of the lake and its watershed is archived in the depth/time profile of the sediments. Incorporated in these sediments is a surprisingly large library of information on the conditions present in the lake (from autochthonous indicators), as well as environmental conditions that existed outside the lake (from allochthonous indicators). Physical, chemical, and biological information is archived in sediments; however, for the purpose of this chapter, we will primarily focus on algal data. Paleolimnology is now widely recognized as a robust environmental management tool. We mainly discuss lake paleoenvironmental studies in this chapter, as most of the research has centered on these systems. However, many paleo approaches can easily be transferred to other aquatic systems such as ponds (Douglas et al., 1994), rivers (e.g., Reavie and Smol, 1987, 1998; Amoros and Van Urk, 1989; Reavie et al., 1998), wetlands (Bunting et al., 1997), estuaries (Cooper, 1999), and marine systems (Anderson and Vos, 1992). After algal attributes from habitats with different levels of human disturbance have been assessed, their response to human disturbance can be characterized. Log-transformation of biomass-related variables (such as chl a, cell density, and biovolume) is recommended to meet the equal variances assumption of regression analysis and sometimes to make patterns more linear. Other data transformations may be necessary to meet assumptions of statistical tests (e.g., Green, 1979; Sokal and Rohlf, 1998) or to manage the sensitivity of metrics. For example, arc-sine transformations of proportional data and log or square-root transformations of relative abundances should increase the normality of the data (Sokal and Rohlf, 1998) and can reduce the importance of highly variable abundant taxa, which increases the precision of some metrics. Both nonparametric and parametric statistical techniques can be used to determine whether algal attributes respond to gradients of human disturbance. The simplest and most direct method is to compare attributes to the gradient of human disturbance by regression or correlation (e.g., Hill et al., 2000), if human disturbance can be quantified on a continuous scale. Alternatively, human disturbance can be categorized as low and high, and ANOVA or Mann–Whitney U tests can be used to test for differences in metrics with differences in human

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disturbance (Green, 1979; Barbour et al., 1992; Sokal and Rohlf, 1998; Barbour et al., 1999).

C. Multimetric Indices Summarizing data in the form of multimetric indices has been a valuable method for communicating results of complex analyses that often involve multiple lines of evidence. This method has been used commonly with fish and invertebrate assemblages as multimetric indices of biotic integrity (IBI) (Karr, 1981; Kerans and Karr, 1992), but it has also been used with periphyton (Kentucky Division of Water, 1993; Hill et al., 2000). Development of a multimetric index calls for selecting 6–10 metrics that describe a diversity of responses of assemblages that will be sensitive to all probable environmental stressors. For example, species richness, percentage of diatoms, percentage similarity to reference assemblages, number of taxa sensitive to pollution, percentage motile diatoms, percentage aberrant diatoms, inferred trophic status, inferred salinity (conductivity), inferred saprobity, and inferred pH could be used in a multimetric index, if they all responded to gradients of human disturbance (i.e., performed as good metrics). The range of each metric should be normalized, for example, to a range of 0–10, so that each metric has equal weight (see Hill et al., 2000). Then values of each metric for a sample can be summed. In an example with 10 metrics and each ranging from 0 to 10 in scale, the multimetric index would then range from 0 to 100. The value of multimetric indices is that they provide a single number as a summary of multiple lines of evidence. Such a summary statistic is highly valuable for communicating information to a lay audience, especially when compared with interpretations of ordination analyses (Karr and Chu, 1999). The disadvantage is that they may mask effects on one or two metrics; however, they can provide a hierarchically decomposable system of metrics for assessing ecological risk and even diagnosing causes or threats to impairment (Stevenson and Pan, 1999).

D. Multivariate Statistics and Hazard Assessment Multivariate statistics are powerful and informative statistical tools for determining the major patterns of change in species composition and relating them to physical, chemical, or other biological characteristics of the habitats studied. We regard a multimetric index of biotic integrity (IBI) and multivariate statistics as complementary tools. Cluster analysis and ordination provide multivariate methods for grouping stations by similarity in assemblage structure, exploring patterns in data, and

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illustrating those patterns (Hill, 1979; Pielou, 1984; Jongman et al., 1995). Cluster analyses can be bottomup, such as UPGMA, or top-down, like TWINSPAN. A recent evaluation comparing these approaches showed that UPGMA, compared with TWINSPAN, grouped artificial assemblages better (developed based on a selected set of assumptions and assignment of species’ relative abundances, based on a probabilistic distribution) (Belbin and McDonald, 1993). However, many researchers use TWINSPAN and find results with actual data to be highly interpretable (e.g., Pan et al., 2000). Ordination (e.g., correspondence analysis and principal components analysis) condenses patterns in assemblage characteristics to axes that explain the covariation in assemblage characteristics among samples. Similarity in species composition among sites and species responses to environmental conditions at sites can be compared by plotting sample scores and species loadings in ordination space (Fig. 5). Environmental factors can then be incorporated into correspondence analyses (CAs) to relate variation in species distributions and sampled sites to variation in environmental conditions among sites. Many canonical ordination techniques can be applied to relating species and environmental variance among sites. If great differences occur in species composition among sites, a U-shaped pattern in ordination scores of sites often occurs; this artifact can be reduced by using detrending techniques (e.g., DCA) (Jongman et al., 1995). Multivariate statistics are valuable for the early stages of environmental programs when initial relationships between changes in assemblages and environmental conditions are being explored to develop metrics and multimetric IBI (e.g., Pan et al., 1996). They can be an important part of any program when it is necessary to reduce the complexity of multivariate data. Even though cluster analysis and ordination group assemblages in classes that are often biologically interpretable, they do not test hypotheses that are directly related to questions of whether a site or a group of test sites is impaired or not. Testing these hypotheses calls for multivariate analysis of variance or discriminate function analysis (Pan et al., 2000). These latter approaches may be valuable for finding thresholds along gradients and being able to establish a probability that sites with specific characteristics were exposed to unacceptable levels of human disturbance and were impaired. Another weakness in multivariate statistics as an endpoint in ERA is that results are often not easily and repeatedly interpretable by audiences that are not trained in the use of multivariate statistics. Reviews of multivariate statistics and their use can be found in Green (1979), Pielou (1984), Jongman et al. (1995), and Birks (1995, 1998).

FIGURE 5 Plots of sample sites and species along ordination axes to indicate the relationship between similarity in species composition among sites and the species that are most important in defining that similarity (from Pan and Stevenson, 1996). (A) Plot of species and environmental variables (arrows) along ordination axes to show which species are most important in defining similarity among assemblages. (B) Plot of sample sites that shows sites with similar species composition located in similar locations along ordination axes with related environmental variables (arrows).

V. EXPOSURE ASSESSMENT: WHAT ARE ENVIRONMENTAL CONDITIONS? Exposure assessment may be as simple as measuring stressors, such as pH or phosphorus concentration, directly, or it may call for using biological indicators of


exposure. Exposure assessment is important for precisely characterizing the level or intensity of environmental conditions that may be affecting valued ecosystem components or services. Often, environmental conditions in a habitat cannot be measured accurately or precisely, particularly in shallow-water habitats like streams and wetlands, where environmental conditions change diurnally and seasonally with biological activity and from day to day with weather. Many algal taxa have long been recognized to be, with varying degrees of specificity, restricted to certain aquatic environments (Kolkwitz and Marsson, 1908); therefore, they can potentially be used as bioindicators of environmental conditions. The simplest of the quantitative stressor or causal indicators are simply the sum of relative abundances of organisms that are either tolerant or sensitive to a specific environmental stressor, such as the relative abundance of motile diatoms or aberrant diatoms, which indicate silt and heavy metal pollution, respectively (Bahls, 1993; McFarland et al., 1997). Alternatively, the relative abundance of organisms adapted to environmental extremes could be used to diagnose stressors, such as high organic contamination, high salinity, low dissolved oxygen, or low pH (Stevenson and Bahls, 1999). More complex quantitative approaches use species composition of algal assemblages and categorical rankings of species environmental preferences, with either weighted average equations (Zelinka and Marvan, 1961) or regression equations (Renberg and Hellberg, 1982) to infer the stressor level. Recently, new accessibilities to personal computers and new statistical techniques (weighted average assessment of species preferences) have enabled the development of more accurate characterizations of species environmental preferences and more accurate and precise biological indicators of stressors in ecosystems (see Birks, 1995, 1998, for reviews). Thus, stressor levels in a habitat can be inferred with weighted average equations with species autecologies that were developed based on a categorical ranking of species environmental preferences or with autecologies determined with weighted average techniques. Simple autecological ranks have been assigned to characterize environmental preferences for many taxa and many environmental characteristics (see van Dam et al., 1994, for a review). These autecological characteristics of taxa have been compiled in several reviews: Lowe (1974), Beaver (1981), Denys (1991), Hofmann (1994), and van Dam et al. (1994). Using a weighted average formula, stressor levels in habitats can be inferred based on the categorical autecological ranks of taxa (often eight or fewer categories) and relative abundances of taxa in samples. For example, a simple autecological index (SAI) for trophic status can be

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developed based on autecological ranks (Θi) of 0–7 for taxa observed to be most abundant in waters classified as ultraoligotrophic, oligotrophic, oligo- to mesotrophic, mesotrophic, meso- to eutrophic, eutrophic, and hypereutrophic, respectively (see van Dam et al., 1994). Then the trophic index can be calculated as SAITI = Σi = 1,SpiΘi where pi is the proportion of the ith species and Θi is the ecological condition in which the highest relative abundances of the ith species are collected. If autecological information is not known for all taxa, valuable information can still be obtained by correcting the index for the proportion of taxa with autecological characterizations. Next, one can redefine the community as the subset of taxa for which autecological characteristics are known by dividing the SAI by the sum of the proportional abundances of taxa with known autecological information. This weighted average approach with simple autecological characterizations of taxa has been used extensively in stream assessments with algae, particularly in Europe (Table II) (see reviews in Whitton et al., 1991; Whitton and Rott, 1996; Prygiel et al., 1999). Software and databases of diatom autecological characteristics (e.g., OMNIDIA; Lecointe et al., 1993) have been developed that can be used to calculate these indices. In tests of these indices, some perform better than others when used in regions other than those for which they were originally developed (e.g., Kwandrans et al., 1998). Regional calibration of these indices may be required to improve performance by reassessment of algal autecological characteristics. Stressor indicators based on accurate weightedaverage assessments of species’ environmental optima are more precise than indicators based on categorical characterizations of species’ autecologies (e.g., ter Braak and van Dam, 1989; Agbeti, 1992). However, acquiring accurate descriptions of species’ autecologies may be more difficult than using categorical characterizations. Weighted-average inference (WAI) models have been widely used to precisely infer environmental characteristics (Birks, 1998). The general principle for characterizing species autecological preferences is that, under most circumstances, the distributions of most algal taxa will exhibit a unimodal, Gaussian response curve (Fig. 6), if the gradient is long enough. The optimum (m) is estimated by the position along the environmental gradient where the taxon is most common, and the tolerance (t) can be estimated by the standard deviation of the curve. Because different taxa will have different optima and tolerances to environmental variables, these data can be used to make quantitative inferences of these variables (Fig. 7).

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FIGURE 6 Idealized pattern in relative abundances of a single algal species along an environmental gradient showing a unimodal curve with optima (µ) and tolerance (t) to environmental gradients.

These approaches, which have been used extensively in both paleo- and neo-environmental assessments, are statistically robust and ecologically sound (reviewed in Charles and Smol, 1994; Birks, 1995, 1998). In general, these quantitative inferences are based on transfer functions that have been derived from paleolimnological studies, typically with the use of surface sediment calibration sets (reviewed by Charles and Smol, 1994) or with the use of present-day algal assemblages from a large suite of sites (e.g., Siver, 1995; Reavie and Smol, 1997, 1998b; Stevenson et al., 1999; Pan et al., 1996).

Some of the largest and most robust ecological calibration sets, or training sets, have been developed by paleolimnologists using diatoms and chrysophytes preserved in the surface (recent) sediments from a set of calibration lakes. Briefly, a calibration set is constructed by choosing a suite of sites that have been well studied (i.e., limnological characteristics are well defined) and span the gradient of interest (e.g., pH, trophic status, etc.). For example, one of the most pressing environmental questions in North America in the 1980s was, “Have lakes acidified because of acid precipitation?” Very little long-term monitoring data were available, so paleolimnological approaches were used to infer past pH and related limnological variables. A major research effort was focused on the lakes in Adirondack Park (New York) (Charles et al., 1990). To develop transfer functions to infer past lakewater acidity levels in a suite of Adirondack lakes, a calibration set of 71 lakes was chosen that ranged in presentday pH from 4.4 to 7.8 (Dixit et al., 1993). From each of these lakes, the surface sediments (e.g., top 1 cm of sediment accumulation, representing the last few years of sediment deposition) were removed with a gravity corer (Glew, 1991). The indicators preserved in these sediments, in this case diatom valves (Dixit et al., 1993) and chrysophyte scales (Cumming et al., 1992a), were analyzed (identified to the species level, counted, and expressed as relative frequencies) from the surface sediments of the calibration lakes. This provides one of the matrices (i.e., the 71 lakes and the percentages of the taxa found in the recent sediments of lakes)

FIGURE 7 Weighted averaging calibration models for inferring lakewater pH (A) and lakewater total phosphorus (B) from diatom assemblages preserved in the surface sediments of 309 lakes in the northeastern United States. Modified from Dixit et al., (1999). The error is estimated by the bootstrapped root mean squared error.


required for the calibration (i.e., the species matrix). The second matrix was composed of the present-day environmental data collected for the 71 calibration lakes (in the above example, 21 limnological variables, such as lakewater pH, monomeric aluminum, dissolved organic carbon, nutrient levels, and depth were recorded). Once these two matrices are constructed, a variety of direct gradient analysis techniques (see reviews in Charles and Smol, 1994; Birks, 1995, 1998), such as Canonical Correspondence Analysis (CCA), can then be used to determine which environmental variables are most closely related to species distributions. Thereafter, weighted averaging calibration and regression (e.g., WACALIB; Line et al., 1994) were used to construct robust inference equations to infer lakewater characteristics (with known errors) from the diatoms or chrysophyte assemblages recorded in the sediments (e.g., Cumming et al., 1992b, 1994). Such approaches (e.g., Fig. 8) have been used in a variety of management

FIGURE 8 Paleolimnological assessment of acidification and recovery in Baby Lake, Sudbury, Ontario (modified from Dixit et al., 1992b). The core has been dated with 210Pb chronology. Closed circles represent inferred pH values from diatom assemblages, open circles represent values from scaled chrysophyte assemblages, and squares represent measured pH data (from Hutchinson and Havas, 1986) collected for the lake from various sources over the last three decades. These paleolimnological data clearly show a marked acidification of the lake in the middle part of this century, as a result of the emissions from the Sudbury smelters. Following the closure of the Coniston smelter in 1972, the fossil algal assemblages track a recovery pattern, which is matched by the measured pH data for this period. As is often the case, chrysophytes typically record more extreme acidification sequences, perhaps because they primarily bloom during spring, when the effects of acidification may be most severe.

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issues (see Smol, 1992, 1995; Anderson and Battarbee, 1994). The above calibration approaches can also be applied, in slightly modified forms, in settings where sediment accrual is not as regular as in lakes (e.g., some rivers and wetlands). For example, Reavie and Smol (1997, 1998) developed inference models from diatom assemblages attached to a variety of substrata in the St. Lawrence River and then used these transfer functions to infer past river conditions from sediment cores taken from fluvial lakes in the river system (Reavie et al., 1998). In situations where environmental conditions are highly variable, weighted averaging inference models have been shown to be better indicators of environmental conditions than one-time sampling and measurement of physical and chemical conditions (Stevenson, 1998). Field travel and sampling is an expensive part of program budgets, so habitats are often only sampled once. Water chemistry (TP concentration, for example) could be estimated based on a single water chemistry sample from a habitat or inferred based on algal species composition and autecological characteristics of those algae in a habitat. The precision of the estimate of mean TP concentration in a stream with a single water sample is the standard deviation of that assessment. The precision of an estimate of mean TP concentration in a stream with a single algal sample is the root mean square error of a weighted average regression model. Hence, 66% of estimates of mean TP concentration in a stream should be within one standard deviation of the measured TP concentration in a water sample and within one root mean square error of an inferred TP concentration from a weighted averaging inference model. Based on estimates of temporal variability in measured concentrations of TP along a wetland P gradient and among streams in a regional assessment, the standard deviation of the measured TP concentration was greater than the root mean square error of inferred TP with a weighted average model (Fig. 9; Stevenson, unpublished data). Although intuitively one might think that it would be imperative to use ecological calibration data taken only from the region of study, and that autecological data are not readily transferable from region to region, experience sugggests that this is not strictly the case. What is most critical in ecological calibration is to capture the range of environmental conditions that one will need to infer from the biological indicators. Although regional calibration data would be more likely to contain analogues, several studies have shown that data from geographically distant regions can also be used effectively. For example, as part of the SWAP acidification program in Europe, diatom calibration

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FIGURE 9 A comparison of total phosphorus (TP) variability estimated by one-time sampling and assay of water chemistry (SD_WQ) and by one-time sampling of diatom assemblages and inference with weighted average models (SE_DI). Variation in TP assessed in water chemistry (SD_WQ) is based on the standard deviation in assessed TP in the Everglades at one location over 2.5 years (provided by Paul McCormick, South Florida Water Management District, West Palm Beach, FL) and in a stream near Louisville, KY (Stevenson, unpublished data). The TP variability is based on the RMSE of diatom inference models from Everglades data (Slate, 1998) and midAtlantic Highlands streams (Pan et al., 1996).

data were effectively pooled from England, Norway, Scotland, Sweden, and Wales (Birks et al., 1990). Bennion et al. (1996) have combined regional diatom calibration sets from England, Wales, Northern Ireland, Denmark, and Sweden to develop robust transfer functions to infer lakewater epilimnetic phosphorus concentrations. As part of the CASPIA project, diatom calibration sets from North America, Africa, Australia, and Europe were pooled (Juggins et al., 1994). Ongoing work in our labs is also showing that calibration data are not as regional as may have once been thought, but that careful attention must be paid to taxonomic consistency and to the development of calibration data sets that effectively capture the necessary range of environmental variables.

VI. STRESSOR–RESPONSE RELATIONS Effects of specific environmental changes on assemblage characteristics can be determined at many temporal and spatial scales and with both observational and experimental approaches. The largest scale employs surveys of large regions (e.g., ecoregions) and correlates changes in environmental conditions and assemblages, which provides observations of potential stressor–response relationships. Correlations between

environmental factors and assemblage responses from surveys may show great changes and precision, and cause–effect relations may be biologically reasonable. However, experimental manipulation of environmental factors and measurement of assemblage response is important for more reliable confirmation of cause–effect relations. Experimental confirmation of stressor–response relations is particularly valuable in large-scale projects where expensive restoration efforts are planned and identification of the principal stressor is critical (e.g., McCormick and O’Dell, 1996; McCormick and Stevenson, 1998; Pan et al., 2000). Experimental approaches are also valuable in smallscale projects, where surveys of a large number of habitats are not practical because of budget or availability of habitats, and in toxicological studies of chemicals when chemicals are not yet widespread in the environment (Hoagland et al., 1993; Belanger et al., 1994). Observation of stressor–response correlations in largescale surveys is useful because the relative importance of multiple environmental factors can be compared. In addition, observation of stressor–response relations in surveys shows that responses occur in the natural setting and that the relation can be expected to hold over the range of conditions studied in the survey. Distinction between stressors and human activities that cause stressors is important in assessing stressor– response relations and in developing management strategies; this is the distinction between direct and indirect relations (U.S. Environmental Protection Agency, 1993; Yoder and Rankin, 1995; Kentucky Natural Resources and Environmental Protection Cabinet, 1997). Ecological responses to human disturbance may be caused directly by changes in physical, chemical, or biological conditions in a habitat (stressors) and indirectly by the human activities that cause those stressors. Distinguishing which stressors are most responsible for undesirable ecological responses and which human activities can be regulated to control those stressors is important for developing a plan to protect or restore environmental conditions. Many human activities (e.g., farming, logging, urbanization, sewage treatment plants) may be the source of a specific stressor (e.g., P enrichment). Many stressors (e.g., N and P enrichment, siltation, organic enrichment, and flow and light regimes) may be altered by a single human activity (e.g., farming). The magnitude and linearity of stressor–response relationships may vary with the attribute tested. Many results indicate that metrics based on higher levels of biological organization (ecosystem/community level: e.g., biomass and productivity) are less sensitive to environmental change than metrics based on lower levels of biological organization (community/popula-


tion: e.g., species composition) (Schindler, 1990; Leland, 1995). Because of high dispersal rates and high species numbers in microbial assemblages, species adapted to altered environmental conditions are probably able to invade and populate a habitat relatively quickly. Thus, impaired populations may be replaced by populations that are adapted to the altered conditions and thereby maintain ecosystem function (Stevenson, 1997). Therefore, biomass and many functional attributes of algal assemblages are often less sensitive to environmental change than changes in species composition (Schindler, 1990). Relating the stressor–response relations between algal species composition and environmental factors to stressor–response relations for ecosystem attributes may help in the understanding of ecosystem dynamics and establishing criteria to protect ecological integrity. Recently, results from the Everglades show punctuated (sudden, discrete) changes in algal species composition along a phosphorus gradient (Pan et al., 2000) (Fig. 10). Along the same gradient, higher-level biological attributes, such as biomass and productivity, change linearly or asymptotically. Thus, punctuated changes in species composition may result in multiple stable states (May, 1974) along an environmental gradient, which result from changes in the factor or factors that are the most important constraints on species composition. Punctuated changes in biomass and productivity may be blurred by species replacement along the phosphorus gradient, and spatial and temporal variability in other factors that have more short-term effects on biomass. Criteria for protecting the ecological integrity of a habitat may be established at thresholds along

FIGURE 10 Ordination of sites based on species composition of algal assemblages at sites along a phosphorus gradient in the Everglades (Pan et al., 2000).

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FIGURE 11 Linear (diagonal dashed line) and nonlinear (solid line) ecological responses along an environmental gradient (e.g., µg TP/L). Setting criteria for protection and remediation of ecological integrity may be facilitated by nonlinear ecological responses. Reference conditions (Ref) are indicated by the vertical arrow. The vertical line indicates a criterion.

gradients where punctuated changes in species composition occur that correspond to undesirable changes in more than one ecological attribute (Fig. 11).

VII. RISK CHARACTERIZATION AND MANAGEMENT DECISIONS Risk characterization relates assessments of exposures to stressor–response relationships to evaluate the level of threat to an unimpaired system or the probable stressors and intensity of stress of impaired systems (Fig. 2) (U.S. Environmental Protection Agency, 1992). Thus, based on a set of metrics and stressor–response relationships, we can predict ecological effects of exposures to specific stressors, assess the likelihood that effects will occur, or assess the likelihood that effects were caused by specific stressors or interactions among multiple stressors. Standard risk characterizations also increase risk ratings with increasing uncertainty in information. Therefore, we are concerned with both underprotecting as well as overprotecting our resources. Quantitatively, risk characterization on a metricby-metric and stressor-by-stressor basis can be conducted to assess the sustainability of ecological conditions and the level of impairment or restorability of impaired conditions (Stevenson, 1998) (Fig. 12). Sustainability can be defined quantitatively as the difference between

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that produce stressors that are causing or threatening environmental impairment. Biological assessments can actually be used to infer the human activities that cause the main ecological stressors (Yoder and Rankin, 1995; Stevenson, 1998), but detailing that approach is beyond the scope of this chapter. Thus, management options for controlling nutrients and deoxygenation in streams include reducing point sources and nonpoint sources of nutrients and BOD to streams. Point sources (e.g., municipal waste treatment plants) are easier to regulate and significantly reduce nutrient loading and related problems in many lakes and streams. However, further reduction in nutrient problems will require addressing nonpoint sources of nutrients. Thus, distinguishing the direct and indirect effects of stressors and human activities provides for a more focused approach to evaluating management options and developing a risk management decision.

FIGURE 12 Relating ecological response to the sustainability of ecological conditions in unimpaired ecosystems and to restorability of conditions in impaired ecosystems. Reference conditions (Ref) are indicated by the vertical arrow. The vertical lines indicate a criterion and assessed conditions at two sites. Assessed conditions at Sites 1 and 2 are unimpaired and impaired, respectively. Site 1 illustrates that the greater the difference between assessed conditions and the criterion for unimpaired sites, the greater is the sustainability of valued ecological attributes at a site. Site 2 indicates that the greater the difference between assessed conditions and the criterion for impaired sites, the less restorable a habitat is.

measured exposure (stressor) levels and the exposure criteria for protecting ecological integrity (or designated use of the ecosystem). Restorability is assessed for impaired ecosystems and is equal to 1/(difference between measured exposure levels and the criteria for protecting ecological integrity). These two measures of risk characterization can provide quantitative assessments of probable effects on ecological systems—the greater the sustainability, the lower the probability of stressor effects on ecological integrity, and the lower the restorability, the greater the probability of stressor effects on ecological integrity. Management decisions can be linked to risk assessment through risk characterizations and the stressor linkage to human activities and management options (Fig. 1). After risk characterizations are completed, management options for protecting or remediating environmental stressors will be considered with the many other factors that affect risk management decisions (e.g., political, economic, and regulatory factors). Management options depend upon the human activities

VIII. CONCLUSIONS Algae have been used successfully for environmental assessment of many streams, larger rivers, lakes, and wetlands around the world. Many different approaches can be used, which vary in the level of technical expertise required and in the amount of time required per sample. In this chapter we have tried not to make specific recommendations for which methods should be used, because different programs may call for different methods. In addition, we would not want to constrain the great promise for further development and linkage of algal methods for environmental assessment by making specific recommendations of methods. However, we have started to develop a framework for relating the different methods of environmental assessment and to explore the specific situations in which different methods should be used. We have linked the algal framework for environmental assessment to a standardized risk assessment framework so that assessments with algae can be better related to assessments with other organisms and other approaches (such as laboratory-based toxicology). Future developments in algal methods for environmental assessment should be directed to the ultimate goal of solving environmental problems. More rigorous hypothesis testing of the precision and sensitivity of algal indicators will be important in documenting the performance of algal indicators and encouraging their application. Transferability of algal indicators among regions should be rigorously evaluated because this allows development of consistent approaches across regions (and perhaps habitat type) and saves costs of developing regional indicators. Current efforts to devel-


op web-based access to autecological characteristics of taxa will be valuable for making this information available to a broader audience, but this approach also requires cautious evaluation of the quality of information on the web. Great challenges also exist for our more consistent application and integration of multiple lines of evidence, which may be facilitated by using a framework, such as the risk assessment framework. In addition, we must learn to communicate the results of our research more effectively and clearly to a broad audience, some of whom have little experience with interpreting mathematically complex information. Algae can cause ecological problems, but can also perform valuable ecological services. In a recent report by the National Research Council of the US (CEIMATE, 2000), total and native species diversity, productivity, trophic status, and nutrient-use efficiency were listed as fundamental ecological indicators for the United States. Algal attributes related to these indicators, as well as other valued ecological attributes of algae, should provide one part of a rationale for assessing algal properties of ecosystems. The second part of that rationale is the extraordinary sensitivity and diagnostic power of algae to detect environmental problems and identify their causes.

ACKNOWLEDGMENTS We acknowledge the contributions of anonymous reviewers and comments by the editors of this book, John Wehr and Bob Sheath. In addition, we are grateful for the openness of interactions among colleagues and students, which has moved our discipline forward so successfully. Stevenson’s efforts were supported by the U.S. Environmental Protection Agency. Smol’s efforts were supported by the Natural Sciences and Engineering Research Council of Canada.

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CONTROL OF NUISANCE ALGAE Carole A. Lembi Department of Botany and Plant Pathology Purdue University West Lafayette, Indiana 47907 I. Introduction II. Problems Associated with Algae A. Microscopic Algae B. Macrophytic Filamentous Algae C. Chara and Nitella

I. INTRODUCTION Algae play many important and beneficial roles in freshwater environments. They produce oxygen and consume carbon dioxide, act as the base for the aquatic food chain, remove nutrients and pollutants from water, and stabilize sediments. Excessive algal growths, however, can cause detrimental effects on aquatic systems, endangering the organisms that live in or depend on these systems and hampering or preventing human uses of the infested waterways. When we refer to the kinds of problems that algae cause, it is helpful to divide algae into three groups according to their growth habits: microscopic algae (primarily phytoplanktonic), filamentous mat-forming algae, and the Chara/Nitella group. Each group poses its own unique problems to aquatic systems. This chapter describes the problems caused by each of these three groups and then covers the control methods that typically are used for these algae.

II. PROBLEMS ASSOCIATED WITH ALGAE Many of the problems that the public associates with algae occur in more or less static bodies of water (i.e., ponds, lakes, and reservoirs) with long residence Freshwater Algae of North America Copyright © 2003, Elsevier Science (USA). All rights of reproduction in any form reserved.

III. Control Methods for Nuisance Algae A. Nutrient Manipulation B. Direct Control Methods Literature Cited

times. Algae also produce excessive or unwanted growths in flowing waters such as streams, rivers, and water delivery systems. Control of algae in these sites is more typically achieved with watershed management techniques that reduce nutrient inputs than with direct control methods. Nuisance algae found in irrigation canals and drainage systems can be and are managed using direct control techniques, but the options are more limited than those used in static systems. Although the algae of flowing waters are discussed, the emphasis in this chapter is on the problems caused by the algae of lakes and ponds.

A. Microscopic Algae The term “bloom” is typically reserved for excessive growths of microscopic, planktonic algae. Any discussion of bloom-forming algae starts with the cyanobacteria (also referred to as blue-green algae) Microcystis, Anabaena, and Aphanizomenon (see Chap. 3) Blooms of these prokaryotic organisms give a characteristic green or yellow-green color to water. Under static conditions, they rise to the surface to form very distinctive films and windrows of greenish scum (Fig. 1) [for a review of mechanisms regulating buoyancy, see Oliver (1994)]. Their notoriety is well deserved. In addition to being indicators of nutrient (particularly 805

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FIGURE 1 A surface scum formed by a cyanobacterial bloom of Microcystis.

phosphorus)-enriched waters, the presence of cyanobacterial blooms is a very visible symptom of deteriorated bodies of water. Population crashes (death) and the microbial decomposition of cyanobacterial cells result in the depletion of dissolved oxygen. Anoxic conditions can cause fish kills in bodies of water as small as prairie potholes (Barica, 1975, 1978) or as large as Lake Okeechobee, Florida (Jones, 1987). The largest cyanobacterial bloom recorded on Lake Okeechobee, which occurred on June 30, 1986, covered 337 km2, almost 20% of the lake surface (Lamon, 1995). The most infamous example of the adverse impact of algal blooms on a large body of water was the deterioration of the western basin of Lake Erie in the 1960s (Beeton, 1969; Rosa and Burns, 1987). Increases in the quantity of algal biomass (Davis, 1964) and shifts in species composition from diatoms to cyanobacterial blooms (Ogawa and Carr, 1969; Munawar and Munawar, 1976; Nicholls et al., 1980; Stoermer, 1988) exacerbated an already deteriorating water quality situation. The resulting oxygen-depleted conditions hastened the demise of native fish species and their replacement by invasive species such as alewife and lamprey. Crashes of cyanobacterial blooms also have an adverse effect on the aquaculture industry. A fish kill in an 8.9 ha aquaculture pond (Boyd et al., 1975) caused by the die-off of a cyanobacterial bloom resulted in oxygen depletion that killed 6800 kg of catfish. At today’s market value, the loss would have been worth between U.S. $11,000 and $19,000. Many other cyanobacteria, as well as planktonic chlorophytes, euglenoids, diatoms, synurophytes, and dinoflagellates can bloom in nutrient-enriched waters. An important phenomenon is the occurrence of red

water and red surface scums, caused by blooms of Oscillatoria (Planktothrix) rubescens, generally in large lakes in early spring (Jaag, 1972; Konopka, 1982a, b), and by species of Euglena and Trachelomonas in static waters in mid-to-late summer (Lackey, 1968). These red water-causing organisms should not be confused with red tides, which are marine, are composed primarily of dinoflagellates, and often produce toxic compounds. Other than being symptoms of highly nutrient-enriched waters, the red color-producing euglenoids of freshwaters are not toxic. The O. rubescens/agardhii complex has been reported to produce hepatotoxins (Carpenter and Carmichael, 1995) and may be responsible for dermatitis or skin irritation when people come in contact with contaminated water (Gorham and Carmichael, 1988), but documented reports of incidents are rare (W. W. Carmichael, personal communication) in comparison to reports of those caused by other cyanobacteria (discussed below). The presence of blooms of microscopic algae has long been associated with eutrophication (Schelske and Stoermer, 1971; Schindler, 1975, 1977; Reavie et al., 1995). As a result, most trophic classification systems (e.g., Carlson, 1977; Wetzel, 1983; EPA, 1990) are based on some measure of algal biomass (e.g., chlorophyll or cell volumes), Secchi disk readings (a measure of transparency that can be affected by algal biomass), and types of algae present. Cooke et al. (1993b) summarized the effects of eutrophication as follows: “Symptoms of eutrophication, such as algal blooms (including surface scums), low transparency, rapid loss of volume in reservoirs, noxious odors, tainted fish flesh, impaired potable water supplies, dissolved oxygen depletions, fish kills, and the development of nuisance or exotic animal populations (e.g., common carp) can bring about economic losses in the forms of decreased property values, high cost treatments of raw drinking water, illness, depressed recreation industries, expenditures for management and restoration, and the need to build new reservoirs.” While these authors describe algal blooms as one of several symptoms of eutrophication, the presence of the algal blooms themselves can lead to low transparency, noxious odors, tainted fish flesh, impaired potable water, dissolved oxygen depletions, and fish kills. From an economic standpoint, the most important problem caused by algal blooms is production of taste and odor in surface water supplies (Raman, 1985; Hawkins and Griffiths, 1987). The potential for significant taste and odor problems exists in the United States because more water for human uses is obtained from surface water than from groundwater. Approximately 65% of the 399 billion gallons of freshwater withdrawn for all purposes in the United States in 1985

24. Control of Nuisance Algae

was obtained from surface water sources (Solley et al., 1988). The cyanobacteria Microcystis, Anabaena, Aphanizomenon, and Pseudanabaena and the golden flagellates Synura, Mallomonas, and Dinobryon are common causes of taste and odor in water supplies, but diatoms, dinoflagellates, and even some green algae also cause problems (Palmer, 1962; Nicholls and Gerrath, 1985; American Public Health Association, 1992). Colonial species of the diatom Stephanodiscus were reported to cause undesirable odors and clog filter runs at municipal water plants on the Great Lakes (Stoermer, 1988). The fishy tastes and odors produced by Synura spp. are frequently reported in softwater lakes in Ontario, and the haptophyte Chrysochromulina breviturrita has been cited as a producer of particularly offensive odors in that area (Nicholls et al., 1982). The tastes and odors produced by cyanobacteria such as Anabaena and Oscillatoria are caused by two compounds: 2-methylisoborneol (2-MIB) at concentrations greater than 12 ng L–1 and geosmin at concentrations greater than 7 ng L–1 (Simpson and MacLeod, 1991). The cost of treating water for taste and odor problems is high. Copper sulfate (CuSO4) is widely used to control or eliminate potential taste- and odor-causing algae in water supply reservoirs. Water treatment with activated carbon is then used to remove undesirable tastes and odors that do occur. Chlorine has no effect on the removal of musty/earthy aromas nor does treatment with ozone or aeration (Maga, 1987). The cost to treat Lake Manatee (Florida) Reservoir water with activated carbon where blooms have occurred has exceeded U.S. $14,000 a day (Clarke et al., 1997). Additional expenses are incurred to replace the activated carbon filters, which frequently become clogged with humic substances, and to treat the lake water with copper sulfate. Taste and odor problems are typical not only of surface waters where blooms occur but also of deeper waters. In a study of six Kansas lakes, Arruda and Fromm (1989) noted that the mostly fishy and grassy tastes in the surface waters were caused by algae and correlated with the trophic status of the lake. The foul taste of the bottom water (musty, sulfurous, and rotten egg-like) was typical of highly organic, anoxic sediments and overlying water. Even this problem was related to algae because it was probably caused by the deposition and slow decomposition of organic matter contributed over time by dying algal blooms. Having to draw from deep waters to avoid infested surface water can lead to other problems such as the deposition of iron and manganese in pipes and on clothing in washing machines. The other major taste and odor problem caused by cyanobacterial blooms is off-flavors in the flesh of

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aquaculture-produced fish (particularly catfish) and other animals (Jüttner et al., 1986; Maga, 1987; Martin et al., 1991; Schrader and Blevins, 1993). Some of the causative organisms are species of Lyngbya, Oscillatoria, Aphanizomenon, Anabaena, and Phormidium. Geosmin (Brown and Boyd, 1982; Lovell et al., 1986) and 2-MIB (Martin et al., 1988; van der Ploeg et al., 1995) are the primary chemicals that produce off-flavors. In general, once fish have been tainted with these compounds, they must be moved to clean water for several weeks so the off-flavor can dissipate before they are marketed. The harvest and transport of the fish to a new site for cleansing are expensive and laborious. Off-flavor problems have been estimated to add $50 million each year to the cost of producing catfish in the United States (Schrader et al., 1997). Various cyanobacteria produce toxins that are harmful to humans and animals [for review, see Carmichael (1997)]. The major genera are Anabaena, Aphanizomenon, Microcystis, Cylindrospermopsis, Nodularia, and Oscillatoria. There are many reported instances of livestock, pets, wild animals, and birds that have died after drinking tainted water, but, in general, the adverse effects of cyanobacterial blooms to humans have been limited to forms of dermatitis and irritation of the mucous membranes. Some evidence that human gastrointestinal disorders were associated with consumption of water from reservoirs with blooms of cyanobacteria can be found (Carmichael et al., 1985; Carmichael and Falconer, 1993), and cyanobacterial toxins were implicated in the deaths of 26 people in Brazil when contaminated water was used for hemodialysis (Jochimsen et al., 1998). An association between toxins in water supplies and primary liver cancer in China has been suggested (Carmichael, 1994). Fortunately, the instances of human poisoning are rare because the unattractiveness and foul odors of water in which an algal bloom occurs usually deter people from using or drinking the water. Algal blooms can have adverse impacts on the health of organisms other than fish and humans. High nutrient concentrations in the water column trigger algal blooms, which reduce light penetration. Light reduction can severely limit the growth of submersed vascular plants (Spence, 1976; Jupp and Spence, 1977; Jones et al., 1983), thus decreasing habitat and shelter available for fish and fish food organisms. As a result, the most eutrophic lakes are those that are dominated by bloom-forming algae, with little or no submersed vascular plant production (Wetzel, 2001). In addition to loss of habitat, cyanobacterial blooms may cause a loss of system-level productivity. Some cyanobacterial species are allelopathic to other algae that are considered to be food sources for

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zooplankton (Keating, 1976, 1977, 1978), and they themselves are not significantly grazed by zooplankton (which is one reason why they can dominate aquatic systems). Cladoceran populations (e.g., Daphnia) decline or disappear when cyanobacteria, particularly the filamentous forms, predominate (Burns, 1968; Keating, 1976; Infante and Abella, 1985; Rothhaupt, 1991). The major reason for lack of predation appears to be a mechanical interference with feeding when the filamentous forms accumulate in the filtering apparatus. The loss in available energy suppresses zooplankton reproduction. Webster and Peters (1978) showed that as densities of filaments of Anabaena spp., Aphanizomenon flos-aquae, Oscillatoria tenuis, or Lyngbya spp. increased, the larger-sized cladocerans filtered at lower rates, increased rejection rates, and decreased brood sizes. The increased energy expenditure of trying to obtain sufficient food appears to increase the respiration rates of cladocerans (Porter and McDonough, 1984), further reducing assimilation efficiency, growth, and reproduction. As few as 50 cyanobacterial filaments per mL of water have an adverse effect on zooplankton feeding rates (Infante and Abella, 1985). The presence of filamentous cyanobacteria can cause a shift from large-bodied to small-bodied zooplankters, which feed on other materials such as small algae, bacteria, and organic debris. Some evidence suggests that the adverse effect of cyanobacteria on cladocerans also is due to toxin production (Infante and Abella, 1985; Fulton and Paerl, 1987; DeMott et al., 1991).

FIGURE 2 Filamentous algal mats (Spirogyra) causing obvious aesthetic problems in a lake.

B. Macrophytic Filamentous Algae The problems caused by macrophytic filamentous algae in aquatic systems are primarily due to their ability to form large mats of vegetation (Fig. 2). These algae are typically found in shallow water where they may be free-floating (e.g., Pithophora, Rhizoclonium, Spirogyra, and Hydrodictyon) or attached (e.g., Cladophora, Ulothrix, Stigeoclonium, and Oedogonium) to substrata, either living (plants and other algae) or nonliving (rocks, cement linings, and sediments). The freefloating forms are generally restricted to static waters such as ponds and the sheltered littoral zones of lakes. Attached forms occupy a much wider range of habitats. They are found in both static and flowing systems, including the wave-scoured edges of lakes (e.g., Cladophora in the Laurentian Great Lakes), fast-flowing streams, and the extensive irrigation systems and aqueducts of the western United States (Fig. 3). The genera listed above are all green algae. Filamentous cyanobacteria also can form free-floating mats. The cells of filaments of Lyngbya wollei (Speziale

FIGURE 3 Filamentous algal mats (Cladophora) in an irrigation canal that have broken away from the sides and are floating downstream. These mats can clog irrigation intakes and pumps. Photo courtesy of Lars Anderson, USDA-Agricultural Research Service.

and Dyck, 1992) are quite large (cell diameter: 25– 64 µm, length: 2–11 µm) for a cyanobacterium, and the dimensions, coarseness, and even color (dark green) of the filaments may cause the untrained observer to think that they are handling a filamentous green alga. The mats are dense and can completely cover ponds and shallow areas of lakes. Many species of Oscillatoria form benthic mats that break free from the bottom and float to the surface (Fig. 4A) when gas bubble accumulation dislodges the mats (Halfen and McCann, 1975). These growths are typically dark bluegreen to black in color and are quite slimy. The mats are often coated with sediments that were deposited on them while they were still associated with the bottom


FIGURE 4 Mats of macrophytic Oscillatoria. (A) Infestation of free-floating mats that have broken loose from the sediments in the shallow cove of a lake. (B) Close-up photo of the free-floating mats. The light color is due to the sediment deposited on the surface of the mats. Some of the mats have been turned upside down and show the natural dark color of the organism. Photos courtesy of Neil Gerber, Aquatic Management, Bluffton, Indiana.

substratum (Fig. 4B). Several species of Phormidium form the “black algae” growths that attach to the cement linings of swimming pools (Fitzgerald, 1959; Adamson and Sommerfeld, 1978). The occurrence of filamentous algae is widespread. The Florida Department of Environmental Protection, Bureau of Aquatic Plant Management, recently surveyed 451 public bodies of water in Florida (Schardt, 1994). Filamentous algae were the only submersed “plant” grouping observed in more than 50% of the water bodies (62%) and were the dominant group in 16% of the waters. They represented one of only five plant groupings that showed an increase over the previous 12 years (increasing by 22%, more than any of the other plants). The two genera that were singled

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out as being predominant were Pithophora spp. and L. wollei. In an unpublished 1995 survey (J. Schardt, personal communication), L. wollei was collected in more than 218 lakes in Florida. An indication of the severity (or perceived severity) of algal problems in the state is the fact that approximately 90% of the calls from individuals or lake associations to aquatic plant management services in Florida seek information on algae control (J. Williams, The Lake Doctors, Winter Springs, FL; personal communication). Extensive survey data are less available for other parts of North America, but filamentous algae are a widespread management problem. In the midwestern United States, approximately 60–70% of the total volume of aquatic plant control chemicals applied is for the control of (primarily) filamentous algae (R. Johnson, Aquatic Control, Seymour, IN; personal communication). Cladophora glomerata, Pithophora spp., and Rhizoclonium spp. are listed as causing problems in the western United States and Canada, and Cladophora and Chara are on the list of the 13 most consistently problematic aquatic weed species in these regions (Anderson, 1993). Excessive growths of mat-forming algae, either alone or in combination with aquatic vascular plants, impair recreational activities such as swimming, fishing, and boating. Swimming beaches fouled with algal mats are not only unappealing but also hazardous when ladders, rocks, and submerged concrete are coated by slime-producing species such as Spirogyra (Bennett, 1971) and cyanobacteria. Cladophora growths in the Great Lakes were noted as posing a potential danger to young and inexperienced swimmers who might become entangled in the mats and drown (Herbst, 1969). When large odiferous masses are washed up on the shore they decay and are aesthetically unpleasant, are a barrier to recreation, and are implicated in taste and odor events in drinking water supplies (Brownlee et al., 1984; Painter and Kamaitis, 1987; American Public Health Association, 1992). The loss of recreational and aesthetic values can have a significant economic impact on waterfront properties. Ormerod (1970) reported that the value of real estate on Lake Erie fronted with Cladophora mats averaged 80–85% of the value of clean frontage. There is some evidence that mat-forming cyanobacteria (e.g., L. wollei, Oscillatoria spp.) also produce toxins similar to those produced by the phytoplanktonic cyanobacteria (Gunn et al., 1992; Carmichael et al., 1997; Onodera et al., 1997). Although there are no reports to date of animals being killed by the toxins produced by L. wollei, dogs were reportedly killed by the toxins from benthic mats of Oscillatoria (Gunn et al., 1992).

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Macrophytic algae restrict and greatly reduce the efficiency of culture and harvest activities in fish culture ponds (Tucker et al., 1983). They may compete with phytoplankton, thus reducing the base of the aquatic food chain (Boyd, 1982). A 50% reduction in fish production in farm ponds was attributed to heavy Pithophora growth and concomitant loss of phytoplankton (Lawrence, 1954). Although reports implicating macrophytic algae as direct causes of fish kills are few (e.g., Robinson and Hawkes, 1986), excessive algal growth must add to the oxygen deficits that result from respiration of submersed plant growth and/or phytoplankton at night, during periods of cloudy weather, or under snow-covered ice. Fish kills in ponds that are dominated by filamentous algae are not uncommon (personal observation). Oxygen deficits can be stressful to fish in other ways by causing declines in food consumption and growth and by making them more prone to bacterial infections (Boyd, 1982). Overpopulation, stunted or reduced growth, and a decline in the capture efficiency of forage species by predatory fish have been attributed to dense vascular plant growth (Colle and Shireman, 1980; Savino and Stein, 1982; Shireman et al., 1983), and algal growth is assumed to contribute to this problem. However, the need for some macrophytic growth to provide protection for young fish and habitat for fish food organisms is also recognized (Barnett and Schneider, 1974; Wiley et al., 1984). The proper balance of macrophyte coverage and density for optimal fisheries is at this point somewhat controversial (Bettoli et al., 1992, 1993; Hoyer and Canfield, 1996a, b; Maceina, 1996), in part because of the variability in study sites and interpretations. The possible beneficial or detrimental roles of macrophytic algae as fish habitat clearly require further study (Hinkle, 1986). Cladophora, Stigeoclonium, Oedogonium, and Ulothrix have been cited as presenting serious problems in irrigation canals because they attach to concrete canal linings, thus reducing both flow rate and capacity. Cladophora can become associated with beds of pondweeds (Elodea spp.) and coontails (Ceratophyllum spp.), which increases resistance to the flow of water (Mitchell et al., 1989). Mats that break away from the linings float downstream where they foul pump inlets, irrigation siphons, trashracks, and sprinkler heads (Hansen et al., 1984). C. glomerata in particular is a major problem in water delivery systems in the western states. For example, in the 620 km length of the Salt River Project Canal, which supplies water and electricity for the cities of Phoenix and Tempe, Arizona, the control of aquatic weeds is a major operation/maintenance task (Corbus, 1982). The annual budget in the late 1980s for aquatic macrophyte

control (primarily Cladophora) in this system was approximately U.S. $1.5 million. The clogging of rivers, canals, and drainage ditches by aquatic plants and algae can prevent adequate drainage so that water backs up, even to the point of causing flooding. Another problem associated with water conveyance or storage systems, particularly in arid parts of the world, is the potential to lose water through evaporation from floating or emergent plant surfaces. Although considerable data are available to show that substantial loss does occur with coverage of vascular plants such as water hyacinth (Brezny et al., 1973), nothing is known of the potential of surfacefloating algae to add to the problem. The impacts of filamentous algae on the dynamics of food webs in natural systems have not been well documented. Most studies that have been conducted have focused on Cladophora. For example, Cladophora is not a major food source for the invertebrates or fish that live in lakes (see reviews by Lembi et al., 1988; Dodds and Gudder, 1992), although it is grazed by fish in river systems (Power, 1990). Cladophora provides an extensive surface area for colonization by periphyton and invertebrates, and the limited grazing that does occur may have more to do with ingesting these associated organisms than with the filamentous alga itself (Dodds and Gudder, 1992). On the other hand, dense growths of Cladophora were reported to reduce invertebrate diversity and to have disrupted shoal spawning by walleye, whitefish, and lake trout in the Great Lakes (Neil, 1975). Filamentous algal mats compete with submersed vascular plants for space and light. Examples include the replacement of angiosperms, such as Najas marina, by Spirogyra (Phillips et al., 1978), diverse macrophytes by C. glomerata (Bolas and Lund, 1974), Elodea by Cladophora and Spirogyra (Simpson and Eaton, 1986), and Potamogeton pectinatus by Cladophora (Ozimek et al., 1991). Phillips et al. (1978) suggested that replacement of submersed vascular plants in lakes undergoing eutrophication may be due more to the shading by epiphytic and filamentous algae than to phytoplankton. The diversity among filamentous algae, for example, the sliminess of Spirogyra (which appears to prevent colonization by periphyton) in contrast to the thick cell walls of Cladophora (which provide an excellent substratum for periphyton), the summer domination by Pithophora in contrast to the spring/fall distribution of Cladophora, and the net-like habit of Hydrodictyon or the unbranched habit of Spirogyra in contrast to the branched habit of Cladophora, suggests that much remains to be learned about the micro-niches that these algae make available to invertebrate and fish communities and their impacts on food webs of static freshwater systems.


C. Chara and Nitella Charophytes are usually viewed as being beneficial components of aquatic systems, and their reestablishment is an important factor in lake restoration (van den Berg et al., 1998b). Chara and Nitella are considered excellent habitats for littoral invertebrates (Rosine, 1955; Quade, 1969; Allanson, 1973; Hargeby et al., 1994) and fish (Fassett, 1957; Schardt, 1994), and they are a major food source for herbivorous waterbirds (Hargeby et al., 1994; van den Berg et al., 1998b). Their ability to form low-growing meadows of vegetation reduces the resuspension of sediments (van den Berg et al., 1998b). These macroalgae, however, can cause problems in shallow water when their growths reach the surface of the water, thereby preventing successful angling, swimming, and boating (Fig. 5). Chara and Nitella are typically named when weedy submersed species (mostly vascular plants, such as Ceratophyllum, Myriophyllum, and Elodea) are

FIGURE 5 A solid stand of Chara infests this pond. Although the vegetation has not formed a surface canopy, the underwater growth limits swimming and fishing success.

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listed. For example, Steward (1993) listed Chara and Nitella as among the plant groups causing weed problems in the eastern United States, and Anderson (1993) cited these genera for the western United States also. Charophytes also produce repellent (allelopathic) materials that exclude certain limnetic species of invertebrates (Pennak, 1966, 1973) and phytoplankton (Gibbs, 1973; Anthoni et al., 1980; Wium-Andersen et al., 1982). The latter finding may provide a partial explanation for the lack of epiphytes and clear water conditions frequently associated with some charophyte species [Crawford, 1979; Wium-Andersen et al., 1982 (but see Chap. 2, Sect. II.F.4)]. Chara is common in regions with hard water (e.g., areas of the Midwest with a limestone bedrock), and Nitella is more characteristic of soft waters (e.g., granitic regions of the Northeast). In a survey of 451 water bodies in Florida (which has regions of both soft and hard waters), Schardt (1994) collected Nitella in 64 bodies, and found that it was dominant in 28 of them. Chara was found in 52 bodies and was dominant in 15 of them. An interesting characteristic of Chara is that it tends to colonize sites in which vascular plants have been controlled (Nichols, 1984). Drawdown, the draining and exposure of shallow or shoreline areas to desiccation, eliminated water shield (Brasenia schreberi), restricted the spread of parrot feather (Myriophyllum brasiliense) and water lily (Nymphaea odorata), but enhanced the infestation of Chara vulgaris in a Louisiana reservoir (Lantz et al., 1964). In a survey of the effects of drawdowns, C. vulgaris increased in 33 cases, decreased in 15 cases, and stayed the same in 44 cases (Cooke et al., 1993b). Invasion or expansion by Chara has also been documented after dredging (Born et al., 1973; Nichols, 1984), mechanical harvesting (Anonymous, 1990), and the application of herbicides for the control of vascular plants (C. A. Lembi, personal observations). Opening of sites disturbed by weed control activities to light is the major reason cited for the invasion by Chara (Born et al., 1973), and recent studies seem to confirm that irradiance is a major factor regulating charoid distribution (Steinman et al., 1997). Although some evidence suggests that as eutrophication proceeds, charophyte populations may be reduced because of their sensitivity to “toxic” levels of phosphorus (P) (Forsberg, 1965), other studies show that increased P levels do not have an adverse effect on charoid growth (Blindow, 1988). Melzer et al. (1977) suggested that increased P concentrations play an indirect role in the disappearance of Chara, primarily by causing an increase in phytoplankton growth and turbidity, which in turn shades out charoid growths. The restoration of

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a Chara community in one system was achieved by reducing P concentrations, which resulted in higher water transparencies (Simons et al., 1994). However, light may not be the only factor critical to Chara establishment, particularly in mixed plant communities. van den Berg et al. (1998a) experimentally observed that Chara was negatively impacted by shading from sago pondweed (Potamogeton pectinatus), but the fact that Chara dominates sago pondweed in some clear water lakes suggests that light is probably not a key factor in that domination. The more efficient use by Chara of carbon (HCO3–) at low concentrations, which are typical within Chara meadows, has been suggested as a possible reason for its dominance (van den Berg et al., 1998b).

III. CONTROL METHODS FOR NUISANCE ALGAE Management practices for nuisance algae are divided into two major categories: nutrient manipulation and direct control techniques. Nutrient manipulation, particularly reduction of nutrient inputs, should be viewed as the best approach for long-term control of algal problems. There are situations for which significant nutrient reduction is impractical or ineffective; under these conditions, direct control of the algal biomass may be the only alternative available. Direct control methods should only be viewed as temporary solutions and should be coupled with longer-term strategies for reducing nutrient inputs.

A. Nutrient Manipulation It has long been known that inputs of nutrients, particularly P, stimulate algal growth. Many studies have shown a strong correlation between total phosphorus (TP) and planktonic algal biomass (Dillon and Rigler, 1974; Jones and Bachmann, 1976; Carlson, 1977; Schindler, 1978; Prepas and Trew, 1983). The positive relationship between chlorophyll-a concentrations (or shallower Secchi disk transparencies) and TP is a commonly used tool to predict water quality and trophic status (Vollenweider, 1969; Dillon and Rigler, 1974; Dillon et al., 1988). Some lakes are nitrogen (N)-limited. For example, a number of Florida lakes are surrounded by rich phosphate-containing deposits and soils and therefore may be N-limited. In a study of 223 Florida lakes, 27% were considered N-limited (Canfield, 1983). Also, studies of lakes in the semiarid and mountainous regions in the western United States indicate that the importance of N may be equal to or greater than that of P in limiting phytoplankton growth (Elser et al.,

1990; Reuter et al., 1993). N limitation, as well as P limitation, has been implicated in the regulation of filamentous algal growth (Spencer and Lembi, 1981; O’Neal et al., 1985; Dodds and Gudder, 1992). In addition to the amount of phytoplankton biomass that is produced with P or N additions, another consideration is species composition, particularly in relation to nutrient ratios. When sufficient silicon (Si) and N are available in relation to P (high Si:P and N:P ratios), diatom growth appears to be favored (Tilman and Kiesling, 1984). These conditions are typical of spring periods in temperate lakes following turnover or when sediment deposition occurs with spring rains. In late spring or early summer, green algae may dominate over diatoms as Si concentrations decrease [lower Si:P ratios (Sommer, 1983)]. From a management standpoint, the most critical nutrient ratios are low N:P or Si:P. These generally occur under conditions of excessive P loading, and it is under these circumstances that N-fixing cyanobacteria (Anabaena, Aphanizomenon, and others), which fix atmospheric N2 when the water becomes N-depleted, become dominant (Schindler, 1977; Smith, 1983), particularly during the summer months. For example, the shift from high Si:P and N:P ratios to low ratios was concomitant with the shift in dominance from diatoms to cyanobacteria in the Great Lakes in the 1960s (Schelske and Stoermer, 1971; Schelske, 1975). For this reason the emphasis on nutrient removal for generally improving water quality has been placed on P rather than on N. Another approach for maintaining a high N:P ratio is to increase N rather than to decrease P. In fact, several researchers (Leonardson and Ripl, 1980; Smith, 1983) suggested that N removal could be counterproductive and that N addition might actually be helpful in increasing populations of green algae or diatoms in relation to cyanobacteria. Barica et al. (1980) added N to ponds with low N:P ratios to see if it could reduce the incidence of cyanobacterial blooms, particularly blooms of Aphanizomenon that were causing fish kills when they crashed. The initial N:P ratios in these ponds were around 4 to 5 [Schindler (1977) found that cyanobacteria dominate when N:P ratios dropped from 15 to 5]. When low amounts of N were added (0.1 g N m–3 d–1) prior to bloom formation (but not during the bloom) or when high amounts (1 g N m–3 d–1) were added during the bloom, a shift from Aphanizomenon to green algae and cryptomonads occurred. The technique worked, but it was not considered to be a realistic approach because N would have to be added over several to many weeks. Although similar results were reported by Stockner and Shortreed (1988), the general consensus is that, when possible, it is much


better to reduce P concentrations than to elevate N concentrations. In fact, increasing the N:P ratio stimulated the growth of non-nitrogen fixing cyanobacteria such as Lyngbya, Oscillatoria, and Chroococcus in mesocosms placed in Lake Okeechobee (Havens and East, 1997). In an analysis of the literature, Elser et al. (1990) suggested that both P and N potentially limited algal growth. They found little support for P alone as a causative factor. However, they recommended that efforts should concentrate on P reduction because it is easier to achieve from a technical standpoint than N reduction. Clearly where N fixation by planktonic cyanobacteria is a response to N reduction, P should be the more reliable means to lower algal biomass. The same general approach is used for the control of filamentous algae and Chara, although the situation regarding specific nutrient ratios and target amounts is far from clear. There are three general approaches for achieving P reduction: decrease external P loading, suppress internal P loading, and increase P output from the system. External inputs of P can be decreased with diversion and advanced wastewater treatment, with detention basins and wetlands, and by the initiation of other watershed management techniques. Internal P loading can be suppressed with alum applications, dredging, and aeration. P-laden waters from the site can be released with hypolimnetic withdrawal.

1. Diversion and Advanced Wastewater Treatment These two techniques (used together) are the most frequently used methods to reduce external loading. Diversion is achieved primarily through sewage collection systems, and the water is then subjected to tertiary treatment in which P is removed by alum (aluminum sulfate), lime (calcium carbonate), or iron (ferric chloride). There have been a number of successes (for case histories, see Cooke et al., 1993b). Probably the best example is Lake Washington in Seattle, Washington (Edmundson and Lehman, 1981; Edmundson, 1994), in which 88% of the lake’s external loading was diverted from 1964 to 1967. TP declined from a mean annual concentration of 64 µg L–1 prior to diversion to 21 µg L–1 5 years after diversion. Chlorophyll-a decreased from 36 to 7 µg L–1 by 1969. Secchi disk depth increased from 1 to 3.1 m. Further reductions in algal biomass were attributed to increased populations of Daphnia [following a decline in planktonic Oscillatoria, which negatively impacts Daphnia feeding (Infante and Abella, 1985)] and a decrease in a planktivorus crustacean (Neomysis mercedis) population. The condition of the lake in the late 1970s was 17 µg L–1 TP, 3 µg L–1 chlorophyll-a, and 7 m Secchi

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disk depth, and it had clearly shifted from a eutrophic to a meso- or oligomesotrophic state. During the 1970s, significant reductions in P loading to Lake Erie also were achieved through legislation that upgraded sewage treatment to include chemical precipitation of P and reduced the allowable levels of phosphates in laundry detergents (Phosphorus Management Strategies Task Force, 1980). Declines in phytoplankton biomass averaged about 5% per year over the period from 1970 to 1985 (Nicholls and Hopkins, 1993) and were correlated with the resultant reduction in P loading (Nicholls et al., 1977, 1980). Interestingly, phytoplankton populations continue to decline due to removal by zebra mussels (Nicholls and Hopkins, 1993). Filamentous algal growths also respond to nutrient diversion. For example, Cladophora biomass and tissue P concentrations at seven sites in Lake Ontario steadily decreased from 1972 to 1983 in response to P control programs introduced in the early 1970s (Painter and Kamaitis, 1987). Diversion and treatment work best where there are distinct point sources of nutrient inputs. They are less successful at sites impacted by nonpoint sources or in which significant concentrations of nutrients have been stockpiled in the sediments and are a major source of internal loading.

2. Detention Basins and Wetlands Discharge of domestic wastewater and urban runoff into detention basins (also called retention ponds) or natural or constructed wetlands is often recommended for improving water quality before release into a river or lake (Mitsch and Gosselink, 1993; Olson, 1993; Etnier and Guterstam, 1997). Many local and some state ordinances now mandate the construction of retention ponds in new housing developments, industrial parks, and similar sites. These ponds mostly serve as settling basins for sediments and associated nutrients and other pollutants (Walker, 1987; Robbins et al., 1991). Of course, these sites themselves become ideal environments for the development of algal blooms and mats and are in large part the cause for the substantial increase in the number of companies offering aquatic plant and algal management services in recent years (C. A. Lembi, personal observation). Although algae and other aquatic plants in retention ponds serve as a filtration system for nutrients, urban residents frequently complain about having to look at scummy water! Highly vegetated wetland areas also act as settling basins; in addition, they provide biological filtering, uptake and storage, and transformation (e.g., denitrification) of nutrients (Mitsch and Gosselink, 1993). Although the P storage capability can be lost in temperate areas in winter when the plants are no longer

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taking up P and nutrients are released, wetlands do tend to store considerable P in the summer, which is the critical time for algal blooms to occur in downstream sites (Cooke et al., 1993b).

3. Watershed Management The importance of a broad watershed management program to reduce both point and nonpoint sources of fertilizers and other pollutants is gaining increased recognition at local, state, and federal levels. In agricultural areas, the promotion of best management practices (BMPs) has resulted in widespread acceptance of practices that reduce erosion of nutrient-laden soils (Scholze, 1994; EPA, 1998). Such practices include no-til and conservation tillage, vegetated filter strips and grass waterways, lowering of fertilizer application rates, and proper handling of animal manures. The adoption of BMPs in the United States is voluntary although cost-sharing programs are available through federal agencies such as the Farm Services Agency and the Natural Resources Conservation Service. Section 303(d) of the Clean Water Act calls for the implementation of total maximum daily loads into streams and lakes that have low water quality, and the Clean Lakes Program provided assistance in watershed management and improving water quality in lakes prior to 1995. Clearly, there is general recognition of the importance of watershed management in improving water quality, and the erosion of sediments into waterways has been considerably lessened. There are, however, still areas where the implementation of programs has been slow. An example is the discharge of animal wastes into rivers in North Carolina and Maryland watersheds, which appears to have resulted in fish-killing blooms of the estuarine dinoflagellate Pfiesteria piscida (Burkholder et al., 1997).

4. Alum In many situations, reduction of external P loading does reduce algal growth. This is particularly true in water where most of the P loading to the photic zone is from external sources and in deep stratified lakes where P released from the anaerobic bottom sediments and hypolimnion does not reach the photic zone. In shallow lakes, on the other hand, significant quantities of P can be released from the sediments and reach the photic zone (Wetzel, 1990; Cooke et al., 1993b). Therefore, reduction of external P loading may not have much of a short-term impact on phytoplankton growth in these sites. Resuspension of sediments is considered a potential source of nutrients for phytoplankton production in many shallow lakes (Carper and Bachmann, 1984; Riley and Prepas, 1984; Hansen et al., 1997; Havens and James, 1997). Stauffer and Lee

(1973) calculated that all of the summer algal blooms in Lake Mendota, Wisconsin, could be accounted for by internal loading of P from the lower waters and sediments to the photic zone. Therefore, steps to reduce internal P cycling in many of these lakes may be more effective in reducing algal growth than the reduction of external P inputs. The methods used to reduce internal P loading include chemical treatment with alum, the removal of sediments by dredging, and aeration. Alum (Al2[SO4]3) is used to lower P availability through P precipitation and to retard P release from the lake sediments (P inactivation). When added to water, alum and P form aluminum phosphate and a colloidal aluminum hydroxide floc to which certain P fractions are bound (Cooke et al., 1993b). The floc settles to the sediment and continues to sorb and retain P within the lattice of the molecule, thereby preventing further release of P. Sodium aluminate (AlNaO2), which is a good buffering material, is added to alum treatments to maintain pH values between 6 and 8 (Kortmann and Rich, 1994) because a severe shift in pH can be detrimental to fish populations. In addition, alum is not recommended for use in waters with an acidic pH or low alkalinity because of the potential for aluminum toxicity to fish at pH values below 5.5. Iron salts also can be used to inactivate P (Kortmann and Rich, 1994), and treatments with calcium salts (Ca(OH)2 and CaCO3) have successfully reduced P loading from bottom sediments in Canadian lakes (Prepas et al., 1990; Babin et al., 1994). There are numerous examples of success with alum treatments in shallow lake systems, and many treatments last from 2 to 15 years (Welch et al., 1988; Smeltzer, 1990; Cooke et al., 1993a; Jacoby et al., 1994; Welch and Cooke, 1995). In some lakes, internal loading has been significantly reduced for up to 20 years (Welch and Cooke, 1999). Holz and Hoagland (1999) reported improved water clarity, decreased chlorophyll-a concentrations, reduced cyanobacterial biomass and abundance, increased Daphnia biomass and abundance, and increased usable fish habitat in a shallow (mean depth = 4 m), alum-treated lake in Nebraska. To ensure success and long-lasting effects, reduction of internal P cycling must be accompanied by a reduction in external P loading. Factors that can lead to failure of an alum treatment include continued high external P loading (Welch et al., 1988; Barko et al., 1990), redistribution of alum floc to the lake center by wind mixing (Garrison and Knauer, 1984), and P recycling from senescing rooted macrophytes or from macrophytes that expand their range due to improved water clarity (Welch et al., 1988; Welch and Cooke, 1999). There also is evidence that cyanobacteria newly


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recruited from the sediments can transport P into the water column, even in alum-treated lakes (Perakis et al., 1996).

5. Dredging Dredging offers a more permanent solution to internal P loading in shallow lakes than alum treament because sediments, the actual source of the P loading, are removed from the system. Dredging, however, is much more expensive than alum. According to Cooke et al. (1993b), dredging costs nearly 30 times more than alum initially although over the long term (repeat alum treatments every 10 years), the cost differential is only 5 times greater if totaled over 50 years. The cost of alum treatment averages about U.S. $700 per ha and the cost of dredging is about U.S. $20,000 per ha (Cooke et al., 1993b). As with all nutrient reduction approaches, external loading must be reduced or eliminated to achieve long-term results.

6. Aeration P is released from sediments under anoxic conditions. The function of aerators (other than to improve habitat for fish) is to oxygenate the water column, or portions of the water column, and the upper layers of the sediments, thereby preventing the occurrence of low-O2 conditions. In theory, oxidized forms of P are not released into the photic zone to encourage phytoplankton blooms. There are two major methods for aerating pond or lake water (Cooke et al., 1993b; Kortmann and Rich, 1994). The first is artificial circulation. This method oxygenates the whole water column. Air is pumped from a compressor on shore through a tube to a weighted diffuser unit that is placed on the bottom. Air bubbles pass from the diffuser into the water and are often visible as a surface “boil” (Fig. 6A). This method destroys or prevents thermal stratification; therefore, it is not feasible in sites where deep cold water is necessary to maintain coldwater fish populations. It is, however, a good solution to potential oxygen depletion problems for warmwater fish species. The second method is termed “hypolimnetic” aeration. This method maintains stratification because the water is removed from the hypolimnion, oxygenated at the surface, and then returned to the bottom. Hypolimnetic aeration is used in deep lakes to overcome anoxia, improve coldwater fisheries habitat, and control sediment P release. The impacts of either type of aeration method on algal blooms have been difficult to document. The sediment–water interface in many shallow lakes may already be oxygenated, in which case aeration will not have an impact. Cooke et al. (1993b) summarized data

FIGURE 6 Aeration. (A) The surface boil from an underwater circulating aerator. (B) A fountain has been attached to the aerator to improve aesthetics. Photos courtesy of Neil Gerber, Aquatic Management, Bluffton, Indiana.

from a number of aerated lakes and observed that phytoplankton content decreased in less than half of the lakes examined. Cyanobacterial blooms, however, decreased and green algal populations increased in the majority of cases. This shift was attributed to several factors. For example, aeration may increase the carbon dioxide concentration in the water, thus lowering the pH and favoring green algal development. The turbulence created may disrupt the ability of the cyanobacteria to form surface scums, which normally shade out other, potentially competitive, algae. In those cases in which cyanobacterial blooms were not affected, water mixing and aeration may have been incomplete. There is presently no evidence to suggest that aeration has an impact on filamentous algae or Chara. Frodge et al. (1991) reported that Pithophora mats growing among vascular plant canopies were associated with high concentrations of P in the surface water. This trend was thought to be the result of the conversion of iron-bound P to OH–-bound P at high pH values (>10),

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so that the P was not precipitated even at high dissolved oxygen levels. Thus, high pH values, often associated with high photosynthetic rates of dense vegetation, can potentially offset aeration effects. The presence of a fountain in a body of water does not mean that the water is being aerated. Aerators are specifically designed pieces of equipment that move air into the water column, not spray water into the air. A fountain can be attached to an aerator for aesthetic purposes (Fig. 6B). Although fountains may cause some surface circulation and aeration, they do little to prevent nutrient cycling or fish kills.

7. Hypolimnetic Withdrawal The principle behind hypolimnetic withdrawal is the pumping or siphoning of bottom waters that have a high P content into receiving waters. The technique has not been used frequently. Although evidence suggests that it can be successful in reducing the P content of lake water (Cooke et al., 1993b), few studies provide convincing data that algal blooms are reduced. Replacement of water to maintain depth must come from a source with a low P content. Another problem with this technique is the potential for damage downstream caused by releasing anoxic, nutrient-laden, polluted waters.

8. Summary of Nutrient Manipulation Methods Nutrient manipulation techniques, particularly those that regulate P inputs or internal cycling, can successfully reduce the incidences and severity of algal blooms. In some instances, a single technique is not sufficient. Water supply lakes for the city of St. Paul, Minnestoa, were infested with blooms of Anabaena and Aphanizomenon (Walker et al., 1989). Chemicals (powdered carbon and potassium permanganate) were added at the water treatment plant to reduce taste and odors, and copper sulfate was applied every week during the growing season. These approaches were unsuccessful. It was only when the lakes were subjected to a multimethod approach that included the reduction of external and internal P concentrations by using iron chloride to inactivate P, the construction of detention ponds to reduce P loadings from runoff from urban watersheds, and hypolimnetic aeration that some success was achieved. The St. Paul example illustrates the complexities involved in nutrient manipulation procedures. Probably the greatest impediment to initiation of a nutrient removal plan is the watershed analysis and water quality testing (and financial outlay). This analysis is necessary to determine which approach or combination of approaches is most likely to succeed. It is important for water management agencies and property owner asso-

ciations to accumulate and allocate sufficient resources for a thorough lake and watershed monitoring program before implementing nutrient manipulation techniques. Once the management approach has been chosen, however, the science is now to the point where successes far outnumber failures. One outcome of nutrient control techniques to reduce algal populations can be increased colonization by submersed vascular plants. Submersed vascular plants obtain the majority of their nutrients from the sediments rather than from the water (Carignan and Kalff, 1980; Barko and Smart, 1980, 1981). Therefore, the techniques described above have little impact on submersed vascular plant growth. When shading by phytoplankton or filamentous algal mats is removed, these plants can become established or reestablished. The presence of submersed plants is advantageous in many instances, but sometimes it has serious consequences. For example, the improved clarity of Lake Washington has led to invasion by Eurasian watermilfoil (Myriophyllum spicatum) in shallow areas, and there are other examples of similar shifts in populations (Spencer and King, 1984; van Donk et al., 1990). Madsen (1996) recorded that Secchi disk values doubled (1.5 to 3 m) in Lake St. Clair, Michigan, between 1967 and 1995, as a result of the introduction of zebra mussels. Macrophytic plant range expanded from 60 to 95% of the lake over this period, and Eurasian watermilfoil range expanded from 20 to 44% of the lake. Hence, management plans may need to consider which is the better alternative: a eutrophic, algal-dominated lake vs. relatively clean water with abundant vascular plant growth. The latter problem is somewhat ameliorated when native vascular species and Chara, which tend to have shorter growth habits and provide valuable fish habitat, colonize an area. It is exacerbated, however, when the colonizer is an invasive species such as Eurasian watermilfoil or hydrilla (Hydrilla verticillata). The stems of these species grow up through the water column to form a canopy of vegetation at the surface (Barko et al., 1986; Smith and Barko, 1990). This growth form not only prevents use of the water but can shade out stands of native vegetation. Such shifts among populations further illustrate the complexity of dealing with algal and aquatic plant management issues.

B. Direct Control Methods The goal of direct control methods is to remove the algal biomass as quickly, efficiently, and cost effectively as possible. Although choices have to be made about which technique to use, the extensive watershed and inlake monitoring that should precede nutrient manipula-


tion does not have to be conducted. On the other hand, the efficacy of direct control methods often causes the user to overlook the need to initiate a long-term, nutrient management program. Certainly direct control techniques have their place in an overall management plan, but they should not be viewed as the only approach to solving noxious algal blooms or extensive filamentous algal mat growth. The major methods of direct control of algal biomass are harvesting, biomanipulation, biological controls, allelochemicals, and algicides.

1. Harvesting Harvesting methods can range from hand-pulling or raking to use of large mechanized harvesting equipment (Fig. 7). The vegetation is gathered and preferably moved away from the site so that it cannot wash back into the water. This is obviously not a technique that will remove phytoplankton, but it can be used with some success for the removal of floating filamentous algal mats and charophytes. Hand harvesting or raking of filamentous algal mats and Chara is considered difficult because these growths fragment very easily. The tremendous amounts of biomass (and associated water) make hand labor exhausting and time-consuming. However, probably more hand harvesting occurs than one would expect. Pond and lake property owners, for example, frequently clean off beach and dock areas with rakes and other handheld devices. The beaches of Lake Ontario have at times been hand raked of Cladophora growth washed up on shore by municipal workers. Hand harvesting sometimes is encouraged prior to algicide treatment. This is particularly true in late summer when large amounts of mat material have accumulated. The death

FIGURE 7 A mechanical harvester. Photo courtesy of United Marine International, Div. of Liquid Waste Technology, Inc., Somerset, Wisconsin.

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of an excessive amount of biomass (which leads to decomposition and associated bacterial growth, which uses the oxygen in the water) can lead to oxygen depletion and fish kills. Removing at least some of the biomass prior to treatment can help prevent severe oxygen depletion situations. Most mechanical harvesting activities are directed at the removal of rooted submersed vascular plants. There are, however, reports of mechanical harvesting used successully for Chara control (Conyers and Cooke, 1982; Cooke et al., 1993b), particularly in shallow water where the harvesting blades can cut at the sediment–water interface. Chara is almost invariably collected along with submersed vascular vegetation when the two grow intermixed. Some evidence suggests that populations of phytoplankton, filamentous mat-forming algae, and Chara can increase after intensive mechanical harvesting of submersed vascular plants (Neel et al., 1973; Nichols 1973; Cooke and Kennedy, 1989; Anonymous, 1990). Although the increases have not been clearly associated with harvesting, the opening up of areas to light and the potential increase in nutrients after harvest may make algal growth more likely to occur. Another method of harvesting has been used in irrigation systems in the western United States. Racks are inserted at intervals along the canal to collect the algal mats that slough off the sides of the canal and float downstream. The racks are removed periodically, cleaned of algae and other debris, and returned to the canal (Fig. 8). A major consideration in harvesting is to ensure that the collected vegetation does not wash back into

FIGURE 8 A rack to collect floating material taken from an irrigation canal in California. The majority of vegetation is Cladophora. Photo courtesy of Lars Anderson, USDA-Agricultural Research Service.

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the body of water. Even though it may appear that the algal mats have dried out once they are exposed to the air and sun, the underlying portions of the mats may still be viable. Akinetes found in Pithophora mats that were exposed to the drying effects of the sun still showed 80% viability 136 days after initial stranding (Lembi et al., 1980). Little use has been made of harvested aquatic vegetation. It generally has little value as food for livestock or humans, and the energy costs to dry and pellet the vegetation can be prohibitive (National Academy of Sciences, 1976; Joyce, 1993). Some research has been conducted on the potential use of filamentous algae such as Pithophora and Cladophora to make paper, and the protein content of Hydrodictyon, Spirogyra, and Pithophora was reported to be 18–26%, which is comparable to the protein content in some cyanobacteria and vegetables (Khan et al., 1996); however, the major (but sporadic) use of harvested algal mats at present is as mulch and fertilizer for gardens.

FIGURE 9 A flowchart illustrating the connections between foodchain levels as determined by the biomass of piscivores. Biomanipulation attempts to decrease the mass of planktivores in order to increase the numbers of large-bodied zooplankton, which, in turn, graze on phytoplankton.

2. Biomanipulation The observation that the relationship between algal growth and P concentrations is not perfect [in fact, in a study of 66 lakes by Schindler (1978), the regression statistic explained only 48% of the variance in algal productivity] led in part to the theories behind biomanipulation. Much research in the past (reviewed by Cooke et al., 1993b) has indicated that the type of zooplankton present in a body of water can have an effect on phytoplankton populations. The type of zooplankton is affected in turn by the types of fish that are present. The potential for zooplankton (and fish) to have an effect on phytoplankton populations, irrespective of P content in the water, may explain why, in some instances, phytoplankton populations are lower or higher than those predicted by the P content of the lake. The term “biomanipulation” was coined by Shapiro et al. (1975). It is also referred to as top-down feeding and involves manipulating the components of the trophic cascade (Paine, 1980; Carpenter et al., 1985). The premise of biomanipulation, as elucidated by Shapiro (1980) and Carpenter et al. (1985, 1987), is that top predators, such as piscivorous fish, can influence the abundance of planktivorous fish, which in turn can determine the abundance, size structure, and productivity of zooplankton and phytoplankton (Fig. 9). For example, planktivorous fish tend to feed on largebodied zooplankton, which results in domination by small-bodied zooplankton. Because it is the largebodied zooplankton (some species of Daphnia) that feed most effectively on algae, their reduction is typically accompanied by a relatively high phytoplankton

biomass. The key to success in biomanipulation is the addition of piscivorous fish, which theoretically should reduce the numbers of planktivorous fish, which in turn enhances the development of populations of largebodied Daphnia species and a decline in phytoplankton populations. The actual manipulation involves the introduction, where necessary, of piscivorous fish and/ or the removal of planktivorous fish. An example would be the removal of planktivorous fish from a site by rotenone treatment and the stocking of piscivorous fish to eliminate any planktivorous fish that might be introduced later. Observations of fish–zooplankton– phytoplankton relationships and the successful manipulation of all or part of the trophic cascade in enclosures, ponds, and lakes (Spencer and King, 1984; Carpenter et al., 1985, 1987; Elser and MacKay, 1989; Gulati, 1990; Mazumder et al., 1990; Quirós, 1995) have provided evidence that the principle is essentially valid. However, the practice of biomanipulation has been marked by inconsistencies that suggest that aquatic systems are much more difficult to manipulate than originally anticipated. Analyses of data in which biomanipulation did not provide the expected results include those cited by McQueen et al. (1989), McQueen (1990), Vanni and Findlay (1990), Badgery et al. (1994), and Noonan (1998). McQueen et al. (1989) suggested that the various trophic levels depend on nutrients and energy flow, which is essentially a bottom-up process rather than top-down. In less productive oligotrophic lakes, a top-down cascade may


extend all the way to phytoplankton, but in nutrient enriched eutrophic systems the results are less clear because bottom-up forces are large relative to topdown forces. In other words, more phytoplankton populations can be supported by high P concentrations than can be effectively grazed by zooplankton. Benndorf (1989) concluded that the long-term success of top-down manipulation requires a reduction of external P loading. Another complication is that even if large-bodied zooplankton populations increase, the gelatinous (Porter, 1973) or large-celled ungrazable or undigestible algae can still proliferate. The conditions under which biomanipulation will work are still unclear. DeMelo et al. (1992) indicated that a careful analysis of the data does not support the eutrophic/oligotrophic differences proposed by McQueen (1990) and others. Differences in the N and P requirements for the growth and reproduction of Daphnia further complicate the situation. For example, Daphnia growth and reproduction are strongly suppressed when they are fed P-limited algae having a high C:P ratio (Sommer, 1992; Urabe et al., 1997; MacKay and Elser, 1998). When zooplankton that have high P needs ingest phytoplankton with high N:P ratios, there is a disproportionate release of unused N to the system, which in turn can affect algal community structure because of changing nutrient ratios in the water (Urabe, 1993; Steinman, 1996). Biomanipulation can have effects on components of the aquatic ecosystem that are not directly involved in the trophic cascade. For example, Spencer and King (1984) reported that zooplankton successfully reduced phytoplankton densities in ponds with no fish or with dense populations of largemouth bass (a piscivore), but the resulting clear water stimulated dense growths of Cladophora spp. and the submersed vascular plants Elodea canadensis and Potamogeton spp. This kind of a shift is frequently the outcome of any control or nutrient manipulation technique that reduces phytoplankton growth. Biomanipulation as a tool is still experimental and should not be recommended without considerable analysis of the composition of the various trophic levels and regulating environmental factors.

3. Biological Controls The use of one organism to control unwanted organisms has been widely studied in aquatic plant management. Most of the research has focused on the control of aquatic vascular plants, but there are studies that suggest potentially useful agents for algae control. Reports that cyanophages (viruses) lyse cyanobacterial cells date to the 1960s (Safferman and Morris, 1963; Safferman et al., 1969). The first described cyanophage was named LPP-1 for its ability to lyse

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cells of Lyngbya, Phormidium, and Plectonema (Safferman and Morris, 1963). Since then, other cyanophages have been isolated and evaluated as possible biocontrol agents (Padan et al., 1971; Stewart and Daft, 1977; Martin and Benson, 1988; Phlips et al., 1990; Monegue and Phlips, 1991). Although most studies have been conducted in the laboratory, a few have shown successful results in experimental field enclosures and ponds (Martin et al., 1978; Desjardins and Olsen, 1983). It is generally agreed that cyanophage application is most effective when it is applied before the host populations are well established (Desjardins and Olsen, 1983; Monegue and Phlips, 1991), but its impact on established cyanobacterial populations is poorly known. Eukaryotic algae, including the green alga Chlorella, are also susceptible to viruses (Van Etten et al., 1991), but their biocontrol potential is unknown. Although cyanobacterial and eukaryotic algal populations clearly are affected by phages in nature and although lysis can be induced in the laboratory and in small-scale tests, no virus has been developed for control purposes. Extensive field testing under a variety of environmental conditions has not been conducted, and the information needed on amount of inoculum, application technique, and the environmental factors conducive for replication and lysis is not available. The selectivity of viruses to one or a few species further limits the broad application of the technique. Other microorganisms that have shown activity on planktonic algae include bacteria (Shilo, 1967; Burnham and Fraleigh, 1983; Walker and Higginbotham, 2000) and fungi (Redhead and Wright, 1978; Canter and Jaworski, 1979; Kudoh and Takahashi, 1990). Clearly, all of these organisms play a role in natural successional patterns in lakes, but their potential as biological controls remains unexplored. Even so, this approach shows promise for the future. A wide variety of insect and other invertebrate grazers, including snails, caddisfly larvae, mayfly larvae, chironomid larvae, and shrimp have reduced benthic algal growths (Fulton, 1988; Steinman, 1996), but their effect on prolific growths of filamentous algae (outside of river and stream systems) is apparently minimal, and none has been investigated as a potential biological control agent. The crayfish Oronectes immunis significantly reduced stands of Chara (and submersed vascular plants) in a New York lake (Letson and Makarewicz, 1994), but the treatment was more expensive and a higher stocking density was required compared with treatments with grass carp (discussed below). When the vegetation was removed, the crayfish themselves became subject to predation. Therefore, it was difficult to maintain sufficient crayfish densities to consume vegetation regrowth without additional stocking.

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A number of fish species have been investigated for their potential to control algae and aquatic vascular plants. The silver carp (Hypophthalmichthys molitrix) and bighead carp (Aristichthys nobilis) consume phytoplankton and zooplankton (Dimitrov, 1984; van der Zweerde, 1993). Many tilapia (Tilapia spp.) are herbivorous (Hauser et al., 1976; Smith, 1985). Some are filter-feeders that consume phytoplankton; others feed on macrophytes including filamentous algae and Chara. The distribution of tilapia is restricted by temperature (they are native to India, Africa, and South America); they do not survive in waters colder than 10°C. Therefore, in the United States their use has been confined to the South and to sites that receive heated discharge (Crutchfield et al., 1992). In addition, their use has many disadvantages, such as the ability to switch to animal food when they have eliminated plant and algal growth, a high reproductive potential, and interference with native fish species. The most successful and widely used biological control agent for aquatic vascular plants and some algae has been the grass carp (Ctenopharyngodon idella; Fig. 10) (see reviews by Cooke et al., 1993b; van der Zweerde, 1993). This fish is native to northern China and was introduced into the United States in the 1960s. It was originally introduced into Arkansas but is now used in at least 35 states for aquatic weed control (Sanders et al., 1991). Because there are no predators in its native range, grass carp do not show typical avoidance behaviors and are extremely susceptible to predation. Therefore, fish must be at least 20 to 25 cm long (~450 g) when stocked to avoid predation by largemouth bass and other native predators. Under ideal conditions grass carp can grow to a weight of at least 23 kg within 5–10 years. Recommended stocking

FIGURE 10 The grass carp (Ctenopharyngodon idella), a widely used biological control agent for certain macrophytic algae and submersed vascular plants.

densities vary from region to region. Recommendations from the Indiana Department of Natural Resources, for example, suggest 37 grass carp vegetated ha–1 if maintenance of some vegetation is desired and 74 fish ha–1 if elimination of vegetation is desired. The grass carp survives in cold water and begins to feed regularly at about 14°C. Feeding peaks at about 20–26°C and decreases when the water temperature reaches about 33°C. The concern about the potential for grass carp to reproduce and crowd out native species led to the development of sterile, triploid grass carp. Even with this precaution, grass carp must not be introduced in areas in which the elimination of vegetation in wetland areas would destroy valuable habitat for waterfowl and other animal life. Young grass carp up to 50 mm (about 2 in) in length feed mostly on zooplankton. After that they shift to a diet of filamentous algae, duckweed, and submersed vascular plants. There has never been any evidence of the fish shifting to an animal diet once they exceed a length of about 100 mm. The fish clearly has preferences for the kinds of plants it eats. Numerous lists of preferred plant species have been published (Fowler and Robson, 1978; Cassani and Caton, 1983; Shireman et al., 1983a; Pine and Anderson, 1991; Sanders et al., 1991; Cooke et al., 1993b), and in almost every case, Chara and Nitella are listed as preferred or highly preferred plants. Bauer and Willis (1990) reported that grass carp introduced at 49 fish ha–1 almost totally eliminated Chara (the dominant macrophyte) in 2 years in two small South Dakota lakes. The effectiveness of grass carp for the control of filamentous algal mats is less clear. Some of the references listed above claim good control of relatively coarse species such as Cladophora and Pithophora; others indicate no or only weak control of filamentous algae. In some cases, only high stocking densities of more than 123 fish ha–1 have succeeded in controlling filamentous algae. Spirogyra seems to be least preferred, probably because its slimy nature prevents effective ingestion; however, it too can be eaten if no other vegetation is available (Lembi et al., 1978). In mixed populations of vegetation, grass carp will clearly consume soft-bodied vascular plants such as pondweeds, elodea, and naiads, and Chara (even though Chara can be coated with a hard coat of calcium carbonate, fish appear to be attracted to it) in preference to filamentous algae. When filamentous algae are the only plant material present, grass carp will feed on it, probably to avoid starvation. Problems with grass carp include lack of consistency and predictibility. Use of grass carp does not work under all circumstances, and visible control may not be


achieved until several years after introduction, particularly in heavily infested areas. Increased turbidity due to increased phytoplankton populations has been noted in some situations (Shireman et al., 1985; Maceina et al., 1992), and overstocking can result in the total elimination of all vegetation, a situation that is not considered beneficial for fish and other animal life in natural bodies of water. In fact, considerable controversy in the sportfishing industry has erupted over the potential for elimination of vegetation cover by grass carp, particularly because the effect of removing weed beds on angling success is still being debated (Bain, 1993; Bettoli et al., 1993; Killgore et al., 1998). Other control methods (e.g., mechanical harvesting or the use of chemicals) can be used for weed beds in certain areas of a lake so that some vegetation can be selectively retained. Unfortunately, there is little ability, once grass carp have been introduced, to dictate where they will graze and how much vegetation they will consume over time. Stocking rates can be reduced to avoid elimination of plant populations, but because the grass carp work so slowly and because it is so difficult to predict their impact on the vegetation, herbicides/algicides or other methods may have to be used if the fish cannot provide adequate control. Finally, grass carp may consume desirable native species (including Chara and Nitella) and leave less desirable species, such as the invasive weed Eurasian watermilfoil (Fowler and Robson, 1978). Another method of biological control is the use of waterfowl, specifically geese or swans. Filamentous algae are consumed by waterfowl, and charophytes are a favored food of herbivorous ducks, coots, and swans (Martin et al., 1961; Hargeby et al., 1994). A pair of swans reportedly will keep a 0.4 ha pond free of submersed vegetation, as will 7–20 geese or ducks ha–1 (Holm and Yeo, 1981). Unfortunately, there are many problems associated with the presence of waterfowl. Birds that are introduced for aquatic weed control are usually rendered flightless. Therefore, their diet of aquatic vegetation must be supplemented to provide adequate nutrition, they must be protected from predators, and lake managers must be willing to tolerate their aggressiveness during the breeding season. As with free-living waterfowl, their waste materials can litter the banks and stimulate phytoplankton blooms. An unusual form of biological control is the use of competitive plants. Doyle and Smart (1998) showed that established plantings of the flowering plants pickerelweed (Pontederia cordata) and American pondweed (Potamogeton nodosus) reduced the biomass of L. wollei by 50% and prevented the formation of floating mats. The effect was attributed to shading and possibly to competition for nutrients in the sedi-

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ments. The main drawback of this method is that it only works in very shallow areas where the flowering plants can root. In a larger sense, the water user must be willing to accept the shift from an algal infestation to dense stands of flowering plants. This fact alone negates the potential benefits of shading by free-floating flowering plant species such as water hyacinth (Eichhornia crassipes) or duckweed (Lemna spp.), both of which tend to be weedy. On the other hand, the premise of using competitive submersed plants, possibly those that have been selected or genetically modified to produce shortened stems with minimal canopy formation, should be explored further. The use of biological agents shows great promise for the control of weedy algae and plants, but it has drawbacks. More research is necessary, particularly on those organisms that might be relatively selective, such as phages. The introduction of nonselective agents, such as grass carp, has been used successfully in some situations, but it also has the potential to cause adverse ecological impacts in others.

4. Allelochemicals Allelopathic chemicals are chemicals produced by plants that have either an adverse or beneficial effect (usually adverse) on other plants (Rice, 1984). Although this area has not received much attention in controlling nuisance algae, there is evidence that allelochemicals may be useful. A number of cyanobacteria have been investigated for their potential to produce allelochemicals that inhibit the growth of other cyanobacteria or algae (Mason et al., 1982; Flores and Wolk, 1986). Allelochemicals from some fungi (Redhead and Wright, 1978) and terrestrial vascular plants (mostly phenylpropanoids) (Della Greca et al., 1992) have been shown to have algicidal activity. Studies have been conducted on allelochemicals produced by aquatic vascular plants, but most of the bioassays have been conducted using vascular plant species, such as duckweed or lettuce seedlings, rather than algae (Elakovich and Wooten, 1989; Sutton and Porter, 1989; Wooten and Elakovich, 1991). One exception is the study by Gross et al. (1996) in which extracts from Eurasian watermilfoil inhibited cyanobacteria. Although some potential algicidal allelochemicals have been identified, the major constraints to further development is the expense of culturing the organisms and extracting sufficient amounts of the allelochemicals for application. An alternative is to synthesize the active chemical, but this is also costly, particularly in view of the relative cheapness and availability of copper sulfate. Thus, the financial incentive for industry to develop these compounds for the aquatic market is lacking.

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A cheaper method of allelopathic control may be the “bale of hay” technique. For years, farmers have applied straw or hay to ponds to reduce algal growth. This method has been tested and substantiated in a series of experimental studies in England (Welch et al., 1990; Gibson et al., 1990; Pillinger et al., 1992, 1994; Newman and Barrett, 1993). In the laboratory, rotting barley (Hordeum) straw inhibited the growth of several planktonic (including Microcystis) and filamentous algae. Additions to a canal over a 3-year period resulted in reduced biomass of C. glomerata after 2 years. The apparent mechanism is through the release of quinone compounds (Pillinger et al., 1994). Attempts to replicate these effects in North American waters have had mixed results. Nicholls (1996) showed decreases in chlorophyll a in ponds treated with barley in Ontario, but tests using barley, wheat (Triticum), and rye (Secale) in ponds with Pithophora in North Carolina were unsuccessful (Kay, 1997). Discrepancies among results may be due to differences in straw concentrations, target species, environmental conditions, and length of exposure. Because barley is not grown in many parts of the United States and thus is not widely available, other forages should be tested. Laboratory and small-scale field testing suggests that alfalfa (Medicago) hay may be effective for filamentous algae control (Marencik and Lembi, 1998), but additional research is needed to determine if its rapid breakdown in water could cause oxygen depletion problems. Further research in this area is warranted.

5. Algicides A common method of controlling algal infestations is the use of chemicals (algicides). Of the algicides, the most commonly used compounds are copper-containing products. Other products, such as diquat and the mono (N,N-dimethylalkylamine) salt of endothall, are registered for algae control, but their use is relatively minor compared to that of the copper-containing products. Of the copper products, copper sulfate (CuSO4) is the most widely used algicide for controlling algal populations in water supplies, recreational lakes, and reservoirs (Elder and Horne, 1978; Effler et al., 1980; Raman, 1985). It has been used since at least 1905 and probably was used earlier (Moore and Kellerman, 1905; Murphy and Barrett, 1993). In the late 1960s and early 1970s, more than 9 million kg of CuSO4 were applied annually in waters in the United States (Fitzgerald, 1971). Approximately 68% of all water area in the United States treated with a chemical product in 1992 was treated with an algicide (unpublished industry data). CuSO4 was applied to 70%

of this area, and the remainder was treated mostly with other copper compounds (chelated copper compounds). Even though algicides were used on 68% of the area treated, they accounted for just 20% of the total sales ($32.5 million) of all aquatic chemicals (herbicides and algicides) due to their relatively low cost. Copper is effective on a wide range of algae (Maloney and Palmer, 1956). The toxic agent is free cupric ion (Cu2+), and toxic cupric ion activities range from greater than 10–6 to 10–11 M for species of diatoms, dinoflagellates, microscopic green algae, and cyanobacteria (McKnight et al., 1983). The fact that cyanobacteria are more sensitive to copper than some of the eukaryotic algae (Whitton, 1973; Swain et al., 1986) accounts for its widespread success and acceptance. Diatoms are probably next in sensitivity followed by the green algae (Swain et al., 1986; Havens, 1994). A copper concentration of 25–40 µg L–1 effectively controlled A. flos-aquae in shallow eutrophic lakes in Manitoba (Whitaker et al., 1978), a dose that is considerably lower than the typical doses of 125–250 µg L–1. Suppression of nitrogen fixation by Anabaena and Aphanizomenon was observed after copper additions of only 5–10 µg L–1 (Horne and Goldman, 1974), leading to the suggestion that maintaining low doses to inhibit N2 fixation would be an alternative to a single large dose. This approach has not been practical because the short residence time of copper in the water column mandates almost continual copper application. Certain microscopic green [e.g., Oocystis (Meador et al., 1993; personal observations)] and euglenoid planktonic (Hawkins and Griffiths, 1987) algae are relatively tolerant to copper treatments. For example, the microscopic green algae Ankistrodesmus, Scenedesmus, and Pandorina may require copper concentrations as high as 500 µg L–1 for control (Copper Sulfate Fine Crystals product label). Among mat-forming green algae, Spirogyra and Oedogonium are very susceptible to copper (Whitton, 1970; Francke and Hillebrand, 1980), whereas Pithophora and Hydrodictyon are considerably more tolerant (Table I). In addition to inherent tolerance or susceptibility, mat structure also may dictate relative tolerance to copper (or other exogenously applied materials). Mat structure appears to be governed in part by branching pattern. Pithophora filaments, which are branched, produce intertwined, tighter mats than filaments of Spirogyra and Oedogonium, which are unbranched (Table I). It may be more difficult for copper to penetrate the extremely dense, massive mats that are produced by Pithophora (Lembi et al., 1984) than the loose, less tangled mats formed by Spirogyra and Oedogonium. Of all mat-forming species, the cyanobacteria are the most tolerant to copper, which


TABLE I Taxa, Filament Morphologies, Mat Structures, and Susceptibilities to Coppera of Filamentous Mat-Forming Algae EC50b

Taxon

Morphology

Mat structure

Chlorophyta Spirogyra

Unbranched

Loose

1

Unbranched

Loose

3

Oedogonium

Hydrodictyon Net-like Pithophora Branched Cyanobacteria Oscillatoria

Unbranched

Lyngbya

Unbranched

a b

Moderate Dense Dense, slime + associated sediment Dense

48 46 290 1630

Lembi, 2000; data on Lyngbya from Hallingse and Phlips (1996). EC50 = concentration of Cu2+ in µg L–1 required to reduce biomass (dry weight) of alga by 50% under laboratory (not field) conditions.

seems unusual given the susceptibility of planktonic cyanobacteria. Both an inherent tolerance (at least 6fold greater than that of Pithophora; Table I) plus the presence of thick slime and a coating of sediment make Oscillatoria mats extremely difficult to control. Likewise, Lyngbya (which also produces sheaths) produces thick, dense mats which probably add to its tolerance. Unfortunately, it is likely (although not well documented) that the elimination of susceptible species (both microscopic and mat-forming) has led to their replacement by tolerant species. The mechanisms by which copper affects algae appear to vary. The list of reported copper effects [taken from Gledhill et al. (1997); see references therein] indicates that it inhibits photosynthesis (see also Kallqvist and Meadows, 1978), disrupts electron transport in photosystem II (see also Cedeno-Maldondo and Swader, 1974), reduces pigment concentrations, affects the permeability of the plasma membrane and induces losses in cations, inhibits nitrate uptake, restricts growth, affects cell motility, and affects the distribution of proteins, lipids, sterols, sterol esters, and free fatty acids in the cell. Although this list was developed primarily for marine algae, there is no reason to think that the same effects should not be expected in freshwater algae. In addition, copper has been reported to inhibit P uptake (Peterson et al., 1984) and to precipitate proteins in the cell (Murphy and Barrett, 1993). All of these presumed modes of action suggest that copper is a general algal cell toxicant. When applied at the recommended dosage (250 µg L–1 copper), copper acts rapidly, usually within a period of hours.

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A number of mechanisms have been proposed to account for the differential tolerance among algae and include (Lage et al., 1996) intracellular accumulation of copper in polyphosphate bodies, storage of copper in membrane-bound vesicles, excretion into the medium of organic compounds that bind copper, intracellular chelation of copper by organic compounds like phytochelatins, and efflux of the copper. In addition, copper accumulation in the cell walls of some algae and higher plants has been reported (Pearlmutter and Lembi, 1986; Allan and Jarrell, 1989). As noted above, mat structure also should be considered a factor in species tolerance. Water chemistry, particularly pH and alkalinity (a measure of bicarbonates, carbonates, and hydroxides), plays an important role in copper toxicity (McKnight et al., 1983). Below neutral pH, Cu2+ is the major copper species; above neutral pH the major forms of copper are the copper carbonate complexes and malachite and tenorite. The various complexes and precipitants formed above neutral pH values effectively prevent Cu2+ from being taken up by target organisms. At high pH and alkalinity, the concentration of soluble Cu2+ in the water is extremely low and possibly ineffective for algal control (Button et al., 1977). In low alkalinity, acidic waters, the recommended dose (250 µg L–1 Cu2+) of CuSO4 will kill algae, but given the relatively high amounts of soluble Cu2+, particularly sensitive fish species, such as trout, can also be killed. This phenomenon is even true for the chelated copper products (discussed below), and all currently registered copper products have a statement similar to the one on the Cutrine-Plus label: “Do not use in water containing trout if the carbonate hardness of the water does not exceed 50 mg L–1.” As total alkalinity increases, so must the CuSO4 dosage to overcome the precipitation problem. Toxicity to fish is essentially nonexistent at high alkalinities because of the low concentrations of Cu2+ in the water [as little as 0.5% of the total dissolved copper has been calculated to be present as free cupric ion (Wagemann and Barica, 1979)]. For example, at the low alkalinity of 18.7 mg L–1 (as CaCO3 + HCO3), the LC50 (concentration that will kill 50% of the population) for bluegill (Lepomis) is 884 µg L–1 copper (3.5 mg L–1 CuSO4) (Herbicide Handbook, 1994). At the moderate to high alkalinity of 166 mg L–1, the LC50 is 7300 µg L–1 copper (29.2 mg L–1 CuSO4). The upper legal limit for copper use in water is 1000 µg L–1 copper (4 mg L–1 CuSO4); this concentration (which is seldom recommended) could kill fish in low alkalinity waters. However, there is almost a 30-fold safety factor between the LC50 and the typical recommended dosage [250 µg L–1 copper (1 mg L–1 CuSO4)] in high alkalinity waters.

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Although direct copper toxicity is seldom a problem to fish, the depletion of oxygen during algal death and decomposition can cause fish kills. Treatments must never be made to bodies of water that have heavy algal infestations. Chelation with organic compounds stabilizes soluble copper and theoretically retards its precipitation and adsorption (McKnight et al., 1983). This principle is the basis for the formulation of the chelated forms of copper as substitutes for copper sulfate. Most commercial chelated formulations are variations of ethanolamine complexes. One of the presumed advantages of using a copper chelate is that a longer persistence of active copper in water should increase algal contact and control. In a comparative study of copper chelates and copper sulfate used at the same copper dose, Masuda and Boyd (1993) found that chelates slowed the loss of total copper (from an initial level of 500 to 100 µg L–1) from the water from 4.3 to 6.3 days, but they concluded that this advantage did not compensate for the greater cost of the chelated formulation. On the other hand, the chelated copper products provide flexibility in application because they are formulated either as granules or as liquids, whereas CuSO4 is packaged as a solid material only. The combination of liquid formulations with other liquid aquatic herbicides is useful for commercial applicators to control a broad spectrum of species that include both algae and vascular plants. Phytoplanktonic blooms are usually treated by pumping the copper compounds (dissolved in water in a tank mounted in the boat or airboat) through a boom and trailing hoses into the water (Fig. 11A). The hoses can be adjusted to deliver the compound to different depths in the water column. For example, hoses that disperse the copper in the upper meter of water (or surface treatments with a spray) are effective for the control of cyanobacteria that have formed surface scums. The hoses can be set lower in the water column to deliver copper to filamentous algal mats that are still lying on the bottom and have not floated to the surface. Once the mats have floated to the surface, spot treatments directly on the mats with a spray can ensure that contact with the cells is maximal (Fig. 11B). This ability to place the treatment directly on or near the target organism allows the applicator to selectively treat some areas and not others, thus reducing both the amount of copper needed and the volume of water that comes into contact with the chemical. Other methods of application include slow dispersal through a burlap sack or drip or single high dose applications in flowing water systems such as irrigation and drainage canals,. The persistence time of copper in water is relatively short. Button et al. (1977) found that 95% of the

FIGURE 11 Algicide applications. (A) A unit set up to deliver chemical through trailing hoses. (B) A unit set up to deliver a spray directly to the algal mats. B courtesy of Neil Gerber, Aquatic Management, Bluffton, Indiana.

copper sulfate distributed over a lake surface dissolved in the upper 1.8 m of the water column, but the total copper concentration was at pretreatment levels within 24 h. Tucker and Boyd (1978) made 10 applications of 0.84 kg ha–1 CuSO4 at 2-week intervals to ponds without causing an appreciable increase in total copper concentration. Wagemann and Barica (1979) reported a half-life of total dissolved copper from 1 to 7 days. Anderson and Dechoretz (1984) reported a half-life of about 1 day with all of the copper gone from the water column at 14–28 days. In other bodies of water, copper has persisted for up to 30 days after treatment (Elder and Horne, 1978; Whitaker et al., 1978; McKnight, 1981; Hawkins and Griffiths, 1987). The implications of a short residence time in water are several. First of all, copper effects, whether on target or nontarget organisms, are temporary. In most cases, algal populations rebound, although not necessarily at the same densities or with the same species. Although this is a problem from the standpoint that


repeated treatments may be needed during a single season to provide adequate control, the short persistence reduces the exposure time of nontarget organisms, including humans. One of the major concerns with the use of copper is its ultimate fate. The copper complexes, as well as decaying copper-containing algae, fall to the bottom where the copper is readily adsorbed onto sediments. Copper, as a heavy metal, persists in the sediment for prolonged periods of time (Frank, 1972; Brown, 1978). The key issue is that copper will accumulate in sediments and be toxic to benthic organisms and then serve as a source of copper to the water after treatment is discontinued. The evidence is somewhat conflicting. Sanchez and Lee (1978) noted that copper-enriched sediments in Lake Monona, Wisconsin (treated over 50 years to 1950), were not interacting with the more recent sediments deposited or with overlying waters. The copper content of the water was no different from that of local hardwater lakes which had not been treated, and they concluded that there were no longterm adverse effects resulting from the copper treatments. Ankley et al. (1993) studied sediment and pore water from Steilacoom Lake, Washington, which had been “grossly contaminated” by copper because of copper sulfate treatments. Extracted copper concentrations in sediments ranged from 0.6 to 3.0 µmol g–1 dry weight, but pore water and overlying water concentrations were less than the analytical detection limit of 7 µg L–1. Toxicity tests showed no effects of the water on the amphipod Hyalella azteca. Probably the most negative report of copper treatment effects on sediments is from a string of interconnected lakes in southern Minnesota that were treated over a period of 58 years (Hanson and Stefan, 1984). Effects included elevated concentrations of copper in the sediments, fish kills due to oxygen depletion or possibly copper toxicity, increased internal P cycling, rapid recovery of algal populations within 7–21 days, shifts of game fish to rough fish, disappearance of macrophytes, and reductions of benthic macroinvertebrates. Some conditions improved when the sediments had been dredged and the use of copper discontinued. It is difficult to evaluate the results of this study because factors other than copper applications may have caused some of these effects. Fortunately, such extreme effects have not been observed in most coppertreated lakes. If they did occur in large numbers, copper should have been banned long ago. In fact, Sanchez and Lee (1978) concluded that 50 years of copper treatments in Lake Monona had not resulted in the loss of the excellent sport fisheries supported by that lake. Nevertheless, the Minnesota study should serve as a warning that a program that includes

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long-term, whole-lake treatments with copper must be scrutinized and continually monitored for potential deleterious effects. An additional concern is the sensitivity of zooplankton to copper. The LC50 for planktonic crustaceans is 60–90 µg L–1 copper; for rotifers it is 1100–1700 µg L–1 (Demayo et al., 1982). Other studies on invertebrates indicate LC50 values ranging from 10 to 130 µg L–1 copper (McIntosh and Kevern, 1974; Winner, 1985; Meador et al., 1993). Concentrations as low as 8 µg L–1 copper have caused significant effects on cladocerans in life cycle toxicity tests (Belanger et al., 1989), and Hedtke (1984) found large reductions in zooplankton biomass (along with that of snails, total macroinvertebrates, and midges) when laboratory microcosms were treated with 30–270 µg L–1 copper. Most of these concentrations are well within the range of normal use dosages. Studies using lake mesocosms also showed reductions in zooplankton populations (Moore and Winner, 1989; Havens, 1994) as well as species shifts (Moore and Winner, 1989; Winner et al., 1990; Havens, 1994). Both phytoplankton and zooplankton communities were more sensitive to copper in the spring than in the summer or fall (Winner et al., 1990), an observation that was attributed to differences in levels of copper-complexing compounds in the water during the various seasons. Effects of copper on zooplankton communities in lakes have been somewhat mixed. Effler et al. (1980) found no effects of a low-level CuSO4 treatment on several zooplankton populations in a lake; however, McKnight (1981) found decreases of populations of Bosmina, Tetramastix, and Keratella following treatment of Mill Pond, Massachusetts. Long-term effects of copper treatments on zooplankton in lakes have not been adequately studied, although the short-term persistence of copper within the water column should allow most zooplankton populations to recover. The loss of zooplankton populations, even if temporary, may explain why algal populations can recover, sometimes to levels higher than original levels. The loss of grazing impact on phytoplankton due to copper effects on zooplankton has been suggested by the work of McKnight (1981), Taub et al. (1989), and Havens (1994). In addition, bacterial biomass has been reported to rapidly recover or even increase following copper treatment (Effler et al., 1980; Havens, 1994; Dionigi and Champagne, 1995), and in some cases this may be due to loss of grazing pressure from zooplankton (Havens, 1994). Copper is a trace element that is required for the survival of many plant and animal species, including humans. The low mammalian toxicity of copper when diluted in water, its short persistence time in water,

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and its lack of bioaccumulation in fatty tissues are the reasons that it is the only one of the algicides/herbicides registered in the United States for which the use of water following treatment at normal doses is not restricted. This includes use of the water for drinking, swimming, fishing, livestock watering, and irrigation, although a 24-hour waiting period after treatment is desirable just to be cautious. Copper-containing compounds have not been reported to induce cancers in humans or experimental animals (Sunderman, 1978). The only warning on EPA-approved products is the potential toxicity to trout under softwater conditions.

6. Summary of Direct Control Methods Methods for the direct control of algae are available, but none of them ensure that algae problems will be solved other than in the short term. Even grass carp, which live for up to 16 years, become less efficient feeders after about 5 years and restocking is required for long-term control. The adverse effect of copper on food webs must be considered, and appropriate long-term ecosystem studies must be undertaken. These concerns are offset in part by the overall safety of copper to humans, by the short persistence of copper in the water column, and by our ability to place copper directly on or in the vicinity of the target algae. However, lakes and reservoirs will be better protected if repeated whole-lake copper treatments can be avoided. A program of watershed and water quality management through nutrient manipulation is the best alternative, particularly for phytoplanktonic blooms. There will still be a role for the use of copper sulfate, at least in the United States. Copper treatments may be needed periodically or at certain sites where filamentous mat-forming algae or Chara are problems in lakes and reservoirs despite nutrient reduction efforts. Residents on small lakes and ponds that receive large inputs of nutrients from nonpoint sources will continue to require direct control techniques, particularly because they may have few financial or political resources to minimize these inputs. The ability to maintain irrigation systems in the western states free of algae and provide maximal water delivery rates to urban and rural users will still require copper applications for the foreseeable future. Regulatory agencies in several regions of the United States have indicated an interest in eliminating the use of copper for algae control. Copper is not used to any great extent in Canada because of that country’s very cautious approach to water quality (H. Vandermeulen, Fisheries & Oceans Canada, personal communication). However, the immediate benefits and favorable economics of copper sulfate usage suggest

that this compound will continue to be used in the United States. Unfortunately, the ready availability and efficacy of the copper compounds reduce incentives needed for development of alternative control methods, such as organically based algicides and allelochemicals or viral and bacterial biological control organisms. The short-term nature of direct control techniques plus the difficulties inherent in regulating nutrient inputs in every situation dictate that algae will pose water quality problems for many years to come. These problems will continue to pose a challenge to all of us who have an interest in maintaining or restoring healthy aquatic ecosystems.

ACKNOWLEDGMENTS I thank Erik C. Brockman for his tremendous assistance in the literature search for this chapter. I also acknowledge the contributions and insights of numerous associates, both academic and those who deal with algal management on a day-to-day basis. The constructive comments of reviewers Alan D. Steinman and Herb Vandermeulen are greatly appreciated.

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van den Berg, M. S., Scheffer, M., Coops, H., Simons, J. 1998b. The role of characean algae in the management of eutrophic shallow lakes. Journal of Phycology 34:750–756. van der Ploeg, M., Dennis, M. E., de Regt, M. Q. 1995. Biology of Oscillatoria cf. chalybea, a 2-methylisoborneol producing bluegreen alga of Mississippi catfish ponds. Water Science and Technology 31:173–180. van der Zweerde, W. 1993. Biological control of aquatic weeds by means of phytophagous fish, in: Pieterse, A. H., Murphy, K. J., Eds., Aquatic weeds. The ecology and management of nuisance aquatic vegetation. Oxford Science, pp. 201–221. van Donk, E., Grimm, M. P., Gulati, R. D., Kline Breteler, J. P. G. 1990. Whole-lake food-web manipulation as a means to study community interactions in a small ecosystem. Hydrobiologia 200/201:291–301. Van Etten, J. L., Lane, L. C., Meints, R. H. 1991. Viruses and viruslike particles of eukaryotic algae. Microbiological Reviews 55:586–620. Vanni, M. J., Findlay, D. L. 1990. Trophic cascades and phytoplankton community structure. Ecology 71:921–937. Vollenweider, R. A. 1969. A manual on methods for measuring primary production in aquatic environments, IBP Handbook 12. Blackwell Science, Oxford, UK. Wagemann, R., Barica, J. 1979. Speciation and rate of loss of copper from lakewater with implications to toxicity. Water Research 13:515–523. Walker, H. L., Higginbotham, L. R. 2000. An aquatic bacterium that lyses cyanobacteria associated with off-flavor of channel catfish (Ictalurus punctatus). Biological Control 18:71–78. Walker, W. W., Jr. 1987. Phosphorus removal by urban runoff detention basins. Lake and Reservoir Management 3:314–326. Walker, W. W., Jr., Westerberg, C. E., Schuler, D. J., Bode, J. A. 1989. Design and evaluation of eutrophication control measures for the St. Paul water supply. Lake and Reservoir Management 5:71–83. Webster, K. E., Peters, R. H. 1978. Some size-dependent inhibitions of larger cladoceran filterers in filamentous suspensions. Limnology and Oceanography 23:1238–1244. Welch, E. B., Cooke, G. D. 1995. Internal phosphorus loading in

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GLOSSARY

accessory pigment pigment capable of capturing radiant energy and transferring it to chlorophyll-a. accumulation body colored (red, orange, or yellow) lipid material in the cytoplasm, but with uncertain function. acicular needle shaped. acidophile (acidophilic) species that typically occurs in dilute waters (soft water) with lower pH values. aerobic oxygenated environment or metabolism requiring oxygen. aerophytic tendency to colonize subaerial or terrestrial habitats, often under conditions of high humidity. aerotope in cyanobacteria, clusters of gas vesicles; the older term is gas vacuole. agarose gelatinous substance (a sulfated polygalactan) produced in the walls of certain species of red algae. agglomeration loosely arranged mass (of cells); usually colonies with no definite shape or cellular arrangement. Note: Glossary terms were defined by individual authors in reference to their chapter. Consequently, terms may actually apply to other taxa and have broader definitions than indicated here.

akinete thick-walled cell produced by members of several algal classes; may be released by vegetative cells or attached to filaments; functions as an asexual resting stage and typically is resistant to harsh conditions (e.g., low temperatures). algal mat thallus composed of tightly interwoven filaments. algicide various chemicals used to control algal growth in lakes and ponds, such as CuSO4 or Diquat. alkalophile (alkalophilic) species that is typically localized in high ion waters of neutral to high pH values. allochthonous in aquatic ecosystems, matter or production formed outside of the water body, such as terrestrial carbon entering a stream from a forest. allophycocyanin type of blue colored phycobilin pigment produced by members of the cyanobacteria and red algae. alternation of generations life history with two alternating multicellular phases, in which the haploid (n) phase (= gametophyte) produces gametes, which later fuse to form a zygote and later form a diploid (2n) phase (sporophyte). Spores are produced by the sporophyte

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Glossary

often through meiosis. The two phases may be isomorphic or heteromorphic; also termed diplohaplobiontic. alum various forms of aluminum sulfate (e.g., Al2[SO4]3) used to precipitate and reduce dissolved phosphorus in lakes. alveolate pitted or with many small cavities or pores. amoeboid type of cell organization that lacks a cell wall and possesses flexible and frequent changes in shape; present in some members of Chrysophyceae. amorphous without a definite shape. amphiesma in dinoflagellates, a cell covering that includes an outer membrane and a layer of vesicles that may contain cellulosic plate material. ampulla (1) flasklike reservoir that covers the cell of some euglenoids; (2) the reproductive region within the cortical cells of some red algae. amylopectin component of true starch that is a branched glucose polymer composed of α-1,4 linkages and α-1,6 branches; similar to glycogen. amylose component of true starch composed of α-1,4 linkages; an unbranched glucose polymer. anaerobic oxygen-free environment or metabolism that either does not require oxygen or requires an absence of oxygen. anastomosing combination of, or communication between separate branches, sheaths, or filaments (as in the joining of tributaries with rivers). androspore zoospore that becomes a dwarf male (in Oedogoniales). anisogamy (anisogamous) sexual reproduction in which the two types of gametes differ in size or appearance but are both typically motile. antheridium (antheridia) male gametangium in a wide variety of algal groups. antherozoid motile male gametes (e.g., Sphaeroplea), spermatozoid. anthropogenic caused by human activities, often referring to disturbances to ecological systems, such as acidic precipitation. apical cell cell positioned at the end of a filament or thallus; often the site of meristematic growth. apical pore field group of pores at one or both poles in freshwater cymbelloid and gomphonemoid diatoms; functions in secretion of mucilage and forms stalks that attach cells to surfaces. apical series plates at the apex of a cell and that may surround an apical pore. aplanogamete nonmotile gamete produced by a member of an algal group that normally produces flagellated gametes (e.g., flagellated green algae). aplanospore nonmotile (nonflagellated) spore produced by divisions of parental cell; may have the potential to produce flagella in some algal groups. araphid diatom a pennate diatom with no raphe system on either valve. archeopyle in dinoflagellates, the exit pore from which a geminating cell is released from the cyst. arcuate arched or crescent shaped; in diatoms, pennate cells that are bent along the apical axis.

areola in diatoms, a perforation or pore through the valve that is bounded by an internal or external sieve membrane; a more specific term than “puncta,” which is a general term for an opening in the valve wall. armor cellulose plates that form a covering in some dinoflagellates. ash free dry mass measure of biomass in algal and limnological studies, based on the difference between dry mass and the ash remaining after combustion in a muffle furnace (generally between 450 and 600°C); used as an estimate of the organic mass of biological materials. assimilative growth phase in which the cell takes in nutrients (either autotrophic or heterotrophic), not reproductive or dispersive. astaxanthin orange–red xanthophyll (carotenoid) accessory pigment present (temporary or permanent) in some algae (e.g., Haematococcus, Euglena), rendering cells a reddish color. Also occurs in the eyespot of several species; termed haematochrome in older literature. athecate in dinoflagellates, lacking cellulose plates (naked). autochthonous in aquatic ecosystems, matter or production (e.g., primary production) formed in situ, within the water body, such as carbon formed by phytoplankton. autocolony colony formed asexually as coenobia, usually within parent cells and often a miniature of the parent colony (e.g., Scendesmus). autospore nonflagellated spore similar in appearance to the vegetative cell that produced it. autotrophic cells capable of synthesizing organic matter; in algae via photosynthesis. auxospore large cell resulting from sexual reproduction or autogamy. In diatoms, they lack the rigid valve construction of other cells; instead, they are covered by delicate siliceous scales or bands (perizonium). auxotrophy nutritional requirement for external sources of vitamins. axial area in diatoms, an unornamented area along the apical axis; includes the central sternum. In the older literature, the axial area is referred to as the pseudoraphe when applied to araphid diatoms. baeocyte spores produced by certain cyanobacteria or green algae via repeated (often rapid) divisions of vegetative cells, yielding smaller daughter cells. band-shaped diatom colony type caused by the attachment of cells along the entire valve face; also ribbon-shaped. basal toward or at the base or point of origin of a thallus or filament. basal siliceous layer first layer of a diatom frustule deposited during its formation. basionym original name given to a genus or species, and which is retained if transferred to another position or grouping (also written basonym). benthic organisms that grow on or are associated with the bottom (e.g., sediments or rocks) zone of a water body. beta-carotene carotenoid pigment (also β-carotene) in several groups of algae. biogeography study of the geographical distribution of organisms, their patterns, origins, movements, and sources.

Glossary

biomass mass of biological material in a specific area (per square meter) or volume (per liter or cubic meter); in algae, expressed as dry mass, ash-free dry mass, chlorophyll-a, or carbon. biovolume apparent cell volume as calculated from external dimensions. bipartite consisting of or divided into two parts. biraphid diatom diatom with raphe systems on both valves. bloom massive or conspicuous growth of algae, typically planktonic and often forming surface scums; often a large percentage of the total cells are one or a few species. brackish slightly saline, a mixture of fresh and marine water; often used to describe salinity conditions in estuaries. brittlewort see stonewort. caespitose clustered, in thick tufts or clumps, forming a turf. calcification process of depositing calcium carbonate (CaCO3), which occurs on algal cell surfaces or in walls, especially in alkaline habitats. calcite form of calcium carbonate with rhombohedral crystals. calyptra thickened or enlarged tip; in filamentous cyanobacteria, occurs at the tip of some trichomes. cameo raised figure; refers to the raised portion of some diatom valves. canal raphe general term for a raphe that opens into a channel or canal (e.g., Epithemia); internal openings (when present) of the canal raphe are called portules. canopeum in diatoms, an external siliceous covering slightly to distinctly elevated from the valve face; widths are variable, sometimes extending to the valve margin; visible with scanning electron microscope only. Found to contain nitrogen-fixing cyanobacteria in several taxa. capitate with a distinct head; knoblike or swollen at the end. carboxysomes granules or inclusions in cyanobacterial cells (may appear like polyhedrons); contain the enzyme ribulose-1,5-bisphosphate carboxylase/oxygenase. carina ridge (dinoflagellates). carotenoids group of lipid-soluble pigments, including carotenes (yellow or orange) and xanthophylls (yellow or golden), that serve as accessory pigments in photosynthesis. carpogonial branch supports the female gametangium (carpogonium) in the red algae; may or may not be differentiated from surrounding vegetative cells. carpogonium female gametangium in red algae, usually consisting of the receptive region (trichogyne) for the male gamete (spermatium) and a base. carposporangium sporangium of many red algae that typically forms diploid spores (carpospores) on tips of the postfertilization stage (carposporophyte). Carpospores germinate into a free-living diploid phase of the life history. carposporophyte postfertilization stage of many red algae, typically consisting of small diploid filaments (gonimoblast) localized on the haploid gametophyte. Gonimoblast filaments form diploid spores (carpospores) at their apices. cell wall typically rigid external structure enclosing the cell membrane; in algae, may consist of cellulose, silica, pectin, or other materials.

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central area in diatoms, an unornamented area in the central or middle part of the valve face, near or surrounding the proximal raphe ends. central nodule internally, the area between the proximal raphe ends that is usually thickened. central sternum in diatoms, a region of solid silica deposited during the initial stages of silica deposition of the valve. This apically oriented region may or may not be perforated by a raphe slit; nonperforated is termed pseudoraphe in older literature. central strutted process in diatoms, a strutted process that occurs on the valve face, as opposed to the valve margin; see also fultoportule. centric diatom diatom that has a valve structure symmetrical around a point, also termed radial symmetry. centroplasm less pigmented, central region of a cell, especially in cyanobacteria; less heavily pigmented than the chromatoplasm. CER chloroplast endoplasmic reticulum; encircles chloroplasts of some algae. chantransia stage diploid life-history phase of members of the red algal order Batrachospermales and Rhododraparnaldia of the Balbianiales. Forms the haploid gametophyte stage directly attached to it by meiosis of an apical cell. character as applied here, specific structure or attribute useful in classification. chlorophyll-a primary photosynthetic pigment and light receptor (maximum absorption about 663 nm) in algae and higher plants (photosystem I) that is in the form of a porphyrin ring with a central magnesium atom; also used as a chemical marker for algal biomass in ecological studies; also associated with PS-II. chlorophyll-b secondary photosynthetic pigment present in higher plants, green algae, prochlorophytes, and euglenophytes (absorption maximum about 645 nm). chlorophyll-c secondary class of photosynthetic pigment, which occurs in chrysophytes, synurophytes, diatoms, cryptophytes, tribophytes, dinoflagellates, and brown algae; includes two components (both forms not found in all algal groups) termed c1 and c2, each has several different absorption peaks. chloroplast membrane-bound organelle that contains photosynthetic thylakoids, chlorophyll-a, and other pigments (occasionally as “plastid”). chromatic adaptation see photoacclimation. chromatophore organelle that contains pigments (syn. plastid). chromatoplasm portion of a cell that contains pigments; in cyanobacteria, typically a peripheral pigmented region. chrysolaminarin polysaccharide storage product, a β-1,3linked glucan; occurs in several algal groups, including chrysophytes, synurophytes, haptophytes, tribophytes, and diatoms. Also termed leucosin. chytrids group of small fungi that parasitize many organisms, including algae; appear as colorless globose cells on various planktonic diatoms, desmids, and other algae. cingulum in dinoflagellates, a transverse groove that encircles the cell (usually) and holds the transverse flagellum in place. In diatoms, another name for all the girdle bands of a diatom cell (elements of girdle region).

838

Glossary

cirque in limnology, a glacially created lake that formed at the head of a glacial valley in mountainous region; amphitheater shaped. clade grouping of taxa that is recognized to have descended from a common ancestor. cladoceran group of mostly microscopic, planktonic crustaceans (e.g., Daphnia), common in freshwater environments. clathrate with irregular perforations or openings; lattice-like. clone cells or organisms with identical genetic complements derived from an single ancestor. coccoid simple cell type that is spherical, subspherical, or rod-shaped. coccolith calcite structure (scale), aggregations of which surround the exterior of coccolithophorid haptophyte cells. coccolithophorid coccolith-bearing haptophyte. coenobium (coenobia) form of colony in which the number of cells is fixed (genetically determined) at an early develop mental stage; cell number does not change during development. coenocyst in some green algae, condition where the cytoplasm divides in multinucleate portions and develops into a thick-walled resting stage (e.g., Protosiphon). coenocytic multinucleate condition and lacking cross walls; some coenocytic taxa produce cross walls infrequently or during reproduction (e.g., Vaucheria). colony group of cells that may be connected or held together by cytoplasmic strands, mucilage, or parent cell wall. confluent growing together or intermingled; sheaths that join together. conjugation form of sexual reproduction in which nonflagellated gametes join by means of a specialized tube or papillae (e.g., Zygnemataceae). conspecific belonging to the same species. contractile vacuole membrane-bound vesicle that expands and contracts to regulate water and/or osmotic conditions within cells; especially in nonwalled forms. copulae another name for girdle bands that make up the cingulum (in diatoms). cordate heart shaped; also termed cordiform. corona ring of cells at the apex of the oogonium of charophyte algae. cortex peripheral layer of a differentiated thallus that is typically photosynthetic and surrounds the inner colorless medulla cortical filament filament that is part of a surrounding layer that covers the main axis of the thallus (e.g., red algae and charophytes). corticated possessing cortical filaments or cells. costa(e) Latin for rib; as used here, refers to linear thickenings in a diatom valve. craticula network of siliceous bars or ribs formed internally in the diatom genus Craticula, usually under environmental conditions of increased solute concentration. crenate with an edge or margin that is notched, wavy, or scalloped. cribra finely poroid siliceous membranes, which occlude puncta.

cricolith heterococcolith (in haptophytes) with two narrow subhorizontal shields connected by a central tube. cruciform cross-shaped; also termed cruciate. crust closely adherent, flat thallus composed of compacted tiers of cells. cryophilic organisms that colonize ice or snow environments. cuboidal shape similar to a cube. cuneate wedge shaped. cyanelle cyanobacterium-like plastid endosymbiotic within certain protist cells (e.g., green algae Chalarodora and Gloeochaete), although other symbiotic cyanobacteria exist as whole cells within other organisms. cyanophage virus that infects cyanobacteria. cyanophycean starch type of starch produced in cyanobacterial cells that is composed of α-1,4 linkages and α-1,6 branches; similar to glycogen and does not react positively to an iodine test. cyclomorphosis repeated or cyclic change in form, or plasticity in the morphology of an organism, such as elongation of spines (e.g., Scenedesmus) or horns (e.g., Ceratium). cyst nonmotile stage that protects the cell during adverse conditions; also termed resting cyst. daughter cells cells derived from a parent by a mitotic division. dendroid tree-like morphology. desmid unicellular and filamentous green algae that give rise to nonflagellated, amoeboid gametes that conjugate. desmokont in dinoflagellates, a flagellar arrangement where both flagella extend longitudinally from the posterior of the cell. detritus dead or partially degraded organic matter. diadinoxanthin carotenoid pigment (euglenophytes, tribophytes, chrysophytes, synurophytes, dinoflagellates, and diatoms); acts as an accessory pigment in photosynthesis. diatomite type of rock formed primarily from diatom frustules; also known as diatomaceous earth. diatoxanthin golden carotenoid pigment produced by many algal groups (euglenophytes, tribophytes, chrysophytes, synurophytes, haptophytes, dinoflagellates, and diatoms, brown algae); acts as an accessory pigment in photosynthesis. dichotomous branch forked branch consisting of two approximately equal branches. dimictic lake lake that mixes twice per year and stratifies in summer and winter. dinokaryon unique nucleus found in dinoflagellates with permanently condensed chromosomes. dinokont in dinoflagellates, a flagellar arrangement with one transverse flagellum that encircles the cell and a second, longitudinal flagellum that extends out at the posterior of the cell. dinospore motile (flagellated) stage in the life cycle of parasitic forms. dioecious literally, two households; organisms in which male and female (or + and –) gametes are produced on different individuals. diplobiontic life cycle in which there are two free-living growth phases.

Glossary

diplontic life cycle in which there is one multicellular, diploid phase, and the haploid phase consists of only gametes. disjunct in biogeography, locations or populations that are widely separated and generally isolated from one another. distal away from the base or center of a thallus or filament or origin (opposite of basal). distal raphe ends in diatoms, the external terminus of raphe at the poles or ends of valves. dorsal back of a cell or thallus. dystrophic waters rich in organic matter (often from allochthonous sources), that have slow rates of organic matter decay and typically are colored yellow to reddish brown; in many, the pH can be low (≤ 4.0). ecdysis shedding of thecal walls (e.g., dinoflagellates). ecomorphotype type of morphological modification caused by or related to certain ecological conditions. ecorticate condition of a main axis or branch that lacks a surrounding layer of smaller filaments (cortical cells). edaphic referring to the soil. ejectisome in cryptomonads, projectile-like structure (often in a spiral) that is discharged from the cell; may serve as an escape mechanism or direct defense against other organisms. emarginate uneven margin or surface; may have notches or concavities. endemism geographic distribution pattern that is restricted to a particular habitat or locality. endolithic (endolithan) growing within rocks or intercrystalline spaces. endophytic (endophyton) growing within plants or algae; may be intra- or intercellular, or within another species’ mucilage (endogloeic). endoplasmic reticulum series of cytoplasmic membranes in cells that function to process and transport materials or cellular components. endospore in cyanobacteria, specialized spores see baeocytes. In conjugating green algae, the innermost layer of the zygospore wall; see also mesosopore. endosymbiont organism that lives within another, resulting in a mutually beneficial and intimate relationship between the two (e.g., cyanobacteria in diatoms and algae in lichens). environmental optimum apparent preference of a species along a controlling environmental gradient. epicingulum all the girdle elements associated with epivalves. epicone in dinoflagellates, the upper or anterior portion of an athecate cell. epidendric (epidendron) growing on wood. epilimnion upper, well-mixed region of stratified lakes, above the thermocline; typically has greater light availability and is isothermal. epilithic (epilithon) growing attached to the surface of rock or stone substrata. epipelic (epipelon) growing on sediments, clays, and silt. epiphytic (epiphyton) growing on plants and other algae. epipsammic (epipsammon) growing attached to or living among grains of sand. epitheca upper or anterior portion of thecate cell; in diatoms and dinoflagellates.

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epivalve in diatoms, the larger and therefore older of two valves of a frustule. epizooic (epizoon) growing attached to animal surfaces (e.g., Colacium on copepods and cladocerans). eucentric centric diatoms that are circular or approximately circular, in valve view. eukaryotic cell type characterized by the presence of membrane-bound organelles, including a nucleus, mitochondria, endoplasmic reticulum, and (in most algae and plants) chloroplasts. euphotic zone upper depths of a water body that receive sufficient light to support photosynthesis. euplanktonic capable of completing an entire life cycle suspended in the water column. eutrophic literally, well nourished; water bodies that have high levels of dissolved nutrients (esp. N and P) and high levels of organic production; in lakes, often shallower with a broad littoral zone, depleted summer hypolimnetic oxygen, and reduced transparency. eutrophication process of becoming eutrophic. evenness in ecological studies, a measure (or index) of the relative proportion of the community (in numbers or biomass) represented by the species present. eversion process by which a colony in some flagellated green algae fold backward and turns inside out; also termed inversion. exiccata slide set permanent microscope slides and accompanying data distributed to subscribers or established museums for the purpose of allowing access to the specimens at multiple locations. exocyte in cyanobacteria, daughter cells that divide off of distinctly polarized cells (e.g., Chamaesiphon); produced singly or successively in rows; also as known as exospore. exospore in conjugating green algae, the outer layer of the zygospore wall; see also mesosopore. eyespot light sensitive, typically red-colored (carotenoids pigments) spot in many algal cells (typically flagellates or motile reproductive stages of multicellular forms), and typically in the anterior region of the cell. FAA common fixative 10:7:2:1 parts of 95% ethanol:distilled water:formalin: acetic acid. facultative heterotroph ability in cells to use photosynthesis and also to obtain external sources of organic matter, such as under low light conditions. false branching condition where branches arise from a break (or cell death) in the main filament and the continued growth of one or both ends; in cyanobacteria, only the sheath splits while the vegetative trichomes separate. fascia literally a band. In the pennate diatoms, an unornamented area across the middle portion of a valve and visible in valve view, generally rectangular and bandshaped. fascicle literally a bundle. In filamentous algae, a cluster of multiple filaments; in some red algae, lateral vegetative branches; in centric diatoms, a group of rows of areolae oriented radially. fasciculate occurring in bundles, as clusters of filamentous cyanobacteria (e.g., Aphanizomenon) or the striae on the diatom Stephanodiscus.

840

Glossary

fenestra in keel-bearing diatoms, such as Surirella and Cymatopleura, part of the valve surface and mantle fuse together to form openings (fenestra or windows). fenestrated window-like; divided into shiny portions. fibulae internal struts that provide structural support to the raised keel that contains the raphe; extend transapically from valve face to valve mantle (e.g., Nitzschia) or under the raphe in the central region (e.g., Denticula). filament common type of thallus in which cells are arranged in a linear series and in which adjacent cells share a common cross wall. filiform thread-like. fimbriate fringed appearance or bordered with hairs or hairlike structures. flagellar transition region zone between the flagellum and its basal body. flagellate nonphyletic term for protists that possess one or more flagella; also the condition of possessing a flagellum (-a). flagellum (flagella) long, threadlike organelle that projects out of the cell and functions in motility; in eukaryotic cells, they consist of a 9 doublet + 2 central singlet array of microtubules. floridean starch carbohydrate storage granules localized in the cytoplasm of red algae; similar in composition to glycogen; consists of glucose polymers with α-1,4 linkages and α-1,6 branches. footpole in a heteropolar diatom, pole by which the cells attach to each other or to substrata. fresh water ill-defined term used to describe largely inland, low-salinity (< 0.1 g L–1) bodies of water; amounts of dissolved salts vary, but generally much less saline than seawater (ca. 35 g of salts L–1). frustule literally, a box. As applied here, the entire siliceous covering of a diatom cell; the collective term for the epivalve and hypovalve, and their associated cingulum elements. fucosan vesicles small, refractive bodies or vesicles present in brown algal cells and that contain fucosan; also termed physodes. fucoxanthin brown-colored carotenoid pigment produced by members of the golden-pigmented algae (chrysophytes, synurophytes, haptophytes, and diatoms) and brown algae (Phaeophyta); acts as an accessory pigment in photosynthesis. fultoportule another term for strutted process (in diatoms). furcate forked or branched, in filaments, spines, or other processes; used to denote characteristic branching number (e.g., 2 = bifurcate; 3 = trifurcate). fusiform spindle-shaped; any elongate morphology that is widest near the middle and tapering at either end. gametangium (gametangia) any structure or feature of a thallus that produces gametes gametophyte multicellular, usually haploid, gamete producing stage in organisms that exhibit alternation of generations. gas vesicles gas containing spherical or cylindrical structure in cyanobacterial cells, often gathered in clusters termed aerotopes (the older term is gas vacuoles) and visible

with light microscopy as red or brownish bodies (due refraction of light, may be mistaken as actual pigmentation). Function to provide buoyancy to planktonic forms (e.g., Microcystis, Anabaena, and Aphanizomenon). gemmae dense (asexual) aggregations of cells formed in cavities in the thallus (e.g., red alga Hildenbrandia); when released, form new thalli. geniculate having joints like a knee; occurs in some filamentous green algae (e.g., Klebsormidium, Sirogonium, and Zygnema). girdle bands in diatoms, another term for elements of the cingulum. In desmids, formed from very short wall segments; there may be several per cell and they may be obvious in some species (e.g., Closterium and Penium). girdle view in diatoms, a frustule oriented so that the girdle bands and valve mantle are visible, opposed to valve view, where the valve surface and, in some cases, a portion of the valve mantle are visible. globose in the form of a globe; completely or nearly spherical. gloeocapsin dark blue pigment present in the sheaths of certain cyanobacteria glomerate in a compact cluster. glycoprotein protein with attached carbohydrates, often in wall or external gelatinous coatings. Golgi body organelle that consists of a series or stack of flattened, membrane-bound sacs, in which materials are assembled or modified and from which are secreted; also termed dictyosome. gone cell cell derived from a germinating zygote in some green algae. gonidium large, nonmotile cell in colonies of volvocalean green algae that can generate a new daughter colony. gonimoblast filament in red algae, a diploid filament that composes the postfertilization stage (carposporophyte) and is localized on the gametophyte. Diploid spores (carpospores) are formed at the filament tips. guild assemblage of co-occurring taxa with similar ecological requirements or functional roles. gullet in some euglenoids, chrysophytes, cryptomonads, and other flagellates, a depression in the anterior region of the cell where the flagellum (-a) emerges. haematochrome see astaxanthin. hair cell typically colorless, thin, and elongate cell at branch apices that increases the thallus surface area for nutrient and gas exchange; in some filamentous cyanobacteria, green algae and red algae, produced under periods of nutrient deficiency. haplobiontic life cycle in which there is one haploid vegetative phase, where zygotes are the only diploid phase. haptobenthon community of organisms living on hard substrata. haptonema (haptonemata) flagellum-like organelle, the base of which is linked to the basal bodies of flagella; present in haptophyte algae; in longer haptonemata, there is a tendency to form tight coils (e.g., Chrysochromulina parva). In contrast to 9 + 2 ultrastructure of a flagellum, the haptonema consists of 6–7 peripheral microtubules enclosed in a cylinder of endoplasmic reticulum (ER), which is confluent with the peripheral ER of the cell.

Glossary

haptophyte any member of the algal division Haptophyta that possess a well-developed or vestigial haptonema. haptophyte scale platelike organic structure with microfibrillar basal structure (and another type with spinelike superstructure in some species), produced in Golgi bodies and released to the cell exterior where they form a layer surrounding the cell. helictoglossa in diatoms, an internal, liplike structure that terminates the distal raphe. heterococcolith in certain haptophytes, a coccolith with an organic base plate constructed within a specialized Golgi vesicle. heterocyst in certain cyanobacteria (e.g., Anabaena, Nostoc, and Calothrix), a thick-walled, multilayered (apparently gas-tight, anaerobic), and weakly pigmented cell; contains the nitrogenase enzyme, which enables fixation of gaseous nitrogen (N2) to ammonium; also termed heterocyte. heterokont possessing flagella of unequal length or different ornamentation. heteromorphic life history life history in which the gametophyte and sporophyte have different morphologies (i.e., alternation of generations). heteropolar asymmetric to the transverse (longitudinal) axis in a filament, diatom valve, cell, or other structure. In cyanobacteria, morphologically differentiated in basal and apical parts. heterothallic two different strains that are compatible for sexual fertilization; self-incompatible. heterotrichous filamentous thallus with two forms of organization; typically with prostrate (creeping) and erect components (e.g., Gongrosira, Heribaudiella, Hildenbrandia, and Trentepohlia) or otherwise dissimilar forms of branching. heterotrophy (heterotrophic) type of nutrition in which organic matter is obtained from external sources; may involve osmotrophy (dissolved) or phagotrophy (particulate). heterovalvar condition when the two valves of a diatom cell have different structure, such as the density, direction, and ultrastructure of the striae. holdfast unicellular or multicellular structure that attaches a thallus or filament to the substratum homologous character evolutionarily derived character, as opposed to an analogous character, which may appear similar, but is the product of convergent evolution. homonym name applied to an organism that has also been used for another different organism (typically the latter name is invalid). hormogonium (hormogonia) in filamentous cyanobacteria, a means of vegetative reproduction (and dispersal) formed via fragmentation of the trichome, forming distinct segments that are often motile (via gliding). hyaline transparent or colorless. hypersaline extremely saline habitats. hypnospore thick-walled resting cyst. hypnozygote in green algae, dinoflagellates, and some other flagellated forms, a thick-walled, nonmotile resting zygote (cyst) that is dormant in sediments under adverse

841

conditions; may germinate under favorable conditions (in dinoflagellates called a dinocyst). hypocingulum girdle elements associated with the hypovalve (in diatoms). hypocone lower or posterior portion of athecate cell (in dinoflagellates). hypolimnion deeper region of a stratified lake, below the thermocline; typically colder, more poorly illuminated, and richer in nutrients. hypotheca lower or posterior portion of thecate cell; in diatoms and dinoflagellates (below the cingulum). hypovalve in diatoms, the smaller and therefore younger of two valves of a frustule (= hypotheca). imbricate overlapping or arranged like scales or roof tiles; an overlapping series. incrassate swollen or thickened in form or appearance. indeterminant growth pattern or structure (e.g., filament or branch) with no genetically determined form or limit to growth. infrageneric taxon that is below the level of a genus, including species, sections, varieties, and forms. intaglio in diatoms, a surface that is impressed; as used here, the opposite of cameo. intercalary growth growth in the middle of the thallus. internal valves in diatoms, some valve form that are not associated with cell division; they are morphologically similar to normal valves and produced inside normal valve walls; a cell may form several internal valves. internode portion of a thallus, filament or axis between two nodes. inversion see eversion. involucral lateral branch of the main branch (carpogonial branch) bearing the female gametangium (carpogonium) in red algae. isodiametric diameters equal; in filamentous forms, the diameter is more or less constant along its length. isogamy (isogamous) sexual reproduction in which the two types of gametes are morphology indistinguishable, although functionally different. isokont flagella of equal size and form (as recognized by light or electron microscopy). isolated puncta puncta set off from others, in striae. isomorphic life history life history in which the gametophyte and sporophyte have similar (or indistinguishable) vegetative morphologies. isopolar symmetric to the transverse (longitudinal) axis of a filament, diatom valve, cell, or other structure. isthmus in desmids, the constriction (narrow region) between semicells; typically the location of the nucleus. ITS internal transcribed spacers of the ribosomal RNA array; there are two, designated ITS-1 and ITS-2. Janus valves diatom cells where the frustule has two distinctly different valve types in species where the two valves usually appear to be identical. keel in diatoms, a raised or elevated ridge that contains the raphe, formed from a folding of the valve wall. Many keel-forming diatoms have fibulae under the raised ridge for structural support. keritomized netlike in appearance; within cells, sometimes

842

Glossary

the result of vacuolation or irregular arrangement of materials. kettle lake glacial lake formed after melting glacier; typically bowl-shaped depressions, also called kettle holes; common across Wisconsin and the prairie provinces of Canada. kleptoplastidy stealing plastids from prey; plastids usually remain functional in the new host. Km Michaelis Menton constant; a measure of the affinity of an enzyme for its substrate. labiate process tubular process, extending through the valve, with a slitlike inner opening that has thickened margins (giving the appearance of lips); the external opening may be an extended tube or a simple circular pore; also called a rimoportula. lacuna opening or cavity. lacustrine referring to lake (standing water) environments. lamellae stack of thylakoids; also refers to any structures that occur in layers or stacks of plates. lamellate layered or arranged in layers. laminarin in brown algae, a food storage polysaccharide composed mainly of β-1,3 linked glucose units that exists in cells outside of chloroplasts, in vacuoles. laminate platelike. lanceolate shape that is wider near the center than the ends; dory shaped. landslide lake lake basin formed when water flows through an existing depression and is blocked by rock or other material (e.g., Spirit Lake, Mount St. Helens, WA, and Mountain Lake, VA). lateral conjugation in members of the Zygnemataceae, a form of conjugation that occurs between two adjacent cells on the same filament. lentic referring to standing waters; ponds, lakes, or marshes. lenticular lenslike or lentil shaped; flattened or thinning toward the ends. leucoplast colorless plastid with few or no thylakoids; may contain starch (= amyloplast); in some green algae and cryptomonads. leucosin see Chrysolaminarin. lichen mutualistic symbiosis between an alga or cyanobacterium and a fungus, in which the two organisms grow together in an intimate organization. The symbiosis typically forms a distinct macroscopic morphology; often occur in terrestrial, epiphytic, xeric, and occasionally aquatic habitats. ligula silica extension and part of the cingula, in diatoms; in some taxa, projects to intersect with neighboring cingula. lime encrusted calcified thallus; see calcification. linear morphologies with generally parallel sides. linking spine in diatoms, a spine modified to bind daughter valves together, thereby effecting colony formation. list in dinoflagellates, a winglike plate extension. littoral region of lakes or large rivers near the shore; often defined as the area from the shore to the maximum depth of rooted macrophytes. LM light microscope; light microscopy. loculate valve walls that have more than one layer of silica

may have a complex, chambered structure. Areolae may also be considered to be chambered, or loculate (the chamber is termed the loculus). locules older term used for the chambers and other components associated with the girdle bands in Mastogloia; see also partecta. longitudinal canal tubelike chamber apically oriented, extending most of or the entire length of the valve, and may be interrupted in the central area; a general term that applies to structures that may not have the same evolutionary origin (not homologous) such as the canals of Neidium and Diploneis. longitudinal lines lines that run along the apical axis, on either side of the axial area; term is used for lines visible in the light microscope, although they may be formed by different structures. lorica protective investment or envelope that surrounds the protoplast (naked, nonwalled cell); present in genera from several flagellate groups (e.g., Dinobryon and Trachelomonas). lotic running waters springs, streams, and rivers. Lugol’s iodine common fixative; a saturated solution of I2KI: 2 g potassium iodide + 1 g resublimed iodine +300 mL distilled water; it can be acidified with 10 mL glacial acetic acid or made neutral or slightly basic with 1 g of sodium acetate; sometimes used to preserve algal samples; often used for quantifying phytoplankton by inverted microscope method. lutein carotenoid pigment present in many algal divisions. macrandrous sexual reproduction where the male filament or thallus is as large as the female thallus (e.g., species of Oedogonium and Bulbochaete). macroalga algae that form macroscopic or plantlike morphologies with a thallus structure that is recognizable with the naked eye; can be important in the benthic communities of streams and lakes. marginal strutted process tubular process through the valve, with two or more satellite pores on the inner valve surface; external expression may be a tube or a simple pore. marl CaCO3 deposits on substrata or other surfaces in hard water or alkaline lakes and streams; appears whitish and may be precipitated on the surface of algal cells during photosynthesis (termed lime in some literature). mastigonemes hairlike appendages on flagella; numbers and arrangement vary among some algal groups. mastigote motile cell. medulla cells of the inner layer of a differentiated thallus; often colorless and surrounded by an outer photosynthetic layer (the cortex). meiospores spores produced from a zygote via meiosis. meromictic lake lake that mixes in the upper layers (mixolimnion), but deeper waters (monimolimnion) do not; generally the result of much greater density or salt concentration. mesokaryotic nucleus of dinoflagellates once thought to be intermediate between prokaryotic and eukaryotic. mesoplankton plankton size category between 200 and 2000 µm; includes only taxa that form very large colonies, macroscopic clumps, or mats in the plankton.

Glossary

mesospore middle layer of the zygospore wall of conjugating green algae (with exospore and endospore); its structure varies from smooth to ornamented, and is used in identification of genera and species. mesotrophic water bodies with intermediate levels of plant nutrients and organic production. metaboly in euglenoids, motility and flexibility in the pellicle (outer cell covering) without the aid of a flagellum. metaphyton microscopic floating community among the littoral zone plants of lakes and ponds; often loosely associated with aquatic plants. microplankton plankton size category between 20 and 200 µm; includes the largest unicellular algae and many colonial taxa. microplasm connecting pore between neighboring cells (at cross walls) in filamentous cyanobacteria. mitochondrion (mitochondria) in eukaryotes, an organelle with internal and external membranes, the former arranged in folds called cristae; responsible for respiration and contains DNA. mixotrophy (mixotrophic) cells with both photosynthetic and heterotrophic (typically phagotrophic) nutrition (e.g., Dinobryon); occasionally spelled myxotrophy. moniliform filamentous morphology that looks like a string of beads. monoecious literally, one household; organisms in which male and female (or + and –) gametes are produced on the same thallus. monomictic lake lakes that mix once per year, typically in the autumn (warm monomictic). monophyletic natural taxonomic group that shares a most recent common ancestor. monoraphid diatom pennate diatom with a raphe system on one valve only. monosporangium asexual sporangium of some red algae that produces one spore (monospore) that germinates into the same life history phase that produced it. monostromatic thallus composed of a single layer of cells; also monolayer. monotypic genus with a single species. morainal lake lake basin formed through glacial activity wherein the resultant valley is closed by glacial deposits (e.g., Finger Lakes of New York). morphometry sizes and shapes of lake basins; also used to describe a suite of morphological features of an organism. morphotype particular recognizable form of a species that has variable morphology. mucocyst saclike vesicle in certain flagellates that may be released by the cell (= muciferous body). multiaxial thallus with the main axis composed of several to numerous parallel filaments (= multiseriate); in some definitions, differs from multiseriate in that the latter is not necessarily in parallel. multinucleate having many nuclei. multipolar as used here, centric diatom in which the valve outline is not circular. multiseriate see multiaxial. myxophycean starch see cyanophycean starch.

843

myzocytosis in dinoflagellates, particle uptake using the peduncle. nannandrous in some species of green algae (e.g., within Oedogonium), a very small (often unicellular) male filament that attaches to a much larger female filament. nanoplankton plankton size category between > 2 and 20 µm; includes many taxa of unicellular algae and small colonies. natural classification taxonomy that takes into account hypotheses of lineage and evolution, leading to monophyletic groups. necridia in cyanobacteria, dead cells that function to aid separation of filaments or pseudofilaments for vegetative reproduction (e.g., hormogonia) or the formation of false branches. net plankton older term that refers to planktonic organisms captured by a standard plankton net with a mesh size of 25, 40, or 64 µm. net primary productivity quantity of primary production in an ecosystem minus respiration costs (gross – respiration); generally the amount available to the next trophic level. neuston (neustonic) planktonic organisms living at the air– water interface, either above the water (epineuston) or just below the surface (hyponeuston). niche hypothetical construct, with several definitions, including (1) an organism’s role in the biotic environment, in relation to its competitors and enemies; and (2) the set of all ecological conditions and resources that a species can exploit effectively for growth and reproduction, expressed as an n-dimensional hypervolume of resource axes. nitrogen fixation process by which cyanobacteria (and other bacteria) convert atmospheric (gaseous) nitrogen (N2) to biologically useful forms (e.g., NH4+). node location along an axis or thallus that produces branches. Nomarski optics type of high resolution optics in light microscopy; also termed differential interference contrast (DIC). nomenclature as used here, formal rules for naming and referring to organisms. nuclear associated organelle (NAO) structure that appears at the poles of the spindle apparatus during the process of mitosis. nucleomorph highly reduced endosymbiont nuclei present in the cells of some protists (e.g., cryptomonads). obovoid inversely ovoid, with the broader end upward or outward; also termed obovate. occluded process tubular process found near the valve surface margin of some centric diatoms, which have no associated internal opening or structure. ocellulimbus in diatoms, a platelike apical pore field on the mantle. ocellus (ocelli) literally, eyes. (1) In dinoflagellates, a lightsensitive organelle; (2) in diatoms, a large group of areolae or porelli that are physically separated from areolae by an unornamented rim or hyaline area. oligotrophic literally, poorly nourished. Water bodies with low levels of nutrients (esp. N and P) and low levels of organic production; often deep and steep sided with high

844

Glossary

transparency, narrow littoral zone, abundant dissolved oxygen with depth, and larger relative hypolimnion volume. oogamy (oogamous) sexual reproduction in which there is a fusion of a smaller flagellated male gamete (e.g., spermatozoid) with a larger, nonflagellated female gamete (e.g., egg or oogonium). oogonium (oogonia) large, single-celled female gametangium that may produce one or more eggs. organelle membrane-bound intracellular structure with a specific function or functions (e.g., nucleus, chloroplast). osmotrophy nutrition involving the absorption of dissolved organic molecules. ovoid egg shaped; rounded with one pole broader than the other. paedogamous conjugation of gametes within a gametangium. pallium feeding veil of extruded cytoplasm from heterotrophic and mixotrophic dinoflagellates that engulfs prey and may serve as the site of extracellular digestion. palmelloid in many algal groups, a nonmotile, colonial stage of indefinite cell number and arrangement; often within a mucilaginous matrix. papilla protuberance or swelling. papillate covered with papillae or granulate. paramylon storage compound in euglenoids and some tribophytes and haptophytes; a polymer of many β-1,3 linked glucans organized in a membrane-bound crystalline structure; appears as distinct rods or disks in the light microscope; does not stain with iodine. paraphyletic taxonomic group that does not include all of the descendents from a specified parent group or ancestor. parenchyma(tous) type of thallus with true tissues, formed from cell division in three planes; often differentiation of cells into an outer photosynthetic cortex and an inner colorless medulla. parietal pertaining to the outer or peripheral surface of a cell or thallus; parietal chloroplasts are localized within plasma membrane near the cell wall. partecta in some diatoms, a loculate chamber that is associated with the valvocopula in Mastogloia. parthenospore spores formed in association with sexual reproduction in conjugating green algae; usually produced in cells that have failed to pair up with a compatible sexual partner. peduncle cytoplasmic extension used as a means of attachment. In dinoflagellates, a feeding structure exerted through the flagellar pore in the sulcus; see also pallium. pelagic open water region of lakes; also termed the limnetic zone. pellicle in some dinoflagellates, the layer beneath the amphiesmal vesicles, which remains upon ecdysis of outer layers. In Euglenoids, the outer proteinaceous surface layer, often in a helical arrangement. penicillate in a small tuft; brushlike. pennate diatom diatom with features symmetrical about a line, termed bilateral symmetry, and produces amoeboid gametes. PER periplastid (surrounding the plastid) endoplasmic reticulum.

pericentral cell formed immediately adjacent to the main axis. peridinin golden brown carotenoid pigment produced by dinoflagellates that acts as an accessory pigment in photosynthesis (esp. in dinoflagellates). periphyton collection of organisms (algae, bacteria, fungi, and protozoa) and detritus attached to surfaces without stipulating substratum type; in common usage refers to algal cells in this habit. periplast in cryptomonads, the outer cell covering, consisting of a series of overlapping plates plus the plasma membrane. phagopod in dinoflagellates, a specialized feeding tube that lacks microtubules and is used to obtain nutrients phagotophically from other organisms. phagotrophy type of heterotrophic nutrition based on ingestion of particulate organic carbon. Particle capture may involve pseudopodia, flagella, or haptonema (haptophytes). photoacclimation ability to vary the amounts of different photosynthetic pigments in response to changes in the light environment; in cyanobacteria this may result in distinctly different colors of the thallus. photoautotroph(ic) organism that synthesizes its own organic compounds through photosynthesis. photoheterotroph pigmented species capable of photosynthesis and metabolizing external dissolved organic molecules to meet its energy needs. phototaxis movement of an organism toward (positive) or away from (negative) a light source. phragmoplast form of cell division in which the microtubules lie perpendicular to the plane of division; cytokinesis thus occurs via centrifugal cell plate formation. Occurs in certain groups of green algae and true plants. phycobiliproteins water soluble pigments of the cyanobacteria, red algae, and cryptomonads; there are three basic types red-colored phycoerythrin, and blue-colored phycocyanin and allophycocyanin. These molecules act as accessory pigments in photosynthesis. phycobilisomes granules (usually spherical or disk shaped) located on the thylakoids, consisting of the phycobiliprotein accessory pigments; in cyanobacteria and red algae. phycocyanin blue pigment phycobiliprotein (water soluble) pigment produced by cyanobacteria, red algae, and cryptomonads. phycoerythrin red pigment phycobiliprotein (water soluble) pigment produced by cyanobacteria, red algae, and cryptomonads. phycoplast form of cell division in which the array of microtubules is oriented parallel to the plain of division; a new set of microtubules lie perpendicular to them to form a new cell wall; occurs in members of the green algal class Chlorophyceae. phylogenetic attributes or studies that recognize the evolutionary relationships among organisms; classification systems based on these relationships. physode see fucosan vesicles. phytoplankton portion of the microscopic floating (plankton) community represented by algae, including cyanobacteria.

Glossary

picoplankton plankton size category between 0.2 and 2 µm; includes several genera of cyanobacteria (e.g., Synechococcus and Synechocystis), a few green (e.g., Nannochloris), eustigmatophyte, and chrysophyte algae. placcoderm desmid members in the class Desmidiaceae; cells with distinct semicells (walls of either half are of different ages), in which there is a median constriction (= sinus), a connecting zone (= isthmus), and pores in their walls. Cells may be solitary, or joined in colonies or filaments. planktonic growing suspended, floating, or drifting in water. planospores motile cells, asexual zoospores or sexual gametes. planozygote motile, thick-walled cyst resulting from sexual reproduction; in some flagellates (e.g., Woloszynskia), may store lipids and/or starch. plaque in diatoms, thin external layers of silica along the girdle bands. plastids organelles bounded by two membranes and typically with thylakoids and photosynthetic pigments (e.g., chloroplasts); also, an alternate term for chloroplast. Others, such as amyloplasts (+ starch) lack pigments. plicate folded or in furrows, giving the appearance of folds. plurilocular sporangium in brown algae, a reproductive structure that becomes subdivided into many compartments (locules), from which single flagellated zoospores (swarmers) are released. pole terminal portions of cells, filaments, pennate diatom valves. polygonal in the shape of a polygon; many sided; angular. polymictic lake shallow lake with frequent or continuous mixing, in tropical and equatorial areas. polymorphic having more than one form. polyphosphate storage granules of condensed polymers of inorganic phosphate that are visible with light microscopy; in several algal groups. polyphyletic artificial taxonomic group; not sharing a recent, common ancestor; members may be more closely related to organisms outside the group. polysiphonous thallus that appears as multiple parallel filaments (or siphons), formed from a series of adjacent filaments; in some genera of red algae (e.g., Polysiphonia). pore field in diatoms, a specialized area of pores unlike the pores of the striae, usually associated with organelles that extrude material from the cell. porelli small, closely packed perforations through the valve wall of diatoms; perforations in ocelli, pseudocelli, and apical pore fields. primary production amount of new organic matter synthesized by autotrophic organisms. prokaryotic cell type that lacks membrane-bound organelles; also lack true (9+2) flagella; also as procaryotic. protist phyletic group (Kingdom) of eukaryotic organisms not classified as members of the Plant, Fungi, or Animal kingdoms; includes all eukaryotic algae, plus non-photosynthetic protists (protozoa). protolichen association of subaerial or soil algae growing in close association with fungi, but not forming an intimate symbiosis; see lichen. protoplast living material inside cells, excluding the organelles.

845

proximal raphe ends raphe ends on the central nodule (internal valve surface) and near the central portion of the valve (external valve surface). pseudocelli group of areolae set off from the pattern of the rest of the valve, which decreases in size from areolae on the main part of the valve. Pseudocelli are not physically separated from areolae by unornamented band or ring. pseudocilia see pseudoflagella. pseudofilament loose chain of cells held together with a common gelatinous matrix, or linked by fibrils or other connections; cells are typically separated from each other. Unlike true filaments, they do not share a common cross wall; also termed a chain. pseudoflagellum giving the appearance of flagella, but not functional in motility (e.g., Tetrapora). pseudonodule in diatoms, a differentiated area or structure on the valve that has a variable form; essentially an area that may resemble an ocellus or pseudocellus, but is structurally different. In freshwater diatoms, this structure is found only in Actinocyclus. pseudoparenchyma(tous) thallus that is tissue-like but composed of compacted or interwoven filaments; resembles parenchyma. Often, the main axis consists of a single filament (uniaxial) or a series of parallel filaments (multiaxial). pseudopyrenoid smaller, naked pyrenoid. pseudoraphe literally, a false raphe; an unornamented linear region in the axial area of some pennate diatom valves. Usually called the central sternum in current terminology. pseudosepta plate or lamina of silica projecting internally from the apical portion of the valve. puncta(e) pore/perforation through the valve when substructure (i.e., sieve membrane) is unknown or lacking. pusule in dinoflagellates, an organelle composed of a series of tubes and thought to function in nutrient uptake and possibly expulsion of excess water. pyrenoid distinct, proteinaceous structure (often spherical), embedded in or associated with chloroplasts of algae; in some, contains the enzyme RuBisCO; associated with starch in some green algae. pyriform pear shaped. quadrate square or rectangular. raphe structure in monoraphid and biraphid diatoms that consists of a slit through the valve face, and associated cytoplasmic structures; usually situated along the apical axis or within a marginal keel; composed of (usually) two branches per valve. The raphe enables a diatom cell to move over substrata. raphe branch single raphe slit, extending from the proximal end to distal ends. rapheless in monoraphid diatoms, an adjective that describes a valve that lacks a raphe system; also araphid. raphid in monoraphid diatoms, an adjective that describes a valve that has a raphe system. RbcL gene encoding the large subunit of the RuBisCO enzyme; sequences are used in phylogenetic analyses. refractive index quantity reflecting the degree to which a substance refracts light. reniform bean or kidney shaped.

846

Glossary

replicate folded back. reticulate netlike or arranged to form a network. rhizoid downward-growing cell or chain of cells that is typically involved in thallus attachment to substrata. rhizoplast striated strand that connects flagellar basal bodies with the nucleus. rhizostyle in cryptomonads, a part of the flagellar apparatus that consists of microtubules that extend from the basal bodies into the cell. rib in diatoms, a linear siliceous thickening on the diatom valve; see costa. In synurophyte algae, a peripheral thickening on siliceous scales. rimoportule(e) in diatoms, another term for labiate process. 16S rRNA gene encoding the small subunit (SSU) of the ribosomal RNA array of prokaryotes, mitochondria and chloroplasts; sequences are used in phylogenetic analyses. 18S rRNA gene encoding the small subunit (SSU) of the nuclear ribosomal RNA array of eukaryotes; sequences are used in phylogenetic analyses. RuBisCO ribulose-1,5-bisphosphate carboxylase/oxygenase; the enzyme in photosynthetic organisms that catalyses (“fixes”) the incorporation of CO2 into carbohydrate. saccate like a sac or balloon. saccoderm desmid members in the family Mesotaeniaceae (older term); mostly simple unicells with no distinct semicells (see placcoderm desmids) and with no pores in their walls; some become linked to form filaments. scalariform conjugation literally, ladder-like; in members of the Zygnemataceae, a type of conjugation, where the conjugation tube forms between two parallel filaments. scrobiculate surface pitted or furrowed; with many small depressions. scytonemin yellow–brown pigment that is present in the sheaths of certain cyanobacteria; may protect against extremes of UV radiation. seasonal succession regular seasonal changes in the species composition of phytoplankton over an annual cycle. SEM scanning electron microscope; scanning electron microscopy. semicell one-half of a cell in members of the Placcoderm desmids. semierect nonrigid thallus that is capable of bending and may exhibit branch reconfiguration to reduce drag in high current velocities. Sensu lato Latin, in the broad sense. Sensu stricto Latin, in the strict sense. separation valve specialized valve with elongated spines that facilitates separation of daughter valves and serves to limit colony size in some centric diatoms. septum (septa) in most algal cells, a cross wall. In diatoms, an invagination into the cell lumen in the valve plane, usually arising from the valvocopulae. seta stiff hair or bristle (e.g., Coleochaete); also an elongate, external projection from the cell wall. In some forms it may be a hollow, narrow extension of the wall. sheath mucilaginous covering over cells, colonies, or filaments (trichomes); may be firm or loose, narrow or broad.

silica deposition vesicle vesicle in which the siliceous diatom cell wall is deposited during cell division; also functions in some chrysophytes and synurophytes to form siliceous spores. silica scale siliceous coverings produced by members of the Chrysophyceae and Synurophyceae. siliceous composed primarily of silicon. sinkhole lake lake formed from the dissolution of (mainly limestone) bedrock by surface and underground waters charged with CO2. sinus constricted region in a cell; the median constriction in desmids (class Desmidiaceae). siphonous type of thallus with large multinucleate cells and few (or no) cross walls, except where reproductive structures occur. smooth flagellum flagellum lacking hairs; see whiplash flagellum. species richness ecological attribute of communities that describes the total number of species present in an area (or volume of water). specific conductance ability of a water sample to conduct electricity; it is the reciprocal of resistance and is measured in units of micromhos per centimeter (µΩ cm–1) or microsiemens per centimeter (µS cm–1). Used as an estimate of total dissolved solids (TSS); the older term is conductivity. spermatangium in red algae, a male gametangium that is typically a colorless, obovoid cell produced at the tips of vegetative branches; each produces a single male gamete (spermatium) that is released to fertilize the female gametangium (carpogonium). Some red algae form spermatangia on specialized stalk cells. spermatium colorless male (nonflagellated) gamete in red algae. spine thin projection, simple or branched, cylindrical or conical, ending in a point or flattened; common in many algal groups. In diatoms, appears as a granule or a tube. spinule minute spinelike or thornlike protuberances. sporangium (sporangia) structure that produces spores. spore specialized (asexual) reproductive structure that germinates without fusion. sporic meiosis meiosis that occurs during spore production. sporophyte typically diploid, spore producing stage in organisms that exhibit alternation of generations. sporopollenin resistant polymer produced in the vegetative walls (e.g., some Chlorococcales) or spore walls (e.g., Charales) of certain green algae that provides added strength and is thought to help prevent desiccation. starch storage polysaccharide; a polymer composed of glucose units with α-1,4 and α-1,6 linkages. Composed of amylose and amylopectin. statospore siliceous resting stage produced by several types of algae (e.g., Chrysophyceae and Synurophyceae). status in diatoms, differing types of frustules regularly formed by some genera, notably Aulacoseira. stauros in diatoms, where the central nodule (more heavily silicified) is expanded to the valve mantle to form a crosslike structure. stellate literally star shaped; describes several features of algal

Glossary

cells, such as chloroplasts or cell shape. In diatoms, a colony type caused by attachment of cells at the poles at one end of each cell (e.g., Asterionella). stenotherm organism with a narrow range of temperature tolerance or preference. sternum silica thickening along the axis of many pennate diatoms along which the valve structure is built during cell division. stichidia inflated, multichambered structures at the tips of vegetative branches (red algae). stigma see eyespot. stigmata in some diatoms, an isolated perforation through the valve face, usually in the central area, where the external opening is rounded (or nearly so) and internal openings may be slitlike or otherwise modified. stigmoids perforation through valve face the external opening of which is similar to puncta of the valve and the internal opening of which is slightly modified from the other puncta. stipulode unicellular outgrowth of branchlets (bractlike) present in some members of the Charales. stomatocyst see statospore. stonewort common name for charophyte algae; refers to the common occurrence of calcification on the thallus in many species. stratification property of lakes in which water forms two or more layers that differ in temperature and/or density, with warmer upper waters and cooler, deeper strata. striae literally a line. In diatoms, an approximately linear array of puncta or areolae. strutted process in diatoms, a tubular process found in some centric diatoms, usually associated with secretion of β-chitin. Internally a tube surrounded by two or more satellite pores; externally either a tube or a simple pore in the valve wall; also as fultoportula. subaerial ecological habit exposed to the air, but usually attached to various substrata; also terrestrial. subarctic regions that are nearly arctic; somewhat south of the Arctic Circle. sulcus in dinoflagellates, a longitudinal groove in the ventral face of the cell (in hypotheca/hypocone) that may extend into epitheca/epicone; holds the longitudinal, whiplash, flagellum. suture in dinoflagellates, the edge of a plate in thecate species, sometimes raised. tabulation system used to classify the plate morphology in thecate dinoflagellates. taxon (taxa) species or a group at any taxonomic level. taxonomy formal system of classification. tectonic lake lake formed by movements of the Earth’s crust, usually continental rifting [e.g., Lake Tahoe (USA), Lake Baikal (Siberia), Lake Chapala (Mexico), and African rift lakes]. TEM transmission electron microscope; transmission electron microscopy. terminal fissure continuation of the raphe on the valve exterior beyond the point where the raphe penetrates to the internal part of the valve.

847

terminal nodules in raphid pennate diatoms, the terminal nodule is a siliceous thickening distal to the internal raphe terminus. tetrachotomous dividing into four equal or nearly equal braches or divisions. tetrasporangium meiotic sporangium of certain red algae that forms four spores (tetraspores) that germinate into the gametophyte. tetraspore haploid spores are formed by meiosis and germinate into the gametophytic stage in red algae. tetrasporophyte diploid, typically free-living stage of some red algae. Forms four haploid spores (tetraspores) by meiosis. These spores germinate into the gametophyte stage. thallus (thalli) general form or body of an alga. theca in dinoflagellates, cellulose plate covering. In diatoms, another term for a diatom frustule. In flagellate green algae, a rigid wall. thermocline region in lakes in which there is an abrupt change of at least 1° m–1; the depth where the rate of temperature change with depth is greatest. thermokarst lake lakes formed from the freezing and thawing action in ice and soil; common in the arctic. thylakoid flattened membranous vesicles (or sacs) that form the photosynthetic membranes and contain photosynthetic pigments; arranged in various patterns in cyanobacterial cells or within plastids in eukaryotic algae. tinsel flagellum flagellum with dense mastigonemes (bristlelike extensions); in some groups they occur in two rows of tripartite hairs (e.g., members of Chrysophyceae, Synurophyceae, and Phaeophyceae). tomont in dinoflagellates, a cyst formed following feeding in parasitic forms. travertine calcium carbonate (concretionary limestone) that is precipitated from alkaline water, often with the participation of algae; common in several species of cyanobacteria. tremalith heterococcolith composed of pentagonal calcite elements fused into a ringlike or tubelike structure (e.g., in the freshwater species Hymenomonas roseola). trichoblasts elongate hair cells, often terminal; occur in cyanobacteria, red and green algae. trichocyst in dinoflagellates and raphidophytes, a proteinaceous ejectile organelle. trichogyne receptive portion of the female gametangium (carpogonium) of many red algae. trichome in cyanobacteria, a term referring to a filament without its sheath. trichotomous dividing into three equal or nearly equal braches or divisions. trophont feeding stage in certain dinoflagellates. true branching branches formed from lateral divisions of the primary axis; in cyanobacteria, where divisions occur longitudinally and resultant cells grow roughly perpendicular to the original trichome. True branches are physiologically connected to the original trichome, unlike false branches. tufts short radiating filaments without a common matrix.

848

Glossary

turbidity degree of cloudiness or opacity of a water sample as influenced by suspended matter (sediment, silt, organisms), and which causes light to be scattered and absorbed; typically measured via nephelometry in standard units (NTUs) against formazin-based reference suspensions. tychoplanktonic species that are predominately planktonic, but capable of prolonged survival on or in sediments. ultrastructure morphological features observable with an electron microscope. uniaxial (= uniseriate) thallus with the main axis composed of a single chain of cells (filament). unilocular sporangium in brown algae, a reproductive structure (sporangium) that becomes enlarged (often spherical or club-shaped) but does not subdivide into compartments, and which later releases many spores, typically the result of meiosis. valve top (epivalve) and bottom (hypovalve) elements of a diatom frustule. valve face portion of the valve visible in valve view, that is, the valve oriented to the valvar plane. valve mantle portion of the valve, differentiated by slope, that is apparent in the girdle view (oriented to the apical plane). valvocopula girdle elements directly attached to valves in diatom frustules. Accessory structures, such as septa, usually arise from valvocopulae. velum in diatoms, a thin plate or flap of silica that covers the openings of loculate areolae; may be perforated with smaller openings (in diatoms). ventral front (in dinoflagellates, sulcal) side of a cell or thallus.

verruca(e) short or wartlike irregular projection(s) on the surface of cell walls or spores. voigt faults irregularities in striae ornamentation bordering one side of the axial area, approximately equidistant from the valve center. These interruptions in the pattern of the striae are the result of valve ontogeny, representing the last areas of silica deposition. volcanic lake lake basin formed via volcanic activity; include maars (volcanic explosion) and caldera (collapsed crater); examples include Crater Lake in Oregon and Lake Nicaragua. whiplash flagellum smooth flagellum with no hairs. whorl type of branching that has several lateral filaments arising at a common position in the main axis. wing in diatoms, a complex type of keel that results from an extensive fold in the valve wall around the raphe. Extreme folding can result in a siliceous fusion (or partial fusion) of the internal valve surface below the raphe. zigzag colony type (e.g., diatoms) caused by the attachments at both ends of cells; also a growth patterns in some filamentous green algae (e.g., Stichococcus). zooplankton portion of the plankton community represented by mainly microscopic (up to a few millimeters) animals and protists (in freshwater: cladocerans, copepods, rotifers, heterotrophic flagellates, ciliates, and fish larvae). zoospore motile (flagellated) spore (“naked swarmer”) formed by vegetative cells or in specialized sporangia; typically the same ploidy as the parental cell. zygospore thick-walled resting cyst formed after fertilization of an oogonium or fusion of gametes in other algal groups. zygotic meiosis meiosis that occurs after maturation (cell division) or germination of the zygote.

Author Index

A Abel, R. A., Olson, D. M., Dinerstein, E., Hurley, P. (2000), 12, 45 Aboal, M., Puig, M. A., Soler, G. (1996), 584, 588 Adamson, R. P., Sommerfield, M. R. (1978), 809, 826 Adler, R. W. (1995), 786, 797 Admiraal, D. M. J. (1993), 38, 45 Admiraal, W., Breebaart, L., Tubbing, D. M. J., Van Zanten, B., de Ruitjer van Steveninck, E. D., Bijerk, R. (1994), 38, 46 Admiraal, W., Mylius, S. D., de Ruitjer van Streveninck, E. D., Tubbing, D. M. J. (1993), 32, 38, 46 Agbeti, M. D. (1992), 791, 797 Agbeti, M. D., Kingston, J.-C., Smol, J. P., Watters, C. (1997), 587–588, 616, 631 Agbeti, M. D., Smol, J. P. (1995), 613, 616, 631 Ahlstrom, E. G. (1937), 477, 488, 498, 503 Ahmadjian, V. (1993), 45–46 Al-Dhaheri, R. S., Willey, R. L. (1996), 408, 416 Al-Thukair, A. A., Golubic, S., Rosen, G. (1994), 107, 110 Albertano, P. Capucci, E. (1997), 65, 110 Albertano, P., Kovácik, L. (1994), 135, 191 Albertano, P., Pinto, G., Pollio, A. (1994), 489, 503 Alcantara, I. I. (1997), 587–588 Allanson, B. R. (1973), 811, 826

Allegre, C. F., Jahn, T. L. (1943), 415–416 Allen, D. L., Jarrell, W. M. (1989), 823, 826 Allen, D. M., Northcote, D. H. (1975), 484, 504 Allen, J. D. (1995), 28, 34, 46 Allen, M. M. (1968), 81, 110 Allen, T. F. H. (1977), 623, 631 Allen, W. E. (1921), 36, 46 Alles, E., Nörpel-Schempp, M., Lange-Bertalot, H. (1991), 665, 666 Allorge, P., Manguin, E. (1941), 763, 772 Aloi, J. E. (1990), 24–25, 46, 780, 797 American Public Health Association (1992), 807, 809, 826 American Public Health Association (1998), 779–780, 783, 797 Amoros, C., Van Urk, G. (1989), 789, 797 Anagnostidis, K. (1961), 120–121, 127, 190, 191 Anagnostidis, K. (2001), 150, 192 Anagnostidis, K., Golubic´, S. (1966), 132, 192 Anagnostidis, K., Komárek, J. (1988), 110, 117–120, 128, 135, 139, 144, 150, 155, 192 Anagnostidis, K., Komárek, J. (1990), 60, 110, 120, 192 Anagnostidis, K., Komárek, J. (2001), 132, 139, 144, 192 Anagnostidis, K., Pantazidou, A. (1991), 66, 110 Anagnostidis, K., Roussonoustakaki, M. (1985), 144, 145, 192 Andersen, R. A. (1982), 480, 504

849

850

Author Index

Andersen, R. A. (1985), 528, 552 Andersen, R. A. (1986), 481, 501, 504 Andersen, R. A. (1987), 471, 474, 504, 523–524, 528, 552 Andersen, R. A. (1989), 513, 519 Andersen, R. A. (1992), 716, 751 Andersen, R. A. (1996), 410, 416 Andersen, R. A., Barr, D. J. S., Lynn, D. H., Melkonian, M., Moestrup, Ø., Sleigh, M. A. (1991), 402, 416 Andersen, R. A., Brett, R. W., Potter, D., Sexton, J. P. (1998), 424–425, 427, 465–466 Andersen, R. A., Brett, R. W., Potter, D., Sexton, J. P. (1998a), 471, 473, 504 Andersen, R. A., Mulkey, T. J. (1983), 474, 504, 528, 552 Andersen, R. A., Potter, D., Bidigare, R. R., Latasa, M., Rowan, K., O’Kelly, C. J. (1998b), 471, 473, 491, 504 Andersen, R. A., Potter, D., Daugberg, N., Bailey, J. C. (1998c), 503–504 Andersen, R. A., Saunders, G. W., Paskind, M. P., Sexton, J. P. (1993), 471, 504 Andersen, R. A., Van de Peer, Y., Potter, D., Sexton, J. P., Kawachi, M., LaJeunesse, T. (1999), 528, 534, 552 Andersen, R. A., van de Peer, Y., Potter, D., Sexton, J. P., Kawachi, M., LaJeunesse, T. (1999), 473–474, 504 Andersen, R. A., Wetherbee, R. (1992), 20, 46, 473, 504 Anderson, E. (1962), 734, 746, 751 Anderson, L. W. J. (1993), 809, 811, 826 Anderson, L. W. J., Dechoretz, N. (1984), 824, 826 Anderson, N. J., Battarbee, R. W. (1994), 789, 793, 797 Anderson, N. J., Ripply, B., Gibson, C. E. (1993), 787, 797 Anderson, N. J., Vos, P. (1992), 789, 797 Andresen, N. A., Stoermer, E. F. (1978), 666 Andrews, H. T. (1970), 483, 501, 504 Andreyeva, V. M. (1998), 307 Angeler, D. G. (2000), 385, 416 Angermeier, P. L., Katt, J. R. (1994), 776, 786, 797 Anita, N. J., Cheng, N. Y., Foyle, F. A., Percival, E. (1979), 736, 751 Anita, N. J., Kalley, J. P., McDonald, T., Bisalputra, T. (1973), 747, 751 Ankley, G. T., Mattson, V. R., Leonard, E. N., West, C. W., Bennett, J. L. (1993), 825, 826 Ann, S. S., Friedl, T., Hegewald, E. (1999), 256–257, 307 Anonymous (1975), 670, 682 Anonymous (1990), 811, 817, 826 Anthoni, U., Christophersen, C., Madsen, J. O., Wium-Andersen, S., Jacobsen, N. (1980), 811, 826 Apt, K. E., Collier, J. L., Grossman, A. R. (1995), 737, 751 Archibald, R. E. M., Barlow, D. J. (1983), 663, 666 Archibald, R. E. M., Schoeman, F. R. (1984), 663, 666 Archiblad, P. (1973), 291, 307 Archiblad, P., Bold, H. C. (1970), 255, 307 Archiblad, R. E. M. (1970), 682 Archiblad, R. E. M. (1972), 784, 797 Ariztia, E. V., Andersen, R. A., Sogin, M. L. (1991), 424, 466 Arnoldi, V. P. (1916), 247–248 Arruda, J. A., Fromm, C. H. (1989), 776, 797, 807, 827 Arvola, L., Ojala, A., Barbosa, F., Heaney, S. I. (1991), 738, 751 Arvola, L., Tulonen, T. (1998), 740, 751 Asaul, Z. I. (1975), 415–416 Ashley, J., Rushforth, S. R., Johansen, J. R. (1985), 605, 631 Asmund, B. (1968), 538, 552 Asmund, B., Hilliard, D. K. (1961), 552 Asmund, B., Kristiansen, J. (1986), 524, 531, 533–534, 551–552 Asmund, B., Takahashi, E. (1969), 552 Auer, M. T., Graham, J. M., Graham, I. E., Kranzfelder, J. A. (1983), 25–26, 46

Autenrieth, R., Bonner, J., Schreiber, L. (1991), 797 Aziz, A., Whitton, B. A. (1988), 39, 46 B Babin, J., Prepas, E. E., Murphy, T. P., Serediak, M., Curtis, P. J., Zhang, Y., Chambers, P. A. (1994), 814, 827 Bachmann, H. (1908), 478, 504 Bachmann, H. (1921), 472, 504 Bachmann, M. D., Carlton, R. G., Burkholder, J. M., Wetzel, R. G. (1986), 45–46 Badgery, J. E., McQueen, D. J., Nichols, K. H., Schaap, P. R. (1994), 818, 827 Bahal, M. Talpasayi, E. R. S. (1972), 119, 192 Bahls, L. L. (1982), 650, 651 Bahls, L. L. (1993), 778, 784, 787, 791, 797 Bahls, L. L., Weber, E. E., Jarvie, J. O. (1984), 604, 631 Bahls, L. Weber,C. I. (1988), 674, 682 Bailey, D., Mazurak, A. P., Rosowski, J. R. (1973), 63, 110 Bailey, J. C., Andersen, R. A. (1998), 429, 466 Bailey, J. C., Bidigare, R. R., Christensen, S. J., Andersen, R. A. (1998), 443, 466, 473, 491, 503–504 Bain, M. B. (1993), 821, 827 Baker, A. L., Baker, K. K. (1979), 37–38, 46, 110 Baker, A. L., Baker, K. K. (1981), 65, 110 Baker, E. R., McLaughlin, J. J. A., Hutner, S., DeAngelis, B., Feingold, S., Frank, O., Baker, H. (1981), 406, 416 Balch, W. M., Holligan, P. M., Ackelson, S. G., Voss, K. J. (1991), 511, 519 Balech, E. (1974), 690, 710 Balech, E. (1980), 690, 710 Bando, T. (1988), 379 Barber, H. G., Haworth, E. Y. (1981), 20, 46, 599, 631, 651 Barber, L. (1980), 385, 416 Barbiero, P. R., McNair, C. M. (1996), 535, 552 Barbiero, R. P., Welch, E. B. (1992), 23, 46 Barbiero, R. P., Welch, H. (1992), 64, 110, 166, 192 Barbour, M. T., Plafkin, J. L., Bradley, B. P., Graves, C. G., Wisseman, R. W. (1992), 789, 797 Barbour, M.T., Gerritsen, J., Snyder, B. D., Stribling, J. B. (1999), 786, 789, 797 Barica, J. (1975), 806, 827 Barica, J. (1978), 806, 827 Barica, J., Kling, H., Gibson, J. (1980), 812, 827 Barker, G. L. A., Handley, B. A., Vacharapiyasophon, P., Stevens, J.R., Hayes, P. K. (2000), 177, 192 Barko, J. W., Adams, M. S., Clesceri, N. L. (1986), 816, 827 Barko, J. W., James, W. F., Taylor, W. D., McFarland, D. G. (1990), 814, 827 Barko, J. W., Smart, R. M. (1980), 816, 827 Barko, J. W., Smart, R. M. (1981), 816, 827 Barnett, B. S., Schneider, R. W. (1974), 810, 827 Barr, D. J. S., Hickman, C. J. (1967), 364, 379 Barsanti, L., Passarelli, V., Walne, P. L., Gualtieri, P. (1997), 402, 416 Bartlett, R., Willey, R. (1998), 408, 416 Bateman, L., Rushforth, S. R. (1984), 604, 631 Batko, A. (1970), 235, 248 Batko, A., Zakrys, B. (1995), 389, 402, 412, 415–416 Battarbee, R. W., Charles, D. F., Dixit, S. S., Renberg, I. (1999), 560, 588, 775, 781, 797 Batterbee, R. W. (1986), 605, 631 Battey, J. F. (1992), 687, 710 Bauer, D. L., Willie, D. W. (1990), 820, 827 Beaumont, P. (1975), 28, 46 Beaver, J. (1981), 561, 572, 588, 638, 651, 791, 797 Beech, P. L. (1990b), 533, 552

Author Index Beech, P. L., Wetherbee, R., (1990a), 528, 552 Beech, P. L., Wetherbee, R., Pickett-Heaps, J. D. (1990), 531, 533, 553 Beeton, A. M. (1969), 806, 827 Beijernick, M. W. (1980), 254, 307 Belanger, S. E., Barnum, J. B., Woltering, D. M., Wowling, J. W., Ventullo, R. M. Schermerhorn, S. D., Lowe, R. L. (1994), 794, 797 Belanger, S. E., Farris, J. L., Cherry, D. S. (1989), 825, 827 Belanger, S. E., Lowe, R. L., Rosen, B. G. (1985), 672, 682 Belay, A., Kato, T., Ota, Y. (1996), 121, 192 Belbin, L., McDonald, C. (1993), 790, 797 Belcher, J. H. (1959), 771–772 Belcher, J. H. (1966), 481, 484, 499, 504 Belcher, J. H. (1968), 484, 504 Belcher, J. H. (1969), 484, 504, 524, 528, 531, 553 Belcher, J. H., Swale, E. M. F. (1976), 502, 504 Belcher, J. H., Swale, E. M. F. (1997), 587, 588 Belcher, J. H., Swale, E. M. F., Heron, J. (1966), 583, 588 Bell, R. A. (1993), 45, 46 Benke, A. C., Hall, C. A. S., Hawkins, C. P., Lowe-McConnell, R. H., Stanford, J. A., Suberkrop, K., Ward, J. V. (1988), 29, 31–32, 46 Benndorf, J. (1989), 819, 827 Bennett, G. W. (1971), 809, 827 Bennett, H. D. (1969), 41–42, 46 Bennion, H., Juggins, S., Anderson, N. J. (1996), 794, 797 Benson, C. E., Rushforth, S. R. (1975), 454, 459, 461, 463, 466 Berg, C. O. (1963), 13, 16, 42, 46 Bergman, B. (2002), 141, 192 Bergquist, A. M., Carpenter, S. R. (1986), 19, 46 Berlyn, G. P., Micksche, J. P. (1976), 365, 379 Berninger, U. G., Caron, D. A., Sanders, R. W. (1992), 23, 46 Berninger, U. G., Finlay, B. J., Canter, H. M. (1986), 45–46 Bessey, C. E. (1899), 655, 666 Bethge, H. (1925), 569, 588 Bettoli, P. W., Maceina, M. J., Noble, R. L., Betsill, R. K. (1992), 810, 827 Bettoli, P. W., Maceina, M. J., Noble, R. L., Betsill, R. K. (1993), 810, 821, 827 Bhattacharya, D., Helmchen, T., Bibeau, C., Melkonian, M. (1995), 737, 751 Bhattacharya, D., Weber, K., An, S. S., Berning-Koch, W. (1998), 228, 248 Bianchi, T. S., Findlay, S., Dawson, R. (1993), 39, 46 Bicuda, C. (1984), 379 Bicudo, C. E. deM., De-Lamonica-Freire, E. M. (1993), 414, 416 Bicudo, C. E. deM., Wolowski, K. (1998), 388, 416 Bidigare, R. R., Ondrisek, M. E., Kennicutt, M. C., Iturriaga, R., Harvey, H. R., Hohman, R. W., Macko, S. A. (1993), 43–44, 46 Biebel, P. (1968), 335, 349 Biggs, B. F., Goring, D. G., Nikora, V. I. (1998), 35–36, 46 Biggs, B. J. F. (1990), 785, 798 Biggs, B. J. F. (1995), 776, 779, 785, 798 Biggs, B. J. F. (1996), 33–35, 46, 779, 782, 798 Biggs, B. J. F., Close, M. E. (1989), 779, 798 Biggs, B. J. F., Kilroy, C., Mulcock, C. M. (1989), 775, 779, 798 Biggs, B. J. F., Ondrisek, M. E., Kennicutt, M. C., Iturriaga, R., Harvey, H. R., Hohman, R. W., Macko, S. A. (1993), 46 Biggs, B. J. F., Price, G. M. (1987), 783, 798 Billen, G., Garnier, J., Hanset, P. (1994), 38, 46 Bird, D. F., Kalff, J. (1986), 473, 504 Bird, D. F., Kalff, J. (1987), 23, 46, 473, 504 Birks, H. J. B. (1995), 785, 790–793, 798 Birks, H. J. B. (1998), 604, 631, 790–793, 798 Birks, H. J. B., Line, J. M., Juggins, S., Stevenson, A. C., ter Braak, C. J. E. (1990), 794, 798

851

Bixby, R. J. (2001), 631 Blackburn, S. I., Jones, G. J. (1995), 177, 192 Blackwell, J. R., Cox, E. J., Gilmour, D. J. (1991), 255, 307 Blanck, H. (1985), 776, 798 Blank, R. J. (1992), 687, 710 Bliding, C. (1963), 349–350 Bliding, C. (1968), 349–350 Blindow, I. (1987), 24, 28, 46 Blindow, I. (1988), 811, 827 Blindow, I. (1992), 28, 46 Blinn, D. W. (1971), 42, 46 Blinn, D. W. (1993), 42–43, 46 Blinn, D. W., Fredericksen, A., Korte, V. (1980), 36, 46 Blinn, D. W., Prescott, G. W. (1976), 210, 221 Blinn, D. W., Shannon, J. P., Benenati, P. L., Wilson, K. P. (1998), 34, 46 Blinn, D. W., Stein, J. R. (1970), 333, 348, 350 Blinn, D. W., Truitt, R. T., Pickart, A. (1989), 25, 46 Bloem, J., Bär Gilissen, M.-J. B. (1988), 514, 519 Blum, J. L. (1951), 463, 466 Blum, J. L. (1956), 31–32, 34, 46 Blum, J. L. (1972), 465–466 Bodyl, A. (1996), 385, 416 Bolas, P. M., Lund, J. W. G. (1974), 810, 827 Bold, H. C. (1958), 349–350 Bold, H. C., MacEntee, F. J. (1973), 403–404, 416 Bold, H. C., Starr, R. C. (1953), 236, 248 Bold, H. C., Wynne, M. J. (1978), 255, 257–258, 307 Bold, H. C., Wynne, M. J. (1985), 225, 230, 232, 235, 238, 245, 248, 415–416, 423–424, 426, 429, 432, 466, 758, 772 Boltovskoy, A. (1975), 703, 710 Boltovskoy, A. (1976), 703, 710 Boltovskoy, A. (1989), 703, 710 Boney, A. D. (1980), 354, 363, 379 Boney, A. D. (1981), 353, 363, 379 Boney, A. D. (1982), 354, 363, 379 Booton, G. C., Floyd, G. L., Fuerst, P. A. (1998), 257, 307, 313, 350 Boraas, M. E., Estep, K. W., Johnson, P. W., Sieburth, J. McN. (1998), 740, 751 Borchardt, M. A. (1996), 782, 798 Borchardt, M. A., Hoffmann, J. P., Cook, P. W. (1994), 35, 46 Born, S. M., Wirth, T. L., Brick, E., Peterson, J. O. (1973), 811, 827 Bornet, E., Flahault, C. (1886-1888), 155, 158, 161, 164, 166, 171, 174, 177, 180–181, 184, 187, 189, 192 Boschker, H. T. S., Dekkers, E. M. J., Pel, R., Cappenberg, T. E. (1995), 39, 46 Bothe, H. (1982), 20, 45, 46, 192 Bott, T. L. (1996), 776, 798 Bott, T. L., Brock, J. T., Cushing, C. E., Gregory, S. V., King, D., Petersen, R. C. (1978), 785, 798 Boucher, P., Blinn, D. W., Johnson, D. B. (1984), 16, 46, 274, 308 Bouck, G. B. (1982), 402, 416 Bouck, G. B., Ngô, H. (1996), 399, 401, 416 Bouck, G. B., Rogalski, A., Valaitis, A. (1978), 402, 416 Bourne, C. E. M. (1992), 569, 585, 588 Bourrelly, P. (1957), 497, 503, 504 Bourrelly, P. (1963), 485, 504 Bourrelly, P. (1966), 110, 126, 192, 268–269, 271–273, 275, 278–279, 282–283, 285–287, 289–290, 292–294, 296–297, 299–300, 302, 304–308 Bourrelly, P. (1968), 707, 710 Bourrelly, P. (1968, 1981), 434, 447, 451, 453, 458–459, 465–466 Bourrelly, P. (1970), 119, 160, 192, 387–388, 411, 415, 416, 708, 710, 716, 741, 746, 751

852

Author Index

Bourrelly, P. (1981), 503–504, 512, 516, 519, 523, 553, 757–759, 765, 770–772 Bourrelly, P. (1985), 12, 40, 46, 63, 100, 102, 104, 110 Bourrelly, P. (1988), 307–308, 313, 345, 349–350 Bourrelly, P., Manguin, E. (1952), 75, 102, 110, 234, 241–242, 248, 412, 415–416 Bovee, E. C. (1982), 404, 416 Bowling, L. C., Baker, P.D. (1966), 776, 783, 798 Boxhorn, J. E., Holden, D. A., Boraas, M. E. (1998), 501, 504 Boyd, C. E. (1982), 810, 827 Boyd, C. E., Prather, E. E., Parks, R. W. (1975), 806, 827 Boyer, C. S. (1926), 561, 588 Boyer, C. S. (1927a), 561–562, 588 Boyer, C. S. (1927b), 561–562, 588 Boyne, A. F. (1979), 734, 752 Braarud, T. (1954), 516, 519 Bradbury, J. P. (1975), 604, 631 Bradbury, J. P. (1987), 604, 631 Bradbury, J. P., Waddington,J. C. B. (1978), 605, 631 Bradbury, J. P., Winter, T. C. (1976), 604, 631 Bradbury, J. P.(1988), 561, 588 Braun, A. (1855), 757, 772 Brett, S. J., Wetherbee, R. (1986), 732, 741, 745, 752 Brezny, D., Mehta, I., Sharma, R. K. (1973), 810, 827 Brezonik, P. L. (1978), 779, 798 Briand, F., Trucco, R., Ramamoorthy, S. (1978), 785, 798 Bright, R. C. (1968), 604, 631 Brinson, M. M. (1993), 779, 798 Broadwater, S. T., Scott, J. L. (1994), 201, 221 Broady, P. (1984), 121, 189, 192 Broady, P., Given, D., Greenfield, L., Thompson, K. (1987), 189, 192 Brock, M. L., Wiegert, R. G., Brock, T. D. (1969), 41, 46 Brock, T. D. (1967), 41, 47 Brock, T. D. (1973), 40, 42, 47 Brock, T. D. (1985a), 12, 19, 47 Brock, T. D. (1985b), 41, 47 Brock, T. D. (1986), 41, 47 Brocklesby, J. (1851), 384, 416 Brönmark, C., Klosiewski, S. P., Stein, R. A. (1992), 25, 47 Brook, A. J. (1981), 354, 361, 363–364, 379 Brook, A. J. (1997), 371, 379 Brook, A. J. (1998), 371, 380 Brook, A. J., Williamson, D. B. (1988), 40, 47 Brooks, A. E. (1966), 243, 248 Brooks, A. E. (1972), 227, 248 Brooks, J. L., Deevey, E. S. (1963), 19, 40, 47 Browder, J. A., Gleason, P. J., Swift, D. R. (1994), 39, 47, 66, 110 Brown, A. W. A. (1978), 825, 827 Brown, C. W., Yoder, J. A. (1993), 511, 519 Brown, D. L, Weier, T. E. (1968), 200, 221 Brown, L. M., Smith, R. J., Shivers, R. R., Day, A. W. (1986), 518–519 Brown, R. M., Herth, W., Franke, W. W., Romanovicz, D. (1973), 484, 504 Brown, S.-D. (1973), 623, 631 Brown, S. W., Boyd, C. E. (1982), 807, 827 Brownlee, B. G., Painter, D. S., Boone, R. J. (1984), 809, 827 Brugam, R. B. (1980), 604, 631 Brugam, R. B., McKeever, K., Kolesa, L. (1998), 604, 631 Brugerolle, G., Bricheux, G. (1984), 531, 553 Bruno, S. F., McLaughlin, J. J. A. (1977), 700, 710 Brunskill, G. J., Ludlam, S. D. (1969), 16, 47 Brunthaler, J., Prowazek, S., von Wettstein, R. (1901), 583, 588 Bryant, D. A. (1994), 61, 110

Buchheim, M. A., Buccheim, J. A., Chapman, R. L. (1997), 234, 248 Buchheim, M. A., Chapman, R. L. (1997), 234, 249 Buchheim, M. A., Lemieux, C., Otis, C., Gutell, R. R., Chapman, R. L., Turmel, M. (1996), 234, 249 Buchheim, M. A., McAuley, M. A., Zimmer, E. A., Theriot, E. C., Chapman, R. L. (1994), 225, 232, 249 Bucka, H., Zurek, R. (1992), 699–700, 710 Buckland-Nicks, J., Reimchen, T. E. (1995), 697, 701, 710 Buckland-Nicks, J., Reimchen, T. E., Garbary, D. J. (1997), 686, 688, 698, 701, 709–710 Budde, H. (1927), 763, 765, 772 Büdel, B. (1985), 103, 110 Büdel, B., Henssen, A. (1983), 103, 110 Büdel, B., Lüttge, U., Stelzer, R., Huber, O., Medina, E. (1994), 110 Büdel, B., Wessels, D. C. J. (1991), 110 Buell, H. F. (1938), 82–83, 110 Buetow, D. E. (1968), 384, 416 Bujak, J. P., Davies, E. H. (1983), 698, 710 Bujak, J. P., Williams, G. L. (1981), 697, 710 Bukhtiyarova, L. N. (1995), 631 Bukhtiyarova, L., Round, F. E. (1996), 597–598, 602, 627, 631 Bunting, M. J., Duthie, H., Campbell, D., Warner, B., Turner, L (1997), 789, 798 Burkholder, J. M. , Wetzel, R. G., Klomparens, K. I. (1990), 674, 683 Burkholder, J. M. (1996), 669, 682 Burkholder, J. M. (1998), 685, 710 Burkholder, J. M., Glasgow, H. B., Jr. (1997), 776, 798 Burkholder, J. M., Malin, M. A., Glasgow, H. B., Larsen, L. M., McIver, M. R., Shank, G. C., Deamer-Melis, N., Briley, D. S., Springer, J., Touchette, B. W., Hannon, E. K. (1997), 814, 827 Burkholder, J. M., Noga, E. J., Hobbs, C. H., Glasgow, H. B. (1992), 685, 710 Burkholder, J. M., Sheath, R. G. (1984), 364, 380 Burkholder, J. M., Wetzel, R. G. (1990), 25, 47 Burnham, J. C., Fraleigh, P. D. (1983), 819, 827 Burns, C. W. (1968), 808, 827 Burns, C. W., Stockner, J. G. (1991), 20, 47, 65, 110 Bursa, A. A. (1969), 688, 698, 706, 710 Bursa, A. A. (1970), 688, 698, 701, 706, 710 Butcher, R. W. (1933), 29, 47 Butcher, R. W. (1947), 775, 798 Butcher, R. W. (1967), 716, 741, 743, 748, 752 Button, K. S., Hostetter, H. P., Mair, D. M. (1977), 823–824, 827 C Cain, J. R., Trainor, F. R. (1973), 785, 798 Calado, A. J., Moestrup, Ø. (1997), 701, 710 Calado, A. J., Rino, J. A. (1994), 524, 526, 528, 553 Caljon, A. (1983), 739, 752 Calkins, G. N. (1926), 385, 416 Callieri, C., Bertoni, R., Amicucci, E. Pinolini, M. A. (1995), 65, 110 Callow, P. (1973a), 26, 47 Callow, P. (1973b), 26, 47 Camburn, K. E. (1982), 434, 459, 466, 561, 588 Camburn, K. E., Kingston, J. C. (1986), 573, 588 Camburn, K. E., Kingston, J. C., Charles, D. F. (1984–1986), 561, 588, 598–599, 601, 628, 631 Camburn, K. E., Lowe, R. L., Stoneburner, D. L. (1978), 604, 631 Camburn, K. E., Warren, M. L., Jr. (1983), 212, 221 Cameron, R. E. (1963), 63, 110 Cameron, R. E., Morelli, F. A., Blank, G. B. (1965), 63, 110 Cameron, W. A., Larson, G. L. (1993), 15, 47 Campbell, C. E., Prepas, E. E. (1986), 43, 47 Campbell, E. O., Sarafis, V. (1972), 349–350 Campeau, S., Murkin, H. R., Titman, R. D. (1994), 39, 47

Author Index Canavan, R. W., Siver, P. A. (1995), 19, 47 Canfield, D. E. (1983), 812, 827 Canter, H. M. (1968), 739, 752 Canter, H. M., Jaworski, G. H. M. (1979), 819, 827 Canter-Lund, H., Lund, J. W. G. (1995), 45, 47, 171, 192, 314, 350, 387, 416, 604, 631 Caraco, N. F., Cole, J. J., Raymond, P. A., Strayer, D. L., Pace, M. L., Findlay, S. E. G., Fischer, D. T. (1997), 37, 47 Carefoot, J. R. (1966), 242, 249 Carignan, R., Kalff, J. (1980), 816, 827 Carlson, R. E. (1977), 775–776, 785, 798, 806, 812, 827 Carmichael, W. W. (1992), 64, 110, 169, 192 Carmichael, W. W. (1994), 786, 798, 807, 827 Carmichael, W. W. (1997), 64, 110, 121, 155, 192, 807, 827 Carmichael, W. W., Evans, W. R., Yin, Q. Q., Bell, P., Moczydlowski, E. (1977), 809, 827 Carmichael, W. W., Ewans, W. R., Yin, Q. Q., Bell, P., Mocauklowski, E. (1997), 121, 155, 192 Carmichael, W. W., Falconer, I. E. (1993), 807, 827 Carmichael, W. W., Jones, C. L. A., Mahmood, N. A., Thiess, W. C. (1985), 807, 818, 827 Carmona-Jimenéz, J., Gold-Morgan, M. (1994), 181–182, 192 Caron, D. A., Lim, E. L., Dennett, M. R., Gast, R. J., Kosman, C., DeLong, E. F. (1999), 473, 504 Caron, D. A., Sanders, R. W., Lim, E. L., Marrase, C., Amaral, L. W., Whitney, S., Aoki, R. B., Porter, K. G. (1993), 23, 47 Carpenter, E. J. Carmichael, W. W. (1995), 806, 828 Carpenter, S. R., Kitchell, J. F. (1993), 776, 798 Carpenter, S. R., Kitchell, J. F., Hodgson, J. R. (1985), 818, 828 Carpenter, S. R., Kitchell, J. F., Hodgson, J. R., Cochran, P. A., Elser, J. J., Elser, M. M., Lodge, D. M., Kretchmer, D., He, X., von Ende, C. (1987), 818, 828 Carpenter, S. R., Lathrop, R. C., Munoz-del-Rio, A. (1993), 19, 47 Carper, G, L., Bachmann, R. W. (1984), 814, 828 Carr, N. G., Whitton, B. A. (1973), 59, 63, 68, 110, 120–121, 192 Carr, N. G., Whitton, B. A. (1982), 59, 111, 121, 192 Carrick, H. J., Lowe, R. L., Rotenberry, J. T. (1998), 605, 631 Carson, J. L., Brown, R. M. (1978), 45, 47 Carty, S. (1986), 693, 698, 702, 710 Carty, S. (1989), 689, 702, 710 Carty, S. (1993), 698, 701–702, 710 Carty, S., Cox, E. R. (1985), 695, 701, 707, 710 Carty, S., Cox, E. R. (1986), 708, 710 Carty, S., Fazio, V. W., III (1997), 701, 710 Cassani, J. R., Caton, W. E. (1983), 820, 828 Cassie, V., Dempsey, G. P. (1980), 587, 588 Castenholz, R. W. (1960), 42, 47 Castenholz, R. W. (1969a), 66, 82, 111, 192 Castenholz, R. W. (1969b), 82, 111, 192 Castenholz, R. W. (1970), 82, 111 Castenholz, R. W. (1976), 66, 111, 129, 192 Castenholz, R. W. (1977), 111 Castenholz, R. W. (1982), 24, 47 Castenholz, R. W. (1992), 60, 62, 111, 117–118, 192 Castenholz, R. W. (2001), 59–60, 111, 192 Castenholz, R. W., Waterbury, J. B. (1989), 59, 111 Castenholz, R. W., Wickstrom, C. E. (1975), 41, 47, 66, 111, 189, 192 Catling, P. M., McKay, S. M. (1980), 347, 350 Cattaneo, A. (1983), 25, 47 Cattaneo, A. (1996), 35, 47 Cattaneo, A., Amireault, M. C. (1992), 25, 47 Cattaneo, A., Galanti, G., Gentinetta, S. Romo, S. (1998), 24–25, 47 Cattaneo, A., Kerimian, T., Roberge, M., Marty, J. (1997), 36, 47 Cavalier-Smith, T. (1981), 383, 416 Cavalier-Smith, T. (1986), 523, 553, 715, 737, 742, 752

853

Cavalier-Smith, T. (1991), 387, 416 Cavalier-Smith, T. (1993), 383, 385, 387, 416, 742, 749, 752 Cavalier-Smith, T. (1998), 383–385, 402, 416 Cavalier-Smith, T. (1999), 385, 387, 402, 416 Cavalier-Smith, T., Chao, E. E. (1996), 423–424, 466 Cavalier-Smith, T., Couch, J. A., Thorssteinsen, K. E., Gilson, P., Deane, J. A., Hill, D. R. A., McFadden, G. I. (1996), 737, 752 Cedeno-Maldondo, A., Swader, J. A. (1974), 823, 828 CEIMATE (2000), 797, 798 Cepak, V. (1993), 61, 111 Chadefaud, M. (1950), 759, 763, 767, 772 Chandler, D. E. (1984), 734, 752 Chapman, A. D., Pfiester, L. A. (1995), 694, 710 Chapman, R. L., Waters, D. A. (1992), 311, 315, 332, 343–344, 348, 350 Charles, D. F. (1985), 604, 631 Charles, D. F., Binford, M. W., Furlong, E. T., Hites, R. A., Mitchell, M. J., Norton, S. A., Oldfield, F., Paterson, M. J., Smol, J. P., Utala, A. J., White, J. R., Whitehead, D. R., Wise, R. J. (1990), 792, 798 Charles, D. F., Smol, J. P. (1988), 537, 553, 787, 798 Charles, D. F., Smol, J. P. (1994), 792–793, 798 Chesnick, J. M., Cox, E. R. (1987), 694, 707, 710 Chesnick, J. M., Cox, E. R. (1989), 694, 707, 710 Chesnick, J. M., Morden, C. W., Schmieg, A. M. (1996), 686, 710 Chessman, B., Growns, I., Currey, J., Plunkett-Cole, N. (1999), 788, 798 Chiavelli, D. A., Mills, E. L., Threlkeld, S. (1993), 408, 416 Chilton, E. W., Lowe, R. L., Schurr, K. M. (1986), 798 Chisholm, S. W., Stross, R. G. (1976a), 406, 417 Chisholm, S. W., Stross, R. G. (1976b), 406, 417 Chodat, R. (1922), 482, 504 Choi, H.-G., Kraft, G. T., Saunders, G. W. (2000), 218, 222 Cholnoky, B. J. (1965), 661–662, 666 Cholnoky, B. J. (1968), 583, 588, 623, 631, 672, 683 Chorus, I., Bartram, J. (1999), 64, 92, 111, 121, 169, 192 Christen. see von Christen Christensen, C. L., Reimer, C. W. (1968), 676, 682–683 Christensen, T. (1962), 471, 504, 511, 519 Christensen, T. (1964), 471, 504 Christensen, T. (1968), 465–466 Christensen, T. (1969), 465–466 Christensen, T. (1978), 757, 772 Christensen, T. (1987a), 465–466 Christensen, T. (1987b), 465–466 Christensen (1987), 379, 380 Christie, C. E., Smol, J. P. (1993), 604, 631 Christoffersen, K., Riemann, B., Hansen, L. R., Klysner, A., Sorensen, H. B. (1990), 20, 47 Cienkowski, C. (1870), 485, 504 Clark, R. L., Rushforth, S. L. (1977), 604, 631 Clarke, K. B. (1994), 584, 588 Clarke, R. A., Stanley, C. D., MacLeod, B. W., McNeal, B. L. (1997), 807, 828 Clasen, J., Bernhardt, H. (1982), 535, 553 Claus, G. (1962), 44, 47 Clay, B. L., Kurgens, P. , Lee, R. E. (1999), 716, 732, 734, 737, 742–743, 746, 752 Clay, B. L., Kurgens, P. (1999a), 716, 732–733, 735, 737, 739, 742, 749, 752 Clay, B. L., Kurgens, P. (1999b), 716, 729, 732, 735, 737, 739, 742, 749, 752 Clay, B. L., Kurgens, P. (1999c), 716, 732, 737, 742, 749, 752 Clayton, C., Häusler, T., Blattner, J. (1995), 386, 417 Clements, F. E., Shantz, H. L. (1909), 86, 111

854

Author Index

Cleve, P. T. (1894), 637, 646–647, 651, 659, 665, 666 Cleve, P. T. (1895), 637, 651 Cleve, P. T., Grunow, A. (1880), 596, 631 Cleve-Euler, A. (1911a), 569, 588 Cleve-Euler, A. (1911b), 569, 588 Cleve-Euler, A. (1951), 562, 588 Cleve-Euler, A. (1952), 562, 588 Cleve-Euler, A. (1953), 599, 613, 628, 631 Cleve-Euler, A. (1953a), 562, 588 Cleve-Euler, A. (1953b), 562, 588 Cleve-Euler, A. (1955), 562, 588 Cmiech, H. A., Leedale, G. F., Reynolds, C. S. (1986), 120, 192 Cochran-Stafira, D. L., Andersen, R. A. (1984), 40, 47 Codd, G. A. (1995), 64, 111, 121, 169, 192 Coesel, P. F. M. (1983), 364, 380 Coesel, P. F. M. (1991), 364, 380 Coesel, P. F. M. (1993), 354, 374, 376, 379–380 Coesel, P. F. M. (1996), 364, 380 Coesel, P. F. M., Delfos, A. (1986), 373, 380 Cohen-Bazire, G., Bryant, D. A. (1982), 61, 111 Cole, G. A. (1963), 15–16, 40–41, 47 Cole, G. A. (1994), 12, 15–16, 40, 42, 47 Cole, J. J., Caraco, N. F., Peierls, B. (1992), 32, 37, 47 Coleman, A. W. (1959), 226–227, 241, 249 Coleman, A. W. (1977), 226–227, 241, 249 Coleman, A. W. (1996), 227, 249 Coleman, A. W. (1999), 242, 249 Coleman, A. W., Goff, L. J. (1991), 473, 504 Coleman, A. W., Suarez, A., Goff, L. (1994), 227, 249 Colle, D. E., Shireman, J. V. (1980), 810, 828 Colletti, P. J., Blinn, D. W. Pickart, A., Wagner, V. T. (1987), 35, 47 Collins, F. S. (1905), 463, 466 Collins, F. S., Holden, I., Setchell, W. A. (1898), 770, 772 Collins, G. B., Kalinsky, R. G. (1977), 604, 631 Colt, L. C. (1974), 429, 433, 461, 463, 466 Colt, L. C. (1985), 433, 463, 466 Colt, L. C., Jr., Saumure, R. A., Jr., Baskinger, S. (1995), 12, 47, 339, 350 Comas, A. (1996), 303, 307–308 Compère, P. (1982), 584, 588 Compère, P. (1996), 376, 379–380 Compère, P. (2001), 631–631 Conforti, V. (1991), 405, 417 Conforti, V. (1999), 415, 417 Conforti, V., Joo, G.-J. (1994), 416, 417 Conforti, V., Perez, M. del C. (2000), 405, 417 Conforti, V., Ruiz, L. (2000), 414, 417 Conforti, V., Walne, P. L., Dunlap, J. R. (1994), 414–417 Conley, D. J., Kilham, S. S., Theriot, E. (1989), 598, 632 Conley, D. J., Kilham, S. S., Theriot, E, C, (1989), 567, 588 Conley, D. J., Schelske, C. L., Stoermer, E. F. (1993), 567, 588 Conn, H. W., Edmondson, C. H. (1918), 385, 417 Conrad, W. (1926), 247, 249 Conrad, W. (1928), 516, 519 Conrad, W. (1934), 415, 417 Conrad, W., van Meel, L. (1952), 415–417 Conyers, D. L., Cooke, G. D. (1982), 817, 828 Cook, R. B., Kreis, R. G., Jr., Kingston, J. C., Camburn, K. E., Norton, S. A., Mitchell, M. J., Fry, B., Shane, L. C. K. (1990), 617, 632 Cooke, G. D., Kennedy, R. H. (1989), 817, 828 Cooke, G. D., Welch, E. B., Martin, A. B., Fulmer, D. G., Hyde, J. B., Schrieve, G. D. (1993a), 814, 828 Cooke, G. D., Welch, E. B., Peterson, S. A., Newroth, P. R. (1993b), 806, 811, 813–818, 820, 828

Cooke, M. C. (1883), 235, 249 Cooksey, K. E., Cooksey, B. (1978), 39, 47 Cooper, S. R. (1999), 789, 798 Copeland, J. J. (1936), 60, 66–67, 72–73, 75–82, 88, 94–95, 104–105, 107, 111, 118, 121, 126, 128, 131–132, 134–137, 139–140, 144, 148, 154–155, 158, 186–189, 192 Corbus, F. G. (1982), 810, 828 Corliss, J. O. (1975), 384, 417 Corliss, J. O. (1984), 383, 385, 417 Corliss, J. O. (1989), 384, 391, 417 Corliss, J. O. (1990), 388, 417 Corliss, J. O. (1991a), 388, 417 Corliss, J. O. (1991b), 385, 417 Corliss, J. O. (1995), 385, 388, 417 Corliss, J. O., Esser, S. C. (1974), 404, 417 Corpe, W. A., Jensen, T. E. (1992), 65, 79, 89, 111 Coste, M., Bosca, C., Dauta, A. (1991), 776, 782, 787, 798 Cotê, R. (1983), 786, 798 Cottingham, K. L., Carpenter, S. R., St. Amand, A. L. (1998), 487, 504 Couch, J. N. (1932), 432, 466 Couté, A. (1983), 480, 504 Couté, A., Sarthou, C. (1990), 202, 222 Couté, A., Tell, G. (1981), 365, 380 Couté, A., Thérézien, Y. (1994), 416–417 Covach, A. P. (1976), 47 Cowles, R. P., Brambel, C. E. (1936), 428, 463, 465–466 Cox, E. J. (1987), 650, 651 Cox, E. J. (1990), 490, 504 Cox, E. J. (1996), 560, 562, 588, 638, 651, 655, 666 Cox, E. R., Bold. H. C. (1966), 337, 349–350 Cox, E. R., Hightower, J. (1972), 45, 48 Cracraft, J. (1989), 569, 588 Craige, J. S. (1974), 757, 772 Craige, J. S. (1990), 200, 222 Cramer, M., Myers, J. (1952), 406, 417 Cranwell, P. A., Robinson, N., Eglinton, G. (1985), 699, 710 Crawford, R. M. (1988), 583, 588 Crawford, R. M., Gardner, C. (1997), 607, 632 Crawford, S. A. (1979), 811, 828 Croasdale, H. (1973), 60, 80, 111, 433, 459, 461, 466 Croasdale, H. T., Bicudo, C., Prescott, G. W. (1983), 374–380 Cronberg, G. (1980), 531, 553 Cronberg, G. (1982), 537, 553 Cronberg, G. (1986), 531, 533, 553 Cronberg, G. (1989), 485, 504, 533, 535–536, 538, 552–553 Cronberg, G. (1996), 485, 504, 526, 538, 553 Cronberg, G., Gelin, C., Larsson, K. (1975), 537, 553 Cronberg, G., Komárek, J. (1994), 67, 89, 111, 135, 192 Cronberg, G., Komárek, J. (2002), 171, 192 Cronberg, G., Kristiansen, J. (1980), 536, 553 Cronberg, G., Lindmark, G., Bjork, S. (1988), 428, 466 Cronk, J. K., Mitsch, W. J. (1994), 39, 48 Croome, R. L. (1988), 537, 553 Croome, R. L., Tyler, P. A. (1985), 535, 537, 553 Croome, R. L., Tyler, P. A. (1987), 709, 711 Crumpton, W. G. (1987), 570, 588 Crumpton, W. G., Wetzel, R. G. (1981), 570, 588 Crumpton, W. G., Wetzel, R. G. (1982), 48 Crumpton, W. G., Wilson, S. E., Hall, R. I., Smol, J. P. (1995), 588 Crutchfield, J. R., Jr., Schiller, D. H., Herlong, D. D., Mallen, M. A. (1992), 820, 828 Cullen, J. J., Ciotiti, A. M., Lewis, M. R. (1997), 783, 798 Cumming, B. F., Davey, K., Smol, J. P., Birks, H. J. (1994), 793, 798

Author Index Cumming, B. F., Smol, J. P., Birks, H. J. B. (1992a), 526, 535–537, 541, 553, 781, 787, 792, 798 Cumming, B. F., Smol, J. P., Kingston, J. C., Charles, D. F., Birks, H. J. B., Camburn, K. E., Dixit, S. S., Uutala, A. J., Selle, A. R. (1992), 604, 632 Cumming, B. F., Smol, J. P., Kingston, J. C., Charles, D. F., BIrks, H. J. B., Camburn, K. E., Dixit, S. S., Uutala, A. J., Selle, A. R. (1992b), 793, 798 Cumming, B. F., Smol, J. P., Kingston, J. C., Charles, D. F., Birks, H. J. B., Camburn, K. E., Dixit, S. S., Uutala, A. J., Selle, A. R. (1992b), 553 Cumming, B. F., Wilson, S. E., Hall, R. I., Smol, J. P. (1995), 604, 630, 632 Cumming, B. F., Wilson, S. E., Hall, R. I., Smol, P. J. (1995), 638–639, 651 Cummins, B. F., Wilson, S. E., Hall, R. I., Smol, J. P. (1995), 560–561, 588 Czarnecki, D. B. (1979), 26, 48 Czarnecki, D. B. (1995), 662, 666 Czarnecki, D. B., Blinn, D. W. (1978), 584, 589, 604, 628, 632, 650, 651 Czarnecki, D. B., Blinn, D. W. (1979), 665, 666 Czarnecki, D. B., Reinke, D. C. (1981), 650, 651 D Dahm, C. N., Grimm, N. B., Marmonier, P. Vallett, H. M., Vervier, P. (1998), 29, 48 Daily, F. K. (1952), 314, 350 Daily, W. A. (1942), 60, 80, 82, 95, 110–111 Daily, W. A. (1943), 139, 192 Daily, W. A. (1946), 60, 111 Damann, K. E. (1945), 461, 466 Danforth, W. F., Ginsburg, W. (1980), 605, 632 Daugbjerg, N., Andersen, R. A. (1997), 424, 429, 466 Daugbjerg, N., Andersen, R. A. (1997a), 473, 504 Daugbjerg, N., Andersen, R. A. (1997b), 473, 504 Davey, M. C. (1987), 565, 589 Davis, B. M. (1904), 432, 466 Davis, C. C. (1964), 20–21, 48, 806, 828 Davis, J. S., Rands, D. G., Hein, M. K. (1989), 605, 632 Davis, L. W., Hoffmann, J. P. Cook, P. W. (1990a), 44, 48 Davis, L. W., Hoffmann, J. P. Cook, P. W. (1990b), 48 Davis, R. B., Anderson, D. S., Norton, S. A., Ford, J., Sweets, P. R., Kahl, J. S. (1994), 617, 632 Davis-Colley, R. J., Vant, W. N. (1988), 779, 799 Dawson, N. S., Dunlap, J. R., Walne, P. L. (1988), 412, 417 Dawson, N. S., Walne, P. L. (1991), 413, 417 Dawson, N. S., Walne, P. L. (1994), 385, 391, 417 Dawson, P. A. (1972), 655, 659, 666 Dawson, P. A. (1973a), 655, 664, 666 Dawson, P. A. (1973b), 655, 664, 666 Dawson, P. A. (1973c), 655, 659, 666 Dawson, P. A. (1974), 655, 666 Dayner, D. M., Johansen, J. R. (1991), 44, 48 de Noyelles, F., Knoechel, R., Reinke, D., Treanor, D., Altenhoifen, C. (1980), 22, 48 de Ruyter van Steveninck, E. D., Admirall, W., Breebaart, L., Tubbingm G, M. J., van Zanten, B. (1992), 37, 48 De Toni, G. (1936), 80, 111 Deane, J. A., Hill, D. R. A., McFadden, G. I. (1998), 734, 752 Dearing, J. A., Håkansson, H., Liedberg-Jönsson, B., Persson, A., Skansjö, S., Windholm, D., El-Dahousy, F. (1987), 799 Deason, T. , Silva, P. C., Watanabe, S., Floyd, G. L. (1991), 256, 308 Deason, T. (1959), 303, 308 Deason, T. R. (1969), 326, 350

855

Deason, T. R. (1971a), 432, 466 Deason, T. R. (1971b), 432, 466 Deason, T. R., Bold, H. (1960), 326, 349–350 Deflandre, G. (1926), 416–417 Deflandre, G. (1930), 414, 417 Della Greca, M., Monaco, P., Pollio, A., Previtera, L. (1992), 821, 828 DeLuca, P., Gambardella, R., Merola, A. (1979), 206, 208, 222 DeLuca, P., Morretti, A. (1983), 41, 48 Demayo, A., Taylor, M. C., Taylor, K. W. (1982), 825, 828 DeMelo, R., France, R., McQueen, D. J. (1992), 819, 828 DeMott, W. R., Zhang, Q.-X., Carmichael, W. W. (1991), 808, 828 Denicola, D. M. (1986), 782, 799 Deniseger, J., Austen, A., Roch, M., Clark, M. J. R. (1986), 587, 589 DeNoyelles, F., O’Brien, W. J. (1978), 537, 553 Denys, L. (1991), 791, 799 Descy, J.-P. (1979), 787, 799 Descy, J.-P., Gosselain, V. (1994), 32, 48 Descy, J.-P., Gosselain, V., Evrard, F. (1994), 32, 37, 48 Descy, J.-P., Servais, P. Smitz, J. S., Billen, G., Everbecq, E. (1987), 37–38, 48 Descy, J.-P., Willems, C. (1991), 587, 589 Desikachary, T. V. (1959), 59, 64, 92, 111, 119–121, 161, 164, 174, 176, 181, 192 Desjardins, P. R., Olsen, G. B. (1983), 819, 828 deVecchi, L., Grilli-Caiola, M. (1986), 118, 192 DeYoe, H. R., Lowe, R. L., Marks, J. C. (1992), 670, 673–674, 683 Diaz, M. M., Lorenzo, L. E. (1990), 513, 519 Diaz, M. M., Pedrozo, F. L., Temporetti, P. F. (1998), 587, 589 DiCastri, E., Younez, T. (1994), 60, 111 Dillard, G. E. , Crider, S. B. (1970), 496, 504 Dillard, G. E. (1967), 454, 463, 466 Dillard, G. E. (1970), 483, 495, 504 Dillard, G. E. (1989), 228, 230–231, 234–236, 238, 241–242, 244–245, 247, 249, 254–255, 270, 272, 280, 284, 288, 291, 298, 307–308, 313, 348, 350 Dillard, G. E. (1999), 253, 258, 307–308, 414, 417, 463, 466 Dillard, G. E. (2000), 388, 400, 410–417 Dillard, G. E., Moore, S. P., Garret, L. S. (1976), 433, 449, 461, 466 Dillon, P. J., Nicholls, K. H., Locke, B. A., de Grosbois, E., Yan, N. D. (1988), 812, 828 Dillon, P. J., Rigler, F. H. (1974), 812, 828 Dimitrov, M. (1984), 820, 828 Dionigi, C. P.,Champagne, E. T. (1995), 825, 828 Diwald, K. (1938), 694, 711 Dixit, A. S., Dixit, S. S., Smol, J. P. (1992a), 775, 781, 799 Dixit, S. S., Cumming, B. F., Kingston, J. C., Smol, J. P., Birks, H. J. B., Uutala, A. J., Charles, D. F., Camburn, K. (1993), 792, 799 Dixit, S. S., Dixit, A. S., Evans, R. D. (1988), 536–537, 553 Dixit, S. S., Dixit, A. S., Smol, J. P. (1989), 535, 553 Dixit, S. S., Smol, J. P. (1994), 775, 781, 799 Dixit, S. S., Smol, J. P., Anderson, D. S., Davies, R. B. (1990), 536, 553 Dixit, S. S., Smol, J. P., Charles, D. F., Hughes, R. M., Paulsen, S. G., Collins, G. B. (1999), 604, 632, 775, 781, 792, 799 Dixit, S. S., Smol, J. P., Kingston, J. C., Charles, D. F. (1992), 604, 632 Dixit, S. S., Smol, J. P., Kingston, J. C., Charles, D. F. (1992b), 793, 799 D’Lacoste, V., Ganesan, E. K. (1987), 202, 222 Dodd, J. D., Stoermer, E. F. (1962), 623, 632 Dodd, J. J. (1987), 561, 589, 604–605, 632, 682–683 Dodd, W. K., Gudder, D. A. (1992), 783, 799 Doddema, H., van der Veer, J. (1983), 490, 504

856

Author Index

Dodds, W. K. (1989), 36, 48, 180, 192 Dodds, W. K., Gudder, D. A. (1992), 35, 48, 340, 350, 810, 812, 828 Dodds, W. K., Gudder, D. A., Mollenhauer, D. (1995), 27, 48, 180, 192 Dodds, W. K., Jones, J. R., Welch, E. B. (1998), 31, 38, 48, 776, 782, 785, 799 Dodge, J. D. (1965), 686, 711 Dodge, J. D. (1966), 686, 711 Dodge, J. D. (1969), 690, 711, 741, 747, 752 Dodge, J. D. (1972), 690, 711 Dodge, J. D. (1975), 690, 711 Dodge, J. D. (1983), 697, 711 Dodge, J. D., Bibby, B. T. (1973), 709, 711 Dodge, J. D., Crawford, R. M. (1970), 690, 711 Dodge, J. D., Hermes, H. B. (1981), 690, 711 Doemel, W. N., Brock, T. D. (1971), 197, 206, 208, 222 Doers, M. P., Parker, D. L. (1988), 111 Dokkulil, M., Mayer, J. (1996), 174, 192 Domozych, D. S. (1989), 235, 249 Domozych, D. S., Nimmons, T. T. (1992), 235, 249 Dop, A. J. (1978), 481, 496–497, 504 Dop, A. J. (1979), 757–758, 760, 763, 766–767, 771–772 Dop, A. J. (1980), 484, 500–502, 505 Dop, A. J., Vroman, M. (1976), 763, 772 Douglas, B. (1958), 36, 48, 767, 772 Douglas, M. S. V., Smol, J. P. (1993), 604, 632 Douglas, M. S. V., Smol, J. P. (1995), 24, 26, 48, 490, 505 Douglas, M. S. V., Smol, J. P., Blake, W., Jr. (1994), 789, 799 Douglas, S. E., Murphy, C. A., Spencer, D. F., Gray, M. W. (1991), 735–737, 752 Dow, C. S., Swoboda, U. K. (2000), 12, 32, 48, 121, 192 Downes, B. J., Lake, P. S., Schreiber, E. S. G., Glaister, A. (1998), 36, 48 Doyle, R. D., Smart, R. M. (1998), 821, 828 Dragos, N., Péterfo, L. S., Popescu, C. (1997), 400, 417 Drew, K. M. (1935), 201, 222 Drews, G. (1959), 119, 192 Drews, G. (1973), 118, 192 Drews, G., Prauser, H., Uhlmann, O. (1961), 79, 111 Drews, G., Weckesser, J. (1982), 61, 111 Drouet, F. (1933), 463, 466 Drouet, F. (1934), 154–155, 193 Drouet, F. (1936a), 60, 111 Drouet, F. (1936b), 60, 111 Drouet, F. (1937), 131–132, 136, 193 Drouet, F. (1938), 60, 111, 148, 193 Drouet, F. (1942), 60, 78, 82, 94, 111, 131, 140, 145, 155, 193 Drouet, F. (1943a), 144, 149–151, 193 Drouet, F. (1943b), 149, 193 Drouet, F. (1954), 461, 463, 466 Drouet, F. (1968), 118, 193 Drouet, F. (1973), 118, 193 Drouet, F. (1981a), 118, 193 Drouet, F. (1981b), 118, 193 Drouet, F., Cohen, A. (1935), 428, 463, 466 Drouet, F., Daily, W. A. (1948), 111 Drouet, F., Daily, W. A. (1952), 111 Drouet, F., Daily, W. A. (1956), 80–81, 88, 96, 101, 111, 118, 193 Drum, R. W., Pankratz, H. S. (1964), 565, 589 Drum, R. W., Pankratz, S. (1965), 670, 673, 683 Duff, K. E., Smol, J. P. (1995), 536–537, 553 Duff, K. E., Zeeb, B. A., Smol, J. P. (1995), 485, 505, 533, 553 Dujardin, F. (1841), 413, 417 Dunlap, J. R., Walne, P. L. (1985), 417

Dunlap, J. R., Walne, P. L. (1987), 406, 417 Dunlap, J. R., Walne, P. L., Bentley. J. (1983), 414, 417 Dunlap, J. R., Walne, P. L., Kivic, P. A. (1986), 403, 414, 417 Dunlap, J.R., Walne, P. L., Preisig, H. R. (1987), 485, 505 Durrell, L. W., Norton, C. (1960), 433–434, 454, 459, 466 Dürrschmidt, M. (1980), 534, 536, 553 Dürrschmidt, M. (1982), 526, 536, 538, 553 Dürrschmidt, M., Croome, R. (1985), 536, 538, 553 Duthie, H. C. (1989), 607, 632 Duthie, H. C., Ostrofsky, M. L. (1975), 411, 417 Duthie, H. C., Ostrofsky, M. L. (1978), 491, 505 Duthie, H. C., Ostrofsky, M. L., Brown, D. J. (1976), 429, 451, 466 Duthie, H. C., Socha, R. (1976), 72, 80–82. 87–88, 92, 95, 111, 118, 132, 135, 141, 155, 158, 161, 164, 166, 169, 171, 174, 177, 180, 184, 193, 230, 232, 234–235, 238–239, 241–242, 245, 247, 249, 411–413, 415, 417, 425, 428–429, 432, 434, 447, 449, 451, 453–454, 463, 466, 491, 505 Duthie, H. C., Sreenivasa, M. R. (1972), 628, 632 Duthie, H., Socha, R. (1976), 698, 711 Dwarte, D., Vesk, M. (1982), 735, 752 Dwarte, D., Vesk, M. (1983), 735, 741, 752 E Eardley-Wilmot, V. L. (1928), 561, 589 Eaton, G. L. (1980), 690, 697, 711 Eaton, J. W., Moss, B. (1996), 27, 48 Echevarria, F., Rodriguez, J. (1994), 700, 711 Eddy, S. (1934), 36, 48 Edgren, R. A., Egren, M. K., Tiffany, L. H. (1953), 314, 350 Edlund, M. B. (1992), 569, 589 Edlund, M. B. (1998), 568, 589 Edlund, M. B., Stoermer, E. F. (1993), 567, 572, 587, 589 Edlund, M. B., Stoermer, E. F. (1997), 560, 568, 589, 638, 652 Edlund, M. B., Stoermer, E. F. (1999), 661, 663, 666 Edlund, M. B., Stoermer, E. F., Taylor, C. M. (1996), 567, 573–574, 589 Edlund, M. B., Taylor, C. M. Schelske, C. L., Stoermer, E. F. (2000), 587, 589 Edmondson, W. T. (1959), 639, 652 Edmondson, W. T. (1963), 12, 16, 48 Edmondson, W. T. (1965), 739, 752 Edmondson, W. T. (1969), 405, 417 Edmondson, W. T. (1977), 23, 48 Edmondson, W. T. (1994), 813, 828 Edmondson, W. T., Lehman, J. T. (1981), 23, 48, 813, 828 Edvardsen, B, Eikrem, W., Green, J. C., Andersen, R. A., Moon-Van der Staay, S. Y., Medlin, L. K. (2000), 511, 515, 519 Edwards, P. (1975), 342, 349–350 Effler, S. W., Litten, S., Field, S. D., Tong-Ngork, T., Hale, F., Meyer, M., Quirk, M. (1980), 822, 825, 828 Effler, S. W., Owens, E. M. (1996), 42, 48 Eguchi, M., Oketa, T., Miyamoto, N. Maeda, H., Kawai, A. (1996), 65, 111 Ehara, M., Hayashi-Ishimaru, Y., Inagaki, Y., Ohama, Y. (1997), 447, 459, 466 Ehrenberg, C. G. (1830), 384, 417 Ehrenberg, C. G. (1831), 384, 411, 417 Ehrenberg, C. G. (1833), 384, 417 Ehrenberg, C. G. (1838), 384, 418, 471, 505 Ehrenberg, C. G. (1841), 595, 632 Ehrenberg, C. G. (1843), 561, 589, 649–650, 652 Ehrenberg, C. G. (1854), 561, 589 Ehrenberg, C. G. (1870), 561, 589 Ekenstam, D., Bozniak, E. G., Sommerfeld, M. R. (1996), 763–764, 772

Author Index Elakovich, S. D., Wooten, J. W. (1989), 821, 828 Elder, J. F., Horne, A. J. (1978), 822, 824, 828 Elenkrn, A. A. (1936, 1938, 1949), 110, 111 Ellis-Adams, A. C. (1983), 478–479, 505 Elmore, C. J. (1992), 561, 589 Eloranta, P. (1989), 505, 534–535, 537, 553 Eloranta, P. (1995), 536, 553 Elser, J. J., MacKay, N. A. (1989), 818, 828 Elser, J. J., Marzolf, E. R., Goldman, C. R. (1990), 812–813, 828 Elster, J., Svoboda, J., Komárek, J., Marvan, P. (1997), 158, 193 Eminson, D., Moss, B. (1980), 24–25, 48 Engstrom, D. R., Swaitn, E. B., Kingston, J. C. (1985), 605, 632 Entwisle, T. J. (1987), 465–466 Entwisle, T. J. (1988a), 465–466 Entwisle, T. J. (1988b), 465–466 Entwisle, T. J. (1989), 33, 48 Entwisle, T. J., Sonneman, J. A., Lewis, S. H. (1998), 226–227, 249 Entwisle, T. J., Sonneman, J., Lewis, S. J. (1997), 768, 772 Entwistle, T. J., Andersen, R. A. (1990), 505 Environmental Protection Agency (1990), 806, 828 Environmental Protection Agency (1998), 814, 828 Erata, M., Chihara, M. (1989), 741, 746, 752 Erikson, R., Pum, M., Vammen, K. Cruz, A., Ruiz, M., Zamora, H. (1997), 18, 48 Eschbach, S., Wolters, J., Sitte, P. (1991), 752 Eskew, D. L., Ting, I. O. (1978), 63, 111 Esser, S. C., Valkenburg, S. D. (1977), 485, 505 Etnier, C., Guterstam, B. (1997), 813, 828 Ettl, H. (1978), 426, 429, 431–432, 434, 440, 442, 444–445, 448, 452, 454–460, 462, 464–466 Ettl, H. (1980), 235, 249 Ettl, H. (1983), 225–228, 230–232, 234–236, 238–239, 241, 243, 245, 247–249 Ettl, H., Gärtner, G. (1995), 254, 268–269, 271–273, 275, 277–279, 283, 287, 290, 292, 297, 300, 304, 306–308, 322, 348, 350 Ettl, H., Moestrup, O. (1980), 739, 752 Ettl, H., Popovsky´, J. (1986), 391, 418 Evans, J. C., Arts, M. T., Robarts, R. D. (1996), 42–43, 48 Evans, J. C., Prepas, E. E. (1996), 42–43, 48 Everitt, D. T., Burkholder, J. M. (1991), 203, 216, 222 Evitt, R. W. (1985), 687, 711 F Faber, Jr., W. W., Preisig, H. R. (1994), 511, 519 Fabry, S., Köhler, A., Coleman, A. W. (1999), 227, 243, 249 Fahnenstiel, G, L., Glime, J. M. (1983), 567–568, 589 Fahnenstiel, G. L., Sicko-Goad, L., Scavia, D., Stoermer, E. F. (1986), 65, 111 Fairchild, E. C., Wilson, D. L. (1967), 63, 111 Fairchild, G. W., Lowe, R. L. (1984), 673–674, 683 Fairchild, G. W., Lowe, R. L., Richardson, W. B. (1985), 605, 632, 674, 683, 786, 799 Fairchild, G. W., Sherman, J. W., Acker, F. W. (1989), 26, 48 Falconer, I. R. (1998), 121, 193 Fallon, R. D., Brock, T. D. (1981), 111 Faridi, M. A. F. (1962), 339, 348, 350 Farmer, J. N. (1980), 386–387, 418 Fassett, N. C. (1957), 811, 828 Faust, M. A. (1974), 732, 741, 752 Faust, M. A., Gantt, E. (1973), 735, 752 Fay, P. (1983), 61, 111, 121, 193 Fay, P., Van Baalen, C. (1987), 63, 111, 117, 121, 193 Feldmann, J. (1958), 111 Felip, M., Sattler, B., Psenner, R., Catalan, J. (1995), 44, 48, 487, 505 Feminella, J. W., Hawkins, C. P. (1995), 35, 48

857

Feminella, J. W., Power, M. E., Resh, V. H. (1989), 35, 48 Fensome, R. A., MacRae, R. A., Moldowan, J. M., Taylor, F. J. R., Williams. G. J. (1996), 687, 711 Fensome, R. A., Taylor, F. J. R., Norris, G., Sarjeant, W. A. S., Wharton, D. L., Williams. G. L. (1993), 687, 697, 711 Ferguson, A. J. D., Thompson, J. M., Reynolds, C. S. (1982), 739, 752 Fields, S. D., Rhodes, R. G. (1991), 686, 690, 711 Findlay, D. L. (1978), 537, 553 Findlay, D. L., Kasian, S. E. M. (1987), 513, 519 Findlay, D. L., Kasian, S. E. M. (1996), 488, 505 Findlay, D. L., Kling, H. J. (1979), 147, 193 Findlay, S., Likens, G. E., Hedin, L., Fisher, S. G., McDowell, W. H. (1997), 31, 48 Fitzgerald, G. P. (1959), 809, 828 Fitzgerald, G. P. (1971), 822, 828 Fjerdingstad, E. (1950), 775, 799 Flechtner, V. R., Boyer, S. L. Johansen, J. R., DeNoble, M. L. (2002), 191, 193 Fleming, R. F. (1989), 254, 308 Fling, E. M. (1939), 463, 466 Flint, L. H. (1955), 34, 48 Flint, L. J. (1970), 197, 222 Floener, L. Bothe, H. (1980), 670, 673, 681, 683 Flores, E., Wolk, C. P. (1986), 821, 828 Florin, M.-B. (1970), 604, 632 Flower, R. J. (1989), 601, 617, 632 Flower, R. J., Battarbee, R. W. (1985a), 601, 632 Flower, R. J., Battarbee, R. W. (1985b), 600, 632 Flower, R. J., Häkasson, H. (1994), 578, 589 Flower, R. J., Jones, V. J. (1989), 627, 632 Flower, R. J., Jones, V. J., Round, F. E. (1996), 598–599, 632 Flower, R. J., Ozornina, S. P., Kuzmina, A., Round, F. E. (1998), 585, 589 Floyd, G. K., Watanage, S., Deacon, T. R. (1993), 257, 308 Foerster, J. W., Schlichting, H. E. J. (1965), 605, 632 Foged, N. (1953), 561, 589 Foged, N. (1955), 561, 589 Foged, N. (1971), 604, 628, 632, 649, 652 Foged, N. (1973), 561, 589 Foged, N. (1981), 561, 589, 649, 652 Fogg, G. E. (1949), 119, 193 Fogg, G. E. (1986), 65, 111 Fogg, G. E., Stewart, W. D. P., Fay, P., Walsby, A. E. (1973), 59, 63, 111, 117, 120–121, 193 Forest, H. S. (1954), 698, 708, 711 Forest, H. S. (1956), 348, 350 Forest, H. S., Weston, C. R. (1966), 63, 111 Forsberg, C. (1965), 811, 828 Fott, B. (1949), 245, 249 Fott, B. (1959), 485, 505 Fott, B. (1963), 238, 248–249 Fott, B. (1967), 245, 249 Fott, B. (1968), 465–466 Fott, B. (1971), 427, 466 Fourreau, J. (1868), 245, 249 Fourtanier, E., Kociolek, J. P. (1999), 613, 627, 632 Fowler, M. C., Robson, T. O. (1978), 820–821, 829 Francke, J. A., Coesel, P. F. M. (1985), 752 Francke, J. A., Hillebrand, H. (1980), 822, 829 Frank, P. A. (1972), 825, 829 Frémy, P. (1930a), 64, 111, 145, 165, 174, 181–183, 193 Frémy, P. (1930b), 111, 145, 165, 174, 181–183, 193 Frémy, P. (1933), 66, 111 Frémy, P. (1949), 72, 111

858

Author Index

French, F. W., Hargraves, P. E. (1980), 567, 589 Fresnel, J. (1994), 512, 519 Frey, D. G. (1963), 12, 48 Frey, L. C., Stoermer, E.F. (1990), 701, 711 Friedl, T. (1995), 257, 308 Friedl, T. (1997), 225, 228, 231, 249, 257, 308 Friedl, T. (1998), 313, 350 Friedl, T., Zeltner, C. (1994), 257, 308 Friedmann, E. I. (1955), 186–187, 193 Friedmann, E. I. (1971), 60, 66, 111 Friedmann, E. I. (1979), 186–187, 193 Friedmann, E. I. (1980), 111 Friedmann, E. I., Ocampo, R. (1985), 66, 112 Friedmann, E. I., Ocampo-Friedmann, R. (1984), 66, 112 Fritsch, F. E. (1922), 45, 49 Fritsch, F. E. (1929), 33, 49, 205, 222, 763, 765, 772 Fritsch, F. E. (1935), 356, 361–364, 380 Fritsch, F. W. (1945), 385, 411, 414–415, 418 Fritz, S. C. (1990), 585, 589, 776, 787, 799 Fritz, S. C., Battarbee, R. W. (1986), 585, 589 Fritz, S. C., Juggins, S., Battarbee, R. W., Engstrom, D. R. (1991), 585, 589, 799 Fritz, S. C., Juggins, S., Batterbee, R. W. (1993), 42–43, 49 Fritz, S. C., Kingston, J. C., Engstrom, D. R. (1993), 604, 632 Frodge, J. D., Thomas, G. L., Pauley, G. B. (1991), 815, 829 Fromentel, E. de (1874), 502, 505 Frost, T. M., Graham, L. E., Elias, J. E., Haase, M. J., Kretchmer, D. W., Kranzfelder, J. A. (1997), 45, 49 Frost, T. M., Williamson, C. E. (1980), 45, 49 Fuller, R. L., Roelofs, J. L., Fry, T. J. (1986), 31, 35, 49 Fulton, A. B. (1978), 226, 249 Fulton, R. S., III (1988), 819, 829 Fulton, R. S., Paerl, H. W. (1987), 64, 112, 808, 829 G Gabor, T. S., Murkin, H. R., Stainton, M. P., Boughen, J. A., Titman, R. D. (1994), 39, 49 Gaines, G., Elbrachter, M. (1987), 701, 711 Galat, D. L. Verdin, J. P., Sims, L. L. (1990), 177, 193 Gálvez, J. A., Niell, F. X., Lucena, J. (1988), 700, 711 Gantt, E. (1971), 732, 741, 747, 752 Gantt, E. (1979), 735, 752 Gantt, E. (1980), 735, 752 Gantt, E., Edwards, M. R., Provasoli, L. (1971), 735, 752 Gantt, E., Scott, J. Lipschultz, C. (1986), 200, 222 Garbary, D. J., Hansen, G. I., Scagel, R. F. (1980), 201, 222 Garcia-Pichel, F., Belnap, J. (1996), 63, 112 Garcia-Pichel, F., Castenholz, R. W. (1991), 61, 112 Garcia Reina, G. (1997), 68, 112 Gardner, N. L. (1906), 112 Gardner, N. L. (1918), 104–105, 107–108, 110, 112 Gardner, N. L. (1927), 60, 66–67, 72, 75, 78, 80, 82, 89, 92–93, 95–97, 99, 102–104, 106–108, 110, 112, 118, 126, 130, 132, 134, 138–139, 143, 145, 148, 150–153, 157, 159– 162, 165, 176, 178, 181, 183, 193 Garric, R. K. (1965), 81, 112 Garrison, P.J., Knauer, D. R. (1984), 814, 829 Gartner, G. (1992), 311, 350 Gartner, G., Ingolic, E. (1989), 315, 348, 350 Garwood, P. E. (1982), 314, 350 Gaufin, A. R., Prescott, G. W., Tibbs, J. F. (1976), 434, 443, 447, 449, 459, 466 Gaul, U., Geissler, U., Henderson, M., Mahoney, R., Reimer, C. W. (1993), 597, 628, 632, 638, 652, 655, 666 Gayral, P., Haas, C., Lepailleur, H. (1972), 485, 505

Geibler, U. (1983), 759, 768, 772 Geissler, U. (1982), 569, 589 Geitler, L. (1925), 60, 77, 91, 101, 112 Geitler, L. (1927), 664, 666 Geitler, L. (1928), 765, 768, 772 Geitler, L. (1932), 59–61, 65, 79, 82, 91, 93, 99–100, 106, 108, 112, 118, 120, 127, 143–145, 147, 151, 154, 158–160, 163, 166, 174–178, 181, 185, 188. 193, 197, 206, 222, 655, 666, 763–765, 772 Geitler, L. (1942), 60–61, 80, 112, 118, 158, 193 Geitler, L. (1951a), 665, 666 Geitler, L. (1951b), 665, 666 Geitler, L. (1951c), 665, 666 Geitler, L. (1956), 664, 666 Geitler, L. (1960), 98, 112, 118, 158, 193 Geitler, L. (1967), 664, 666 Geitler, L. (1973), 560, 589, 638, 652 Geitler, L. (1973a), 655, 666 Geitler, L. (1973b), 665, 666 Geitler, L. (1973c), 665, 666 Geitler, L. (1975), 662, 667 Geitler, L. (1977), 623, 627, 632, 670, 673, 676, 683 Geitler, L. (1979), 623, 627, 632 Geitler, L. (1980), 623, 627, 632 Geitler, L. (1981), 659, 667 Geitler, L. (1982), 118, 193 Geitler, L., Mack, B. (1953), 664, 667 Geitler, L., Ruttner, F. (1935), 67, 82, 104–105, 112 Gektidis, M., Golubic, S. (1996), 107, 112 Gelin, F., Boogers, I., Noordelos, A. A. M., Sinnighe Damsté, J. S., Riegman, R., de Leeuw, J. W. (1997), 254, 308 Gelin, F., Boogers, I., Noordelos, A. A. M., Sinnighe Damsté, J. S., Riegman, R., de Leeuw, J. W. (1997), 312, 350 Genkal, S. I., Håkansson. (1990), 568, 589 Genkal, S. I., Kiss, K. T. (1998), 568, 583–584, 589 Genkal, S. I., Makarova, A., Goncharov, A. A. (1998), 583, 589 Gensemer, R. W. (1991), 786, 799 Gensemer, R. W. (1991a), 607, 632 Gensemer, R. W. (1991b), 607, 632 Genter, R. B. (1996), 782, 799 Gerber, S., Häder, D.-P. (1993), 407, 418 Gerloff, J. (1967), 771–772 Germain, H. (1981), 628, 632 Gerrath, J. F. (1970), 354, 374, 380 Gerrath, J. F. (1993), 363–365, 380 Gersonde, R., Harwood, D. M. (1990), 570, 589 Ghosh, M. Gaur, J. P. (1990), 785, 799 Gibbs, G. W. (1973), 811, 829 Gibbs, S. P. (1978), 385, 402, 418 Gibbs, S. P. (1981), 385, 402, 418, 472, 505 Gibson, C. E. (1975), 129, 193 Gibson, C. E., Smith, R. V. (1982), 64, 112 Gibson, K. N., Smol, J. P., Ford, J. (1987), 535, 553 Gibson, M. T., Welch, I. M., Barrett, P. R. F., Ridge, I. (1990), 822, 829 Gibson, M. T., Whitton, B. A. (1987), 204, 222, 758, 772 Gillot, M. (1990), 715, 734, 736, 752 Gillot, M., Gibbs, S. P. (1980), 734–735, 752 Gillot, M., Gibbs, S. P. (1983), 734–735, 752 Glazer, A. N., Appell, G. S. (1977), 735, 737, 752 Glazer, A. N., Wedemeyer, G. J. (1995), 737, 752 Gledhill, M., Nimmo, M., Hill, S. J., Brown, M. T. (1997), 823, 829 Glew, J. R. (1991), 792, 799 Glew, J. R. (1998), 780, 799 Glime, J. M., Wetzel, R. G., Kennedy, B. J. (1982), 40, 49

Author Index Goff, L. J., Stein, J. R. (1978), 45, 49, 291, 308 Gojdics, M. (1953), 412, 415, 418 Gold-Morgan, M., Montejano, G., Komárek, J. (1994), 60, 106–107, 112, 151, 193 Gold-Morgan, M., Montejano, G., Komárek, J. (1996), 98, 100–102, 112 Golden, J. W., Robinson, S. J., Haselkorn, R. (1985), 119, 193 Goldman, C. R. (1988), 19, 49 Goldsborough, L. G., Robinson, G. G. C. (1996), 38–40, 49, 66, 112, 669, 683 Goldstein, A. K., Manzi, J. J. (1976), 425, 427, 433, 449, 463, 466 Goldstein, M. (1964), 226, 239, 241, 247–249 Golecki, J. R., Drews, G. (1974), 119, 193 Golubic, S. (1965), 60, 92, 112 Golubic, S. (1967a), 60, 63, 65, 92, 112, 121, 166, 193 Golubic, S. (1967b), 60, 92, 112, 121, 166, 193 Golubic, S. (1980), 60, 66, 112, 120–121, 193 Golubic, S., Focke, J. W. (1978), 193 Golubic, S., Friedmann, E. I., Schneider, J. (1981), 66–67, 112, 121, 193 Golubic, S., Hernandez-Mariné, M., Hoffmann, L. (1996), 119, 193 Golubic, S., Perkins, R. D., Lukas, K. J. (1975), 67, 107, 112, 121, 193 Golubic, S., Yun, Z., Campbell, S. E. (1985), 62, 112 Gomont, M. (1882), 142–144, 148–149, 193 Gomont, M. (1892), 59, 112 Gomont, M. (1896), 757, 772 Good, R. H., Chapman, R. L. (1978), 312, 350 Goodrich, S. G. (1859), 385, 418 Goodwin, T. W. (1974), 20, 49 Goodwin (1974), 757, 772 Gorham, E., Eisenreich, S. J., Ford, J., Santelmann, M. V. (1985), 40, 49 Gorham, P. R., Carmichael, W. W. (1988), 92, 112, 806, 829 Gorham, P. R., McLachlan, J., Hammer, U. T., Kim, W. K. (1964), 64, 112 Gosse, P. H. (1859), 383, 385, 418 Gosse, P. H. (1896), 385, 418 Gosselain, V., Descy, J.-P., Everbecq, E. (1994), 38, 49 Gosselain, V., Viroux, L., Descy, J.-P. (1998), 38, 49 Gough, S. B., Woelkerling, W. J. (1976), 24, 49 Graham, J. M., Kranzfelder, J. A., Auer, M. T. (1985), 26, 49 Graham, L. E., Graham, J. M., Wujek, D. E. (1993), 526, 528, 531, 533–534, 537, 539, 551, 553 Graham, L. E., Wilcox, L. W. (2000), 1, 5, 9, 254–255, 308, 312–313, 350, 384, 387, 399, 418, 423–424, 427, 432, 465, 467, 757–759, 772 Graham, M. D., Vinebrooke, R. D. (1998), 24, 26, 49 Graham, T. P., McCoy, J. J. (1974), 406, 418 Grain, J., Mignot, J. P., Puytorac, P. (1988), 734, 752 Green, J. (1953), 411, 418 Green, J. C., Leadbeater, B. S. C. (1994), 512, 519 Green, J. C., Piennar, R. N. (1977), 512, 519 Green, R. H. (1979), 778, 789–790, 799 Greenberg, A. E. (1964), 36–37, 49 Greene, J. C., Miller, W. E., Shiroyama, T., Soltero, R. A., Putnam, K. (1976), 785, 799 Greenwood, A. D. (1959), 432, 467 Greenwood, A. D., Griffiths, H. B., Santore, U. S. (1977), 741, 752 Greenwood, J., Clason, T., Lowe, R. L., Belanger, S. E. (1999), 674, 683 Gretz, M. R., Sommerfeld, M. R., Wujek, D. E. (1979), 552–553 Gretz, M. R., Sommerfield, M. R., Athey, P. V. Aronson, J. M. (1991), 200, 222 Gretz, M. R., Wujek, D. E. Sommerfeld, M. R. (1983), 552–553

859

Grim, J. N., Staehelin, L. A. (1984), 732, 734–735, 741, 746, 752 Gromov, B. V., Mamkayeva, K. A., Bobina, V. D. (1988), 489, 505 Gross, E. M., Meyer, H., Schilling, G. (1996), 821, 829 Gross, F., Zeuthen, E. (1948), 567, 589 Grote, M. (1977), 364, 380 Gruendling, G. K. (1971), 27, 49 Grzebyk, D., Sako, Y., Berland, B. (1998), 709, 711 Gualtieri, P. (1993), 402, 418 Gualtieri, P., Pelosi, P., Passarelli, V., Barsanti, L. (1992), 402, 418 Guglielmi, G., Cohen-Bazire, G. (1982a), 118, 193 Guglielmi, G., Cohen-Bazire, G. (1982b), 118, 193 Guillard, R. R. L. (1975), 739, 752 Guillard, R. R. L., Lorenzen, C. J. (1972), 424, 467 Gulati, R. D. (1990), 818, 829 Gunn, G. J., Raferty, A. G., Rafferty, G. C., Cockburn, N., Edwards, C., Beatie, K. A., Codd, G. A. (1992), 809, 829 Guo, M., Harrison, P. J., Taylor, F. R. J. (1996), 512, 519 Gurnee, J. (1994), 44, 49 Gutowski, A. (1989), 536, 538, 553 Gutowski, A. (1996), 538–539, 553 Gutowski, A. (1997), 536, 553 H Haberyan, K. A., Umana, G., Collado, C., Horn, S. P. (1995), 13, 49, 234–235, 249, 412, 415, 418, 698, 711 Hadas, O., Malinsky-Rushansky, N., Pinkas, R., Cappenberg, T. E. (1998), 20, 49 Häder, D.-P. (1974), 119, 193 Häder, D.-P. (1987), 407, 418 Haffen, L. M., McCann, M. T. (1975), 808, 829 Häkansson, H., Jones, V. J. (1994), 561, 589 Häkansson, H., Kling, H. (1989), 565, 589 Häkansson, H., Kling, H. (1990), 561, 565, 581, 585, 589 Häkansson, H., Kling, H. (1994), 561, 565, 590 Häkansson, H., Mahood, A. (1993), 586, 590 Häkansson, H., Stoermer, E. F. (1984), 567, 569, 586, 590 Häkansson, H., Stoermer, E. F. (1987), 585–586, 590 Halfen, L. N. (1979), 119, 193 Halingse, M. W., Phlips, E. J. (1996), 823, 829 Hall, R. I., Smol, J. P. (1992), 799 Hall, R. I., Smol, J. P. (1996), 781, 799 Hallegraeff, G. M. (1993), 799 Hallegraeff, G. M., Anderson, D. M., Cembella, A. D. (1995), 685, 711 Hällfors, G., Hällfors, S. (1988), 486, 505 Hällfors, G., Munsterhjelm, R. (1982), 98, 112 Hallick, R. B., Hong, L., Drager, R. G., Favreau, M. R. Monfort, A., Orsat, B., Spielmann, A., Stutz, E. (1993), 387, 418 Hambrook, J. A., Sheath, R. G. (1987), 205, 222 Hambrook, J. A., Sheath, R. G. (1991), 202–204, 222, 763, 772 Hamel, G. (1931–1939), 759, 772 Hamilton, P. B., Douglas, M. S. V., Fritz, S. C., Pienitz, R., Smol, J. P., Wolfe, A. P. (1994), 561, 590, 638, 649, 652 Hamilton, P. B., Edlund, S. A. (1994), 45, 49 Hamilton, P. B., McNeely, R., Poulin, M. (1996), 638, 649, 652 Hamilton, P. B., Poulin, M., Charles, D. F., Angell, M. (1992), 604, 632 Hamilton, P. B., Poulin, M., Taylor, M. C. (1990), 638, 649, 652 Hamilton, P. B., Poulin, M., Walker, D. (1995), 638, 649, 652 Hammer, U. T. (1981), 43, 49 Hammer, U. T. (1986), 42, 49 Hammer, U. T., Shamess, J., Haynes, R. C. (1983), 42, 49 Hanagata, N. (1998), 257, 308 Hann, B. J. (1991), 39, 49 Hanninen, O., Ruuskanen, J., Oksanen, J. (1993), 45, 49

860

Author Index

Hansen, G. W., Oliver, F. E., Otto, N. E. (1984), 829 Hansen, L. R., Kristiansen, J., Rasmussen, J. V. (1994), 512–513, 519 Hansen, P. (1995), 526, 554 Hansen, P. (1996), 485, 505 Hansen, P. S., Phlips, E. J., Aldridge, F. J. (1997), 814, 829 Hanson, J. M., Leggett, W. C. (1982), 799 Hanson, M. J., Stefan, H. G. (1984), 810, 825, 829 Hansson, L.-A., Rudstam, L. G., Johnson, T. B., Soranno, P., Allen, Y. (1994), 23, 49 Happey, C., Moss, B. (1967), 490, 505 Happey-Wood, C. M. (1976), 490, 505 Happey-Wood, C. M. (1988), 258, 308, 364, 380 Harding, J. P. C., Whitton, B. A. (1981), 205, 222 Hardwick, G. G., Blinn, D. W., Usher, H. D. (1992), 34, 49 Hargeby, A., Andersson, G., Blindow, I., Johansson, S. (1994), 811, 821, 829 Hargreaves, J. W., Lloyd, E. J. H., Whitton, B. A. (1975), 41–42, 49, 363, 380 Hargreaves, J. W., Whitton, B. A. (1976), 407, 418 Hargreaves, J. W., Whitton, B. A. (1976a), 42, 49 Hargreaves, J. W., Whitton, B. A. (1976b), 42, 49 Harper, D. (1992), 782, 799 Harris, D. O. (1964), 433, 463, 467 Harris, D. O., Starr, R. C. (1969), 227, 241, 248–249 Harris, E. H. (1989), 226, 249 Harris, K. (1953), 533, 554 Harris, K., Bradley, D. E. (1957), 533, 538, 554 Harris, K., Bradley, D. E. (1960), 533, 554 Harrison, S. S. C., Hildrew, A. G. (1998), 26, 49 Hartmann, H., Steinberg, C. (1989), 526, 536, 554 Harvey, R. S., Patrick, R. (1968), 799 Harvey, W. H. (1836), 312, 350 Harwood, D. M. (1999), 561, 590 Harwood, D. M., Gersonde, R. (1990), 570, 590 Harwood, D. M., Nikolaev, V. A. (1995), 559, 570, 590 Haselkorn, R. (1986), 119, 193 Hasle, G. R. , Evensen, D. L. (1975), 565, 585, 590 Hasle, G. R. , Evensen, D. L. (1976), 565, 585, 590 Hasle, G. R. , Lange, C. B. (1989), 587, 590 Hasle, G. R. (1973), 655–656, 665, 667 Hasle, G. R. (1977), 572, 575, 590 Hasle, G. R. (1978), 587, 590 Haüber, M. M., Müller, S. B., Maier, U.-G. (1994), 385, 402, 418 Haughey, A. (1970), 406, 418 Haupt, W. (1972), 354, 380 Haupt, W., Schönbohm, W. (1970), 354, 380 Hauschild, C. A., McMurrer, H. J. G., Pick, F. R. (1991), 65, 112 Hauser, W. J., Legner, E.F., Medved, R. A., Platt, S. (1976), 820, 829 Hausmann, K. (1978), 403, 418, 686, 690, 711 Hausmann, K., Walz, B. (1979), 752 Havens, K. E. (1994), 822, 825, 829 Havens, K. E., Bull, L. A., Warren, G. L., Crisman, T. L., Phlips, E. J., Smith, J. P. (1996), 24, 49 Havens, K. E., East, T. (1997), 813, 829 Havens, K. E., III (1989), 428, 467 Havens, K. E., James, R. T. (1997), 814, 829 Hawes, I., Schwarz, A.-M. (1996), 24, 28, 49 Hawkes, H. A. (1975), 31, 49 Hawkins, P. R., Griffiths, D. J. (1987), 806, 822, 824, 829 Hawley, G. R. W., Whitton, B. A. (1991), 20, 49 Haworth, E. Y. (1972), 604, 610, 632 Haworth, E. Y., Hurley, M. A. (1986), 583, 590 Hayaski, M., Toda, K., Kitaoka, S. (1993), 387, 418 Hayden, A. (1910), 463, 467

Hayes, P. K., Barker, G. L. A., Walsby, A. E. (1997), 177, 193 Hazen, T. E. (1902), 325, 348, 350 Healey, F. P., Hendzel, L. L. (1979), 785, 799 Healey, F. P., Hendzel, L. L. (1980), 785, 799 Healy, F. P. (1982), 61, 112 Heaney, S. I., Chapman, D. V., Morison, H. R. (1983), 696, 711 Heaney, S. I., Furnass, T. I. (1980), 20, 49, 700–701, 711 Heaney, S. I., Lundm J. W. G., Canter, H., Gray, K. (1988), 20, 50 Heaney, S. I., Talling, J. F. (1980), 23, 49, 699–700, 711 Hecky, R. E., Hesslein, R. H. (1995), 26 Hecky, R. E., Kilham, P. (1988), 23, 50, 776, 785, 799 Hedley, S., Patterson, D. J. (1992), 418 Hedtke, S. F. (1984), 825, 829 Hegewald, E. (1976), 434, 447, 449, 459, 467 Hegewald, E., Schmidt, A. (1987), 255, 308 Hegewald, E., Schmidt, A. (1991), 255, 308 Hegewald, E., Silva, P. C. (1988), 254–256, 308 Hegner, R. W. (1922), 412, 415, 418 Hegner, R. W. (1923), 412, 415, 418 Hein, M. K. (1981), 599, 605, 613, 633 Heinonen, P. (1980), 488, 505 Henderson, R. J., Mackinlay, E. E. (1989), 738, 753 Henson, E. B. (1984), 314, 350 Hepperle, D. (1997), 236, 249 Hepperle, D., Krientiz, L. (1996), 238, 249 Hepperle, D., Nozaki, H., Hohenberger, S., Huss, V. A. R., Morita, E., Krienitz, L. (1998), 236, 249 Herbicide Handbook (1994), 823, 829 Herbst, D. B., Blinn, D. W. (1998), 43, 50 Herbst, D. B., Bradley, T. J. (1989), 43, 50 Herbst, D. B.., Castenholz, R. W. (1994), 333, 350 Herbst, R. P. (1969), 809, 829 Heribaud, J. (1903), 633 Herth, W., Barthlott, W. (1979), 567, 590 Herth, W., Kuppel, A., Brown, R. M. (1975), 484, 505 Herth, W., Kuppel, A., Schnepf, E. (1977), 501, 505 Herth, W., Zugenmaier, P. (1979), 484, 505 Herzog, M., Maroteaux, L. (1986), 686, 711 Herzog, M., von Boletzky, S., Soyer, M.-O. (1984), 686, 711 Hesse, L. W., Schmulbach, J. C., Carr, J. M., Keenlyne, K. D., Unkenholz, D. G., Robinson, J. W., Mestl, G. E. (1989), 28, 50 Heynig, H. (1963), 513, 519 Heywood, P. (1973), 465, 467 Heywood, P. (1978a), 427, 467 Heywood, P. (1978b), 427, 467 Heywood, P. (1980), 427–428, 467 Heywood, P. (1988), 735, 753 Heywood, P. (1989), 427, 467 Heywood, P. (1990), 427–428, 463, 465, 467 Hibberd, D. J. (1976), 523, 528, 554 Hibberd, D. J. (1976a), 471, 473–474, 505, 511, 519 Hibberd, D. J. (1976b), 498, 505, 512, 519 Hibberd, D. J. (1977), 485, 505, 735, 738, 753 Hibberd, D. J. (1978), 528, 554 Hibberd, D. J. (1980), 429, 432, 465, 467 Hibberd, D. J. (1981), 424, 465, 467 Hibberd, D. J. (1983), 499, 505, 512, 519 Hibberd, D. J. (1985), 512, 519 Hibberd, D. J. (1990), 384, 387, 418 Hibberd, D. J. (1990a), 429, 431–432, 434, 446, 467 Hibberd, D. J. (1990b), 424–425, 427, 443, 465, 467 Hibberd, D. J., Greenwood, A. D., Griffiths, H. B. (1971), 732, 741, 753 Hibberd, D. J., Leedale, G. F. (1970), 424, 467 Hibberd, D. J., Leedale, G. F. (1971a), 424, 467

Author Index Hibberd, D. J., Leedale, G. F. (1971b), 424, 432, 467 Hibberd, D. J., Leedale, G. F. (1972), 424, 467 Hickel, B. (1981), 73, 112 Hickel, B. (1988), 694, 711 Hickel, B., Häkansson, H. (1991), 568, 590 Hickel, B., Maass, I. (1989), 536, 538, 554 Hickman, M. (1978), 27, 50 Hickman, M., White, J. M. (1989), 604, 633 Hietala, J., Lauren-Maatta, C., Walls, M. (1997), 64, 112 Highfill, J. F., Pfiester, L. A. (1992a), 693–694, 708, 711 Highfill, J. F., Pfiester, L. A. (1992b), 693–694, 708, 711 Hilenski, L. L., Walne, P. L. (1983), 403, 418 Hill, B. H., Herlihy, A. T., Kaufmann, P. R., Stevenson, R. J., McCormick, F. H., Johnson, C. B. (2000), 789, 799 Hill, B. H., Lazorchak, J. M., McCormick, F. H., Willingham, W. T. (1997), 776, 785, 799 Hill, D. J. (1969), 193 Hill, D. J. (1970a), 174, 180, 194 Hill, D. J. (1970b), 174, 180. 194 Hill, D. J. (1972), 194 Hill, D. J. (1976a), 169, 170, 194 Hill, D. J. (1976b), 169, 170, 194 Hill, D. J. (1976c), 169, 170, 194 Hill, D. R. A. (1990), 716, 732, 734, 741–742, 747, 753 Hill, D. R. A. (1991a), 715, 735, 741–742, 744, 747, 749, 753 Hill, D. R. A. (1991b), 715–716, 732, 734–735, 741–742, 745, 747, 749, 753 Hill, D. R. A. (1991c), 716, 734–735, 744, 749, 753 Hill, D. R. A., Rowan, K. S. (1989), 735, 741, 753 Hill, D. R. A., Wetherbee, R. (1986), 715–716, 732, 734, 736, 741, 746, 753 Hill, D. R. A., Wetherbee, R. (1988), 715–716, 732, 741, 753 Hill, D. R. A., Wetherbee, R. (1989), 715–716, 732, 736, 741–742, 753 Hill, M. O. (1979), 790, 799 Hill, W. (1996), 489, 505 Hill, W. R., Boston, J. L. (1991), 785, 799 Hillebrand, H. (1983), 780, 799 Hillebrand, H., Dürlsen, C. D., Kirschtel, D., Pollingher, U., Zohary, T. (1999), 783, 799 Hilliard, D. K. (1966), 478, 487, 499, 505 Hilliard, D. K. (1967), 487, 499, 501, 505 Hilliard, D. K. (1968), 488, 498, 505 Hilliard, D. K. (1971a), 477, 495, 498, 505 Hilliard, D. K. (1971b), 505 Hilliard, D. K., Asmund, B. (1963), 477, 484, 486, 498, 505 Hindák, F. (1963), 349–350 Hindák, F. (1981), 447, 467 Hindák, F. (1984), 81, 112 Hindák, F. (1985), 135, 194 Hindák, F. (1996), 81, 112, 345, 350 Hindák, F. (2002), 120, 194 Hindák, F. Hindakova, A. (1992), 301, 308 Hindák, F., Moustaka, M. P. (1988), 81, 112 Hinkle, J. (1986), 810, 829 Hirose, H., Hirano, M. (1981), 82, 112 Hirsch, A., Palmer, C. M. (1958), 432, 459, 467 Hoagland, K. D., Carder, J. P., Spawn, R. L. (1996), 782, 800 Hoagland, K. D., Drenner, R. W., Smith, J. D., Cross, D. R. (1993), 786, 794, 800 Hoagland, K. D., Peterson, C. G. (1990), 24–25, 50, 613, 633 Hoagland, K. D., Roemer, S. C., Rosowski, J. R. (1982), 25, 50 Hoagland, K. D., Rosowski, J. R., Gertz, M. R., Roemer, S. C. (1993), 567, 590 Hoek. see van den Hoek

861

Hoffman, L. R. (1967), 349, 350 Hoffman, L. R., Vesk, M., Pickett-Heaps, J. D. (1986), 499, 505 Hoffmann, J. P. (1998), 12, 44, 50 Hoffmann, L. (1989), 197, 206, 222 Hoffmann, L. (1996), 12, 50 Hoffmann, L., Demoulin, V. (1985), 161, 194 Hoffmann, L., Willie, E. (1992), 535, 554 Hofmann, C. J. B., Rensing, S. A., Haeuber, M. M., Martin, W. F., Mueller, S. B., Couch, J., McFadden, G. I., Igloi, G. L., Maier, U. G. (1994), 753 Hofmann, G. (1994), 791, 800 Hoham, R. W. (1973), 315, 350 Hoham, R. W. (1974a), 234, 249 Hoham, R. W. (1974b), 234, 249 Hoham, R. W. (1975), 44, 50 Hoham, R. W. (1980), 226, 249 Hoham, R. W., Blinn, D. W. (1979), 43–44, 50, 407, 418 Hoham, R. W., Mohn, W. W. (1985), 44, 50 Hoham, R. W., Mullet, J. E., Roemer, S. C. (1983), 235, 249 Hoham, Roemer, S. C., Mullet, J. E. (1979), 235, 249 Hohman, R. W., Mullet, J. E. (1977), 43, 50 Hohn, M. H. (1959), 665, 667 Hohn, M. H. (1969), 572, 590 Hohn, M. H., Hellerman, J. (1963), 561, 590, 604, 616, 633, 652 Holcomb, G. E. (1986), 343, 350 Holdway, P. A., Watson, R. A., Moss, B. (1978), 512, 519 Holen, D. A., Boraas, M. E. (1995), 534, 554 Hollenberg, G. J. (1939), 94–95, 107, 109, 112 Holm, L. G., Yeo, R. (1981), 821, 829 Holm, N. P., Armstrong, D. E. (1981), 607, 633 Holmes, N. T. H., Whitton, B. A. (1975), 763, 765, 767, 770–772 Holmes, N. T. H., Whitton, B. A. (1977), 30, 33–34, 50 Holmes, N. T. H., Whitton, B. A. (1977a), 763–766, 772 Holmes, N. T. H., Whitton, B. A. (1977b), 763–766, 772 Holmes, N. T. H., Whitton, B. A. (1977c), 763, 765–766, 772 Holmes, N. T. H., Whitton, B. A. (1981), 34, 50, 764, 772, 779, 783, 800 Holmes, R. W. (1985), 623, 633 Holmgren, S. K. (1984), 513, 519 Holmquist, E., Willén, T. (1993), 512, 519 Holomuzki, J. R., Short, T. M., (1988), 783, 800 Holopainen, I. J. (1992), 699, 711 Holt, J. R., Pfiester, L. A. (1981), 700–701, 711 Holz, J. C., Hoagland, K. D. (1999), 814, 829 Hooper, C. A. (1981), 40, 50 Hooper-Reid, N. M., Robinson, G. G. C. (1978), 39, 50 Hoops, H. J. (1984), 242, 249 Hoops, H. J., Floyd, G. L. (1982), 245, 249 Horecká, M., Komárek, J. (1979), 173–174, 194 Hori, H., Osawa, S. (1987), 387, 418 Horne, A., Goldman, C. R. (1994), 21, 50 Horne, A. J., Goldman, C. R. (1974), 822, 829 Horner, R. R., Welch, E. B., Veensstra, R. B. (1990), 35, 50 Hörtzel, G., Croome, R. (1994), 65, 112 Hoshaw, McCourt, R. M., Wang, J. C. (1990), 353, 362–363, 380 Hoshaw, R. W. (1968), 364–365, 380 Hoshaw, R. W., McCourt, R. M. (1988), 363–365, 380 Hoshaw, R. W., Rosowski, J. R. (1973), 409, 418, 740, 753 Hoshaw, R. W., Wang, J. C., McCourt, R. M., Hull, H. M. (1985), 371, 380 Hoshaw, R. W., Wells, C. V., McCourt, R. M. (1987), 371, 380 Hosiaisluoma, V. (1975), 363, 380 Howard, R. V., Parker, B. C. (1980), 203, 210, 221–222 Howarth, R. W., Cole, J. J. (1985), 23, 50 Howe, M. A. (1924), 89, 112

862

Author Index

Howe, M. A. (1932), 81, 112 Howell, E. T., South, G. R. (1981), 363, 380 Hoyer, M. V., Canfield, D. E., Jr. (1996a), 810, 829 Hoyer, M. V., Canfield, D. E., Jr. (1996b, 810, 829 Hua, M. S., Friedmann, E. I., Ocampo-Friedmann, R., Campbell, S. (1989), 1112 Huang, T. C., Grobbelaar, N. (1989), 45, 50 Huber, A. L. (1984), 120, 177, 194 Huber-Pestalozzi, G. (1938), 92, 113 Huber-Pestalozzi, G. (1941), 503, 505 Huber-Pestalozzi, G. (1942), 572, 590 Huber-Pestalozzi, G. (1950), 716, 741–742, 748, 753 Huber-Pestalozzi, G. (1955), 387–388, 401, 415–416, 418 Huber-Pestalozzi, G. (1961), 247, 249 Hudon, C., Paquet, S., Jarry, V. (1996), 37, 50 Hufford, T. L., Collins, G. B. (1972a), 664, 667 Hufford, T. L., Collins, G. B. (1972b), 664, 667 Hufford, T. L., Collins, G. B. (1976), 34, 50 Hughes, E. O. (1948), 434, 441, 454, 459, 461, 467 Hughes, R. M. (1995), 786, 788, 800 Humm, H. J., Wicks, S. R. (1980), 113, 151, 194 Humphrey, K. P., Stevenson, R. J. (1992), 785, 800 Hunt, M. E., Floyd, G. L., Stout, B. B. (1979), 45, 50, 63, 113 Hurlbert, S. H. (1971), 784, 800 Huss, V. A. R., Frank, C., Hartmann, E. C., Hirmer, M., Kloboucek, A., Seidel, B. M., Wenzler, P., Kessler, E. (1999), 253, 257, 308 Huss, V. A. R., Sogin, M. L. (1990), 257, 308 Hustedt, F. (1926), 656, 667 Hustedt, F. (1927), 604, 633 Hustedt, F. (1927–1930), 562, 590 Hustedt, F. (1930), 596, 604, 628, 633, 639, 652, 655, 661, 667, 682–683 Hustedt, F. (1930b), 562, 573, 583–584, 586, 590 Hustedt, F. (1931–1959), 562, 590, 641, 652 Hustedt, F. (1937–1939), 628, 633 Hustedt, F. (1939), 536, 554 Hustedt, F. (1942), 672, 683 Hustedt, F. (1952), 633, 656, 667 Hustedt, F. (1957), 586, 590 Hustedt, F. (1959), 596–597, 604, 628, 633 Hustedt, F. (1961–1966), 637, 652 Hutchinson, G. E. (1957), 12–13, 15–16, 19, 41, 50 Hutchinson, G. E. (1961), 19, 50 Hutchinson, G. E. (1967), 12, 16, 19, 50, 583, 590, 691, 711 Hutchinson, G. E. (1969), 405, 418 Hutchinson, G. E. (1975), 12, 17, 24–25, 27, 50, 314, 339, 350 Hutchinson, T., Havas, M. (1986), 793, 800 Hymes, B. J., Cole, K. M. (1983), 201, 222 Hynes, H. B. N. (1970), 28–32, 34, 50, 203, 222 I Ibelings, B., Admiraal, W., Bijker, R., Letswaart, T., Prins, H. (1998), 776, 800 Iglesias-Prieto, R. (1996), 690, 711 Ikävalko, J., Kristiansen, J., Thomsen, H. A. (1994), 474, 502, 505 Inagaki, Y., Hayashi-Ishimaru, Y., Ehara, M., Igarashi, I., Ohama, T. (1997), 418 Infante, A., Abella, S. E. B. (1985), 808, 813, 829 Islam, A. K. M. N. (1961), 332, 348, 350 Islam, A. K. M. N. (1963), 349–350 Islam, A. K. M. N., Khondker, M. (1994), 465, 467 Israelson, G. (1938), 758, 763–764, 766, 772 Israelson, G. (1942), 204, 222 Israelson, G. (1949), 35, 50, 363, 380 Ito, H. (1989), 513, 519

Ito, H., Takahashi, E. (1982), 487, 505 Iyengar, M. O. P. (1925), 432, 467 Iyengar, M. O. P., Desikachary, T. V. (1981), 226–228, 230, 232, 235, 238, 247–249 J Jaag, O. (1941), 113 Jaag, O. (1945), 61–64, 92, 113 Jaag, O. (1972), 806, 829 Jackim, E., Gentile, J. (1968), 121, 194 Jackson, A. E. (1997), 45, 50 Jackson, J. E., Castenhloz, R. W. (1975), 189, 194 Jackson, L. J., Stockner, J. G., Harrison, P. J. (1990), 587, 590 Jacobs, D. L. (1946), 688, 697, 701, 706, 711 Jacobs, J. E. (1968), 434, 441, 454, 467 Jacobs, J. E. (1971), 447, 459, 461, 467, 491, 505 Jacobsen, B. (1985), 486, 506 Jacobsen, B. A. (1985), 537–538, 554 Jacobson, D. M., Anderson, D. M. (1986), 701, 711 Jacoby, J. M., Gibbons, H. L., Stoops, K. B., Bouchard, D. D. (1994), 814, 829 Jagg, O. (1945), 120, 158, 194 Jahn, T. L. (1946), 399, 404, 407, 413, 415, 418 Jahn, T. L. (1951), 404, 412, 418 Jahn, T. L., Bovee, E. C. (1968), 401, 418 Jahn, T. L., Bovee, E. C., Jahn, F. F. (1979), 407, 418 James, T. L., de la Cruz, A. (1989), 512, 519 James, W. F., Taylor, W. D., Barko, J. W. (1992), 699–701, 711 Jao, C.-C. (1941), 759, 763, 770–772 Jao, C.-C. (1943), 763, 765, 771–772 Jao, C.-C. (1944), 763, 765, 771–772 Jarosch, R. (1970), 528, 554 Jasby, A. D., Goldman, C. R., Reuter, J. E. (1995), 26, 50 Jeffrey, S. W. (1989), 474, 506 Jeffrey, S. W., Sielicki, M., Haxo, F. T. (1975), 690, 711 Jensen, T. E. (1984), 61, 114 Jensen, T. E. (1985), 61, 113 Jensen, W. A. (1962), 539, 554 Jewson, D. J. (1992a), 568, 590 Jewson, D. J. (1992b), 568, 590 Jimenez, J. C. (1999), 208, 222 Jochimsen, E. M., Carmichael, W. W. (1998), 807, 829 Johansen, J., Cognata, S. L., Kociolek, J. P. (1990), 682–683 Johansen, J. R. (1993), 45, 50, 113 Johansen, J. R. (1999), 45, 50 Johansen, J. R., Ashley, J., Rayburn, W. R. (1993), 113 Johansen, J. R., Doucette, G. J., Barclay, W. R., Bull, J. D. (1988), 43, 50, 512, 514–515, 519 Johansen, J. R., Rushforth, S. R. (1981), 604, 628, 633 Johansen, J. R., Rushforth, S. R. (1985), 113 Johansen, J. R., Stray, J. C. (1998), 648, 652 Johanson, J. R., Barclay, W. R., Nagle, N. (1990), 574, 590 Johanson, J. R., Rushforth, S. R. (1985), 574, 590 Johanson, J. R., Theriot, E. C. (1987), 587, 590 John, D. M. (1994), 311–312, 350 John, D. M., Johnson, L. R. (1987), 336, 350 John, D. M., Johnson, L. R. (1989), 349, 351 John, D. M., Moore, J. A. (1985), 314, 351 Johnson, L. M. Rosowski, J. R. (1992), 584, 590 Johnson, L. P. (1944), 391, 404–405, 408, 412, 418 Johnson, L. P. (1968), 404, 418 Johnson, L. P., Jahn, T. L. (1942), 412, 418 Johnson, L. R., John, D. M. (1990), 348, 351 Jones, B. L. (1987), 806, 829 Jones, H. L. J., Leadbeater, B. S. C., Green, J. C. (1994), 514, 520

Author Index Jones, J. R. Bachmann, R. W. (1976), 812, 829 Jones, R. C., Walti, K., Adams, M. S. (1983), 807, 829 Jones, R. I., Rees, S. (1994), 489, 506 Jongman, R. H. G., ter Braak, C. K. F., Van Tongeren, O. F. R. (1995), 790, 800 Jordan, R. W., Chamberlain, A. H. L. (1997), 512, 520 Jordan, R. W., Green, J. C. (1994), 512, 520 Jordan, R. W., Kleinjne, A., Heimdal, B. R., Green, J. C. (1995), 511–512, 516, 520 Joyce, J. C. (1993), 818, 829 Jügensen, M. F. (1973), 63, 113 Juggins, S., Battarbee, R., Fritz, S., Gasse, F. (1994), 794, 800 Juggins, S., ter Braak, C. J. F. (1992), 800 Julius, M. L. Estabrook, G. F., Edlund, M. B., Stoermer, E. F. (1997a), 568–569, 590 Julius, M. L., Stoermer, E. F., Colman, S. M., Moore, T. C. (1997b), 568, 583, 590 Julius, M. L. Stoermer, E. F., Taylor, C. M., Schelske, C. L. (1998), 586, 590 Junk, W. J., Bayley, P. B., Sparks, R. E. (1989), 32, 50 Jupp, B. P., Spence, D. H. N. (1997), 807, 830 Jürgens, U. J., Weckesser, J. (1985), 118, 194 Jüttner, F. (1987), 64, 113 Jüttner, F., Höfflacher, B., Wurster, K. (1986), 807, 830 Jüttner, I., Rothfritz, H., Omerod, S. J. (1996), 784, 800 K Kaczmarczyk, D., Sheath, R. G. (1991), 203, 222 Kaczmarczyk, D., Sheath, R. G., Cole, K. M. (1992), 203, 214, 216, 221–222 Kaczmarska, I., Rushforth, S. R. (1983), 604–605, 628, 633 Kadlubowska, J. Z. (1972), 362, 369–372, 379–380 Kalff, J., Knoechel, R. (1978), 19, 50 Kalinsky, R. G. (1984), 665, 667 Kallqvist, T., Meadows, B. S. (1978), 823, 830 Kann, E. (1941), 25, 50 Kann, E. (1945), 763–764, 772 Kann, E. (1959), 26, 50 Kann, E. (1966), 763, 772 Kann, E. (1972), 101, 113 Kann, E. (1973), 65, 101, 113 Kann, E. (1976), 767, 772 Kann, E. (1978), 25, 33, 50 Kann, E. (1978a), 757, 763–764, 767, 772 Kann, E. (1978b), 763, 772 Kann, E. (1982), 765, 768, 772 Kann, E. (1993), 758, 763–764, 772 Kantz, T. S., Theriot, E. C., Zimmer, E. A., Chapman, R. L. (1960), 257, 308 Karayeva, N. I., Maggerramova, N. R., Rhazeva, S. G. (1984), 663, 667 Karim, A. G., Round, F. E. (1967), 484, 506 Karlström, U. (1978), 36, 50 Karol, K. G., McCourt, R. M., Cimino, M. T., Delwiche, C. F. (2001), 228, 249 Karr, J. R. (1981), 786, 800 Karr, J. R., Chu, E. W. (1999), 782, 786, 789, 800 Karr, J. R., Dudley, D. R. (1981), 777, 786, 789, 800 Karsten, G. (1928), 655, 667 Kato, S. (1991), 428, 467 Kato, S. (1994), 415., 418 Kato, T., Watanabe, M. (1993), 63, 113 Kawabata, Z., Banba, D. (1993), 696, 712 Kawabata, Z., Zagawa, H. (1988), 699–701, 712 Kawachi, M., Inouye, I. (1995), 512, 520

863

Kawai, H., Inouye, I. (1989), 513, 520 Kawecka, B. (1980), 33, 45, 50 Kawecka, B. (1981), 32, 51 Kawecka, B. (1990), 35, 51 Kay, S. H. (1997), 822, 830 Keating, K. I. (1976), 808, 830 Keating, K. I. (1977), 808, 830 Keating, K. I. (1978), 808, 830 Keithan, E. D., Lowe, R. L. (1985), 604, 633 Kelley, I., Pfiester, L. A. (1990), 694, 712 Kelly, M. G., Cazaubon, A., Coring, E., Dell’Uomo, A., Ector, L., Goldsmith, B., Guasch, H., Hürlimann, J., Jarlman, A., Kawecka, B., Kwandrans, J., Laugaste, R., Lindstrrm, E.-A., Leitao, M., Marvan, P., Padisák, J., Pipp, E., Pyrgiel, J., Rott, E., Sabater, S., van Dam, H., Viznet, J. (1998), 775–776, 779, 784, 800 Kelly, M. G., Hornberger, G. M., Cosby, B. J. (1974), 785, 800 Kelly, M. G., Penny, C. J., Whitton, B. A. (1995), 800 Kelly, M. G., Whitton, B. A. (1989), 800 Kelly, M. G., Whitton, B. A. (1995), 776, 782, 787, 800 Kelmer, A., Barko, J. (1991), 700, 712 Kempner, E. S., Miller, J. H. (1972), 407, 418 Kentucky Division of Water (1993), 775, 782, 784, 789, 800 Kentucky Natural Resources and Environmental Protection Cabinet (1997), 794, 800 Keough, J. R., Hagley, C. A., Ruzycki, E., Sierszen, M. (1998), 39, 51 Kerans, B. L., Karr, J. R. (1994), 786, 789, 800 Kesler, D. H. (1981), 25, 51 Kessler, E., Huss, V. A. R. (1992), 307–309 Kessler, E., Schäfer, M. Hümmer, C., Kloboucek, A., Huss, V. A. R. (1997), 256, 308 Khan, M. A. (1993), 384, 412, 418 Khan, M. A. (1995), 584, 591 Khan, M. A. R., Begum, Z. N. T., Rahim, A. T. M. A., Salamatullah, Q. (1996), 818, 830 Khursevich, G. K., VanLandingham, S. L. (1993), 587, 591 Kiener, W. (1944), 407, 418 Kies, L., Berndt, M. (1984), 538, 554 Kilham, P., Tilman, D. (1979), 487, 506 Kilham, S. S., Kilham, P. (1978), 22, 51, 487, 506, 607, 613, 633 Killgore, K. G., Kirk, J. P., Foltz, J. W. (1998), 821, 830 Kim, J. T., Boo, S. M. (1998), 406, 418 Kindle, E. M. (1934), 314, 351 King, J. M. (1983), 501, 506 King, J. M. (1984), 497, 506 King, J. M., Ward, C. H. (1977), 45, 51, 63, 113 Kingsley, J. S. (1888), 384, 418 Kingston, J. C. (1978), 664, 667 Kingston, J. C. (1980), 623, 633 Kingston, J. C. (1982), 40, 51, 605, 633 Kingston, J. C. (1984), 604, 633 Kingston, J. C. (1986), 605, 633 Kingston, J. C. (1997), 598, 633 Kingston, J. C. (2000), 596, 613, 627, 633 Kingston, J. C., Birks, H. J. B. (1990), 604, 633 Kingston, J. C., Birks, H. J. B., Uutala, A. J., Cumming, B. F., Smol, J. P. (1992), 604, 633, 787, 800 Kingston, J. C., Lowe, R. L. (1979), 604, 633 Kingston, J. C., Lowe, R. L., Stoermer, E. F. (1980), 656, 667 Kingston, J. C., Lowe, R. L., Stoermer, E. F., Ladewski, T. B. (1983), 27, 51, 604, 633 Kingston, J. C., Sherwood, A. R., Bengtsson, R., (2001), 631, 633 Kirjakov, I. K. (1983), 416, 419 Kirjakov, I. K. (1998), 413, 419 Kirk, D. L. (1998), 226, 242, 250

864

Author Index

Kirk, D. L., Birchem, R., King, N. (1986), 226, 250 Kirkby, S. M., Hibberd, D. J., Whitton, B. A. (1972), 759, 763–764, 771, 773 Kiselev, I. A. (1947), 80, 113 Kiss, K. T. (1984), 587, 591 Kiss, K. T. (1994), 38, 51 Kivic, P. A., Vesk, M. (1972), 399, 419 Kivic, P. A., Vesk, M. (1974), 399, 401, 419 Kivic, P. A., Walne, P. L. (1984), 385, 419 Klaveness, D. (1977), 738, 753 Klaveness, D. (1981), 735, 742, 748, 753 Klaveness, D. (1982), 739, 753 Klaveness, D. (1984), 739, 753 Klaveness, D. (1985), 716, 734, 741, 745, 753 Klaveness, D. (1988a), 715, 736, 738, 753 Klaveness, D. (1988b), 715, 738, 753 Klebs, G. (1883), 401, 419 Klebs, G. (1893a), 471, 484, 506 Klebs, G. (1893b), 471, 484, 506 Klemm, D. J., Lazorchak, J. M. (1994), 778, 780, 800 Kling, H. (1975), 171, 194 Kling, H., Findlay, D. L. Komárek, J. (1994), 171–172, 194 Kling, H., Holmgren, S. (1972), 129, 147, 194, 537, 554 Kling, H. J. (1981), 514, 518–520 Kling, H. J. (1992), 567–569, 591 Kling, H. J., Kristiansen, J. (1983), 500, 503, 506 Kling, H., Kristiansen, J. (1983), 552, 554 Klut, M. E., Bisalputta, T., Antia, N. J. (1987), 690, 712 Knaust, R., Urbig, T., Li, L., Taylor, W., Hastings, J. W. (1998), 687, 712 Kobayashi, H. (1997), 623, 628, 633 Kobayashi, H., Ando, K., Nagumo, T. (1981), 665, 667 Koch, W. J. (1951), 432, 467 Kociolek, J. P. (1997), 560, 591 Kociolek, J. P. (1998), 597–598, 633, 659, 667 Kociolek, J. P. (2000), 664–665, 667 Kociolek, J. P., Herbst, D. B. (1992), 43, 51 Kociolek, J. P., Kingston, J. C. (1999), 605, 633, 655, 659, 661, 665, 667 Kociolek, J. P., Lamb, M. A., Lowe, R. L. (1983), 584, 591 Kociolek, J. P., Rhode, K. (1998), 595, 633 Kociolek, J. P., Rhode, K., Williams, D. M. (1997), 655, 661, 664, 667 Kociolek, J. P., Rosen, B. H. (1984), 655, 659, 665, 667 Kociolek, J. P., Spaulding, S. A., Kingston, J. C. (1998), 12, 51, 637, 652 Kociolek, J. P., Stoermer, E. F. (1986), 604, 633, 655, 665, 667 Kociolek, J. P., Stoermer, E. F. (1987a), 656, 664, 667 Kociolek, J. P., Stoermer, E. F. (1987b), 655, 664–665, 667 Kociolek, J. P., Stoermer, E. F. (1988a), 560, 591, 655, 659, 662, 667 Kociolek, J. P., Stoermer, E. F. (1988b), 570, 591, 655, 667 Kociolek, J. P., Stoermer, E. F. (1988c), 655, 659, 664, 667 Kociolek, J. P., Stoermer, E. F. (1989), 597, 633, 665, 667 Kociolek, J. P., Stoermer, E. F. (1991), 655, 659, 665, 667 Kociolek, J. P., Stoermer, E. F. (1993a), 655, 659, 667 Kociolek, J. P., Stoermer, E. F. (1993b), 667 Kociolek, J. P., Stoermer, E. F., Edlund, M. A. (1995), 659, 665, 667 Kociolek, J. P., Stoermer, E. F., Sicko-Goad, L. (1994), 664, 667 Kociolek, J. P., Theriot, E. C., Williams, D. M. (1989), 597, 633 Kociolek, J. P., Williams, D. M. (1987), 559–560, 591 Kociolek, J. P., Williams. D. M. (1987), 595, 633 Kociolek, J. P., Yang, J.-R., Stoermer, E. F. (1988), 661, 665, 667 Kofoid, C. A. (1899), 227, 240–241, 250 Kofoid, C. A. (1903), 36, 51 Kofoid, C. A. (1908), 36, 51

Kofoid, C. A. (1909), 690, 712 Kofoid, C. A., Michener, J. R. (1912), 693, 712 Kofoid, C. A., Swezy, O. (1921), 690, 712 Kohl, J.-G., Dudel, G., Schlangstedt, M., Kuhl, H. (1985), 120, 194 Kohl, J.-G., Nicklisch, A. (1981), 126, 129, 194 Kohl, J.-G., Schlangstedt, M., Dudel, G. (1987), 119, 194 Köhler, J. (1993), 37, 51 Köhler, J. (1995), 38, 51 Kol, E. (1942), 355, 363–364, 369, 380 Kol, E. (1944), 363, 369, 380 Kol, E. (1964), 363, 369, 380 Kol, E. (1968), 43–44, 51 Kolbe, R. W. (1956), 665, 667 Kolkwitz, R., Marson, M. (1908), 560, 591, 775, 790, 800 Komagata, K. (1987), 121, 194 Komárek, J. (1958), 67, 83, 90, 113, 120, 127, 141, 145, 167, 172, 194 Komárek, J. (1969), 68, 76, 113 Komárek, J. (1970), 113 Komárek, J. (1975), 98, 113 Komárek, J. (1976), 60, 68, 113 Komárek, J. (1984), 89, 113, 142, 163–164, 194 Komárek, J. (1985), 104, 107, 109, 113 Komárek, J. (1989), 76, 82–83, 86–87, 99, 102, 113, 131, 146, 150, 152, 170, 173–174, 184–185, 194 Komárek, J. (1991), 410, 419 Komárek, J. (1993), 62, 64, 92–93, 113 Komárek, J. (1994), 60, 93, 95, 113, 126, 163–164, 194 Komárek, J. (1995), 72, 77, 79, 83, 87, 113 Komárek, J. (1999), 60–61, 65, 113, 120, 194 Komárek, J. (2001), 129, 145, 174, 194 Komárek, J., Albertano, P. (1994), 147, 194 Komárek, J., Anagnostidis, K. (1986), 60, 62, 101, 113 Komárek, J., Anagnostidis, K. (1989), 117–121, 161, 171, 177, 180, 194 Komárek, J., Anagnostidis, K. (1995), 113 Komárek, J., Anagnostidis, K. (1998), 60, 62–63, 65, 67, 72, 74, 79, 81–82, 87–89, 91–92, 95–96, 101–102, 104, 107, 110, 113 Komárek, J., Cáslavaská, J. (1991), 118, 194 Komárek, J., Cepak, V. (1998), 77, 79, 113 Komárek, J., Cronberg, G. (2001), 135, 141, 194 Komárek, J., Fott, B. (1983), 253–256, 302, 306–308 Komárek, J., Hindák, F. (1975), 103–105, 113 Komárek, J., Hindák, F. (1988), 87–88, 113 Komárek, J., Hindák, F. (1989), 113 Komárek, J., Hübel, H. _amarda, J. (1993), 120, 177, 194 Komárek, J., Kling, H. (1991), 133, 135, 194 Komárek, J., Komárková-Legnerová, J. (1992), 64–65, 67, 83, 85–87, 89, 113 Komárek, J., Komárková-Legnerová, J. (1993), 113 Komárek, J., Komárková-Legnerová, J. (2002), 60, 80, 87, 113 Komárek, J., Komárková-Legnerová, J., Sant’Anna, C. L., Azevedo, M. T. P., Senna, P. A. C. (2002), 92, 113 Komárek, J., Kopecky, J., Cepák, V. (1999), 79, 113 Komárek, J., Ludvik, J., Pokorny, V. (1975), 61, 113 Komárek, J., Lukavsky, J. (1988), 66–67, 113 Komárek, J., Montejano, G. (1994), 60, 67, 96–97, 113 Komárek, J., Novelo, E. (1994), 94–95, 113 Komárek, J., Watanabe, M. (1990), 158, 160, 194 Komáreková, J. (1998), 174, 194 Komáreková, J. (2001), 120, 194 Komáreková, J., Ladares-Silva, R., Senna, P. A. C. (1999), 174, 194 Komáreková-Legnerová, J., Tavera, R. (1996), 135, 174, 180, 194 Komárková, J. (2001), 65, 114 Komárková-Legnerová, J. (1991), 74, 113

Author Index Komárková-Legnerová, J., Cronberg, G. (1994), 74, 113 Komárková-Legnerová, J., Tavera, R. (1996), 82, 113 Kondrateva, N. V. (1961), 118, 194 Kondrateva, N. V. (1968), 63, 113, 133, 138, 146, 149, 154, 159, 163, 165–168, 175, 179, 194 Kondrateva, N. V. (1972), 120, 194 Kondrateva, N. V., Kovalenko, O. V., Prichod’kova, I. P. (1984), 75, 113 Konopka, A. (1982a), 806, 830 Konopka, A. (1982b), 806, 830 Koppen, J. D. (1975), 601, 617, 621–622, 633 Koppen, J. D. (1978), 601, 604, 617, 633 Korch, J. E., Sheath, R. G. (1989), 201, 222 Korner, H. (1971), 600, 607, 633 Korshikov, A. A. (1924), 247, 250 Korshikov, A. A. (1925), 235, 247, 250 Korshikov, A. A. (1928), 245–247, 250 Korshikov, A. A. (1929), 481, 506, 528, 554 Korshikov, A. A. (1942), 479, 506 Korte, V. L., Blinn, D. W. (1983), 605, 633 Korte, V. L., Blinn, D. W. (1993), 36, 51 Kortmann, R. W., Rich, P. H. (1994), 814–815, 830 Koryak, M. (1978), 699, 712 Kosinskaja, E. K. (1948), 93, 100, 114, 151, 195 Kouwets, F. A. C. (1980), 477, 506 Kouwets, F. A. C. (1995), 256, 308 Kouwets, F., Coesel, P. (1984), 373, 380 Kovácik, L. (1988), 89, 114 Krammer, K. (1979), 663, 667 Krammer, K. (1981), 663, 667 Krammer, K. (1982), 656, 659, 663–664, 667 Krammer, K. (1990), 601, 623, 633 Krammer, K. (1992a), 650, 652 Krammer, K. (1992b), 650, 652 Krammer, K. (1997a), 561, 591, 656, 659, 663–664, 667 Krammer, K. (1997b), 561, 591, 656, 659, 663–664, 667 Krammer, K., Lange-Bertalot, H. (1986), 560–563, 591, 637, 639, 646, 652, 661–664, 668 Krammer, K., Lange-Bertalot, H. (1988), 560–563, 591, 669, 682–683 Krammer, K., Lange-Bertalot, H. (1991), 639, 652, 661–662, 665, 668 Krammer, K., Lange-Bertalot, H. (1991a), 560–563, 579, 585–586, 591, 596, 599–600, 613, 616, 628, 633 Krammer, K., Lange-Bertalot, H. (1991b), 560–563, 591, 596–597, 601, 623, 628, 633 Krebs, W. N., Bradbury, J. P., Theriot, E. C. (1987), 568, 591 Kreis, R.G., Jr., Stoermer, E. F. (1979), 627–628, 633 Krejci, M. E., Lowe, R. L. (1987a), 604, 634 Krejci, M. E., Lowe, R. L. (1987b), 604, 634 Kremer, B. (1983), 203, 222 Krieger, W. (1933), 361, 380 Krienitz, L., Hehmann, A., Caspar, S. J. (1997), 488, 506 Krienitz, L., Hepperle, D., Stich, J., Weiler, W. (2000), 425, 427, 467 Krienitz, L., Takeda, H., Hepperle, D. (1999), 257, 308 Kristiansen, J. (1969), 484, 497, 506 Kristiansen, J. (1971), 513, 520 Kristiansen, J. (1972), 484, 497, 506 Kristiansen, J. (1975), 538, 552, 554 Kristiansen, J. (1980), 536, 554 Kristiansen, J. (1981), 536–537, 554 Kristiansen, J. (1985), 536, 554 Kristiansen, J. (1986), 474, 506, 535–538, 554 Kristiansen, J. (1988), 487, 506 Kristiansen, J. (1988a), 524, 554

865

Kristiansen, J. (1988b), 536, 554 Kristiansen, J. (1990), 471, 506 Kristiansen, J. (1992), 486, 506, 535, 552, 554 Kristiansen, J. (1995a), 471, 506 Kristiansen, J. (1995b), 491, 506 Kristiansen, J., Andersen, R. A. (1983), 503, 506 Kristiansen, J., Cronberg, G. (1996), 503, 506 Kristiansen, J., Cronberg, G., Geissler, U. (1989), 503, 506 Kristiansen, J., Takahashi, E. (1982), 536, 554 Kristiansen, J., Tong, D. (1989), 536, 554 Kristiansen, J., Tong, D. (1995), 486, 506 Kristiansen, J., Vigna, M. S. (1996), 486, 506 Krogmann, D. W., Buttala, R., Sprinkle, J. (1986), 37, 51, 65, 114 Krolikowska, J. (1997), 27, 51 Kudoh, S., Takahashi, M. (1990), 819, 830 Kugrens, P. (1999), 744, 746, 753 Kugrens, P., Clay, B. L., Lee, R. E. (1999), 725, 749, 753 Kugrens, P., Lee, R. E. (1986), 716, 723–724, 729, 732, 745–746, 753 Kugrens, P., Lee, R. E. (1988), 736, 741, 746, 753 Kugrens, P., Lee, R. E. (1990), 748, 753 Kugrens, P., Lee, R. E. (1991), 716, 728, 732–735, 740–741, 744, 746–749, 753 Kugrens, P., Lee, R. E., Andersen, R. E. (1986), 716, 734, 741, 745–747, 749, 753 Kugrens, P., Lee, R. E., Andersen, R. E. (1987), 716, 732, 734, 741, 745–746, 753 Kugrens, P., Lee, R. E., Corliss, J. O. (1994), 732, 734–735, 749–750, 753 Kuhn, D. L., Plafkin, J. L., Cairns, J. J., Lowe, R. L. (1981), 598, 634 Kumano, S. (1978), 202, 222 Kumano, S., Hirose, H. (1959), 759–760, 767, 773 Kumano, S., Necchi, O., Jr. (1987), 202, 222 Kusel-Fetzmann, E. L. (1996), 758–760, 763–767, 771, 773 Kusel-Fetzmann, E. L., Schagerl, M. (1992), 760, 767, 773 Kützing, T. F. (1849), 62, 114 Kuzinicki, L., Mikolajczyk, E., Walne, P. L. (1990), 391–392, 419 Kwandrans, J., Eloranta, P., Kawecka, B., Wojtan, K. (1998), 783–784, 791, 800 L Lackey, J. B. (1938), 42, 51, 487, 506 Lackey, J. B. (1939), 512, 520 Lackey, J. B. (1942), 428, 434, 459, 467 Lackey, J. B. (1968), 405–407, 419, 806, 830 Laessle, A. M. (1961), 45, 51 Lage, O. M., Parente, A. M., Vasconcelos, M. T. S. D., Gomes, C. A. R., Salema, R. (1996), 823, 830 Lagerheim, G. (1989), 432, 467 Lair, N., Reyes-Marchant, P. (1997), 32, 51 Laird, K., Fritz, S. C., Cumming, B. F. (1998), 604, 634 Lakshminarayana, J. S., Devi, J. S. (1975), 499, 506 Lamb, M. A., Lowe, R. L. (1987), 605, 634 Lamberti, G. A. (1996), 776, 783, 800 Lamberti, G. A., Gregory, S. V., Ashkenas, L. R., Steinman, A. D., McIntire, C. D. (1989), 34–35, 51 Lamberti, G. A., Steinman, A. D. (1993), 786, 800 Lamberti, G. A., Steinman, A. D. (1997), 32, 51 Lamon, E. C., III (1995), 806, 830 Lang, N. J., Fay, P. (1971), 119, 195 Lange, T. R., Rada, R. G. (1993), 38, 51 Lange-Bertalot, H. (1976), 682–683 Lange-Bertalot, H. (1979), 775, 782, 787, 800 Lange-Bertalot, H. (1980), 613, 634, 659, 668, 682–683

866

Author Index

Lange-Bertalot, H. (1989), 599, 616, 634 Lange-Bertalot, H. (1993), 561, 591, 665, 668 Lange-Bertalot, H. (1995), 665–666, 668 Lange-Bertalot, H. (1997a), 598–599, 613, 634 Lange-Bertalot, H. (1997b), 597–598, 602, 627, 634 Lange-Bertalot, H. (1999), 598, 602, 634 Lange-Bertalot, H., Compère, P. (2001), 631, 634 Lange-Bertalot, H., Krammer, K. (1989), 596, 601, 634 Lange-Bertalot, H., Krammer, K. (1993), 669, 683 Lange-Bertalot, H., Metzeltin, D. (1996), 561–562, 591, 638, 647, 649, 652, 665, 668 Lange-Bertalot, H., Moser, G. (1994), 638, 646, 652 Lange-Bertalot, H., Ruppel, M. (1980), 602, 634 Lantz, K. E., Davis, J. T., Hughes, J. S., Schafer, H. E. (1964), 811, 830 LaRivers, I. (1978), 429, 433–434, 451, 459, 463, 467 Larkum, T. (1996), 686, 712 Larsen, A., Eikrem, W., Paasche, E. (1993), 512, 520 Larsen, J., Patterson, D. J. (1991), 387, 404, 419 Larson, A., Kirk, M. M., Kirk, D. L. (1992), 242, 250 Larson, G. K. (1989), 15, 51 Larson, G. L., McIntire, C. D., Hurley, M. Buktenica, M. W. (1996), 15, 51 Lasenby, D. C. (1975), 776, 800 Lavau, S., Saunders, G. W., Wetherbee, R. (1997), 524, 526, 534, 548, 554 Lawrence, J. M. (1954), 810, 830 Lawry, N. H., Simon, R. D. (1982), 61, 114 Leadbeater, B., Dodge, J. D. (1967), 690, 712 Leadbeater, B. S. C. (1986), 531, 554 Leadbeater, B. S. C. (1990), 531, 554 Leadbeater, B. S. C., Barker, D. A. N. (1995), 524, 531, 554 League, E. A., Greulach, V. A. (1955), 433, 465, 467 Leander, B. S., Farmer, M, A, (2000), 399, 403, 419 Leavitt, P. R. (1993), 781, 800 LeCampion-Alsumard, T., Golubic, S. (1985), 62, 109, 114 Lecointe, C., Coste, M., Prygiel, J. (1993), 604, 634, 791, 800 Lee, J. J., Capriulo, G. M. (1990), 383, 419 Lee, K., Round, E. F. (1987), 663, 668 Lee, K., Round, E. F. (1988), 663, 668 Lee, R. E. (1989), 424, 429, 432, 467, 757, 759, 767–768, 773 Lee, R. E. (1999), 353, 380, 384, 399, 419 Lee, R. E., Kugrens, P. (1986), 716, 734, 741–742, 745, 753 Lee, R. E., Kugrens, P. (1991), 716, 733, 735, 744, 749–750, 753 Lee, R. E., Kugrens, P. (1998), 474, 506, 738, 753 Lee, R. E., Kugrens, P. (2000), 738, 753 Lee, R. E., Kugrens, P., Mylnikov, A. P. (1991), 716, 749–750, 753 Lee, R. E., Miller-Hughes, C., Kugrens, P. (1993), 753 Lee, Y.-K., Soh, C.-W. (1991), 45, 51 Leedale, G. F. (1964), 386, 399, 403, 419 Leedale, G. F. (1967a), 406, 415, 419 Leedale, G. F. (1967b), 383–384, 387–388, 397, 399, 401–402, 404–405, 407, 410, 413, 415, 419 Leedale, G. F. (1982), 402–403, 419 Leeper, D. A., Porter, K. G. (1995), 501, 506 Lefébure, P., Cheneviére, E. (1938), 245, 250 Lefèvre, M. (1932), 690, 707, 709, 712 Leitch, A. R., John, D. M., Moore, J. A. (1990), 312, 351 Leland, H. V. (1995), 795, 800 Lembi, C. A. (1975), 235, 250 Lembi, C. A. (1980), 226, 234, 250 Lembi, C. A. (2000), 823, 830 Lembi, C. A., O’Neal, S. W., Spencer, D. F. (1988), 782–783, 800, 810, 830 Lembi, C. A., Pearlmutter, N. L., Spencer, D. L. (1980), 818, 830

Lembi, C. A., Ritenour, B. G., Iverson, E. M., Forss, E. C. (1978), 820, 830 Lembi, C. A., Spencer, D. F., O’Neal, S. W. (1984), 822, 830 Lemieux, C., Otis, C., Turmel, M. (2000), 228, 250 Lemmermann, E. (1899), 89, 114 Lemmermann, E. (1905), 91, 114 Lentin, J. K., Williams,G. L. (1989), 687, 712 Leonardson, L., Ripl, W. (1980), 812, 830 Leopold, L. B., Wolman, M. G., Miller, J. P. (1964), 28, 51 Lepisto, L., Antikainen, S., Kivinen, J. (1994), 428, 467 Lepistö, L. Rostenström, U. (1998), 489, 506 Letson, M. A., Makarewicz, J. C. (1994), 819, 830 Leukart, P., Hanelt, D. (1995), 203, 222 Levin, E. D., Schmechel, D. E., Burkholder, J. M., Glasgow, H. B., Jr., Deamer-Melia, N. J., Moser, V. C., Harry, G. J. (1997), 685, 712 Lewandowski, K., Ozimek, T. (1997), 28, 51 Lewin, R., Robinson, X. (1979), 89, 114 Lewis, I. F., Zirkle, C., Patrick, R. (1933), 461, 463, 467 Lewis, L. A. (1997), 256, 308 Lewis, L. A., Wilcox, L. W., Fuerst, P. A., Floyd, G. L. (1992), 256, 308 Lewis, W. M. (1984), 560, 568, 591 Lewitus, A. J., Caron, D. A. (1991), 740, 753 Lewitus, A. J., Glasgow, H. B., Burkholder, J. M. (1999), 690, 712, 738–739, 753 Lhotsky, O., Komárek, J. (1981), 68, 114 Li, R., Watanabe, M. M. (1998), 117, 195 Li, R., Watanabe, M., Watanabe, M. M. (2000), 195 Lichti-Federovich, S. (1979), 613, 627, 634 Likens, G. E. (1985), 19, 51 Likens, G. E., Bormann, F. H., Pierce, R. S., Eaton, J. S., Johnson, N. M. (1997), 28, 51 Lim, E. E., Amaral, L., Caron, D. A., DeLong, E. F. (1993), 473, 506 Lim, E. E., Caron, D. A., DeLong, E. F. (1996), 473, 506 Lim, E. E., Caron, D. A., Dennett, M. R. (1999), 473, 506 Lin, C. K., Blum, J. L. (1977), 26, 51, 205, 222 Lindeman, R. L. (1941a), 40, 51 Lindeman, R. L. (1941b), 40, 51 Lindeman, R. L. (1942), 40, 51 Lindström, K. (1991), 699, 712 Line, J. M., ter Braak, C. J. F., Birks, H. J. B. (1994), 792, 800 Ling, H. U. (1996), 226, 250 Ling, K. H., Sin, Y. M., Lam, T. J. (1993), 701, 712 Linnaeus, C. (1753), 312, 351 Linnaeus, C. (1758), 225, 250 Linne von Berg, K.-H., Kowallik, K. V. (1996), 12, 51 Linton, E. W., Hittner, D., Lewandowski, C., Auld, T., Triemer, R. E. (1999), 385, 387, 391, 419 Linton, E. W., Triemer, R. E. (1999a), 387, 391, 401, 419 Linton, E. W., Triemer, R. E. (1999b), 385, 391, 419 Livingstone, D., Khoja, T. M., Whitton, B. A. (1983), 118, 195 Livingstone, D., Whitton, B. A. (1984), 39, 51 Livingstone, D.A. (1963), 14, 51 Lobban, C. S., Mann, D. G. (1987), 682–683 Lock, M. A., Wallace, R. R., Costerton, J. W., Ventulloa, R. M., Charlton, S. E. (1984), 32, 51 Lock, M. A., Williams, D. D. (1981), 28, 32, 51 Lodge, D. L., Kershner, M. W., Aloi, J. E., Covich, A. P. (1994), 25, 51 Lodge, D. M. (1986), 24–25, 51 Loeb, S. L., Reuter, J. E., Goldman, C. R. (1983), 25, 51 Loeblich, A. R., III (1969), 690, 712 Loeblich, A. R., III (1976), 697, 712 Loeblich, A. R., III (1980), 708, 712 Loeblich, A. R., III (1982), 697, 712

Author Index Loginova, E. I. (1988), 582, 591 Lohman, K. E., Andrews, G. W. (1968), 561, 591 Lokhorst, G. M. (1978), 349, 351 Lokhorst, G. M. (1991), 345, 351 Lokhorst, G. M. (1992), 465, 467 Lokhorst, G. M. (1996), 346, 351 Lokhorst, G. M. (1999), 341, 351 Lokhorst, G. M., Rongen, G. P. J. (1994), 335, 351 Lom, J. (1981), 706, 712 Lorenzen, C. J. (1967), 783, 801 Losee, R. F., Wetzel, R. G. (1983), 24, 52 Lott, A. M., Siver, P. A., Marsicano, L. J., Kodama, K. P., Moeller, R. E. (1994), 537, 554 Lovell, R. T., Lelana, I. Y., Boyd, C. E., Armstrong, M. S. (1986), 807, 830 Lowe, C. W. (1927), 432–434, 458, 461, 467 Lowe, R. L. (1974), 561, 591, 638, 652, 661–662, 668, 775, 791, 801 Lowe, R. L. (1975), 582, 591 Lowe, R. L. (1996), 25, 52, 672, 683 Lowe, R. L., Busch, D. E. (1975), 587, 591 Lowe, R. L., Collins, G. B. (1973), 605, 634 Lowe, R. L., Kociolek, J. P. (1984), 659, 665, 668 Lowe, R. L., Pan, Y. (1996), 34, 52, 670, 683, 776, 801 Lowe, R. L., Rosen, B. H., Kingston, J. C. (1982), 26, 52, 605, 634 Lucas, I. A. N. (1970a), 741, 753 Lucas, I. A. N. (1970b), 735, 741, 753 Lucas, I. A. N. (1982), 742, 753 Ludwig, M., Gibbs, S. P. (1985a), 735–736, 754 Ludwig, M., Gibbs, S. P. (1985b), 736, 754 Ludwig, M., Gibbs, S. P. (1989), 735–736, 754 Lukas, K. G. Golubic, S. (1981), 62, 114 Lukas, K. G. Golubic, S. (1983), 60, 107, 114 Lukas, K. G., Hoffman, E. J. (1984), 107, 114 Lukesová, A. (1993), 63, 114 Lund, J. W. G. (1942), 408, 413, 419, 514, 520 Lund, J. W. G. (1949), 607, 634 Lund, J. W. G. (1950), 607, 634 Lund, J. W. G. (1955), 146–147, 195 Lund, J. W. G. (1962), 754 Lund, J. W. G. (1964), 20, 52 Lund, J. W. G., Kipling, C., LeCren, E. D. (1958), 490, 506, 783–784, 801 Lund, J. W. G., Reynolds, C. S. (1982), 23, 52 Lupikina, E. G., Khursevich, G. K. (1992), 587, 591 Luther, A. (1899), 432, 467 Luther, H. (1954), 764, 773 M Maceina, M. J. (1996), 810, 830 Maceina, M. J., Cichra, M. F., Betsill, R. K. Bettoli, P. W. (1992), 821, 830 MacEntree, F. J., Schreckenberg, S. G., Bold, H. C. (1972), 63, 114 MacFarlane, J. J., Raven, J. A. (1985), 203, 222 MacKay, N. A., Elser, J. A. (1998), 819, 830 MacRae, R. A., Fensome, R. A., Williams, G. L. (1996), 687, 712 Madsen, J. D. (1996), 816, 830 Maga, J. A. (1987), 807, 830 Mahmood, N. A., Carmichael, W. W. (1986), 121, 195 Mahood, A. D. (1981), 585–586, 591 Mahood, A. D., Fryxell, G. A., McMillan, M. (1986), 587, 591 Mahood, A. D., Thomas, R. D., Goldman, C. R. (1984), 583, 591 Main, S. (1988), 616, 634 Makarewicz, J. C. (1993), 21, 52, 587, 591 Makarewicz, J. C., Baybutt, R. I. (1981), 22, 52 Makarewicz, J. C., Bertram, P. (1991), 587, 591

867

Malin, G., Liss, P. S., Turner, S. M. (1994), 511, 520 Maloney, T. E., Palmer, C. M. (1956), 822, 830 Maltais, M.-J., Vincent, W. F. (1997), 24, 26, 52 Mann, D. G. (1977), 682–683 Mann, D. G. (1980a), 682–683 Mann, D. G. (1980b), 682–683 Mann, D. G. (1981), 682–683 Mann, D. G. (1982a), 659, 668 Mann, D. G. (1982b), 659, 668 Mann, D. G. (1984), 604, 634, 663, 668 Mann, D. G. (1984a), 637–638, 652 Mann, D. G. (1984b), 638, 652 Mann, D. G. (1984c), 649, 652 Mann, D. G. (1985), 637, 652 Mann, D. G. (1986), 682–683 Mann, D. G. (1989), 638, 650, 652 Mann, D. G. (1993), 560, 591, 652 Mann, D. G., Stickle, A. J. (1991), 644, 646–647, 652 Mann, N. H., Carr, N. G. (1992), 63, 114 Manny, B. A., Edsall, T. A., Wujek, D. F. (1991), 212, 222 Manton, I., Peterfi, L. S. (1969), 516, 519–520 Mantoura, R. F. C., Llewellyn, C. A. (1983), 783, 801 Marencik, J., Lembi, C. A. (1998), 822, 830 Margalef, R. (1947), 454. 467 Margalef, R. (1978), 568, 591 Margulis, L. (1974), 385, 419 Margulis, L., Corliss, J. O., Melkonian, M., Chapman, D. J. (1990), 383, 385, 387–388, 419 Marin, B., Klingberg, M., Melkonian, M. (1998), 737, 742, 754 Marino, R., Howarth, R. W., Shamess, J., Prepas, E. (1990), 23, 52 Marks, J. C., Lowe, R. L. (1993), 25–26, 52, 672, 683 Martin, A. C., Zim, H. S., Nelson, A. L. (1961), 821, 830 Martin, E., Benson, R. (1988), 819, 830 Martin, E. L., Leach, J. E., Kuo, K. J. (1978), 819, 830 Martin, J. F., Izaguirre, G., Waterstrat, P. (1991), 807, 830 Martin, J. F., McCoy, C. P., Greenleaf, W., Bennett, L. (1988), 807, 830 Martin, T. C., Wyatt, J. T. (1974), 118–120, 195 Martin, W., Sommerville, C. C., Loiseaux-de Goer, S. (1992), 737, 754 Marzoulf, E. R., Mulholland, P. J., Steinman, A. D. (1994), 785, 801 Mason, C. P., Edwards, K. R., Carlson, R. E., Pignatello, J., Gleason, F. K., Wood, J. M. (1982), 821, 830 Massalski, A., Leedale, G. F. (1969), 432, 467 Masuda, K., Boyd, C. E. (1993), 824, 831 Mataloni, G., Tell, G. (1996), 40, 52 Mattox, K. R., Stewart, K. D. (1984), 225, 250, 257, 308 Mattox, K. R., Stewart, K. D. (1985), 313, 351 Matula, J. (1992), 348, 351 Matvienko, A. M. (1941), 524, 554 Matvienko, A. M. (1954), 480, 503, 506 Maxwell, C. D. (1991), 63, 114 May, R. M. (1974), 795, 801 Mayama, S., Kobayashi, H. (1990), 665, 668 Mayama, S., Kobayashi, H. (1991), 665, 668 Mayer, M. S., Likens, G. E. (1987), 31, 52 Mazumder, A., Taylor, W. D., McQueen, D. J., Lean, D. R. S. (1990), 818, 831 McBride, S. A., Edgar, R. K. (1988), 569, 591 McCormick, P. V., Carins, J., Jr. (1994), 782, 786, 801 McCormick, P. V., O’Dell, M. B. (1996), 783, 786, 794, 801 McCormick, P. V., Rawlik, P. S., Lurding, K., Smith, E. P., Sklar, F. H. (1996), 785, 801 McCormick, P. V., Stevenson, R. J. (1998), 39, 52, 66, 114, 776, 783, 785, 794, 801

868

Author Index

McCourt, R. M. (1995), 313, 351 McCourt, R. M., Hoshaw, R. W., Wang, J. C. (1986), 363, 380 McCourt, R. M., Karol, K. G., Kaplan, S., Hoshaw, R. W. (1995), 353, 380 McFadden, G. I. (1993), 735–736, 754 McFadden, G. I., Gilson, P. R., Douglas, S. E., Cavalier-Smith, T., Hofmann, C. J. B., Maier, U.-G. (1997), 735–737, 754 McFadden, G. I., Gilson, P. R., Hill, D. R. A. (1994), 736, 754 McFarland, B. H., Hill, B. H., Willingham, W. T. (1997), 791, 801 McGrory, C. B., Leadbeater, B. S. C. (1981), 531, 554 McInteer, B. B. (1930), 461, 463, 467 McInteer, B. B. (1939), 461, 463, 467 McIntire, C. D., Phinney, H. K., Larson, G. L., Buktenica, M. (1994), 567, 584, 591 McIntire, C. D., Tinsley, I. J., Lowry, R. R. (1969), 35, 52 McIntosh, A. W., Kevern, N. R. (1974), 825, 831 McKay, R. M. L., Gibbs, S. P. (1990), 200, 222 McKenzie, C., Deibel, D., Paranjape, M., Thompson, R. J. (1995), 489, 506 McKenzie, C., Kling, H. (1989), 486, 506, 552, 554 McKerracher, L., Gibbs, S. P. (1982), 735–736, 754 McKnight, B. K., Niem, A., Kociolek, J. P., Ranne, P. (1995), 561, 591 McKnight, D. (1981), 825, 831 McKnight, D. M., Chisholm, S. W., Harleman, D. R. F. (1983), 822–824, 831 McLachlan, J. L., Boalch, G. T., Jahn, R. (1997), 709, 712 McLachlan, J. L., Curtis, J. M., Boutilier, K. Keusgen, M., Seguel, M. R. (1999), 408, 419 McLachlan, J. L., Seguel, M. R., Fritz, L. (1994), 388, 401, 408, 419 McNabb, C. D. (1960), 490, 506 McQueen, D. J. (1990), 818–819, 831 McQueen, D. J., Johannes, M. R. S., Post, J. R., Stewart, T. J., Lean, D. R. S. (1989), 818, 831 McQuoid, M. R., Hobson, L. A. (1995), 587, 591 Meador, J. P., Taub, F. B., Sibley, T. H. (1993), 822, 825, 831 Medlin, L. K., Elwood, D. J., Stickel, S., Sogin, M. L. (1991), 570, 591 Medlin, L. K., Kooistra, W. H. C. F., Gersonde, R., Sims, P. A., Wellbrock, U. (1997), 570, 592 Medlin, L. K., Kooistra, W. H. C. F., Gersonde, R., Wellbock, U. (1996), 563, 570, 583, 584, 592 Medlin, L. K., Kooistra, W. H. C. F., Potter, D., Saunders, G. W., Andersen, R. A. (1997), 473, 506 Medlin, L. K., William, D. M., Sims, P. A. (1993), 570, 592 Meffert, M. E. (1987), 128–129, 195 Meffert, M. E. (1988), 128–129, 195 Meffert, M. E., Krambeck, H. J. (1977), 129, 195 Meijer, M.-L., Hosper, H. (1997), 28, 52 Melkonian, M. (1990), 336, 351 Melkonian, M. (1996), 387, 419 Melkonian, M., Robenek, H., Reize, I. B., Preisig, H. (1987), 473, 506 Melkonian, M., Surek, B. (1995), 257, 308 Melzer, A., Haber, W., Kohler, A. (1977), 811, 831 Mensinger, A. F., Case, J. F. (1992), 687, 712 Mercado, A. (1977), 180, 195 Metzeltin, D., Lange-Bertalot, H. (1995), 659, 668 Meulemans, J. T., Heinis, F. (1983), 39, 52 Meyer, K. (1929), 664, 668 Meyer, R. L. (1969), 433–434, 446–447, 453, 459, 461, 467 Meyer, R. L. (1971), 478, 499, 501, 507 Meyer, R. L. (1995), 443, 467 Meyer, R. L., Brook, A. J. (1969), 412, 419, 491, 507, 512, 520, 698, 709, 712

Meyer, R. L., Wheeler, J. H., Brewer, J. R. (1970), 429, 432–434, 447, 449, 454, 459, 461, 463, 467 Meyer, S. R., Pienaar, R. N. (1981), 736, 754 Meyer, S. R., Pienaar, R. N. (1984a), 747, 754 Meyer, S. R., Pienaar, R. N. (1984b), 736, 747, 754 Meyer-Harms, B., Pollehne, F. (1998), 690, 712 Mickle, A. M., Wetzel, R. R. (1978), 39, 52 Mignot, J. P. (1965), 740, 744, 754 Mignot, J. P. (1967), 427–428, 468 Mignot, J. P. (1976), 427–428, 468 Mignot, J. P., Brugerolle, G. (1982), 528, 531, 554 Mignot, J. P., Brugerolle, G., Bricheux, G. (1987), 404–405, 419 Miller, A. R., Lowe, R. L., Rosenberry, J. T. (1987), 604, 634 Miller, P. E., Scholin, C. A. (1998), 257, 308 Miller, S. R., Wingard, C. E., Castenholz, R. W. (1998), 66, 114 Millie, D. R., Pearl, H. W., Hurley, J. P. (1993), 783, 801 Milliman, J. D., Meade, R. H. (1983), 28, 52 Mills, E. L., Leach, J. H., Carlton, J. T., Secor, C. L. (1993), 587, 592 Minshall, G. W. (1978), 29, 32, 52, 776, 801 Minshall, G. W., Petersen, R. C., Cummins, K. W., Bott, T. L., Sedell, J. R., Cushing, C. E., Vannote, R. L. (1983), 29, 31–32, 52 Mitchell, D.S., Pieterse, A. H., Murphy, K. J. (1989), 810, 831 Mitsch, W. J., Gosselink, J. G. (1993), 38, 52, 813, 831 Moeller, R. E., Burkholder, J. M., Wetzel, R. G. (1988), 25, 39, 52 Moestrup, Ø. (1982), 432, 468 Moestrup, Ø. (1991), 225, 228, 230, 250 Moestrup, Ø. (1994), 512, 514, 520 Moestrup, Ø. (1995), 484, 507 Moestrup, Ø., Andersen, R. A. (1995), 474, 507 Moestrup, Ø., Thomsen, H. A. (1980), 491, 507, 514, 520 Moestrup, Ø., Thomsen, H. A. (1995), 512, 520 Moestrup, Ø (1995), 524, 528, 533, 554 Moewus, F. (1940), 432, 468 Moewus, F. (1959), 235, 250 Moll, R. A., Stoermer, E. F. (1982), 567, 592 Mollenhauer, D. (1970), 180, 195 Mollenhauer, D., Mollenhauer, R. (1996), 180, 195 Mollenhauer, D., Mollenhauer, R., Kluge, M. (1996), 180, 195 Momeu, L., Péterfi, L. S. (1979), 524, 533, 554 Monastersky, R. (1998), 384, 419 Monegue, R. L., Phlips, E. J. (1991), 819, 831 Montegut-Felkner, A. E., Triemer, R. E. (1997), 387, 391, 401, 419 Montejano, G., Gold, M., Komárek, J. (1993), 100, 106–107, 114 Montejano, G., Gold, M., Komárek, J. (1997), 62, 99, 102, 114 Montoya, H.T., Golubic, S. (1991), 66, 114 Moore, B. N.(1937), 561, 592 Moore, G. T., Carter, N. (1923), 82, 114 Moore, G. T., Carter, N. (1926), 434, 451, 468 Moore, G. T., Kellerman, K. F. (1905), 822, 831 Moore, J. K., Villareal, T. A. (1996), 567, 592 Moore, J. W. (1974), 27, 52 Moore, J. W. (1979), 407, 419 Moore, J. W. (1980), 605, 634 Moore, J. W. (1981), 699–701, 712 Moore, M. V., Winner, R. W. (1989), 825, 831 Morabito, G., Curradi, M. (1997), 587, 592 Moraczewski, I. R., Zakrys, B. (1992), 419 Morgan, K., Kalff, J. (1975), 738, 754 Morisawa, M. (1968), 28, 52 Morita, E., Abem T., Tsuzuki, M., Fujiwara, S., Sato, M., Hirata, A., Sonoike, K., Nozaki, H. (1998), 234, 250 Morita, E., Abem T., Tsuzuki, M., Fujiwara, S., Sato, M., Hirata, A., Sonoike, K., Nozaki, H. (1999), 235, 250 Morrall, S., Greenwood, A. D. (1980), 734, 754 Morrall, S., Greenwood, A. D. (1982), 735, 754

Author Index Morrill, L. C., Loeblich, A. R., III (1981), 690, 712 Morris, I., (1980), 485, 507 Moser, G., Lange-Bertalot, H., Metzeltin, D. (1998), 643, 652 Moss, B. (1973), 407, 419 Moss, B., Beklioglu, M., Carvalho, L., Klinic, S., McGowan, S., Stephen, D. (1997), 23, 44, 52 Moss, D., Wright, J. F., Furse, M. T., Clarke, R. T. (1999), 788, 801 Moss, M. O., Carter, J. R. (1982), 627, 634 Moss, M. O., Gibbs, G., Gray, V., Ross, R. (1978), 665, 668 Mosser, J. L., Mosser, A. G., Brock, T. D. (1977), 23, 43, 52 Mosto-Cascallar, P. (1987), 270, 308 Mrozinska, T. (1995), 348, 351 Mühling, M., Scott, M., Harris, N., Whitton, B. A. (1997), 132, 195 Mulholland, P. J. (1996), 776, 801 Müller, D. G. (1979), 768, 773 Müller, D. G., Geller, W. (1978), 758–760, 765, 767–768, 771, 773 Müller, K. M., Sheath, R. G., Vis, M. L., Crease, T. J., Cole, K. M. (1998), 12, 34, 52, 199, 206, 210, 221–222 Müller, K. M., Sherwood, N. R., Rueschel, C. M., Gutell, R. R., Sheath, R. G. (2002), 221–222 Müller, K. M., Vis, M. L., Chiasson, W. B., Whitick, A., Sheath, R. G. (1997), 203, 205, 221–222 Müller, O. (1903), 569, 592 Müller, O. (1906), 569, 592 Müller, O. F. (1786), 471, 495, 507 Munawar, M. (1972), 406, 419 Munawar, M., Bistricki, T. (1979), 716, 734, 741, 744, 754 Munawar, M., Munawar, I. F. (1976), 806, 831 Munawar, M., Munawar, I. F. (1978), 454, 461, 468 Munawar, M. Munawar, I. F. (1981), 230, 234, 238, 250 Munawar, M., Munawar, I. F. (1982), 513, 520 Munawar, M., Munawar, I. F. (1996), 19, 21, 52 Munawar, M., Munawar, I. F. (2000), 19, 52 Munawar, M., Talling, J. F. (1986), 19, 52 Munch, C. S. (1980), 605, 634 Mundie, J. R. (1929), 432, 468 Mur, L., Bejsdorf, R. O., (1978), 23, 52 Murkin, H. R., Stainton, M. P. Boughen, J. A., Pollard, J. B., Titman, R. D. (1991), 39, 52 Murphy, K. J., Barrett, P. R. F. (1993), 823, 831 Murtaugh, P. A. (1996), 786, 801 Muylaert, K., Sabbe, K. (1996), 587, 592 N Nägeli, C. (1849), 62, 114 Nagy, J. P. (1965), 197, 208, 222 Nakamura, H. (1994), 385, 419 Nakayama, T., Watanabe, S. Inouye, I. (1996a), 230, 250 Nakayama, T., Watanabe, S. Inouye, I. (1996b), 226, 250 Nakayama, T., Watanabe, S., Mitsui, K., Uchida, H., Inouye, I. (1996), 257, 308 Nakazawa, A., Krienitz, L., Nozaki, H. (2001), 235, 248, 250 National Academy of Sciences (1976), 818, 831 National Academy of Sciences (1987), 43, 52 Nauwerck, A. (1979), 478, 507 Neale, P. J., Talling, J. F., Heaney, S. I., Reynolds, C. S., Lund, J. W. G. (1991), 20, 52 Necchi, O., Jr. (1993), 202, 204, 222 Necchi, O., Jr. (1995), 202, 222 Necchi, O., Jr. (1997), 203, 222 Necchi, O., Jr., Entwisle, T. J. (1990), 215–216, 223 Necchi, O., Jr., Sheath, R. G., Cole, K. M. (1993a), 201, 203–204, 211–213, 221, 223 Necchi, O., Jr., Sheath, R. G., Cole, K. M. (1993b), 201, 212, 221, 222

869

Necchi, O., Jr., Sheath, R. G., Cole, K. M. (1993c), 214–215, 221, 222 Neel, J. K., Peterson, S. A., Smith, W. L. (1973), 817, 831 Neely, R. K (1994), 24, 39, 52 Neil, J. H. (1975), 810, 831 Neilan, B. A., Jacobs, D., De Lot, T., Blackall, L. L., Hawkins, P. R., Cox, P. T., Goodman, E. (1997), 63, 114 Neill, C., Cronwell, J. C. (1992), 801 Newbold, J. D., Elwood, J. W., O’Neill, R. V., Van Winkle, W. (1981), 31, 52 Newman, J. R., Barrett, P. R. F. (1993), 822, 831 Newman, S., Aldridge, F. J., Phlips, E. J., Reddy, K. R. (1994), 776, 801 Nicholls, K. H. (1978), 491, 507 Nicholls, K. H. (1979), 490, 507 Nicholls, K. H. (1981a), 489, 507 Nicholls, K. H. (1981b), 478, 495, 497, 500, 507 Nicholls, K. H. (1981c), 503, 507 Nicholls, K. H. (1984a), 484, 500, 503, 507 Nicholls, K. H. (1984b), 500, 503, 507 Nicholls, K. H. (1984c), 497, 503, 507 Nicholls, K. H. (1984d), 507 Nicholls, K. H. (1985), 500, 507 Nicholls, K. H. (1987), 478, 503, 507 Nicholls, K. H. (1988), 500, 507 Nicholls, K. H. (1989), 503, 507 Nicholls, K. H. (1990), 480, 501, 503, 507 Nicholls, K. H. (1995), 487, 490, 507 Nicholls, K. H. (1996), 822, 831 Nicholls, K. H. (2000), 477, 498, 503, 507 Nicholls, K. H., Beaver, J. L., Estabrook, R. H. (1982), 807, 831 Nicholls, K. H., Carney, E. C., Robinson, G. W. (1977), 486, 507 Nicholls, K. H., Gerrath, J. F. (1985), 807, 831 Nicholls, K. H., Hopkins, G. J. (1993), 813, 831 Nicholls, K. H., Nakamoto, L., Keller, W. (1992), 488, 507 Nicholls, K. H., Standen, D. W., Hopkins, G. J. (1980), 806, 813, 831 Nicholls, K. H., Standen, D. W., Hopkins, G. J., Carney, E. C. (1977), 813, 831 Nichols, H. W. (1973), 740, 754 Nichols, H. W., Bold, H. C. (1965), 349, 351 Nichols, H. W., Nichols, M. S., Thomas, C. M., Deacon, J. S., Veith, M. (1991), 301, 309 Nichols, K. H. , Beaver, J. L., Estabrook, R. H. (1982), 512–513, 520 Nichols, K. H. , Nakamoto, L., Keller, W. (1992), 513, 520 Nichols, K. H. (1978), 518–520, 539, 554 Nichols, K. H. (1979), 512, 520 Nichols, K. H. (1982), 552, 555 Nichols, K. H. (1987), 535, 555 Nichols, K. H. (1988a), 524, 535, 552, 555 Nichols, K. H. (1988b), 535, 555 Nichols, K. H. (1995), 535–537, 555 Nichols, K. H. (1998), 693, 698, 701, 708, 712 Nichols, K. H., Beaver, J. R., Estabrook, R. H. (1982), 23, 52 Nichols, K. H., Gerrath, J. F. (1985), 526, 528, 531, 533, 535, 541, 551–552, 555 Nichols, S. A. (1973), 817, 831 Nichols, S. A. (1984), 811, 831 Niederlehner, B. R., Cairns, J. C., Jr. (1994), 801 Niiyama, Y., Watanabe, M., Umezaki, I. (1993), 141, 147, 195 Nipkow, F. (1927), 568, 592 Nipkow, F. (1950), 573, 592 Nixdorf, B., Mischke, U., Le_mann, D. (1998), 489, 507 Noonan, T. A. (1998), 818, 831

870

Author Index

Nordin, R. N., Stein, J. R. (1980), 177, 195 Norris, R. E. (1967), 114 Norris, R. E. (1980), 225, 227, 230, 250 Norris, R. E., Munch, C. S. (1970), 526, 541, 555 Northcote, T. G., Larkin, P. A. (1963), 13, 40, 52 Norton, T. A., Melkonian, M., Andersen, R. A. (1996), 410, 419 Novarino, C. (1991a), 734, 736, 741, 754 Novarino, C. (1991b), 734, 736, 741, 754 Novarino, C. (1993a), 734, 754 Novarino, C. (1993b), 734, 754 Novarino, C., Lucas, I. A. N. (1993), 734, 741–743, 754 Novarino, C., Lucas, I. A. N., Morrall, S. (1994), 734, 741, 748–749, 754 Nozaki, H. (1981), 240–241, 250 Nozaki, H. (1982), 240, 242, 250 Nozaki, H. (1983), 226, 243, 248, 250 Nozaki, H. (1986), 227, 245, 247–248, 250 Nozaki, H. (1988), 227, 241–242, 250 Nozaki, H. (1989a), 238–239, 243, 250 Nozaki, H. (1989b), 247–248, 250 Nozaki, H. (1993), 257, 309 Nozaki, H. (1994), 234, 250 Nozaki, H. (1995), 227, 240, 245, 250 Nozaki, H., Aizawa, K., Watanabe, M. M. (1994a), 233–234, 247–248, 250 Nozaki, H., Aizawa, K., Watanabe, M. M. (1994b), 227, 250 Nozaki, H., Ito, M. (1994), 226, 238, 243, 250 Nozaki, H., Ito, M., Sano, R., Uchida, H., Watanabe, M. M., Kuroiwa, T. (1995a), 226, 239, 242, 250 Nozaki, H., Ito, M., Sano, R., Uchida, H., Watanabe, M. M., Kuroiwa, T. (1996a), 234, 250 Nozaki, H., Ito, M., Sano, R., Uchida, H., Watanabe, M. M., Kuroiwa, T. (1996b), 244, 251 Nozaki, H., Ito, M., Sano, R., Uchida, H., Watanabe, M. M., Kuroiwa, T. (1997a), 226, 239, 241–243, 247, 250 Nozaki, H., Ito, M., Sano, R., Uchida, H., Watanabe, M. M., Kuroiwa, T. (1997b), 226, 239, 250 Nozaki, H., Ito, M., Sano, R., Uchida, H., Watanabe, M. M., Kuroiwa, T. (1997c), 226, 251 Nozaki, H., Krienitz, L. (2001), 251 Nozaki, H., Kuroiwa, H., Kuroiwa, T. (1994b), 227, 251 Nozaki, H., Kuroiwa, H., Mita, T., Kiroiwa, T. (1989), 226, 239, 241, 248, 251 Nozaki, H., Kuroiwa, T. (1990), 242, 248, 251 Nozaki, H., Kuroiwa, T. (1991), 241, 248, 251 Nozaki, H., Kuroiwa, T. (1992), 238–239, 241–242, 248, 251 Nozaki, H., Misawa, K. Kajita, T., Kato, M., Nohara, S., Watanabe, M. M. (2000), 239, 243–244, 251 Nozaki, H., Ohta, N., Morita, E., Watanabe, M. M. (1998b), 227, 233–234, 247, 251 Nozaki, H., Ohta, N., Takano, H., Watanabe, M. M. (1999), 239, 242–244, 251 Nozaki, H., Ohtani, S. (1992), 227, 245, 248, 251 Nozaki, H., Onishi, K., Morita, E. (2002a), 226, 235, 242, 251 Nozaki, H., Song, L.-R., Liu, Y.-D., Hiroki, M., Watanabe, M. M. (1998a), 242, 247, 251 Nozaki, H., Takahara, M., Nakazawa, A., Kita, Y., Yamada, T., Takano, H., Kawano, S., Kato, M. (2000b), 251 Nozaki, H., Watanabe, M. M. Aizawa, K. (1995b), 234, 251 Nudelman, M. A., Lombardo, R., Conforti, V. (1998), 414, 419 Nultsch, W. (1974), 119, 195 Nygaard, G. (1978), 472, 486, 503, 507, 526, 555 Nygaard, G. (1984), 139, 195 Nyström, P. Brönmark, C., Granéli, W. (1996), 26, 53

O Oakley, B. R., Bisalputra, T. (1977), 736, 754 Oakley, B. R., Dodge, J. D. (1973), 736, 754 Oakley, B. R., Dodge, J. D. (1976), 736, 754 Oakley, B. R., Heath, I. B. (1978), 736, 754 Oakley, B. R., Santore, U. J. (1982), 732, 754 Odum, E. P., Finn, J. T., Franz, E. H. (1979), 784–785, 801 O’Farrell, I. (1994), 586, 592 Ogawa, R. E., Carr, J. F. (1969), 806, 831 O’Grady, K., Brown, L. M. (1989), 513, 520 Ohtani, S. (1986), 363, 380 Ohtani, S. (1990), 379–380 O’Kelly, C. J. (1989), 432, 468 O’Kelly, C. J. Watanabe, S., Floyd, G. I. (1994), 257, 309 Oldfield, F., Appleby, P. G. (1984), 780, 801 Oliver, R. L. (1994), 805, 831 Oliver, R. L., Ganf, G. G. (2000), 12, 53, 120, 195 Olli, K. (1996), 404, 408, 419 Olrik, K. (1998), 489, 507 Olsen, J. L. (1990), 473, 507 Olsen, Y., Vadstein, O., Andersen, T., Jensen, A. (1989), 22, 53 Olson, R. K. (1993), 813, 831 Oltmanns, F. (1895), 432, 468 O’Neal, S. W., Lembi, C. A., Spencer, D. L. (1985), 812, 831 Onodera, H., Satake, M., Oshima, Y., Yasumoto, T., Carmichael, W. W. (1997), 809, 831 Ormerod, G. K. (1970), 809, 831 Ortega, M. M. (1984), 227, 232, 234–235, 241–242, 251, 411–413, 415, 419, 472, 507, 698, 712 Osorio-Tafall, B. F. (1942), 701, 712 Ostrofsky, M. L., Duthie, H. (1975), 488, 507 Ott, D. W., Brown, R. M. (1974), 432, 468 Ott, D. W., Brown, R. M. (1978), 468 Ott, D. W., Hommersand, M. H. (1974), 432, 463, 468 Ott, F. D. (1972), 206, 221, 223 Ott, F. D. (1976), 203, 208, 221, 223 Owens, J. L., Crumpton, W. G. (1995), 37, 53 Owens, K. J., Farmer, M. A., Triemer, R. E. (1988), 385, 401, 411, 415, 419 Ozimek, T., Pieczynska, E., Hankiewicz, A., (1991), 810, 831 P Paasche, E., Johansoon, S., Evenson, D. L. (1975), 565, 569, 592 Pace, M. L., Findlay, S. E. G., Links, D. (1992), 38, 53 Padan, E., Rimon, A., Ginzburg, D., Shilo, M. (1971), 819, 831 Padisák, J. (1985), 699–701, 712 Padisák, J. (1990), 174, 195 Padisák, J. (1997), 174, 195 Paerl, H. W., Bowles, N. D. (1987), 37, 53 Paerl, H. W., Priscu, J. C., Brawner, D. L. (1989), 20, 53 Paerl, H.W. , Bowles, N. D. (1987), 65, 114 Paerl, H.W. (1996), 64, 114 Paerl, H.W., Millie, D. F. (1996), 64, 114 Paine, R. T. (1980), 818, 831 Painter, D. S., Kamaitis, G. (1987), 809, 813, 831 Palamar-Mordvinsteva, G. (1976), 378, 380 Palinska, K. A., Leisack, W., Rhiel, E., Krumbein, W. E. (1996), 66, 114 Palmer, C. M. (1962), 44, 53, 776, 801, 807, 831 Palmer, C. M. (1969), 406, 419, 673, 683, 775, 787, 801 Palmer, C. M. (1980), 405–406, 419 Palmer, J. D. (1995), 686, 712 Palmer, J. D. (1996), 686, 712 Palmer, J. D., Round, F. E. (1965), 23, 53, 405, 407–408, 412, 419 Palmer, T. C. (1902), 415, 420

Author Index Pan, Y., Stevenson, R. J. (1996), 39, 53, 787, 801 Pan, Y., Stevenson, R. J., Hill, B. H., Herlihy, A. T., Collins, G. B. (1996), 784, 787, 790, 792, 794, 801 Pan, Y., Stevenson, R. J., Vaithiyanathan, P., Slate, J., Richardson, C. J. (2000), 783, 786, 790, 794–795, 801 Pandian, T. J., Marian, M. P. (1986), 35, 53 Pankow, H. (1976), 166, 195 Papenfuss, G. F. (1951), 757, 759, 773 Pappas, J. L., Stoermer, E. F. (2001a), 600, 631, 634 Pappas, J. L., Stoermer, E. F. (2001b), 600, 631, 634 Parducz, B. (1967), 716, 754 Park, H.-D., Hayashi, H. (1992), 694, 696, 712 Park, H.-D., Hayashi, H. (1993), 694, 699, 712 Park, N. E., Karol, K. G., Hoshaw, R. W., McCourt, R. M. (1996), 353, 380 Parke, M., Lund, J. W. G., Manton, I. (1962), 513, 517, 520 Parke, M., Martin, I., Clarke, S. (1955), 511, 514, 519–520 Parker, B. C., Preston, R. E., Fogg, G. E. (1963), 431, 468 Parker, B. C., Samsel, G. E., Prescott, G. W. (1973), 488–489, 499, 507 Parker, B. C., Wenkert, L. J., Parson, M. J. (1991), 18, 53 Parker, B. C., Wolfe, H. E., Howard, R. V. (1975), 15, 53 Parker, D. L. (1982), 60, 114 Pascher, A. (1910), 480, 507 Pascher, A. (1914), 312, 351, 471, 507 Pascher, A. (1917), 482, 507 Pascher, A. (1925), 481, 483–484, 507, 524, 555 Pascher, A. (1927), 233, 246, 251 Pascher, A. (1929), 483, 507 Pascher, A. (1931), 312, 351, 388, 420, 484, 507 Pascher, A. (1937-1939), 429, 432, 465, 468 Pascher, A. (1939), 503, 507 Pascher, A. (1940a), 478–479, 507 Pascher, A. (1940b), 479, 507 Patrick, R. (1945), 604, 634 Patrick, R. (1949), 775, 801 Patrick, R. (1959), 638, 652 Patrick, R. (1961), 561, 592 Patrick, R. (1973), 784, 801 Patrick, R. (1976), 605, 634 Patrick, R., Freese, L. (1961), 561, 592, 604, 628, 634 Patrick, R., Hohn, M., Wallace, J. H. (1954), 655, 658, 668, 775–776, 780, 784, 801 Patrick, R., Reimer, C. W. (1966), 561–562, 592, 596–601, 605, 628, 634, 637–639, 646, 649, 652, 655, 661–662, 664–665, 668, 670, 676, 683 Patrick, R., Reimer, C. W. (1975), 561–562, 592, 639, 650, 652, 655–656, 661–662, 668, 681–683 Patrick, R., Roberts, N. A. (1979), 561, 592 Patterson, D. J. (1981), 736, 754 Patterson, D. J. (1989), 423, 468 Patterson, D. J., Hedley, S. (1992), 385, 410, 420 Paulsen, S. G., Larsen, D. P., Kaufmann, P. R., Whittier, T. R., Baker, J. R., Peck, D. V., McGue, J., Hughes, R. M., McMullen, D., Stevens, D., Stoddard, J. L., Larzorchak, J., Kinney, W., Selle, A. R., Hjort, R. (1991), 788, 801 Pavoni, M. (1963), 480, 507 Pearlmutter, N., Lembi, C. A. (1986), 823, 832 Pearsall, W. H. (1929), 691, 703, 712 Pejler, B. (1977), 739, 754 Pennak, R. W. (1966), 811, 832 Pennak, R. W. (1973), 811, 832 Pennak, R. W. (1989), 385, 388, 420 Pennick, D. L. (1981), 734, 742, 754 Penno, S., Campbell, L., Hess, W. R. (2000), 384, 420

871

Pentecost, A. (1982), 44, 53 Pentecost, A. (1991), 364, 380 Pentecost, A., Whitton, B. A. (2000), 44, 53, 119, 121, 195 Perakis, S. S., Welch, E. B., Jacoby, J. M. (1996), 815, 832 Perasso, L., Brett, S. J., Wetherbee, R. (1993), 736, 754 Perez, C. C., Roy, S., Levasseur, M., Anderson, D. M. (1998), 696, 712 Perez, M. C., Bonilla, S., deLe\’f3n, L., \’8amarda, J., Komáek, J. (1999), 177–178, 195 Pernthaler, J., Simek, K., Sattler, B., Schwarzenbacher, A., Bobkova, J. Psenner, R. (1996), 20, 53 Perrin, C. J., Bothwell, M. L., Slaney, P. A. (1987), 604, 634 Perty, M. (1849), 413, 420 Perty, M. (1852), 413, 420 Péterfi, L. S. (1969), 477, 507 Péterfi, L. S. (1996), 555 Péterfi, L. S., Momeu, L. (1977), 533–534, 555 Peters, M. C., Andersen, R. A. (1993), 480, 484, 503, 508 Petersen, J. B., Hansen, J. B. (1956), 526, 533, 552, 555 Petersen, J. B., Hansen, J. B. (1958), 526, 533–534, 555 Peterson, C. G. (1996), 36, 53, 605, 634 Peterson, C. G., Grimm, N. B. (1992), 674, 683 Peterson, C. G., Hoagland, K. D. (1990), 623, 634 Peterson, C. G., Stevenson, R. J. (1989), 37, 53 Peterson, C. G., Stevenson, R. J. (1992), 623, 634, 779, 801 Peterson, H. G., Healey, F. P., Wagemann, R. (1984), 823, 832 Pettersson, K., Herlitz, E., Istvanovics, V. (1993), 23, 53 Petts, G., Calow, P. (1996), 28, 53 Pfiester, L. A. (1975), 694, 712 Pfiester, L. A. (1976), 694, 712 Pfiester, L. A. (1977), 694, 712 Pfiester, L. A. (1984), 694, 712 Pfiester, L. A., Anderson, D. M. (1987), 694, 712 Pfiester, L. A., Highfill, J. F. (1993), 697, 707, 713 Pfiester, L. A., Holt, J. R. (1978), 739, 754 Pfiester, L. A., Lynch, R. A., Skvarla, J. J. (1980), 694, 696, 708, 713 Pfiester, L. A., Popovsky, J. (1979), 696, 708–709, 713 Pfiester, L. A., Skvarla, J. J. (1979), 694, 713 Pfiester, L. A., Skvarla, J. J. (1980), 694, 699, 713 Pfiester, L. A., Timpano, P., Skvarla, J.J., Holt, J. R. (1984), 694, 713 Philipose, M. T. (1982), 384, 402, 404–407, 412–413, 420 Philipose, M. T. (1984), 413, 420 Philipose, M. T. (1988), 416, 420 Phillips, D., Boyne, A. E. (1984), 734, 754 Phillips, G. L., Eminson, D., Moss, B. (1978), 27, 53, 810, 832 Phlips, E. J., Monegue, R. L., Aldridge, F. J. (1990), 819, 832 Phosphorous Management Strategies Task Force (1980), 813, 832 Pick, F. R., Cuhel, R. L. (1986), 535, 555 Pick, F. R., Lean, D. R. S. (1987), 64, 114 Pick, F. R., Nalewajko, C., Lean, D. R. S. (1984), 17, 53, 535, 555 Pickett-Heaps, J. D. (1975), 257, 309, 313, 351 Pickett-Heaps, J. D., Staehelin, L. (1975), 254, 309 Pielou, E. C. (1984), 788, 790, 801 Pienaar, R. N. (1976), 739, 754 Pienitz, R., Lortie, G., Allard, M. (1991), 604, 634 Pienitz, R., Smol, J. P. (1992), 604–605, 634 Pierce, S. (1987), 15, 53 Pillinger, J. M., Cooper, J. A., Ridge, I. (1994), 822, 832 Pillinger, J. M., Cooper, J. A., Ridge, I., Barrett, P. R. F. (1992), 822, 832 Pillsbury, R. W., Kingston, J. C. (1990), 607, 634 Pinder, A. W., Friet, S. C. (1994), 45, 53 Pine, R. T., Anderson, L. W. J. (1991), 820, 832 Pipes, L. D., Leedale, G. F. (1992), 526, 548, 555 Pipes, L. D., Leedale, G. F., Tyler, P. A. (1991), 524, 528, 555

872

Author Index

Pipp, E., Rott, E. (1994), 204, 223 Pithart, D., Pechar, L., Mattsson, G. (1997), 428, 468 Pizarro, H. (1995), 270, 309, 465, 468 Plafkin, J. L., Barbour, M. T., Porter, K. D., Gross, S. K., Hughes, R. M. (1989), 786, 801 Platt, T., Li, W. K. W. (1986), 60, 65, 114 Playfair, G. (1921), 415, 420 Playfair, G. I. (1914), 247, 251 Pochmann, A. (1942), 416, 420 Pocock, M. A. (1954), 242–243, 251 Pocock, M. A. (1955), 243–244, 251 Pocock, M. A. (1960), 231–232, 251 Pohlman, J. W., Iliffe, T. M., Cifuentes, L. A. (1997), 44, 53 Pollinger, U. (1981), 738, 754 Pollinger, U. (1986), 513, 520 Pollingher, U. (1987), 20, 53, 699–700, 713 Pollingher, U., Burgi, H. R., Ambühl, H. (1993), 694, 699–700, 713 Pollingher, U., Hickel, B. (1991), 699–700, 713 Polunin, N. (1954), 433, 463, 468 Popovsky, J. (1970), 698, 702, 713 Popovsky, J. (1982), 708, 713 Popovsky, J., Pfiester, L. A. (1990), 698, 708, 713 Porter, K. G. , McDonough, R. (1984), 808, 832 Porter, K. G. (1973), 819, 832 Porter, K. G. (1977), 783, 801 Porter, K. G. (1988), 20, 23, 53 Porter, K. G., Cuffney, T. F., Gurtz, M. E., Meador, M. R. (1993), 779–780, 784, 801 Potter, D., Saunders, G. W., Andersen, R. A. (1997), 424, 468 Potter, M. C. (1888), 351 Poulin, M. E. (1990), 604, 634 Poulin, M., Hamilton, P. B., Proulx, M. (1995), 587, 592, 698, 713 Poulton, E. M. (1930), 424–425, 429, 432–433, 441, 459, 461, 463, 468 Power, M. E. (1990), 674, 683, 776, 783, 801, 810, 832 Power, M. E., Stewart, A. J. (1987), 36, 53 Power, M. E., Stout, R. J., Cushing, C. E., Harper, P. P., Hauer, F. R., Matthews, W. J., Moyle, P. B., Statzner, B., Wais De Badgen, I. R. (1988), 32, 53 Prasad, A. K. S. K., Nienow, J. A., Livingston, R. J. (1990), 565, 592 Prát, S. (1929), 121, 141, 144, 195 Preisendorfer, R. W. (1996), 783, 801 Preisig, H. R. (1986), 485, 508 Preisig, H. R. (1989), 432, 468 Preisig, H. R. (1995), 471, 508 Preisig, H. R. (1999), 257, 309 Preisig, H. R., Anderson, O. R., Corliss, J. O., Moestrup, Ø., Powell, M. J., Roberson, R. W., Wetherbee, R. (1994), 399, 420 Preisig, H. R., Hibberd, D. J. (1982a), 503, 508, 533, 555 Preisig, H. R., Hibberd, D. J. (1982b), 503, 508, 533, 555 Preisig, H. R., Hibberd, D. J. (1983), 523, 555 Preisig, H. R., Hibberd, D. J. (1986), 523, 555 Preisig, H. R., Hibberd, D. J. (1987), 503, 508 Preisig, H. R., Melkonian, M. (1984), 231, 247, 251 Preisig, H. R., V\’f8rs, N., Hällfors, G. (1991), 474, 508 Prepas, E. E., Murphy, R. P., Crosby, J. M., Walty, D. T., Lim, J. T., Babin, J., Champers, P. A. (1990), 814, 832 Prepas, E. E., Trew, D. O, (1983), 812, 832 Prescott, G. W. (1931), 412, 415, 420, 429, 432–434, 439, 449, 454, 458, 461, 463, 468 Prescott, G. W. (1942), 244, 251 Prescott, G. W. (1944), 432, 441, 468 Prescott, G. W. (1951), 126, 137, 141, 151, 155, 177, 195, 323, 325, 327–328, 348, 351, 688, 693, 708, 713 Prescott, G. W. (1953), 433, 463, 468

Prescott, G. W. (1955), 239, 251, 391, 404, 412, 420 Prescott, G. W. (1962), 6, 9, 12, 40, 53, 60, 63, 67, 72, 80–82, 87–88, 92, 95, 114, 118, 139, 141, 150, 155, 171, 174, 180–181, 184, 189, 195, 307, 309, 313, 348, 351, 369, 380, 397– 399, 405, 413, 415, 420, 429, 442, 457, 468, 496, 499, 503, 508, 562, 592, xvi Prescott, G. W. (1963), 463, 468 Prescott, G. W. (1978), 253, 258, 268–269, 271, 275, 279, 285–286, 293–294, 297, 299–300, 305, 307, 309, 384, 388, 405, 407–415, 420, 425–429, 432, 434, 439–441, 443, 445, 450– 452, 458, 463, 468 Prescott, G. W. (1983), 315, 325–326, 329, 332, 345–346, 348, 351 Prescott, G. W. (1984), 388, 420 Prescott, G. W., Bicudo, C., Vinyard, W. C. (1982), 377–379, 381 Prescott, G. W., Croasdale, H. T. (1937), 86, 114 Prescott, G. W., Croasdale, H. T. Vinyard, W. C. (1972), 353, 363, 369–372, 379–380 Prescott, G. W., Croasdale, H. T. Vinyard, W. C. (1975), 372–373, 375, 378–379–380 Prescott, G. W., Croasdale, H. T. Vinyard, W. C. (1977), 375–377, 379–380 Prescott, G. W., Croasdale, H. T. Vinyard, W. C., Bicudo, C. (1981), 373–374, 376–377, 379–380 Prescott, G. W., Dillard, G. E. (1979), 443, 451, 454, 458, 461, 463, 468, 561, 592, 604, 634 Prescott, G. W., Vinyard, W. C. (1965), 89, 114, 432–433, 443, 446, 453–454, 458–459, 461, 468 Pringle, C. M., Naiman, R. J., Bretschko, G., Karr, J. R., Oswood, M. W., Webster, J. R., Welcomme, R. L., Winterbourn, M. J. (1988), 32, 53 Pringle, C. M., Rowe, G., Triska, F. J., Fernandez, J. F., West, J. (1993), 41–42, 53 Pringle, C. W., Bowers, J. A. (1984), 801 Pringsheim, E. G. (1946), 68, 114 Pringsheim, E. G. (1948), 391, 399, 420 Pringsheim, E. G. (1953a), 391, 414–416, 420 Pringsheim, E. G. (1953b), 391, 414–416, 420 Pringsheim, E. G. (1955), 481, 508 Pringsheim, E. G. (1956), 388, 391, 402–403, 406, 412, 415, 420 Pringsheim, E. G. (1960), 247, 251 Pringsheim, E. G. (1963), 387, 420 Pringsheim, E. G. (1968), 738, 742, 754 Pringsheim, E. G., Wiessner, W. (1960), 227, 251 Prinsep, M. R., Caplan, F. R., Moore, R. E., Patterson, G. M. L., Honkanen, R. E., Boynton, A. L. (1992), 121, 195 Printz, H. (1964), 348, 351 Pröschold, T., Marin, B., Schlösser, U. G., Melkonian, M. (2001), 235, 248, 251 Provasoli, L. (1958), 406, 420 Provasoli, L. (1961), 420 Provasoli, L. (1968), 465, 468 Provasoli, L. (1969), 406, 420 Provasoli, L., Pintner, I. J. (1953), 406, 420 Prowse, G. A. (1959), 24, 53 Prowse, G. A. (1962), 526, 534, 555 Prygiel, J., Coste, M., Bukowska, J. (1999a), 776, 791, 802 Prygiel, J., Whitton, B. A., Bukowska, J. (1999b), 776, 791, 802 Pueschel, C. M. (1990), 41, 53, 200, 223 Pueschel, C. M., Sanders, G. W., West, J. A. (2000), 201, 223 Pueschel, C. M., Stein, J. R. (1983), 757–759, 765, 770, 773 Pueschel, C. M., Sullivan, P. G., Titus, J. E. (1995), 33, 53, 218, 223 Putt, M. (1990), 739, 754 Puytorac, P., Mignot, J. P., Grain, J., Groliere, C. A., Bonner, L., Couillard, P. (1972), 526, 541, 555

Author Index Q Quade, H. W. (1969), 811, 832 Quirós, R. (1995), 818, 832 R Radwan, S., Kowalik, W., Kowalczyk, C. (1990), 785, 802 Rae, R., Vincent, W. F. (1998), 37–38, 53 Raman, R. K. (1985), 806, 822, 832 Ramanthan, K. R. (1964), 342, 349, 351 Ranch, D. C. (1981), 347, 351 Raschke, R. L. (1993), 784, 802 Rashash, D. M. C., Dietrich, A. M., Hoehn, R. C., Parker, B. C. (1995), 487, 508 Raven, J. (1992), 314, 351 Raven, J. A. (1993), 202, 223 Raven, J. A., Johnston, A. M., Newman, J. R., Scrimgeour, C. M. (1994), 204, 223 Rawlence, D. J. (1988), 604, 634 Rawson, D. S. (1956), 454, 461, 468 Rawson, D. S., Moore, A. J. (1944), 42, 53 Rayburn, W. R., Starr, R. C. (1974), 242, 248, 251 Reavie, E. D., Smol, J. P. (1997), 789, 792–793, 802 Reavie, E. D., Smol, J. P. (1998), 789, 792–793, 802 Reavie, E. D., Smol, J. P., Carignan, R., Lorrain, S. (1998), 787, 789, 793, 802 Reavie, E. D., Smol, J. P., Carmichael, N. B. (1995), 604, 635, 806, 832 Redhead, K., Wright, S. J. L. (1978), 819, 821, 832 Reed, R. H., Warr, S. R. C., Kerby, N. W., Stewart, W. D. P. (1986), 65, 114 Reichardt, E. (1999), 655, 659, 661, 668 Reichardt, E., Lange-Bertalot, H. (1991), 665, 668 Reid, M. A., Tibbey, J. C., Penny, D., Gell, P. A. (1995), 776, 802 Reimer, C. W. (1954), 682–683 Reimer, C. W. (1959), 638, 649, 652 Reimer, C. W. (1961), 638, 652 Reimer, C. W. (1966), 635 Reimer, C. W. (1990), 605, 635 Reinersten, H. (1982), 513, 520 Reinhard, E. G. (1931), 36, 53, 65, 114 Reinke, D. C. (1979), 415, 420 Rejmánková, E. Roberts, D. R., Manguin, S., Pope, K. O., Komárek, J., Post, R. A. (1996), 66, 114 Renberg, I. (1976), 599, 613, 635 Renberg, I. (1977), 599, 613, 635 Renberg, I., Hellberg, T. (1982), 791, 802 Rengefors, K., Anderson, D. M. (1998), 696, 713 Reuter, J. E., Loeb, S. L., Goldman, C. R. (1986), 26, 53 Reuter, J. E., Rhodes, C. L., Lebo, M. E., Klotzman, M., Goldman, C. R. (1993), 43, 53, 812, 832 Reynolds, C. S. (1980), 738, 754 Reynolds, C. S. (1982), 738, 755 Reynolds, C. S. (1984), 258, 309, 595, 604, 607, 635, 739, 755 Reynolds, C. S. (1984a), 19–20, 23, 54 Reynolds, C. S. (1984b), 19–20, 54 Reynolds, C. S. (1988), 36–38, 54 Reynolds, C. S. (1995), 37–38, 54 Reynolds, C. S. (1996), 32, 54 Reynolds, C. S., Descy, J.-P. (1996), 36–38, 54, 65, 114 Reynolds, C. S., Jawarski, G. H. M., Roscoe, J. V., Hewitt, D. P., George, D. G. (1998), 487, 508 Reynolds, C. S., Jaworski, G. H. M., Cmiech, H. A., Leedale, G. F. (1981), 64, 114 Reynolds, C. S., Walsby, A. E. (1975), 59, 64, 92, 114 Rhee, G. Y., Goldman, I. J. (1980), 22, 54

873

Rhodes, R. G., Stofan, P. E. (1967), 429, 468 Rhodes, R. G., Terzis, A. J. (1970), 461, 468 Rhodes, T. E., Davis, R. B. (1995), 587, 592 Rice, E. L. (1984), 821, 832 Richards, C. B. (1962), 709, 713 Rieth, A. (1980), 429, 431, 465, 468 Riley, E. T., Prepas, E. E. (1984), 814, 832 Rindi, F., Guiry, M. D. Barbiero, R. (1999), 342, 349, 351 Rindi, F., Guiry, M. D., Barbiero, R. P., Cinelli, F. (1991), 45, 54 Rintoul, T. C. Sheath, R. G., Vis, M. L. (1999), 212, 221, 223 Rippka, R., Deruelles, J. B., Waterbury, J. B., Herdman, M., Stanier, R. Y. (1979), 60, 62, 114, 117–118, 120, 195 Robarts, R. D. Evans, M. S., Arts, M. T. (1992), 43, 54 Robbins, R. W., Glicker, D. M., Bloem, D. M., Niss, B. M. (1991), 813, 832 Roberts, D. A., Boylen, C. W. (1988), 27, 54 Roberts, D. A., Boylen, C. W. (1989), 27, 54 Roberts, K. R. (1984), 734, 744–745, 755 Roberts, K. R., Stewart, K. D., Mattox, K. R. (1981), 734–735, 746, 755 Robinson, G. G. C. (1983), 25, 54 Robinson, G. G. C., Gurney, S. E., Goldsborough, L. G. (1997), 38, 54 Robinson, P.K., Hawkes, H. A. (1986), 810, 832 Roijackers, R. M. M. (1983), 487, 508 Roijackers, R. M. M., Kessels, H. (1986), 489, 508, 536, 538, 555 Rojo, C., Cobelas, M. a., Arauzo, M. (1994), 36, 54 Röpstorf, P., Hülsmann, N., Hausmann, K. (1994), 708, 713 Rosa, R., Burns, N. M. (1987), 806, 832 Rosemarin, A. S. (1975), 36, 54 Rosemond, A. D. (1993), 205, 223 Rosen, B. H. (1995), 776, 779, 802 Rosen, B. H., Lowe, R. L. (1981), 599, 616, 635 Rosen, G. (1981), 364, 381 Rosenberg, M. (1941), 508 Rosgen, D. L. (1994), 779, 802 Rosine, W. N. (1955), 811, 832 Rosowski, J. R. (1977), 401–404, 420 Rosowski, J. R., Couté, A. (1996), 415, 420 Rosowski, J. R., Glider, W. V. (1977), 402, 404, 406, 420 Rosowski, J. R., Hoagland, K. D., Roemer, S. C., Lee, K. W. (1981), 414, 420 Rosowski, J. R., Hoshaw, R. M. (1970), 410, 420 Rosowski, J. R., Hoshaw, R. M. (1971), 402, 420 Rosowski, J. R., Kugrens, P. (1973), 393, 403–405, 408, 411, 415, 420 Rosowski, J. R., Langenberg, W. G. (1994), 415, 420 Rosowski, J. R., Lee, K. W. (1978), 384, 396, 399, 411–412, 420 Rosowski, J. R., Vadas, R. L. Kugrens, P. (1975a), 414, 420 Rosowski, J. R., Walne, P. L., West, L. K. (1975b), 414, 420 Rosowski, J. R., Wiley, R. L. (1975), 388, 394, 404, 408, 411, 420 Rosowski, J. R., Wiley, R. L. (1977), 392, 403, 420 Ross, R. (1954), 432–433, 454, 461, 468 Ross, R. (1983), 561, 592 Ross, R., Cox, E. J., Karayeva, N. L., Mann, D. G., Paddock, T. B. B., Simnsen, R., Sims, P. A. (1979), 592 Rother, J. A., Fay, P. (1977), 120, 195 Rothhaupt, K. O. (1991), 808, 832 Rott, E. (1983), 738, 755 Rott, E., Pfister, P. (1988), 29, 54 Rott, E., Pipp, E. (1999), 121, 195 Round, F. E. (1963), 353, 381 Round, F. E. (1971), 21, 54, 353, 381 Round, F. E. (1972), 22, 26, 54, 568, 592 Round, F. E. (1981), 11–12, 19–20, 24, 32, 43–44, 54, 433, 468, 485, 508, 578, 592, 669, 683

874

Author Index

Round, F. E. (1984), 253, 309, 351, 407, 420 Round, F. E. (1991), 776, 802 Round, F. E. (1997), 597, 635 Round, F. E. (1998), 596, 627, 635 Round, F. E., Basson, P. W. (1997), 598, 603, 627, 635 Round, F. E., Bukhtiyarova, L. (1996), 561, 592 Round, F. E., Bukhtiyarova, L. (1996a), 597–598, 602–603, 623, 626–628, 635 Round, F. E., Bukhtiyarova, L. (1996b), 597, 602, 604, 635 Round, F. E., Crawford, R. M., Mann, D. G. (1990), 559–565, 568, 572, 575, 583–584, 586–587, 592, 596–598, 600–601, 603–604, 607, 635, 637–638, 645–646, 648–649, 652, 655, 662–663, 665, 668, 669–670, 672, 676, 681–683 Round, F. E., Häkanson, H.. (1992), 584, 592 Round, F. E., Maidana, N. I. (2001), 600, 631, 635 Round, F. E., Mann, D. G. (1981), 645, 652 Round, F. E., Palmer, J. D. (1966), 405, 407–408, 412, 420 Round, F. E., Sims, P. A. (1981), 563, 592 Roussomoustakaki, M., Anagnostidis, K. (1991), 61, 66, 114 Rout, J., Gaur, J. P. (1990), 779, 802 Rowell, H. C. (1996), 42–43, 54 Rudi, K., Skulberg, O. M., Jakobsen, K. S. (1998), 62–63, 115, 117, 195 Rudi, K., Skulberg, O. M., Skulberg, R., Jakobsen, K. S. (2000), 117, 195 Rumeau, A., Coster, M. (1988), 787, 802 Rumpf, R. R., Vernon, d., Schreiber, D., Birky, C. W., Jr. (1996), 235, 251 Ruse, L., Love, A. (1997), 37, 54 Ruse, L. P., Hutchings, A. J. (1996), 37–38, 54 Rushforth, S. R., Johansen, J. R. (1986), 575, 592 Ruzicka, J. (1977), 353, 361, 364, 381 S Sabater, S., Gregory, S. V., Sedell, J. R. (1998), 36, 54, 314, 351 Safferman, R. S., Morris, M. E. (1963), 819, 832 Safferman, R. S., Schneider, I. R., Steare, R. L., Morris, M. E., Diener, T. O. (1969), 819, 832 Saha, L. C., Wujek, D. E. (1990), 485, 508, 536, 538, 555 Sako, Y., Ishida, Y., Kadota, H., Hata, Y. (1984), 699, 713 Sako, Y., Ishida, Y., Kadota, H., Hata, Y. (1985), 694, 696, 713 Sako, Y., Ishida, Y., Kadota, H., Hata, Y. (1987), 694, 699, 713 Sala, S. E., Guerrero, J. M., Ferrario, M. E. (1993), 659, 668 Salonen, K., Jokinen, S. (1988), 489, 508, 534, 555 Salonen, K., Jones, R., Arvola, L. (1984), 23, 54 Sanchez, I., Lee, G. F. (1978), 825, 832 Sand-Jensen, K., Pedersen, O., Gertz-Hansen, O. (1997), 45, 54 Sand-Jensen, K., Søndergaard, M. (1981), 24, 54 Sanders, L., Hoover, J. J., Killgore, K. J. (1991), 820, 832 Sanders, R. W., Porter, K. G., Caron, D. A. (1990), 473, 508 Sanderson, B. L., Frost, T. M. (1996), 700, 713 Sandgren, C. D. (1980a), 485, 508 Sandgren, C. D. (1980b), 485, 508 Sandgren, C. D. (1981), 485, 508, 531, 555 Sandgren, C. D. (1983), 485, 488, 508 Sandgren, C. D. (1988), 19, 54, 485, 490, 508, 531, 535–539, 555 Sandgren, C. D. (1989), 533, 555 Sandgren, C. D. (1991), 531, 533, 555 Sandgren, C. D., Carney, H. J. (1983), 533, 555 Sandgren, C. D., Flanagin, J. (1986), 485, 508, 531, 539, 555 Sandgren, C. D., Smol, J. P., Kristiansen, J. (1995), 503, 508 Sandgren, C. D., Walton, W. E. (1995), 535, 537–538, 555 Santore, U. J. (1977), 734, 741, 744, 755 Santore, U. J. (1978), 736, 755 Santore, U. J. (1982a), 741, 755

Santore, U. J. (1982b), 741, 755 Santore, U. J. (1982c), 735, 742, 755 Santore, U. J. (1983), 741, 755 Santore, U. J. (1984), 735–736, 741, 744, 746, 755 Santore, U. J. (1987), 735, 741–742, 755 Santore, U. J., Greenwood, A. D. (1977), 735, 755 Santos, L. M. A. (1996), 424, 465, 468 Santos, L. M. A., Leedale, G. F. (1993), 538, 555 Santos, L. M. A., Melkonian, M., Kreimer, G. (1996), 424, 468 Sarjeant, W. A. S. (1978), 687, 713 Sarma, P. (1986), 351 Sarnelle, O. (1993), 739, 755 Sarojini, Y. (1994), 406, 421 Saunders, G. W., Hill, D. R. A., Sexton, J. P., Andersen, R. A. (1997), 697, 713 Saunders, G. W., Potter, D., Paskind, M. P., Andersen, R. A. (1995), 473, 508 Saunders, R. D., Dodge, J. D. (1984), 701, 713 Savino, J. E., Stein, R. A. (1982), 810, 832 Say, P. J., Burrows, J. G., Whitton, B. A. (1990), 785, 802 Say, P. J. Whitton, B. A. (1980), 407, 412, 421 Schagerl, M., Angeler, D. (1998), 244, 251 Schalles, J. F., Shure, D. F. (1989), 17, 40, 54 Schardt, J. D. (1994), 809, 811, 832 Schelske, C. L. (1975), 812, 832 Schelske, C. L., Carrick, H. J., Aldridge, F. J. (1995), 23, 54 Schelske, C. L., Stoermer, E. F. (1971), 567, 584, 592, 806, 812, 832 Schelske, C. L., Stoermer, E. F. (1972), 567, 584, 592 Schiller, J. (1933-1937), 709, 713 Schiller, J. (1956), 66, 93, 95, 115 Schindler, D. W. (1975), 806, 832 Schindler, D. W. (1977), 23, 54, 806, 812, 832 Schindler, D. W. (1978), 23, 54, 812, 818, 832 Schindler, D. W. (1985), 23, 54 Schindler, D. W. (1990), 795, 802 Schindler, D. W., Frost, V. E., Schmidt, R. V. (1973), 26, 54 Schindler, D. W., Kling, H., Schmidt, R. V., Prokopowich, J., Frost, V. E., Reid, R. L., Capel, M. (1973), 486, 508, 537, 555 Schlichting, H. E., Jr. (1960), 407, 421 Schlichting, H. E., Jr. (1964), 407, 421 Schloesser, R. E. (1977), 759, 765, 773 Schloesser, R. E., Blum, J. L. (1980), 758–759, 763, 765, 767, 769, 771, 773 Schmid, A. M. (1994), 565, 567, 569, 592 Schmid, A. M. (1997), 596, 613, 635 Schmidle, W. (1902), 481, 508 Schmidt, A. (1994), 37, 54 Schmidt, M. (1899), 664, 666, 668 Schneider, C. W., Lane, C. E. (2000), 463, 468 Schneider, C. W., Lane, C. E., Norland, A. (1999), 433, 465, 468 Schneider, C. W., MacDonald, L. A., Cahill, J. F., Heminway, S. W. (1993), 433, 465, 468 Schnepf, E., Elbrächer, M. (1992), 739, 755 Schnepf, E., Melkonian, M. (1990), 739, 755 Schnepf, E., Niemann, A., Christian, W. (1996), 427, 449. 468 Schnepf, E., Winter, S., Mollenhauer, D. (1989), 739, 755 Scholz, R. J. (1994), 814, 832 Schönbohn, W. (1972), 354, 381 Schoonoord, M. P., Ellis-Adams, A. C. (1984), 485, 508 Schrader, K. K., Blevins, W. T. (1993), 807, 832 Schrader, K. K., de Regt, M. Q., Tucker, C. S., Duke, S. O. (1997), 807, 833 Schultz, M. E., Trainor, F. R. (1968), 569, 592 Schulz, V. P. (1929), 587, 592

Author Index Schulz-Baldes, M., Lewin, R. A. (1976), 115 Schumacher, G. J., Bellis, V. J., Whitford, L. A. (1963), 432–434, 453–454, 457, 459, 468 Schumacher, G. J., Kim, Y. C., Whitford, L. A., Dillard, G. E. (1966), 424–425, 427, 446, 454, 458–461, 468 Schumacher, G. J., Whitford, L. A. (1965), 203, 223 Schumm, S. A. (1977), 29, 54 Schuster, F.L. (1968), 744, 755 Schuster, F.L. (1970), 734, 746, 755 Schwartzbach, S. D. Osafune, T., Löffelhardt, W. (1998), 385, 391, 421 Schwarz, A.-M., Hawes, I. (1997), 27, 54 Scott, J. (1983), 201, 223 Seckbach, J. (1991), 41, 54, 201, 221, 223 Sedell, J. R., Riley, J. E., Swanson, F. J. (1989), 32, 54 Segal, S. (1969), 45, 54 Selva, J. (1981), 664, 668 Senna, P. A. C., Komárek, J. (1998), 147–148, 195 Senna, P. A. C., Peres, A. C., Komárek, J. (1998), 83, 115 Serieyssol, K. K. (1981), 582, 592 Serieyssol, K. K., Garduno, I. I., Gasse, F. (1998), 587, 592 Serieyssol, K. K., Theriot, E. C., Gasse, F. (1996), 584 Serruya, C., Pollingher, U. (1983), 15, 54 Seshadri, C. V., Jeeri-Bai, N. (1992), 121, 195 Sespenwol, S. (1973), 735, 741, 746, 755 Setchell, W. A. (1924), 110, 115 Setchell, W. A., Gardner, N. L. (1905), 115 Setchell, W. A., Gardner, N. L. (1918), 150, 195 Setchell, W. A., Gardner, N. L. (1919), 78, 110, 115 Setchell, W. A., Gardner, N. L. (1924), 96, 115, 150, 195 Setchell, W. A., Gardner, N. L. (1930), 101, 107, 115 Seto, R. (1977), 202, 223 Shannon, C. F. (1948), 784, 787, 802 Shannon, E. E., Brezonik, P. L. (1972), 16, 54 Shapiro, J. (1980), 818, 833 Shapiro, J., LaMarra, V., Lynch, M. (1975), 818, 833 Sheath, R. G. (1984), 4, 9, 197–203, 214–215, 217, 219, 223 Sheath, R. G. (1986), 14, 19, 54 Sheath, R. G. (1987), 204, 206, 223 Sheath, R. G., Burkholder, J. M. (1985), 779, 783–784, 802 Sheath, R. G., Burkholder, J. M., Hambrook, J. A. Hogeland, A. M., Hoy, E., Kane, M. E., Morison, M. O., Steinman, A. D., Van Alstyne, K. L. (1986), 205, 223 Sheath, R. G., Cole, K. A. (1992), 339, 342, 351 Sheath, R. G., Cole, K. M. (1980), 26, 34, 54 Sheath, R. G., Cole, K. M. (1984), 200, 205, 223 Sheath, R. G., Cole, K. M. (1990), 200, 223 Sheath, R. G., Cole, K. M. (1992), 33–34, 45, 55, 141, 195, 206, 223, 765–766, 773 Sheath, R. G., Hambrook, J. A. (1988), 200, 203, 223 Sheath, R. G., Hambrook, J. A. (1990), 4–5, 9, 33, 55, 197–200, 202–206, 215–216, 223, 758, 773 Sheath, R. G., Hambrook, J. A., Nerone, C. A. (1988), 206, 223 Sheath, R. G., Havas, M., Hellebust, J. A., Hutchinson, T. C. (1982), 42, 55 Sheath, R. G., Hellebust, J. A. (1978), 424, 427, 429, 433, 454, 459, 461, 463, 468 Sheath, R. G., Hellebust, J. A., Sawa, T. (1979), 200, 223 Sheath, R. G., Hymes, B. J. (1980), 203, 210, 221, 223 Sheath, R. G., Kaczmarczyk, D., Cole, K. M. (1993a), 203–204, 218, 221, 223 Sheath, R. G., Morrison, M. O. (1982), 25, 55, 205–206, 223 Sheath, R. G., Morrison, M. O., Korch, J. E., Kaczmarczyk, D., Cole, K. M. (1986), 461, 469 Sheath, R. G., Müller, K. M. (1997), 34, 55, 202, 223, 433, 461, 469

875

Sheath, R. G., Müller, K. M., Colbo, M. H.., Cole, K. M. (1996a), 205, 224 Sheath, R. G., Müller, K. M., Larson, D. J., Cole, K. M. (1995), 205, 224 Sheath, R. G., Müller, K. M., Sherwood, A. R. (2000), 218, 224 Sheath, R. G., Müller, K. M., Vis, M. L., Entwisle, T. K. (1996b), 217, 221, 224 Sheath, R. G., Steinman, A. D. (1981), 434, 454, 459, 461, 463, 469 Sheath, R. G., Steinman, A. D. (1982), 80, 82, 87–88, 92, 94, 101, 115, 118, 132, 135, 137, 144, 150, 155, 158, 161, 164, 166, 169, 171, 174, 177, 180, 184, 195, 230, 232, 234, 239, 241–242, 251, 412, 415, 421 Sheath, R. G., Van Alstyne, K. L., Cole, K. M. (1985), 34, 55 Sheath, R. G., Vis, M. L. (1995), 223 Sheath, R. G., Vis, M. L., Cole, K. M. (1992), 201, 203, 213, 221, 223 Sheath, R. G., Vis, M. L., Cole, K. M. (1993b), 200, 203–204, 217–219, 221, 223 Sheath, R. G., Vis, M. L., Cole, K. M. (1993c), 202, 204, 218–219, 220–221, 223 Sheath, R. G., Vis, M. L., Cole, K. M. (1993d), 221, 223 Sheath, R. G., Vis, M. L., Cole, K. M. (1994), 40, 55 Sheath, R. G., Vis. M. L., Cole, K. M. (1994a), 213, 221, 223 Sheath, R. G., Vis, M. L., Cole, K. M. (1994b), 203, 223 Sheath, R. G., Vis, M. L., Cole, K. M. (1994c), 199, 221, 224 Sheath, R. G., Vis, M. L., Hambrook, J. A., Cole, K. M. (1996), 34, 55, 433, 461, 463, 469 Sheath, R. G., Whittick, A.., Cole, K. M. (1994d), 201–203, 212–213, 221, 224 Sheath, R., Munawar, M. (1974), 486, 508 Sheldon, S. P., Wellnitz, T. A. (1998), 36, 55 Sherman, B. J., Phinney, H. K. (1971), 605, 635 Sherwood, A. R., Garbary, D. J., Sheath, R. G. (2000), 342, 351 Sherwood, A. R., Sheath, R. G.(1999a), 203, 224 Sherwood, A. R., Sheath, R. G.(1999b), 218, 221, 224 Sherwood, A. R., Sheath, R. G.(2000a), 218, 224 Sherwood, A. R., Sheath, R. G.(2000b), 202, 218, 221, 224 Shi, Z. (1995), 388, 416, 421 Shi, Z. (1996a), 388, 407, 421 Shi, Z. (1996b), 421 Shi, Z. (1998), 387, 421 Shi, Z. (1999), 387, 391, 421 Shi, Z., Zao, C. (1998), 416, 421 Shilo, M. (1967), 819, 833 Shimmel, S. M., Darley, W. M. (1995), 63, 115 Shin, W., Boo, S. M. (1999), 384, 421 Shireman, J. V., Haller, W. T., Colle, D. E., DuRant, D. F. (1983a), 810, 820, 833 Shireman, J. V., Hoyer, M. V., Maciena, M. J., Canfield, D. E. (1985), 821, 833 Shireman, J. V., Rottmann, R. W., Aldridge, F. J. (1983b), 833 Shoeman, F. R., Archibal, R. E. M. (1980), 569, 593 Shubert, L. E. (1975), 256, 309 Shubert, L. E. (1998), 258, 309 Shubert, L. E., Trainor, F. R. (1974), 256, 309 Sicko-Goad, L., Schelske, C. L., Stoermer, E. F. (1984), 568, 593 Sicko-Goad, L., Stoermer, E. F., Fahnenstiel, G. (1986), 573, 593 Sicko-Goad, L., Stoermer, E. F., Kociolek, J. P. (1989), 567–568, 572–573, 593 Sicko-Goad, L., Stoermer, E. F., Ladewski, B. G. (1977), 783, 802 Sieburth, J. M., Smetacek, V., Lenz, J. (1978), 20, 55 Sigworth, E. A. (1957), 776, 802 Silva, H., Sharp, A. J. (1944), 461, 469 Silva, P. C. (1959), 247, 251 Silva, P. C. (1972), 247, 251

876

Author Index

Silva, P. C., Papenfuss, G. F. (1953), 226, 251 Simms, R. W., Freeman, P. Hawksworth, D. L. (1988), 348, 351 Simon, N., Brenner, J., Edvardsen, B., Medlin, L. K. (1997), 515, 520 Simons, J., Ohm, M., Daalder, R., Boers, P., Rip, W. (1994), 812, 833 Simons, J., van Beem, A. P., de Vries, P. J. R. (1986), 334, 337, 351 Simonsen, R. (1979), 570, 593, 628, 635 Simonsen, R. (1987), 562, 593, 628, 635 Simonsen, S., Moestrup, Ø. (1997), 512, 520 Simpson, E. H. (1949), 784, 802 Simpson, M. R., MacLeod, B. W. (1991), 807, 833 Simpson, P. S., Eaton, J. W. (1986), 810, 833 Sinclair, C., Whitton, B. A. (1977), 61, 115, 759, 773 Singh, K. P. (1956), 416, 421 Siver, P. A. (1977), 24, 55 Siver, P. A. (1987), 503, 508, 528, 533, 538, 551–552, 555 Siver, P. A. (1988a), 489, 502, 508, 538, 541, 555 Siver, P. A. (1988b), 503, 508, 531, 533, 552, 555 Siver, P. A. (1988c), 533, 555 Siver, P. A. (1988d), 535, 555 Siver, P. A. (1989), 535–536, 556 Siver, P. A. (1991a), 524, 530–531, 533, 535–536, 538, 551–552, 556 Siver, P. A. (1991b), 556 Siver, P. A. (1991c), 533, 556 Siver, P. A. (1992), 556 Siver, P. A. (1993), 487, 508, 535, 537, 556 Siver, P. A. (1994), 556 Siver, P. A. (1995), 535–538, 556, 792, 802 Siver, P. A. (1999), 604, 635 Siver, P. A., Chock, J. S. (1986), 536–538, 556 Siver, P. A., Glew, J. R. (1990), 524, 527, 531, 556 Siver, P. A., Hamer, J. S. (1989), 534–538, 556 Siver, P. A., Hamer, J. S. (1990), 539, 556 Siver, P. A., Hamer, J. S. (1992), 488, 508, 526, 535, 541, 556 Siver, P. A., Hamer, J. S., Kling, H. (1990), 535, 556 Siver, P. A., Hinsch, J. (2000), 631, 635 Siver, P. A., Kling, H. (1997), 565, 593 Siver, P. A., Lott, A. M., Cash, E., Moss, J., Marsicano, L. J. (1999), 535, 537–538, 556 Siver, P. A., Marsicano, L. J. (1993), 556 Siver, P. A., Marsicano, L. J. (1996), 535, 538, 556 Siver, P. A., Skogstad, A. (1988), 535, 539, 556 Siver, P. A., Smol, J. P. (1993), 536, 556 Siver, P. A., Vigna, M. S. (1997), 534, 536, 538, 545–546, 556 Siver, P. A., Wujek, D. E. (1993), 535–538, 552, 556 Siver, P. A., Wujek, D. E. (1999), 552, 556 Siver, P. A.,Smol, J. P. (1993), 802 Skácelová, O., Komárek, J. (1989), 141, 196 Skogstad, A. (1984), 533, 556 Skogstad, A., Reymond, O. L. (1989), 474, 480, 485, 508 Skovgaard, A. (1998), 739, 755 Skuja, H. (1938), 199, 224 Skuja, H. (1939), 75, 115, 741, 755 Skuja, H. (1948), 73, 81, 115, 479, 502, 508, 716, 741, 755 Skuja, H. (1950), 531, 556 Skuja, H. (1956), 524, 556 Skuja, H. (1964), 80, 115 Skulberg, O. M., Carmichael, W. W., Codd, G. A., Skulberg, R. (1993), 141, 169, 174, 196 Skulberg, O. M., Skulberg, R. (1985), 141, 147, 196 Skvortzow, B. W. (1937), 628, 635 Skvortzow, B. W., Meyer, K. (1928), 659, 668 Slàdecek, V. (1973), 775, 787, 802 Slàdecek, V. (1986), 787, 802

Sládecková, A. (1962), 24–25, 55 Sládecková, A., Marvan, P., Vymazal (1983), 44, 55 Slate, J. E. (1998), 794, 802 Slobodkin, L. B. (1964), 45, 55 Sluiman, H. J., Guihal, C. (1999), 275, 309 Smajs, D., Samarda, J. (1999), 60, 65, 115 Smarda, J. (1991), 61, 115 Smarda, J., Smajs, D., Komrska, J. (1996), 61, 115 Smarda, J., Smajs, D., Komrska, J., Krzyzanek, V. (2002), 61, 115 Smayda, T. J. (1970), 20, 55 Smayda, T. J. (1997), 699–700, 713 Smeltzer, E. (1990), 814, 833 Smilauer, P. (1992), 604, 635 Smith, A. J. (1982), 61, 115 Smith, C. S., Barko, J. W. (1990), 816, 833 Smith, D. W. (1985), 820, 833 Smith, D. W., Brock, T. D. (1973), 206, 224 Smith, G. M. (1916), 429, 434, 461, 469 Smith, G. M. (1918), 453, 469 Smith, G. M. (1920), 60, 74, 76, 78, 83, 86, 90, 92, 94, 115, 118, 132, 142, 147, 196, 425, 432, 458, 465, 469 Smith, G. M. (1925), 60, 115 Smith, G. M. (1933), 60, 115, 323–330, 333, 344, 348, 351 Smith, G. M. (1944), 227, 241–242, 248, 251 Smith, G. M. (1950), 5–6, 9, 11–12, 33, 55, 60, 63, 74, 78, 80, 83, 85, 90, 96, 99–101, 104, 107, 109–110, 115, 118, 127–129, 132–133, 135, 137–139, 141–147, 149, 154–156, 158–160, 164, 166, 168–169, 171–175, 177–180, 182, 184, 187, 189, 196, 225, 227–228, 230–232, 234–236, 238–239, 241–242, 244, 247, 251, 255, 307, 309, 315, 344, 348, 351, 353–354, 363, 370, 381, 385, 387–388, 399, 410–413, 415, 421, 426, 429, 432, 457, 464, 469, 502, 508, 562, 593, 688, 709, 713, 758, 770, 773, xv, xvi Smith, G. M. (1955), 239, 251 Smith, T. E., Stevenson, R. J., Caraco, N. F., Cole, J. J. (1998), 37, 55 Smith, V. H. (1983), 812, 833 Smol, J. P. (1988), 604, 635 Smol, J. P. (1990), 561, 593, 655, 668, 789, 802 Smol, J. P. (1992), 776, 781, 786, 789, 793, 802 Smol, J. P. (1995), 535–539, 556, 775, 789, 793, 802 Smol, J. P., Charles, D. F., Whitehead, D. R. (1984), 536, 556 Smol, J. P., Glew, J. R. (1992), 780, 802 Soballe, D. M., Kimmel, B. L. (1987), 37, 55 Sokal, R. R., Rohlf, F. J. (1998), 788–789, 802 Solley, W. B., Merk, C. F., Pierce, R. R. (1988), 807, 833 Solomon, J. A., Walne, P. L., Dawson, N. S., Wiley, R. L. (1991), 385, 401, 421 Sommaruga, R. (1995), 20, 55 Sommer, J. R. (1965), 399, 421 Sommer, U. (1982), 755 Sommer, U. (1983), 812, 833 Sommer, U. (1984), 19, 55 Sommer, U. (1988), 20, 55 Sommer, U. (1992), 819, 833 Sommer, U. (1996), 568, 593 Sommer, U., Gliwicz, Z. M., Lampert, W., Duncan, A. (1986), 700, 713 Sommer, U., Kilham, S. S. (1985), 22, 55 Sournia, A. (1978), 703, 713 Sovereign, H. E. (1958), 604, 628, 635 Sovereign, H. E. (1963), 605, 628, 635 Soyer-Gobillard, M.-O. (1996), 686, 713 Spamer, E. E., Theriot, E. C. (1997), 586, 593 Sparks, R. E. (1995), 28, 55 Spaulding, S. A., Kociolek, J. P., Wong, D. (1999), 638, 649, 652 Spaulding, S. A., Stoermer, E. F. (1997), 638, 649, 652

Author Index Spector, D. L. (1984), 687, 690, 713 Spence, D. H. N. (1976), 807, 833 Spencer, C. N., King, D. L. (1984), 816, 818–819, 833 Spencer, D. L., Lembi, C. A. (1981), 812, 833 Spencer, L. B. (1971), 465, 469 Spero, H. J. (1985), 701, 713 Speziale, B. J., Dyck, L. A. (1992), 808, 833 Squires, L. E., Rushforth, S. R., Brotherson, J. D. (1979), 605, 635 Squires, L. E., Rushforth, S. R., Endsley, C. J. (1973), 488, 508 Squires, L. E., Whiting, M.C., Brotherson, J. D., Rushforth, S. R. (1979), 701, 713 Sreenivasa, M. R. (1971), 628, 635 Sreenivasa, M. R., Duthie, H. C. (1973), 604, 628, 635 St. Clair, L. L., Rushforth, S. R., (1976), 44, 55 St. Clair, L. L., Rushforth, S. R., Allen, J. V. (1981), 605, 635 Stal, L. (2000), 119, 121, 196 Stanier, R. Y., Cohen-Bazire, G. (1977), 59, 62, 115 Starks, T. L., Shubert, L. E. (1979), 254, 309 Starks, T. L., Shubert, L. E., Trainor, F. R. (1981), 45, 55, 63, 115, 258, 309 Starling, M. B., Chapman, V. J., Brown, J. M. A. (1974), 27, 55 Starmach, K. (1957), 144, 146, 196 Starmach, K. (1966), 91, 98, 115, 128, 136, 147–148, 176–177, 196 Starmach, K, (1969), 202, 224 Starmach, K. (1972), 347–348, 351 Starmach, K. (1973), 77, 115 Starmach, K. (1974), 709, 713 Starmach, K. (1977), 763, 771, 773 Starmach, K. (1983), 387–388, 399, 415, 421 Starmach, K. (1985), 501, 503, 508, 523, 556 Starr, R. C. (1955), 256, 309 Starr, R. C. (1962), 242, 251 Starr, R. C. (1970), 242, 248, 251 Starr, R. C., Zeikus, J. A. (1993), 234, 247, 251, 409–410, 421 Stauffer, R. E., Lee, G. F. (1973), 814, 833 Steidinger, K. A., Burkholder, J. M., Glasgow, H. B., Hobbs, C. H., Garrett, J. K., Truby, E. W., Noga, E. I., Smith, S. A. (1996a), 685–686, 713 Steidinger, K. A., Landsberg, J. H., Truby, E. W., Blakesley, B. A. (1996b), 698, 703, 713 Steil, W. N. (1944), 349, 351 Stein, F. (1878), 411, 415, 421, 471, 502, 509 Stein, J. R. (1958a), 226, 242–243, 251 Stein, J. R. (1958b), 226, 243, 252 Stein, J. R. (1959), 244–245, 248, 252 Stein, J. R. (1965), 244, 252 Stein, J. R. (1973), 258, 309 Stein, J. R. (1975), 180, 196, 247, 252, 363, 370–371, 373, 377, 381, 425–429, 433–434, 443, 446–447, 451, 454, 458–459, 461, 463, 469, 477, 483, 509 Stein, J. R., Amundsen, C. C. (1967), 43–44, 55 Stein, J. R., Borden, C. A. (1978), 425, 428–429, 434, 443, 446, 449, 451, 453–454, 458–459, 461, 463, 469 Stein, J. R., Borden, C. A. (1979), 72, 80–82, 87–88, 92, 95–96, 101, 115, 118, 126, 132, 135, 137, 141, 147, 150–151, 155, 158, 161, 164, 166, 169, 171, 174, 177, 180, 184, 189, 196, 232, 234–236, 238–239, 241–242, 244–245, 252, 411–413, 415, 421, 491, 509, 698, 713 Stein, J. R., Gerrath, J. F. (1969), 373, 381, 429, 433–434, 451, 453–454, 459, 463, 469 Steinkötter, J., Bhattacharya, D., Semmerlroth, I., Bibeau, C., Melkonian, M. (1994), 230, 252 Steinman, A. D. (1996), 32, 55, 819, 833 Steinman, A. D., McINtire, C. D. (1986), 205, 224 Steinman, A. D., McIntire, C. D., Lowry, R. R. (1988), 35, 55

877

Steinman, A. D., Meeker, R. H., Rodusky, A. J., Davis, W. P., Hwang, S.-J. (1997), 28, 55, 811, 833 Steinman, A. D., Sheath, R. G. (1984), 33, 55 Steinmüller, K., Kaling, M., Zetsche, K. (1983), 41, 55 Stemberger, R. S., Gilbert, J. J. (1985), 739, 755 Stephens, D. W., Gillespie, D. M. (1976), 43, 55 Sterrenberg, F. A. S. (1994), 586, 593 Stevenson, R. J. (1983), 36, 55 Stevenson, R. J. (1984), 784, 788, 802 Stevenson, R. J. (1990), 779, 802 Stevenson, R. J. (1996), 766, 773, 776, 779, 782–783, 802 Stevenson, R. J. (1996a), 24, 33, 55 Stevenson, R. J. (1996b), 35, 55 Stevenson, R. J. (1997), 34, 55, 784, 795, 802 Stevenson, R. J. (1998), 776, 783, 793, 795–796, 802 Stevenson, R. J. (2001), 777, 802 Stevenson, R. J., Bahls, L. L. (1999), 775–776, 778–779, 782–784, 786–787, 791, 802 Stevenson, R. J., Bothwell, M. L., Lowe, R. L. (1996), 24, 55, 604, 635, 672, 683 Stevenson, R. J., Glover, R. (1993), 785, 802 Stevenson, R. J., Hashim, S. (1989), 780, 802 Stevenson, R. J., Lowe, R. L. (1986), 776, 784, 803 Stevenson, R. J., Pan, Y. (1999), 776, 782, 784, 789, 803 Stevenson, R. J., Singer, R., Roberts, D. A., Boylen, C. W. (1985), 783, 803 Stevenson, R. J., Stoermer, E. F. (1981), 27, 55, 780, 803 Stevenson, R. J., Stoermer, E. F. (1982), 623, 635 Stevenson, R. J., Sweets, P. R., Pan, Y., Schultz, R. E. (1999), 778, 792, 803 Steward, K. K. (1993), 811, 833 Stewart, A. J., Blinn, D. W. (1976), 699, 713 Stewart, K. D., Mattox, K. R. (1975), 313, 351 Stewart, K. D., Mattox, K. R. (1978), 313, 351 Stewart, W. D. P. (1972), 119, 196 Stewart, W. D. P. (1980), 119, 121, 196 Stewart, W. D. P., Daft, M. J. (1977), 819, 833 Stiller, J. W., Hall, B. D. (1997), 387, 421 Stockner, J. G. (1967), 41, 55 Stockner, J. G. (1988), 65, 115 Stockner, J. G., Armstrong, F. A. J. (1971), 26, 55 Stockner, J. G., Benson, W. W. (1967), 605, 635 Stockner, J. G., Callieri, C., Cronberg, G. (2000), 20, 56, 120, 196 Stockner, J. G., Lund, J. W. G. (1970), 573, 593 Stockner, J. G., Shortreed, K. S. (1988), 22, 56, 812, 833 Stockner, J. G., Shortreed, K. S. (1991), 65, 115 Stoecker, D. K. (1998), 490, 509, 701, 713 Stoecker, D. K., Michaels, A. E., Davis, L. H. (1987), 755 Stoecker, D. K., Silver, M. W. (1990), 739, 755 Stoecker, D. K., Silver, M. W., Michaels, A. E., Davis, L. H. (1988/89), 755 Stoermer, E. F. (1963), 650, 652 Stoermer, E. F. (1967), 560, 569, 593 Stoermer, E. F. (1968), 598, 635 Stoermer, E. F. (1975), 605, 635 Stoermer, E. F. (1978), 585, 587, 593 Stoermer, E. F. (1980), 596, 605, 627–628, 635 Stoermer, E. F. (1988), 806–807, 833 Stoermer, E. F., Andersen, N. A., Schelske, C. L. (1992), 568, 593 Stoermer, E. F., Edlund, M. B. (1998), 568, 593 Stoermer, E. F., Emmert, G., Julius, M. L., Schelske, C. L. (1996), 583, 593 Stoermer, E. F., Emmert, G., Schelske, C. L. (1989), 586, 593 Stoermer, E. F., Häkansson, H. (1983), 581, 593 Stoermer, E. F., Häkansson, H., Theriot, E. C. (1987), 578, 581, 593

878

Author Index

Stoermer, E. F., Julius, M. L. (2002), 670, 683 Stoermer, E. F., Kingston, J. C., Sicko-Goad, L. (1979), 567, 569, 586, 593 Stoermer, E. F., Kociolek, J. P., Cody, W. (1990), 583, 593 Stoermer, E. F., Kociolek, J. P., Schelske, C. L., Andresen, N. A. (1991), 605, 635 Stoermer, E. F., Kociolek, J. P., Schelske, C. L., Conley, D. J. (1985c), 561, 593 Stoermer, E. F., Kreis, R. G., Jr. (1978), 561, 593 Stoermer, E. F., Kreis, R. G., Jr., Andresen, N. A. (1978a), 635 Stoermer, E. F., Kreis, R. G., Jr., Andresen, N. A. (1999), 598, 604, 635, 638, 653 Stoermer, E. F., Ladewski, B. G., Schelske, C. L. (1978b), 613, 635 Stoermer, E. F., Ladewski, T. B. (1976), 583, 593 Stoermer, E. F., Ladewski, T. B. (1982), 560, 593 Stoermer, E. F., Ladewski, T. B., Kociolek, J. P. (1986), 560, 593 Stoermer, E. F., Qi, Y., Ladewski, T. B. (1986), 664, 668 Stoermer, E. F., Sicko-Goad, L. (1977), 512, 520 Stoermer, E. F., Smol, J. P. (1999), 10, 56, 560, 571, 593, 595, 597, 604, 635, 776, 782–783, 788, 803 Stoermer, E. F., Wolin, J. A., Schelske, C. L., Conley, D. J. (1985a), 568, 586, 593 Stoermer, E. F., Wolin, J. A., Schelske, C. L., Conley, D. J. (1985b), 568–569, 573, 593 Stoermer, E. F., Yang, J. J. (1969), 583, 593, 595, 604–605, 628, 635 Stoermer, E. F., Yang, J. J. (1970), 595, 598, 605, 616, 635 Stoermer, E. F., Yang, J. J. (1971), 656, 661, 663, 668 Stokes, A. C. (1885), 477, 502, 509 Stokes, A. C. (1886), 472, 498, 509 Stokes, P. M. (1986), 26, 39, 56 Stokes, P. M., Yung, Y. K. (1986), 672, 682–683 Stoneburner, D. L., Smock, L. A. (1980), 699, 713 Stosch. see von Stosch Strahler, A. N. (1957), 28, 56 Straub, F. (1985), 627, 635 Straub, F. (1990), 627, 635 Stross, R. G., Sokol, R. C., Schwarz, A.-M., Howard-Williams, C. (1995), 27, 56 Suda, S., Watanabe, M. M., Inouye, I. (1989), 228, 230, 252 Sugawara, H., Miyazaki, S. (1999), 68, 115, 121, 196 Sulli, C., Fang, Z. W., Muchhal, U., Schwartzbach, S. D. (1999), 385, 421 Sullivan, M. J. (1979), 613, 635 Sunderman, F. W. (1978), 826, 833 Surek, B.. Beemelmanns, U., Melkoniam, M., Bhattacharya, D. (1994), 351 Surek, B., Melkonian, M. (1986), 385, 401, 421 Sutherland, J. M., Reaston, J., Stewart, W. D. P., Herdman, M. (1985a), 120, 135, 196 Sutherland, J. M., Reaston, J., Stewart, W. D. P., Herdman, M. (1985b), 120, 196 Suttle, C. A., Harrison, P. J. (1988), 65, 115 Suttle, C. A., Stockner, J. G., Harrison, P. J. (1987), 20, 22, 56 Suttle, C. A., Stockner, Shortreed, K. S., J. G., Harrison, P. J. (1988), 20, 56 Sutton, D. L., Porter, K. M. (1989), 821, 833 Suxena, M. R. (1955), 415, 421 Suykerbuyk, R. E. M., Roijackers, R. M. H., Houtman, S. S. J. (1995), 487, 509 Svedelius, N. (1930), 759, 765, 770, 773 Swain, E. B., Monson, B. A., Pillsbury, R. W. (1986), 822, 833 Sweets, P. R. (1992), 787, 803 Sze, P. (1986), 381 Sze, P. (1998), 1, 5, 9 Sze, P., Kingsbury, J. M. (1972), 42–43, 56

Szymanska, H., Spalik, K. (1993), 332, 348, 351 Szymanska, H., Zakrys, B. (1990), 763–764, 766, 773 T Taft, C. E. (1964), 347, 349, 352 Taft, C. E. (1978), 702, 714 Taft, C. E., Taft, C. W. (1970), 139, 180, 196 Taft, C. E., Taft, C. W. (1971), 415, 421, 698, 701, 714 Takahashi, E. (1978), 524, 526, 528, 536, 539, 551–552, 556 Takahashi, E., Hayakawa, T. (1979), 536, 556 Tanaka, J., Kamiya, M. (1993), 202, 224 Tani, Y., Tsumura, H. (1989), 406, 421 Tarapchak, S. J. (1972), 424–427, 429, 432–434, 441–453, 456–459, 469 Targett, N. M., Arnold, T. M. (1998), 765, 773 Taub, F. B., Kindig, A. C., Meador, J. P., Swartzman, G. L. (1989), 825, 833 Tavera, R., Komárek, J. (1996), 60, 67, 83, 87, 96–97, 101, 103–104, 107, 115, 134, 177, 196 Taylor, F. J. R. (1980), 697, 714 Taylor, F. J. R. (1987), 687, 714 Taylor, F. J. R. (1999), 697, 714 Taylor, J. F. R. (1990), 388, 421 Taylor, W. D., Hern, S. C., William, L. R., Lambou, V. W., Morris, M. K., Morris, F. A. (1979), 738, 755 Taylor, W. D., Sanders, R. W. (1991), 385, 421 Taylor, W. D., Wee, J. L., Wetzel, R. G. (1986), 539, 556 Taylor, W. R. (1928), 115, 462, 469 Taylor, W. R. (1934), 429, 469 Teiling, E. (1941), 90, 115 Teiling, E. (1948), 379, 381 Teiling, E. (1950), 361, 381 Teiling, E. (1952), 362, 381 Teiling, E. (1957), 376, 381 Teiling, E. (1967), 379, 381 Tell, G., Conforti, V. (1984), 416, 421 Tell, G., Conforti, V. (1986), 388, 414–416, 421 ter Braak, C. J. F. (1988), 604, 635 ter Braak, C. J. F., van Dam, H. (1989), 787, 791, 803 Tett, P., Gallegos, C., Kelly, M. G., Hornerger, G. M., Cosby, B. J. (1978), 803 Thérézien, Y. (1999), 388, 414, 416, 421 Theriot, E. C. (1987), 568–570, 586, 593 Theriot, E. C. (1990), 570, 593 Theriot, E. C. (1992), 568–570, 586, 593 Theriot, E. C., Bradbury, J. P. (1987), 570, 584, 593 Theriot, E. C., Fritz, S. C., Gresswell, R. E. (1997), 41, 56 Theriot, E. C., Häkansson, H., Kociolek, J. P., Round, F. E., Stoermer, E. F. (1987), 570, 578, 593 Theriot, E. C., Håkansson, H., Stoermer, E. F. (1988), 567 Theriot, E. C., Kociolek, J. P. (1986), 570 Theriot, E. C., Serieyssol, K. (1994), 570 Theriot, E. C., Stoermer, E. F. (1984a), 568–569, 586 Theriot, E. C., Stoermer, E. F. (1984b), 560, 568, 586 Theriot, E. C., Stoermer, E. F. (1986), 567–569, 586 Thirb, H. H., Benson-Evans, K. (1982), 203, 224 Thomas, W. H., Duval, B. (1995), 44, 56 Thompson, R. H. (1938), 83, 86, 115, 412, 421, 433, 461, 463, 469 Thompson, R. H. (1947), 688, 698, 702, 706, 714 Thompson, R. H. (1949), 688, 698, 709, 714 Thompson, R. H. (1950), 688–689, 702, 709, 714 Thompson, R. H. (1954), 241–242, 252 Thompson, R. H. (1972a), 315, 352 Thompson, R. H. (1972b), 330, 352 Thompson, R. H. (1975), 765, 771, 773

Author Index Thompson, R. H., Halicki, P. J. (1965), 513, 520 Thompson, R. H., Wujek, D. E. (1989), 232, 248, 252 Thompson, R. H., Wujek, D. E. (1996), 345, 349, 352 Thompson, R. H., Wujek, D. E. (1997), 45, 56, 315, 343, 348, 352 Thompson, R. H., Wujek, D. E. (1998a), 483, 503, 509 Thompson, R. H., Wujek, D. E. (1998b), 496–498, 509 Thomsen, H. A., Zimmerman, B., Moestrup, Ø., Kristiansen, J. (1981), 503, 509 Thorp, J. H., Black, A. R., Haag, K. H., Wehr, J. D. (1994), 28, 32, 38, 56 Thorp, J. H., Black, A. R., Jack, J. D., Casper, A. F. (1996), 786, 803 Thorp, J. H., Delong, M. D. (1994), 31–32, 56 Threlkeld, S. T., Chiavelli, D. A., Willey, R. L. (1993), 408, 411, 421 Threlkeld, S. T., Willey, R. L. (1993), 408, 421 Tiffany, L. H. (1930), 38, 341, 352 Tiffany, L. H. (1934), 83, 115 Tiffany, L. H. (1936), 381 Tiffany, L. H. (1937), 325, 348, 352, 429, 433, 461, 469 Tiffany, L. H. (1944), 348, 352 Tiffany, L. H., Britton, M. E. (1944), 433–434, 459, 462–463, 469 Tiffany, L. H., Britton, M. E. (1952), 118, 158, 161, 164, 166, 169, 174, 177, 196, 325, 330, 348, 352, 710, 714 Tiftickjian, J. D., Rayburn, W. R. (1986), 364, 381 Tilden, J. (1910), 60, 91, 96, 115, 117, 132, 135, 137, 139, 141, 144, 147, 150–151, 155, 158, 161, 164, 166, 169, 171, 174, 180–181, 184, 189, 196 Tillett, D., Parker, D. L. Neilan, B. A. (1999), 63, 115 Tilman, D. (1977), 22, 56 Tilman, D. (1982), 22, 56 Tilman, D., Kiesling, R. L. (1984), 812, 833 Tilman, D., Kiesling, R., Sterner, S. S., Johnson, F. A. (1986), 487, 509 Tilman, D., Kilham, S. S. (1976), 487, 509 Tilman, D., Kilham, S. S., Kilham, P. (1976), 607, 636 Tilman, D., Kilham, S. S., Kilham, P. (1982), 487, 509, 568, 604, 636 Timpano, P. (1978), 771, 773 Timpano, P. (1980), 759, 771, 773 Tippet, R. (1970), 25, 56 Titman, D. (1976), 568, 585 Tokuyasu, K., Scherbaum, O. H. (1965), 403, 421 Tomas, R. N., Cox, E. R. (1973), 686, 707, 714 Tomaselli, L., Palandri, M. R., Tredici, M. R. (1996), 132, 196 Toriumi, S., Dodge, J. D. (1993), 690, 714 Trainor, F. R. (1978), 255–256, 309 Trainor, F. R. (1991), 255, 309 Trainor, F. R. (1998), 254–256, 309 Trainor, F. R., Cain, J. R., Shubert, L. E. (1976), 256, 309 Trainor, F. R., Egan, P. (1990a), 255, 309 Trainor, F. R., Egan, P. (1990b), 255, 309 Trainor, F. R., Egan, P. (1990c), 255, 309 Trainor, F. R., Morales, E. A. (1999), 256–257, 309 Trainor, F. R., Shubert, L. E. (1973), 785, 803 Transeau, E. (1913), 424–425, 463, 469 Transeau, E. (1916), 364, 381 Transeau, E. (1917), 429, 459, 469 Transeau, E. (1925), 369, 381 Transeau, E. (1926), 381 Transeau, E. (1933), 372, 381 Transeau, E. (1951), 355, 362–363, 369–372, 379, 381 Tranvik, L. J., Porter, K. G., Sieburth, J. (1989), 740, 755 Trelease (1889), 171, 196 Trench, R. K., Thinh, L. V. (1995), 687, 714 Triemer, R. E. (1992), 405, 421 Triemer, R. E., Farmer, M. A. (1991a), 385, 387, 391, 401, 421

879

Triemer, R. E., Farmer, M. A. (1991b), 387, 391, 401, 405, 421 Triemer, R. E., Lewandowski, C. L. (1994), 385, 401, 408, 421 Tschermak-Woess, E., Kasel-Fetzman, E. (1992), 509 Tschermak-Woess, W. (1980), 482, 509 Tuchman, M. L., Stevenson, R. J. (1980), 36, 56 Tuchman, M. L., Stoermer, E. F., Carney, J. J. (1984), 607, 636 Tuchman, M. L., Theriot, E. C., Stoermer, E. F. (1984), 560, 569 Tuchman, M., Stevenson, R. J. (1980), 780, 803 Tuchman, N. C. (1996), 673, 683 Tucker, C. S., Boyd, C. E. (1978), 824, 833 Tucker, C. S., Busch, R. L., Lloyd, S. W. (1983), 810, 833 Tupa, D. D. (1974), 322–323, 332, 348, 352 Turner, M. A., Howell, E. T., Robinson, G. G. C., Brewster, J. F., Sigurdson, L. J., Findlay, D. L. (1995), 370, 372, 381 Turner, M. A., Howell, E. T., Robinson, G. G. C., Campbell, P., Hecky, R. E., Schindler, E. U. (1994), 26, 56 Turner, M. A., Schindler, E. U., Findlay, D. L., Jackson, M.B., Robinson, G. G. C. (1995), 26, 39, 56 Turpin, D. H., Harrison, P. J. (1979), 19, 56 Twist, H., Edwards, A. C., Codd, G. A. (1997), 786, 803 Tyler, P. A. (1996), 12, 56 Tyler, P. A., Pipes, L. D., Croome, R. L., Leedale, G. F. (1989), 524, 526–527, 533–534, 548, 551–552, 556 U U. S. Environmental Protection Agency (1978), 803 U. S. Environmental Protection Agency (1992), 776–777, 795, 803 U. S. Environmental Protection Agency (1993), 794, 803 U. S. Environmental Protection Agency (1996), 776–777, 786, 803 U. S. Environmental Protection Agency (1998), 776–777, 788, 803 Ueyema, S., Kobayashi, H. (1988), 665, 668 Umana-Villalobos, G. (1993), 15, 56 Umezaki, I. (1974), 147, 196 Urabe, J. (1993), 819, 833 Urabe, J., Clasen, J., Sterner, R. W. (1997), 819, 833 Utermöhl, H. (1958), 67, 115 V Van Baalen, C. (1965), 68, 115 Van Baalen, C. (1987), 121, 196 Van Baalen, C., O’Donnell, R. (1972), 88, 115 Van Dam, H., Mertenes, A., Sinkeldam, J. (1994), 791, 803 van den Berg, M. S., Coops, H., Noordhuis, R., van Schie, J., Simons, J. (1997), 28, 56 van den Berg, M. S., Coops, H., Simons, J. (1998b), 811–812, 834 van den Berg, M. S., Coops, H., Simons, J., DeKeizer, A. (1998a), 812, 834 van den Hoek, C. (1982), 340, 348, 350 van den Hoek, C., Mann, D. G., Jahns, H. M. (1995), 1, 5, 9, 225, 252, 312–313, 322, 340–341, 345–347, 350, 353, 381, 384, 387, 399, 401, 405, 421, 423–425, 427, 429, 432, 469, 474, 509, 560, 567, 757–759, 773 van der Ploeg, M., Dennis, M. E., de Regt, M. Q. (1995), 807, 834 van der Zweerde, W. (1993), 820, 834 van Donk, E., Grimm, M. P., Gulati, R. D., Kline Breteler, J. P. G. (1990), 816, 834 Van Etten, J. L., Lane, L. C., Meints, R. H. (1991), 819, 834 Van Everdingen, R. P. (1970), 41, 56 Van Heukelem, L., Lewitus, A. J., Kana, T. M. (1992), 783, 803 Van Heurck, H. (1880), 562 Van Heurck, H. (1881), 562, 621, 636 Van Heurck, H. (1882), 562 Van Heurck, H. (1883), 562 Van Heurck, H. (1884), 562 Van Heurck, H. (1885), 562

880

Author Index

Van Landingham, J. I. (1978), 669, 683 Van Landingham, S. L. (1964) Van Landingham, S. L. (1969), 656, 665, 668 Van Landingham, S. L (1967), 665, 668 Van Landingham, S. L (1978), 659, 668 van Leeuwenhoek, A. (1700), 225, 252 Van Niewenhuyse, E. E., Jones, J. R. (1996), 38, 56 Vanni, M. J., Findlay, D. L. (1990), 818, 834 Vanni, M., Layne, C. D. (1997), 22, 56 Vannote, R. L., Minshall, G. W., Cummins, K. W., Sedell, J. R., Cushing, C. E. (1980), 31, 56, 779, 803 Vazquez, G., Moreno-Casasola, P., Barrera, O. (1998), 45, 56 Venkataraman, G. S. (1961), 465, 469 Venkataraman, G. S. (1969), 68, 115 Verb, R. G., Vis, M. L., Ott, D. W., Wallace, R. L. (1999), 463, 469 Vesk, M. Hoffman, L. R., Pickett-Heaps, J. D. (1984), 499, 509 Vickerman, K. (1990), 385, 421 Vickerman, K., Brugerolle, G., Mignot, J.-P. (1991), 385, 422 Vigna, M. S., Kristiansen, J. (1996), 486, 509 Villeneuve, V., Vincent, W. F., Komárek, J. (2001), 166, 196 Vincent, W. F. (2000), 121, 196 Vinebrook, R. D. (1996), 27, 56 Vinebrook, R. D., Hall, R. I., Leavitt, P. R., Cumming, B. F. (1998), 43, 56 Vinebrooke, R. D., Graham, M. D. (1997), 803 Vinyard, W. C. (1955), 314, 352 Vinyard, W. C. (1958), 434, 454, 459, 461, 469 Vinyard, W. C. (1966), 458–459, 469 Viroux, L. (1997), 38, 56 Vis, M. L, Carlson, T. A., Sheath, R. G. (1991), 203, 224 Vis, M. L, Entwisle, T. J. (2000), 215, 224 Vis, M. L., Saunders, G. W., Sheath, R. G., Dunse, K., Entwisle, T. J. (1998), 218, 221, 224 Vis, M. L, Sheath, R. G. (1992), 203–204, 216–217, 221, 224 Vis, M. L, Sheath, R. G. (1993), 204, 208, 210, 221, 224 Vis, M. L, Sheath, R. G. (1996), 203, 213, 221, 224 Vis, M. L, Sheath, R. G. (1997), 221, 224 Vis, M. L, Sheath, R. G. (1998), 221, 224 Vis, M. L, Sheath, R. G. (1999), 216, 221, 224 Vis, M. L, Sheath, R. G., Cole, K. M. (1992), 203–204, 212, 221, 224 Vis, M. L, Sheath, R. G., Cole, K. M. (1996a), 203, 213–214, 221, 224 Vis, M. L, Sheath, R. G., Cole, K. M. (1996b), 213, 221, 224 Vis, M. L., Sheath, R. G., Hambrook, J. A., Cole, K. M. (1994), 4, 9 Vogel, S. (1984), 200, 224 Vollenweider, R. A. (1969), 514, 520, 812, 834 Vollenweider, R. A. (1976), 775, 782, 803 von Christen, H. R. (1961), 706, 709–710 von Daday, E. (1905), 711, 714 von Stosch, H. A. (1965), 694, 713 von Stosch, H. A. (1972), 694, 713 von Stosch, H. A. (1973), 694, 713 Vonshak, A. (1997), 121, 196 Vørs, N. (1992a), 716, 735, 749–750, 755 Vørs, N. (1992b), 716, 735, 749–750, 755 Vørs, N. Johansen, B., Havskum, H. (1990), 486–487, 509 Vymazal, J. (1984), 785, 803 Vymazal, J. (1994), 776, 803 Vyverman, W., Compere, P. (1991), 649, 653 Vyverman, W., Sabbe, K., Mann, D. G., Kilroy, C., Vyverman, R., Vanhutte, K., Hodgson, D. (1998), 656, 668 W Waern, M. (1938), 764, 773

Waern, M. (1952), 758, 763–767, 770–771, 773 Wagemann, R., Barica, J. (1979), 824, 834 Wales, G. H. (1934), 710, 714 Walker, H. L., Higginbotham, L. R. (2000), 819, 834 Walker, I. R., Paterson, C. G. (1986), 617, 636 Walker, W. W., Jr. (1987), 813, 834 Walker, W. W., Jr., Westberg, C. E., Schuler, D. J., Bode, J. A. (1989), 816, 834 Wall, D., Dale, B. (1968), 687, 714 Wallace, J. H. (1960), 661, 665–666, 668 Wallace, J. H., Patrick, R. M. (1950), 665, 668 Walne, P. L. (1971), 391–392, 422 Walne, P. L., Dawson, N. S. (1993), 387, 402, 422 Walne, P. L., Gualtieri, P. (1994), 402, 422 Walne, P. L., Kivic, P. A. (1990), 387, 406, 422 Walne, P. L., Möestrup, P., Norris, R. E., Ettl, H. (1986), 402, 413, 422 Walne, P. L., Passarelli, V., Lenzi, P., Barsanti, L., Gualtieri, P. (1998), 392, 402, 422 Walsby, A. E. (1972), 61, 115 Walsby, A. E. (1978), 61, 115 Walsby, A. E. (1981), 61, 115 Walsby, A. E., Xypolyta, A. (1977), 567, 587 Walters, C. J., Krause, E., Neill, W. E., Northcote, T. G. (1987), 19, 56 Walton, L. B. (1915), 415, 422 Wang, J. C., Hoshaw, R. W., McCourt, R. M. (1986), 371, 381 Ward, A. K., Dahm, C. N., Cummins, K. W. (1985), 36, 56, 180, 196 Ward, D. M., Castenholz, R. W. (2000), 41, 56, 121, 196 Ward, J. V., Stanford, J. A. (1983), 31, 56 Ward, K. A., Willey, R. L. (1981), 411, 422 Warner, R. W. (1971), 42, 56 Watanabe, M. (1971), 120, 196 Watanabe, M. (1999), 60, 115 Watanabe, M., Furuya, M. (1982a), 738, 755 Watanabe, M., Furuya, M. (1982b), 738, 755 Watanabe, M., Komárek, J. (1989), 118, 150, 156, 196 Watanabe, M., Miyoshi, M., Furuya, M. (1976), 738, 755 Watanabe, S., Floyd, G. (1989), 253, 256, 309 Watanabe, T., Asai, K., Houki, A., Tanaka, S., Hizuka, T. (1986), 787, 803 Waterbury, J. B. (1979), 79, 115 Waterbury, J. B. (1989), 115 Waterbury, J. B., Rippka, R. (1989), 63, 116 Waterbury, J. B., Stanier, R. Y. (1977), 62, 116 Waterbury, J. B., Stanier, R. Y. (1978), 60, 62, 79, 99, 110, 116 Wawrik, F. (1979), 533, 556 Wawrzyniak, L. A., Andersen, R. A. (1985), 552, 556 Weber, C. I. (1970), 585 Weber, F. I. (1973), 775–776, 803 Weber van Bosse, A. A. (1925), 110, 116 Webster, K. E., Peters, R. h. (1978), 808, 834 Webster, P. (1989), 401, 422 Wee, J. L. (1982), 523–524, 526, 528, 530, 533, 551–552, 556 Wee, J. L. (1983), 491, 509, 539–540, 556 Wee, J. L. (1996), 473, 509 Wee, J. L. (1997), 530–531, 533–534, 556 Wee, J. L., Booth, D. J., Bossier, M. A. (1993), 526, 535–536, 541, 552, 556 Wee, J. L., Gabel, M. (1989), 535–536, 556 Wee, J. L., Harris, S. A., Smith, J. P., Dionigi, C. P., Millie, D. F. (1994), 535, 556 Wee, J. L., Hinchey, J. M., Nguyen, K. X., Hurley, D. L. (1996), 473, 509

Author Index Wehr, J. D. (1981), 34, 56 Wehr, J. D. (1989), 20, 57, 65, 116 Wehr, J. D. (1990), 20, 57, 65, 116 Wehr, J. D. (1991), 20, 57 Wehr, J. D. (1992), 65, 116 Wehr, J. D. (1993), 22, 57 Wehr, J. D., Brown, L. M. (1985), 23, 57, 513, 520 Wehr, J. D., Brown, L. M., O’Grady, K. (1985), 513, 520 Wehr, J. D., Brown, L. M., O’Grady, K. (1987), 23, 57, 513, 520 Wehr, J. D., Descy, J.-P. (1998), 28, 32, 37, 57, 65, 116, 776, 803 Wehr, J. D., Lonergan, S. P., Thorp, J. H. (1997), 38, 57 Wehr, J. D., Stein, J. R. (1985), 33, 57, 342, 352, 758, 763–766, 770, 773 Wehr, J. D., Thorp, J. H. (1997), 32, 37, 57, 65, 116 Wehr, J. D., Whitton, B. A. (1983), 42, 57 Wei, Y. (1996), 513, 520 Weisse, T. (1993), 20, 57, 65, 116 Weisse, T., Kirchhoff (1997), 701, 714 Weitzel, R. L. (1979), 766, 773 Welch, E. B., Cooke, G. D. (1995), 814, 834 Welch, E. B., Cooke, G. D. (1999), 814, 834 Welch, E. B., DeGasperi, C. L., Spyridakis, D. E., Belnick, T. J. (1988), 814, 834 Welch, I. M., Barrett, P. R. F., Gibson, M. T., Ridge, I. (1990), 822, 834 Welcomme, R. L. Ryder, R. A., Sedell, J. A. (1989), 31, 57 Wemmer, D. E., Wedemeyer, G. J., Glazer, A. N. (1993), 737, 755 Wenrich, D. H. (1923), 412, 415, 422 Wenrich, D. H. (1924), 412, 415, 422 West, G. S. (1904), 463, 469 West, J. A. (1990), 765–766, 773 West, J. A., Kraft, G. T. (1996), 757, 759–760, 766–767, 771, 773 West, W., West, G. S. (1904), 379, 381 West, W., West, G. S. (1905), 379, 381 West, W., West, G. S. (1908), 379, 381 West, W., West, G. S. (1912), 379, 381 West, W., West, G. S., Carter (1923), 379, 381 Westbroek, P., Brown, C. W., van Bleijswijk, J., Brownlee, C., Brummer, G., Conte, M., Egge, J., Fernandéz, E., Jordan, R., Knappertsbusch, M., Stefels, M., Velduis, M., van der Wal, P., Young, J. (1993), 511, 520 Wetherbee, R., Andersen, R. A. (1992), 473, 509 Wetherbee, R., Hill, D. R. A., Brett, S. J. (1987), 732, 741, 755 Wetherbee, R., Hill, D. R. A., McFadden, G. I. (1986), 732, 741, 755 Wetherbee, R., Ludwig, M., Koutoulis, A. (1995), 531, 557 Wetherbee, R., Platt, S. J., Beech, P. L., Pickett-Heaps, J. D. (1988), 473, 509 Wettstein, R. (1924), 60, 116 Wetzel, R. G. (1964), 43, 57 Wetzel, R. G. (1975), 32, 57 Wetzel, R. G. (1983), 538, 557, 776, 783, 803 Wetzel, R. G. (1983a), 12, 18–19, 23–24, 40, 42, 57 Wetzel, R. G. (1983b), 24, 57 Wetzel, R. G. (1983c), 24, 57 Wetzel, R. G. (1990), 16, 57, 814, 834 Wetzel, R. G. (1996), 776, 803 Wetzel, R. G. (2001), 806–807, 834 Wetzel, R. G., Likens, G. E. (1991), 514, 520, 779–780, 783, 785, 803 Wetzel, R. G., Ward, A. K. (1992), 32, 57 Whale, G. F., Walsby, A. E. (1984), 119, 196 Whatley, J. M. (1993), 387, 422 Wheeler, B. D., Whitton, B. A. (1971), 764, 773 Whelden, R. M. (1941), 99, 116, 459, 469 Whelden, R. M. (1947), 82, 96, 98, 101, 116, 118, 134, 137, 155,

881

158, 161, 164, 166, 169, 180, 184, 196 Whitaker, J., Barica, J., Kling, H., Buckley, M. (1978), 822, 824, 834 Whitehead, D. R., Charles, D. F., Goldstein, R. A. (1990), 803 Whitford, L. A. (1960), 35, 57, 203, 224 Whitford, L. A. (1969), 314, 352, 496, 509 Whitford, L. A. (1970), 483, 509 Whitford, L. A. (1977), 758, 773 Whitford, L. A. (1979), 428, 434, 443, 447, 451, 469, 512, 514, 520 Whitford, L. A., Schumacher, C. J. (1984), 363, 370, 381 Whitford, L. A., Schumacher, G. J. (1963), 196 Whitford, L. A., Schumacher, G. J. (1969), 63, 67, 72, 79–80, 82, 88, 92, 95–96, 116, 118, 126, 132, 137, 139, 141, 144, 147, 150–151, 155, 158, 161, 164, 166, 169, 171, 174, 180–181, 184, 189, 196, 315, 335, 348, 352, 413, 415, 422, 424, 428–429, 433–434, 443, 449–453, 458–459, 461, 465, 469, 477–482, 496, 500, 509 Whitford, L. A., Schumacher, G. J. (1973), 503, 509 Whitford, L. A., Schumacher, G. J. (1984), 239, 252, 408, 422, 698, 714, xvi Whitford, L. A., Shumacher, G. J. (1984), 295, 303, 306, 309 Whiting, M. C., Brotherson, J. D., Rushforth, S. R. (1978), 699–700, 714 Whitmore, T. J. (1989), 787, 803 Whittaker, R. H. (1975), 38, 40, 57 Whitton, B. A. (1970), 340, 352, 782–783, 803, 822, 834 Whitton, B. A. (1973), 822, 834 Whitton, B. A. (1975), 11–12, 28, 32–35, 57, 203–204, 224 Whitton, B. A. (1977), 61, 116 Whitton, B. A. (1984), 28, 57, 121, 196, 785, 803 Whitton, B. A. (1987), 118, 164, 196 Whitton, B. A. (1992), 61, 66–67, 116 Whitton, B. A. (2000), 121, 196 Whitton, B. A., Burrows, I. G., Kelly, M. G. (1989), 785, 803 Whitton, B. A., Carr, N. G. (1982), 116 Whitton, B. A., Kelly, M. G. (1995), 776, 803 Whitton, B. A., Peat, A. (1969), 129, 196 Whitton, B. A., Potts, M. (1982), 66, 116 Whitton, B. A., Potts, M. (2000), 19, 57, 63–64, 116, 117, 120, 196 Whitton, B. A., Rott, E. (1996), 681, 684, 775–776, 791, 803 Whitton, B. A., Rott, E., Friedrich, G. (1991), 681, 684, 776, 791, 804 Whitton, B. A., Shehata, F. H. A. (1982), 785, 803 Whitton, B. A., Yelloly, J. M., Christmas, M., Hernádez, I. (1998), 785, 804 Wiedner, C., Nixdorf, B. (1998), 487, 509 Wilce, R. T. (1966), 757–758, 764, 766, 770–771, 773 Wilce, R. T., Weber, E. E., Sears, J. R. (1970), 766, 771, 773 Wilcox, L. W., Lewis, L. A., Fuerst, P. A., Floyd, G. L. (1992), 257, 309 Wilcox, L. W., Wedemayer, G. J. (1984), 686, 714 Wiley, M. J., Gordon, R. W., Waite, S. W., Powless, T. (1984), 810, 834 Wilken, L. R., Kristiansen, J., Jürgensen, T. (1995), 486, 509 Willén, E. (1992), 364, 381 Willen, E., Hajdu, S., Pejler, Y. (1990), 583 Willen, E., Oke, M., Gonzalez, F. (1980), 755 Willey, R. L. (1972), 408, 411, 422 Willey, R. L. (1980), 411, 415, 422 Willey, R. L. (1982), 411, 415, 422 Willey, R. L., Cantrell, P. A. (1990), 408, 422 Willey, R. L., Durbin, E. M., Bowen, W. R. (1973), 404, 422 Willey, R. L., Threlkeld, S. T. (1993), 408, 422 Willey, R. L., Walne, P. L., Kivic, P. (1988), 387, 391, 422 Willey, R. L., Ward, K., Russin, W., Wibel, R. G. (1977), 405, 410, 422

882

Author Index

Willey, R. L., Wibel, R. G. (1985a), 385, 401, 422 Willey, R. L., Wibel, R. G. (1985b), 385–386, 391, 401, 422 Willey, R. L., Willey, R. B., Threlkeld, S. T. (1993), 408, 411, 422 Williams, D. M. (1985), 570, 598, 600–601, 605, 616, 636 Williams, D. M. (1986), 598, 600, 636 Williams, D. M. (1989), 598, 601, 636 Williams, D. M. (1990a), 596–598, 600, 636 Williams, D. M. (1990b), 596, 598, 600, 613, 636 Williams, D. M. (1990c), 596, 599, 613, 636 Williams, D. M. (1993), 601, 636 Williams, D. M. (1994), 601, 604, 623, 636 Williams, D. M. (1996), 601, 605, 623, 636 Williams, D. M. (1997), 601, 636 Williams, D. M. (2001), 631, 636 Williams, D. M., Round, F. E. (1986), 561, 570 Williams, D. M., Round, F. E. (1987), 561, 596, 598–599, 613, 616, 628, 636 Williams, D. M., Round, F. E. (1988a), 598–599, 636 Williams, D. M., Round, F. E. (1988b), 596, 598, 600, 636 Williams, L. G. (1964), 585 Williams, L. G. (1972), 585 Williams, W. D. (1996), 42, 57 Wilmotte, A., Stam, W. T. (1984), 116 Wilson, S. E., Cumming, B. F., Smol, J. P. (1994), 604, 636 Wilson, S. E., Cumming, B. F., Smol, J. P. (1996), 42–43, 57 Winkenbach, F., Wolk, C. P. (1973), 119, 196 Winner, J. M. (1975), 38, 57 Winner, R. W. (1985), 825, 834 Winner, R. W., Owen, H. A., Moore, M. V. (1990), 825, 834 Witkowski, A., Lange-Bertalot, H., Stachura, K. (1998), 637, 653 Witt, F. G., Stöhr, C., Ullrich, W. R. (1999), 700, 714 Wium-Andersen, S., Anthoni, U., Christophersen, C., Houen, G. (1982), 811, 834 Woelkerling, W. J. (1976), 40, 57 Woelkerling, W. J. (1990), 197, 224 Wojciechowski, I. (1971), 90, 116 Wolk, C. P. (1973), 119, 121, 196 Wolk, C. P. (1982), 119, 121, 196 Wolken, . J., Palade, G. E. (1952), 501, 509 Wolle, F. (1890), 561 Wolowski, K. (1992), 416, 422 Wolowski, K. (1993), 402, 422 Wolowski, K., Walne, P. L. (1997), 387, 410, 415, 422 Wong, S. L., Clark, B., Kirby, M., Kosciuw, R. F. (1978), 782, 804 Wood, H. C. (1869), 92, 116 Wood, H. C. (1872), 116 Wood, R. D. (1948), 348, 352 Wood, R. D. (1950), 27, 57 Wood, R. D. (1967), 348, 352 Wood, R. D., Imahori, D. (1964), 331, 348, 352 Woodhead, N., Tweed, R. D. (1960), 604, 613, 636 Woodson, B. R. (1962), 433, 469 Woodson, B. R., Afazal, M. (1976), 433, 469 Woodson, B. R., Holoman, V. (1964), 429, 433–434, 461, 463, 469 Woodson, B. R., Wilson, W. (1973), 461, 463, 469 Wooten, J. W., Elakovich, S. D. (1991), 821, 834 Wootton, J. T., Parker, M. S., Power, M. E. (1996), 32, 57 Wright, A.-D. G., Lynn, D. H. (1997a), 686, 714 Wright, A.-D. G., Lynn, D. H. (1997b), 686, 714 Wu, J.-T., Chou, J.-W. (1998), 699–701, 714 Wujek, D. (1971), 335, 349, 352 Wujek, D. E. (1967), 499, 509 Wujek, D. E. (1976), 503, 509 Wujek, D. E. (1983), 500, 509 Wujek, D. E. (1984), 536, 552, 557

Wujek, D. E. (1996), 474, 509 Wujek, D. E. (1999), 474, 509 Wujek, D. E., Bicudo, C. E. (1993), 526, 538, 557 Wujek, D. E., Bland, R. G. (1988), 503, 509 Wujek, D. E., Bland, R. G. (1991), 526, 536, 541, 557 Wujek, D. E., Gardiner, W. E. (1985), 503, 509, 514, 518–519, 521 Wujek, D. E., Graebner, M. (1980), 575 Wujek, D. E., Gretz, M., Wujek, M. G. (1977), 552, 557 Wujek, D. E., Hamilton, R. (1972), 552, 557 Wujek, D. E., Hamilton, R. (1973), 552, 557 Wujek, D. E., Hamilton, R., Wee, J. (1975), 552, 557 Wujek, D. E., Igoe, M. J. (1989), 541, 557 Wujek, D. E., Kristiansen, J. (1978), 531, 557 Wujek, D. E., Saha, L. C. (1991), 513, 521 Wujek, D. E., Saha, L. C. (1995), 485, 509 Wujek, D. E., Saha, L. C. (1996), 485, 509 Wujek, D. E., Thompson, R. H. (2001), 502–503, 509 Wujek, D. E., Thompson, R. H. (2002), 499, 503, 509 Wujek, D. E., Thompson, R. H., Timpano, P. (1996), 758, 765, 771, 773 Wujek, D. E., Timpano, P. (1984), 524, 557 Wujek, D. E., Timpano, P. (1986), 197, 201, 210, 221, 224 Wujek, D. E., Wee, J. L. (1983), 524, 526, 534, 551–552, 557 Wujek, D. E., Wee, J. L. (1984), 453, 458, 469 Wujek, D. E., Weis, M. M. (1984), 552, 557 Wujek, D. E., Welling, M. I. (1981), 586 Wunsam, S., Schmidt, R., Klee, R. (1995), 583 Wurtsbaugh, W. A., Berry, T. S.(1990), 43, 57 Wynn-Williams, D. D. (2000), 121, 196 X Xavier, M. B., Mainardes-Pinto, C. S. R., Takino, M. (1991), 408, 422 Xie, P., Iwakuma, T., Fujii, K. (1998), 700, 714 Y Yamagishi, T., Couté, A. (1995), 416, 422 Yan, N. D., Stokes, P. (1978), 699, 714 Yang, J.-R., Duthie, H. C. (1995), 604, 636 Yang, J.-R., Pick, F. R., Hamilton, P. B. (1996), 587 Yin-Xin, W. J., Kristiansen (1994), 538, 557 Yoder, C. O., Rankin, E. T. (1995), 794, 796, 804 Yoneda, Y. (1949), 763, 773 Yong, Y. Y. R., Lee, Y.-K. (1991), 45, 57 Yoshida, T. (1959), 202, 224 Yoshikawa, T., Takishita, K., Ishida, Y., Uchida, A. (1997), 686, 699, 714 Yoshizaki, M., Iura, K. (1991), 763, 773 Yoshizaki, M., Miyaji, K., Kasaki, H. (1984), 758, 760, 771, 773 Youngs, H. L., Gretz, M. R., West, J. A., Sommerfield, M. R. (1998), 200, 224 Yung, Y.-K., Sawa, T., Stokes, P. M. (1986), 349, 352 Yung, Y.-K., Stokes, P., Gorham, E. (1986), 40, 57, 205, 224, 363, 381 Z Zacharias, O. (1898), 36, 57 Zakrys, B. (1986), 388, 399–400, 402–405, 407, 409, 412, 415, 422 Zakrys, B. (1994), 388, 403, 422 Zakrys, B. (1997a), 388, 407, 422 Zakrys, B. (1997b), 388, 407, 422 Zakrys, B., Kucharski, R., Moraczewski, I. (1997), 407, 422 Zakrys, B., Moraczewski, I., Kucharski, R. (1996), 404, 407, 422 Zakrys, B., Walne, P. L. (1994), 388, 390–391, 394–395, 397, 399, 402–405, 407, 410, 412–416, 422

Author Index Zakrys, B., Walne, P. L. (1998a), 402, 422 Zakrys, B., Walne, P. L. (1998b), 403, 422 Zalessky, M. M. (1926), 74, 116 Zeeb, B. A., Smol, J. P. (1991), 537, 557 Zeeb, B. A., Smol, J. P. (1995), 537, 557

Zeimann, H. (1991), 787, 804 Zelinka, M., Marvan, P. (1961), 775, 787, 791, 804 Zimmerman, W. (1928), 768, 771, 773 Zohary, T., Breen, C. M. (1989), 64, 116 Zohary, T., Robarts, R. D. (1990), 64, 116

883


Subject Index

Accumulation body (-ies), 690–691 Acid (-ic), Acid environment (also acidification), 6, 7, 40–42, 66, 81, 87–88, 184, 204, 227, 284, 363–364, 370–379, 407, 424, 428, 487–489, 513, 535–537, 552, 560–561, 602, 604, 627, 640, 647–650, 661–662, 664–665, 676, 682, 699, 701, 792–793, 814, 823 Acid mine drainage, 41–42, Acid precipitation (or deposition), 39, 44, 513, 571, 607, 661, 792–793 Acid soils, 206, 369 Acid spring (stream), 41, 80, 208, 498 Acidic (lower) pH, 7, 40–41, 44, 204, 206, 208, 280–281, 341, 363, 371, 406, 407, 412, 427–428, 433, 488–489, 513, 535–536, 552, 602, 607, 617, 627, 650, 665, 672, 676, 682, 699, 739, 814, 823 Adirondack Mountains, 44, 526, 536–537, 541, 552, 792 Aerotope (also Gas vacuole), 20, 61, 70, 72,

81–82, 87, 89, 92, 122–123, 125–126, 129, 132, 139, 141, 144, 147, 151, 155, 158, 164, 166, 169, 171, 174, 177, 180–181, 184 Akinete – see Cyst, Akinete Alabama, 210, 303, 345, 459, 463 Alaska, 13, 41, 80, 89, 101, 129, 137, 141, 150, 203, 212, 365, 377, 379, 425, 427–428, 433–434, 439, 441, 443, 446–447, 449, 451, 453–454, 458–459, 461, 463, 487–488, 552, 561, 585, 628, 649 Alberta, 552 Algae, Acidophilic, 27, 41–42, 80, 206, 208, 227, 363, 375, 378, 489, 536–537, 607, 617, 673 Aerophytic (also aerophilic), 64, 67, 277, 284, 288, 623, 638, 643, 646–649, 661 Alkalophilic, 66, 72, 79, 86, 129, 158, 187, 204, 338, 537, 602 Auxotrophic, 383, 385, 387, 401, 406, 409, 701

Benthic, 5, 8, 9, 23–28, 30–36, 38–39, 42–43, 65–66, 68, 79–80, 82, 89, 124–126, 129, 132, 139, 147, 150–151, 155, 161, 169, 173–174, 177, 180–181, 199, 205, 210, 316, 363, 371, 489–490, 565, 567, 583, 601–606, 608, 610, 613, 616, 646–648, 650, 661–663, 664, 666, 672, 676, 681–682, 757–758, 762–764, 766, 778, 780, 782–783, 785–786, 808–809, 819, 825–826 Cryophilic, 44 Culture (laboratory), 59–60, 63, 67–68, 81, 99, 121, 126, 131, 141, 216, 131, 141, 216, 227, 229, 235, 240, 243–246, 248, 254–258, 291, 315–316, 333, 335–338, 340–341, 345–346, 365, 374, 388, 393–394, 402–411, 428, 432–433, 461–465, 487–489, 497, 501, 512–514, 569, 605, 694, 699, 700, 709, 735, 739–740, 746, 748–749, 751, 758, 760–761, 763, 765–767, 785–786, 806–807, 810 Edaphic – See Algae, soil,

885

886

Subject Index

Algae (continued) Endolithic, 6, 66–68, 71–72, 104, 107–109, 314–315 Endophytic, 45, 72, 95, 107, 180, 259, 274, 295, 298, 314–316, 319–321, 330–332, 342–343, 435, 441, 461, 770 Epilithic, 24–26, 35–36, 43, 65, 67, 71, 101–102, 107, 158, 164, 169, 174, 180–181, 187, 205, 208, 210, 212, 346, 424, 433, 441, 454, 456, 458, 461, 500, 681, 763, 765, 767, 779 Epipelgic, 27, 38, 40, 43, 65, 80, 88, 120, 137, 139, 180, 406–408, 433, 453, 463, 603, 638, 644, 646–648, 650, 669–670, 674–676, 681–682 Epiphytic, 24–28, 34, 38, 39, 40, 45, 65, 67, 71, 76, 79, 81, 87, 96–97, 101–104, 107–109, 120, 122, 124, 132, 150, 158, 161, 164, 166, 169, 205, 208, 210–12, 271–271, 274, 284, 291, 298, 303, 312, 315–316, 319–338, 341–345, 347, 424–435, 432–433, 435–437, 441–444, 446, 450–456, 458–459, 461–463, 482–483, 492, 495–500, 603, 662, 672, 674, 681, 707–709, 767, 779, 810 Epizooic, 12, 88, 171, 212, 271, 303, 319, 339–340, 384–385, 388, 393–394, 411, 433 Filamentous, 2–4, 6, 8, 20, 22, 24–28, 3234, 36, 39–43, 61–63, 65–70, 81, 101–102, 107, 117–190, 199–221, 260–261, 264, 267, 270, 272, 288, 298, 311–349, 353–355, 357, 359–360, 362–374, 376–379, 429–433, 438, 441–443, 446, 451, 454, 458–463, 473, 488, 491–494, 496–497, 499, 502, 674, 696, 703, 709, 758, 767, 783, 805, 808–810, 812–824, 826 Gelatinous, 1–3, 60–61, 69–71, 79–81, 89, 91–92, 95–97, 101–104, 110, 124–125, 141, 144, 151, 166, 177, 180, 184, 187, 189, 198–200, 232–236, 239–249, 259, 262, 264–267, 270, 280, 284, 288, 291–292, 294–295, 297, 301, 304, 321, 332–333, 344, 361, 363, 365, 367, 369, 371, 375–377, 429, 435, 437, 441, 443–444, 453–455, 458, 474, 476, 481–483, 488, 493–503, 643, 707, 767, 770, 819 Lithogenic, 67, 81, 96, 121 Mat, 6, 26, 40–41, 44, 60, 63, 65–67, 79, 104, 119–126, 129, 132, 135, 137, 139, 141, 144, 147–151, 155, 158, 161, 164, 166, 169, 171, 174, 177, 180, 184, 189, 198–199, 314–316, 319, 337, 340, 344, 363, 365, 429, 463, 779, 805, 809, 817, 822–823, 816 Neustonic, 291, 423–433, 441, 490, 493, 496, 501 Planktonic, 5–9, 19–23, 26, 32, 36–38, 42–43, 59, 61, 64–65, 67–68, 70–76, 79–89, 92–95, 119–125, 129–132, 135, 139–144, 147, 151, 155, 166–167, 169–172, 174–177, 180, 227, 257–260, 267–306, 314–317, 323, 344–346,

363–365, 372, 376, 378, 383, 406–409, 412–413, 424–428, 432, 443, 446–454, 458–459, 461, 473, 478, 484–490, 492–503, 512–518, 541, 548, 567, 570, 585–586, 595, 598, 606–607, 617, 650, 672, 681, 694, 700, 702, 707–709, 738–739, 805–806, 812–813, 819, 822–823, 825 Snow, 43–44, 81, 130, 226–227, 235–236, 306, 315–316, 318, 346, 364, 367, 369, 407, 433, 810 Soil, 6, 11, 45, 60, 63, 72, 79, 82, 89, 95, 120–121, 126, 129, 135, 137, 141, 144, 150, 158, 166, 177, 180–181, 184, 198–199, 206, 208, 210, 253, 255, 258, 270, 272, 274, 276, 280, 282, 284, 291, 294–295, 298, 301, 303, 315–322, 332–337, 341–342, 344–347, 363, 369, 372, 404, 407–408, 424–425, 429, 433, 451, 453, 463, 605, 638, 648, 672, 676, 812, 814 Subaerial, 6, 60, 63–64, 70, 79–80, 82, 92, 104, 119, 121–122, 135, 137, 139, 144, 151, 158, 161, 166, 169, 174, 177, 181–184, 187–189, 258, 267, 272, 274–276, 289–301, 306, 315–316, 319, 321–322, 343, 345–346, 363, 369–374, 490, 584, 607, 646 Unicellular (or solitary), 1–2, 6, 8, 9, 24, 44, 60–62, 65, 68, 70–84, 92–96, 129, 199–201, 205, 207–209, 225–239, 253–259, 268–270, 274–276, 284–291, 294, 297–298, 317, 322, 354–363, 372, 379, 383, 388, 410–415, 424, 427–429, 436, 439–459, 464–465, 473–475, 477–478, 480, 482, 491–501, 524, 541, 548, 559, 600–601, 606, 623, 627, 676, 681–682, 685, 715, 740, 757 Algicide, 817, 821–822, 824, 826 Alkaline (higher) pH, 41–42, 63, 189, 204, 208, 210, 212, 218, 221, 364, 370, 412, 428, 433, 461, 489, 535–536, 552, 584, 602–603, 617, 622, 627, 639, 640, 643–644, 646, 650, 661 Alkaline environments, 40–41, 66–67, 72, 79, 86–87, 95–96, 104, 129, 161, 164, 166, 171, 177, 184, 187, 204, 212, 218, 220–221, 338, 369–370, 372, 374, 407, 535, 552, 584, 602–603, 617, 622, 627, 639–640, 643–644, 650, 661–662, 674, 699, 701, 738, 763 Alkaline phosphatase, 25 Alkalinity, 7, 485, 488, 536, 617, 814, 823 Allelochemical (-chemistry, -pathy), 817, 821, 826 Allochthonous (organic) matter, 31–32, 789 Allophycocyanin – see Pigment, Allophycocyanin Alternation of generations, 201, 202, 256, 312, 741 Alum (Al2[SO4]3), 813–815 Amoeboid stage (cell), 353, 362, 374, 429–430, 432, 434–436, 439–442, 450–451, 476, 480, 482, 485, 494–496, 501, 688, 696, 706, 708–709

Amylopectin, 5, 254, 736 Amylose, 5, 254, 736 Anaerobic environment, 18, 119, 227, 248, 409, 814 Anisogamy (-ous), 226, 233, 235–236, 240, 254, 312, 347, 366, 432, 485, 602, 694 Anoxic – See Anaerobic environment Antarctic, 18, 189, 227, 246, 369, 485–486, 561, 649 Antheridium (-a), 331, 338–342, 344, 432, 464–465 Antherozoid, 342 Anthropogenic effect, 584–585, 587, 661, 781 Apical pore field, 586, 600–601, 606, 611, 613, 616–617, 656, 658–659, 663–665, 690–692, 694–695, 705, 707–708 Aplanogamete, 235 Aplanospore – see Spore, aplanospore Aquaculture, 387, 512, 807 Araphid diatom, 7, 9, 560–562, 595–623, 625, 627, 629 Archeopyle – see Cyst, archeopyle Arctic, 12–14, 18–19, 26–27, 33–34, 38, 82, 98, 101, 118, 134, 137, 155, 158, 161, 164, 166, 169, 171, 180, 184, 197, 231, 276, 280, 284, 288, 295, 299, 303, 306, 342, 347, 364, 369, 407, 424, 433, 443, 446, 453–454. 458–459, 461, 463, 486, 490, 535, 552, 561, 584, 604–605, 613, 627, 649, 688, 700–701, 704, 738, 764 Areola (-ae), 572, 574, 584, 599, 600–603, 606–607, 613, 616, 623, 627, 638, 640–641, 643–644, 648–650, 656, 659 Arizona, 40, 44, 139, 208, 210, 439, 552, 628, 810 Arkansas, 41, 203, 425–428, 433–434, 441, 443, 446–447, 449, 451, 453–454, 458–459, 461, 463, 478, 820 Artificial substratum (-a), 789 Clay tile, 26–27, 36 Glass slide, 25, 313, 315–316, 332, 394, 411, 463, 763, 771 Plexiglas, 25 Styrofoam, 25 Ash free dry mass, 782–783 Astaxanthin – see Pigment, Astaxanthin Athecate dinoflagellate, 691, 704, 706, 708–709 Aufwuchs, 24, 314 Australia, 7, 65, 218, 316, 503, 524, 526, 537, 541, 548, 552, 709, 759, 765–766, 768, 794 Autospore – see Spore, autospore Autotrophic (growth or metabolism), 20, 37, 63, 66, 120–121, 202, 254, 387, 406, 489, 514, 534, 685–686, 701, 739–740 Auxospore – see Spore, Auxospore Auxotrophic metabolism – see Algae, Auxotrophic Axial area, 606–607, 624, 627, 639–640, 643, 645–646, 648–650, 662–664

Bacteria (Eubacteria), 59, 61–62, 81, 384, 406, 668, 697, 739–740, 808, 810, 825–826

Subject Index Chemoautotrophic, 18, 41 Epiphytic, 415 Heterotrophic, 20, 23–24, 36, 41, 44, 406, 409, 489, 490, 497, 499, 817 Symbiotic (also endosymbiotic), 490, 492, 494, 497, 499, 697, 739, 744, 747, 749–751 Bacterivory (see also Phagotrophy, Mixotrophy), 20, 23, 41, 383, 441, 473, 489, 697, 740 Bahamas, 89, 96, 107 Beetle, 41, 205 Belize, 66, 72, 80, 212, 218, 702, 707 Benthic, Diatom, 7, 26–27, 36, 39, 490, 565, 583, 595, 601–602, 604–606, 608, 610, 613, 616, 646–648, 650, 661–664, 666, 672, 676, 681–682 Habitat, 37, 24–28, 32–37, 80, 120, 139, 161, 181, 565, 659, 663, 567, 648, 659, 664, 670, 782 Invertebrate, 25–26, 31, 34–36, 39, 783, 825 Bermuda, 89, 107, 151, 459, 481 Beta-carotene (also b-carotene), 254, 474, 560, 690, 735, 757 Bicarbonate (HCO3–), 41–42, 204, 738–739, 812, 823 Big Soda Lake (Nevada), 15, 42 Bioassay, 22, 39, 68, 785–786, 821 Biodiversity, 6, 12, 19–20, 32–34, 62–63, 118, 199–200, 225–226, 311, 354, 387–388, 424, 427, 429, 473, 512, 561, 598, 779, 786, 788 Biogeography, 12, 19, 32–34, 60, 63, 67, 203–205, 486–487, 597, 605, 765–766 Biomanipulation, 22, 817–819 Biomass, 19–20, 22–27, 31–32, 34–40, 43–44, 63–66, 120, 155, 204–205, 423, 487–488, 513, 535–538, 605, 664, 670, 699–701, 776, 779–780, 782–783, 785–786, 789, 794–795, 806, 812–814, 816–818, 822–823, 825 Biovolume (also Cell volume), 24, 36, 598, 670, 776, 782–784, 789 Biraphid diatom, 9, 595–597, 604, 613, 637–650, 655–666 Bird bath, 45 Bloom, 19, 23, 37, 43, 53, 60, 64–65, 68, 72, 81, 89, 92, 117, 120, 141, 147, 169–174, 177, 227, 254, 274, 315, 335, 364, 370, 372, 405–407, 409, 411, 415, 428, 473, 487, 489, 503, 511–512, 513–514, 535–537, 572, 585, 587, 607, 617, 699–701, 783, 793, 805–807, 812–817, 821, 824, 826 Bog, 14, 19, 38, 40, 66, 88–89, 189, 198, 205, 210, 215, 267, 276, 280–281, 284, 291, 295, 298, 301, 303, 306, 335, 339–341, 344–345, 363, 369–373, 375, 378, 407, 424–429, 433, 441, 443, 446–447, 449, 451, 453–454, 458, 463, 488, 490, 498, 537, 605, 613, 639, 646, 661–662, 665, 672, 709 Boreal, 34, 171, 205, 215–216, 433, 441,

446, 489, 552, 567–568, 583, 605, 613, 616, 661, 765 Brackish, 60, 64, 72, 82, 88, 132, 151, 166, 171, 177, 199, 202, 205, 208, 220–221, 311, 338, 347, 413, 432, 463, 485, 512, 514, 515, 548, 586, 607, 638, 646–648, 650, 676, 701, 707, 757, 763–764, 768, 770–771 Brazil, 84, 138, 140–141, 202, 204, 807 British Columbia, 13, 17, 42–43, 95, 101, 132, 137, 147, 174, 181, 184, 189, 233, 235–237, 239, 243–244, 267, 270, 276, 280, 284–285, 298, 301, 339, 363, 370, 412–413, 415, 425, 428–429, 443, 446–447, 449, 451, 453–454, 458–459, 461, 463, 552, 561, 702, 709, 762, 766 Bryophyte, 4, 45, 205, 432, 441, 443, 461, 584, 662 Buoyancy, 20, 23, 488, 539, 567, 690, 805

C:N ratio, 39 C:P ratio, 819 Caddisfly (Trichoptera), 26, 105, 819 Calcification, 44, 137, 147, 152, 169, 239, 366, 513–515, 767, 770–771 Calcium (Ca2+), 239, 363, 364, 366, 374, 376–377, 497, 763–764, 766, 814 Calcium carbonate (CaCO3), 27, 39, 44, 59, 66, 69, 79–80, 89, 119, 137, 147, 158, 166, 169, 187, 206, 237, 311, 316, 319–320, 332, 334–335, 338–339, 759, 763–764, 767, 813–814, 819–820, 823 California, 42, 78, 80–81, 89, 92, 94–95, 100, 102, 104–105, 108–110, 139–140, 149, 201, 216, 246, 364, 377, 434, 459, 463, 561–562, 630, 707, 767, 817 Canadian Shield, 171, 363, 488, 513 Canopeum (-a), 647 Carbon (source), 31–32, 39, 44, 204, 406, 604, 647, 738–740, 793, 807, 812 Carboxysome, 61, 200 Caribbean, 66, 72, 80, 87, 95, 104, 150, 174, 212, 218, 220, 267, 270, 274, 276, 280, 284–285, 289, 291, 295, 298, 302–303, 306, 332, 707 Carotenoid – see Pigment, Carotenoid Carp, 806, 814, 819–821, 826 Carpogonial branch, 202–213, 215–216, 218 Carpogonium, 201–202, 205, 210–216, 218–219 Carposporangium, 211–216, 218 Carpospore, 199, 201–202, 212, 215–218 Carposporophyte, 201–201, 207–208, 211–218 Catfish, 739, 806–807 Cave, 6, 11, 44, 125, 187, 197, 208, 605 Cedar Bog Lake (Minnesota), 14, 40 Cell wall, 1–9, 26, 59, 61–62, 118–120, 125–126, 129, 132, 135, 137, 139, 144, 147, 151, 158, 161, 174, 177, 180–181, 200, 226, 228–229, 231–236, 242, 246, 248, 255, 257, 260–261, 263–267, 270, 274, 276–277, 280–282, 284, 288–289, 291, 295, 298–299, 301, 303, 306,

887

311–312, 318, 322, 353–354, 357, 362, 365–369, 372–376, 378, 388, 425, 427, 430–432, 434–438, 441, 332, 445–447, 449, 451, 453–454, 457–458, 461, 494–495, 500, 503, 559–561, 597–599, 601, 604–605, 669, 736, 757, 775, 783, 823 Cellulose, 2, 5–6, 200, 226, 255, 311–312, 343, 431, 484, 498, 686–687, 690–691, 698, 703, 706, 757 Central America, 60, 67, 81, 90, 94, 100, 102, 147, 164, 171, 267, 270, 765 Central Canada, 73, 80, 85, 89, 96, 156, 164, 171, 267, 270 Centroplasm, 61, 135 Chantransia stage, 201–202, 212–216, 218, 765 China, 387, 513, 536, 759, 763, 765, 771, 807, 820 Chironomid (Midge), 205, 319, 819, 825 Chitin, 255, 411, 473, 501, 565, 567, 584, 587 Chloride (Cl–), 514 Chlorophyll-a – see Pigment, Chlorophyll-a Chlorophyll-b – see Pigment, Chlorophyll-b Chlorophyll-c – see Pigment, Chlorophyll-c Chloroplast (see also Plastid), 5–8, 41, 197–200, 202, 207–210, 212, 215–216, 218, 225–226, 229, 231–237, 239–244, 246, 248, 254, 256–257, 259–267, 270–272, 274, 276–277, 280–282, 284–285, 288–289, 291–292, 295, 298–299, 301–303, 306, 311–312, 317–322, 330, 332–347, 353–355, 357, 361–362, 365–379, 384–391, 394–397, 401–404, 411–415, 472, 474, 491, 493–503, 512, 515–518, 524–525, 528, 531, 533, 548, 559–560, 567, 569, 572–573, 575, 576, 585, 608, 617, 646, 690–691, 697, 701, 704, 707–709, 717, 719, 724, 726, 731, 735–739, 742–748, 750–751, 757, 759–760, 761–762, 767, 768–771 Chromatic adaptation – see Photoacclimation Chromatophore – see Chloroplast Chromatoplasm, 61, 69, 72, 79, 81, 87–88, 101, 121, 135, 147 Chrysolaminarin – see Storage product, chrysolaminarin Chytrid, 364, 379, 739 Ciliate, 20, 45, 272, 385, 701, 738–739, 749–750 Cilium (-ia), 312, 385, 512 Cingulum, 599–603, 612–623, 648–650, 681, 686–687, 690–692, 694–695, 703–704, 706–707 Cladistic, 239, 244, 388, 597–598, 604, 659, 742 Cladocera (-an), 16, 20, 38–39, 408, 411, 700, 808, 825 Classification Brown algae, 757–759 Chrysophytes, 471–473, 484–485 Cryptomonads, 715–716, 740–743, 749 Cyanobacteria, 62–63

888

Subject Index

Classification (continued) Diatoms, 559–561, 563–565, 595–604, 637–638, 655–661, 669–670, 675 Dinoflagellate, 685–686, 697–699 Euglenoids, 383–385, 387 Eustigmatophyte, 423–424 Green algae, 228–229, 232–233, 236–237, 239–240, 243–246, 253–254, 312–313, 353–354 Haptophyte, 511–513 Major groups, 5–9 Raphidophyte, 427–428 Red algae, 197–199 Synurophyte, 523–524, 533–534 Tribophyte, 423–424, 429–432 CO2 (carbon dioxide), 15, 39, 59, 204, 365, 474, 805 Coastal (habitat), 14, 16, 18, 40, 64, 66–77, 72, 78, 80–82, 88, 95–96, 101–102, 107–108, 110, 150, 155, 158, 177–178, 203, 258, 315, 332, 335, 338, 363, 463, 489, 512, 552, 583–584, 586, 661,768 Coccolith, 511–516, 518–519 Coccolithophorid, 43, 511–513 Coenobium (-ia), 237–240, 246, 261–263, 267, 274, 276, 284, 288, 295 Coenocyte (-cytic), 7, 254, 266–267, 276–277, 298, 311, 319, 344, 429, 431–434, 463–464 Colony (-ial), 1–3, 3, 6, 8–9, 20, 27, 37, 60, 69–72, 79–110, 119, 122–125, 132, 135–139, 144, 158, 161–169, 171–173, 177, 179, 180–181, 207–208, 225–229, 232–233, 239–248, 253–306, 315–316, 325, 332, 335, 344, 354, 356, 360, 368, 374–376, 388, 405, 410–411, 424–425, 429, 434–437, 443–435, 450–464, 473–485, 489–503, 524–528, 531, 533–535, 539, 541, 548, 565, 567, 571–574, 583–587, 598, 605–608, 613–614, 616–617, 622–623, 638, 647, 661, 664–665, 673–677, 681, 707, 739, 744, 757–759, 762–768, 770, 777, 784, 807, 810–811, 816 Amorphous (irregular), 60, 69, 70–72, 79, 81, 88–89, 92, 95–96, 102, 104, 107, 119, 122, 141, 147, 155, 177, 180, 187, 208, 260, 263–264, 274, 276, 284, 288, 298, 295, 301, 303, 320, 322, 332, 334, 336–337, 340, 343, 354, 443, 452–453, 455, 498, 768 Autocolony, 226, 239, 246 Chain (chainlike), 81, 199, 282, 341, 431, 437, 452–453, 494, 496, 501, 601, 616–617, 623, 640, 676 Coenobium, 239–240, 246, 261–263, 267, 274, 276, 284, 288, 295 Cube-like, 89, 264, 295 Daughter colony, 226, 241, 243–245 Hemispherical, 72, 96, 107, 124, 137, 151, 166, 330, 332, 334, 499, 770 Moniliform, 102, 121–122, 125, 139, 177, 184, 187–189 Plate-like (flat), 69, 87, 280, 317 Radial (radiating), 69–71, 81–82, 87, 89,

96, 104, 107, 124, 135, 144, 166, 169, 199, 240, 243–244, 264–265, 267, 270, 284, 298, 301, 314, 337, 340, 488, 496, 770 Saccate (sacklike), 125, 207, 209–210, 246, 259, 303 Spherical, 69–70, 72, 79, 81–82, 87, 88–89, 92, 95–96, 119, 123–125, 147, 180, 187, 233, 242–243, 258, 284, 288, 291–292, 295, 301, 303, 306, 320, 330, 340, 435, 437, 443, 452–454, 493, 496, 526, 528, 541, 548 Zigzag, 3, 318 Colorado (CO), 17, 86, 454, 459, 647, 702, 738, 746, 748–749, 751, 763, 766, 770 Colorado River, 34 Condensed chromosomes, 7, 686, 690 Conjugation – see Reproduction, conjugation Connecticut (CT), 80, 158–159, 425, 427, 459, 461, 463, 489, 563, 552, 770, 772 Contractile vacuole, 226, 229, 231–237, 239–240, 242–244, 246, 248, 254, 272, 274, 292, 295, 301, 332, 392, 401, 409, 427–430, 434, 439, 441, 443, 472, 485, 494–503, 514–518, 716–717, 719, 724, 731, 736, 747 Copepod, 20, 39, 212, 411 Copper sulfate (CuSO4), 807, 816, 821–826 Core sample, 571 Cortex, 4, 210, 212, 216, 218, 220, 338 Cortical filament (or cells), 3–4, 207–208, 210–212, 215–217, 220 Costa Rica, 13, 40–42, 210, 212, 218, 221, 235, 343, 415 Costae, 357, 367, 372, 571–572, 583, 586, 600, 602, 616, 623, 640, 643, 648, 681 Crater Lake (Oregon), 12–15 Cricolith, 511, 515, 518 Crust (algal), 4, 24–25, 33, 36, 63, 80, 107, 110, 136–137, 181, 199–200, 204, 206–207, 215, 218, 314, 316, 320, 328, 340, 364–366, 376, 458, 488, 605, 757–760, 763–765, 767–768, 813 Crustacean (micro-), 260, 271, 393, 404, 410–411, 454, 490, 813 Cryophilic algae – see Algae, cryophilic Cuba, 66, 72, 76–66, 79–84, 86, 93, 99, 101, 103–105, 107, 109, 131, 142, 145, 150, 156, 163, 175, 184–185, 189, 291, 303, 307, 454, 702, 707 Culture (studies) – see Algae, culture Current velocity, 28–30, 34–37, 200, 203–204, 206, 210, 212, 215–216, 218, 433, 473, 764, 766, 779 Cyanelle, 263, 266, 270, 284, 385 Cyanophage, 819 Cyanophycean starch – see Storage product, cyanophycean starch Cyclomorphosis, 255, 488 Cyprinoid fish, 205, 212 Cyst (see also Spore) Akinete, 19, 119–121, 124–125, 139, 151, 155, 158, 161, 164, 166, 169, 171, 174, 176–177, 180–181, 184, 189, 210, 246, 295, 312, 320–321, 325, 327–328, 330,

332–341, 343–347, 355, 362, 372, 432, 442, 453, 462–463, 818 Coenocyst, 344 Hypnospore, 43–44, 245–246, 463 Hypnozygote, 226, 240, 243–244, 694 Planozygote, 248, 694 Tomont, 1 Zygospore, 19, 330, 355–356, 362–363, 369–379

Dam, 16–17, 28, 31, 34 Darling River, 65 Daughter cell – see Reproduction, daughter cell Delta Marsh (Manitoba), 38 Denmark, 536, 794 Desert, 6, 29, 32, 40, 42, 60, 66, 121, 141, 203 Desiccation, 26, 39, 45, 123, 206, 255, 363, 734 Desmid, 2, 6, 8, 354, 356–379, 458, 776 Desmokont – see Flagellum, desmokont Detritus, 24, 26, 31, 35, 38–39, 132, 169, 205, 378, 409, 674 Diadinoxanthin – see Pigment, Diadinoxanthin Diatomite, 573, 584–585, 666 Diatoxanthin – see Pigment, Diatoxanthin Dictyosome – see Golgi body Dinokaryon, 697 Dinokont – see Flagellum, dinokont Dinospore, 697 Disjunct, 12, 276, 433, 764–766 Dispersal, 33, 70, 202, 407, 486, 760, 765, 795, 824 Diversity (see also Biodiversity), 12, 19–20, 24–25, 32–36, 39–44, 62–67, 118, 197–199, 225–226, 254, 311, 354, 387–388, 424, 427–431, 473–474, 512–513 Evenness, 782, 784 Shannon index, 784, 787 Species richness, 40, 776, 779, 782, 784 Drinking water, 6, 16, 685, 806, 809 Dystrophic – see Lake, dystrophic

Eastern Canada, 96, 126, 270, 280, 284–285, 288, 291, 295, 298, 302–303, 306, 365, 377, 485, 561 Eastern North America, 12, 40, 42, 137, 169, 203, 205, 210, 216, 280–281, 332, 377, 424, 459, 488, 513, 560–561, 650, 661 Eastern United States, 92, 189, 210, 270, 274, 276, 280–281, 284–285, 288–289, 291, 295, 298, 301–303, 306, 315, 332, 340, 344, 363–364, 377, 459, 461, 485, 561, 665, 792, 811 Ecdysis, 691, 694, 703, 708 Ecological risk assessment (ERA), 776–777 Ecomorphotype (also Morphotype), 67, 89, 104, 135, 189, 406, 410, 461, 485, 569 Ecorticate, 317, 338–339

Subject Index Ejectisome, 715–722, 724, 726–727, 731–732, 734–735, 741–742, 744, 746, 748–751 El Salvador, 15, 41, 206 Electron microscopy (EM) – see Microscopy, electron Elevation, 15, 204, 565, 638, 647, 649, 728 Ellesmere Island, 144, 166, 203, 303 Encrusting algae – see Algae, encrusting Endemic species, 12, 16, 66, 72, 80, 82, 87, 89, 101, 227, 365, 377, 568, 656 Endogloeic algae – see Algae, endogloeic Endolithic algae – see Algae, endolithic Endophytic algae – see Algae, endophytic Endosymbiont (-biosis, also Symbiont), 11, 36, 45, 63, 180, 226, 258, 263, 271, 284, 306, 311, 315, 385, 387–388, 402, 490, 492, 497, 499, 670, 673–676, 681, 686–687, 697, 735–737, 739 English Lake District, 21, 487, 513, 696 Environmental optima (also Ecological optima), 23, 26, 44, 206, 487, 513, 597, 617, 784, 791–792 Environmental Protection Agency (EPA), 776–777, 786, 788, 826 Epilimnion, 17–18, 23, 258, 535, 567, 699, 738, 794 Epilithic algae – see Algae, epilithic Epipelic algae – see Algae, epipelic Epiphytic algae – see Algae, epiphytic Epipsammic algae – see Algae, epipsammic Epitheca, 687, 689, 690, 693–694, 704, 706, 708 Epizooic algae – see Algae, epizooic Eubacteria – see Bacteria, Eubacteria Euphotic zone (also Photic zone), 18, 20, 258, 539, 814–815 Eutrophic environment (see Lake, eutrophic) Eutrophication, 19, 21, 23, 27, 64, 129, 571, 586, 662, 775, 781–782, 806, 810–811 Everglades (Florida), 39, 66, 72, 131, 133–134, 140, 154, 158, 162, 173, 184, 434, 794–795 Eversion (also Inversion), 228, 239, 242, 244 Evolutionary (relationships), 41, 60–62, 226, 257, 313, 362, 384–385, 387, 391, 401–402, 405, 559, 567–570, 582, 656–657, 659, 674, 690, 697, 699, 737 Exocyte – see Reproduction, exocyte Exospore – see Spore, exospore Experimental Lakes Area (ELA), 26, 129, 434, 486, 488, 537 Eyespot, 226, 229–233, 235–237, 239–240, 242–244, 246, 248, 301, 332, 383–385, 388, 391–392, 397, 399, 401–402, 411–4388, 391–392, 397, 399, 401–402, 411–413, 439, 472, 494–503, 686, 690–691, 706–709, 742

FAA – see Fixative, FAA Facultative heterotroph, 254, 406 False branching, 3, 119, 123–126, 132, 141, 153, 155, 158–161, 164–165, 168, 184, 189, 207–209

Fatty acid, 35, 147, 387, 408, 823 Fibulae, 670, 672, 675–676, 681 Filament (true; see Algae, filamentous) Finger Lakes (New York), 12–14, 18, 42 Finland, 486, 489, 699 Fish, 6, 22–23, 31, 38, 43, 171, 205, 313–314, 335, 387, 408, 514, 685, 687, 697, 701–703, 708, 778, 786–787, 789, 806–807, 810–811, 814–816, 818, 820–821, 824–825 Fish kill, 685, 806, 810, 812, 814, 816–817, 823, 825 Fixative FAA, 259, 316, 365, 491 Formaldehyde (formalin), 67–68, 206, 227, 259, 316, 365, 491, 514, 539, 738, 767 Glutaraldehyde, 67–68, 206, 227, 259, 409, 463, 465, 491, 539, 702, 703, 767 Lugol’s iodine, 67, 206, 259, 409, 463–465, 491, 514 M3, 259 Nissenbaum’s solution, 539 Osmic acid, 514 Transeau’s solution, 491 Flagellate, 2, 5–8, 40, 45, 225–248, 255, 257, 383–416, 423–428, 439–454, 471, 490–503, 511–519, 524–528, 685–709, 715–751, 806–807, 814, 822 Flagellum (-a), 1–2, 5–6, 8, 20, 23, 197, 227, 229–230, 235, 237, 254, 316, 339, 341, 387, 393, 401–402, 405–406, 408, 410, 412, 424, 427–429, 441–443, 472–473, 484, 490–491, 501, 512, 514, 518, 524–525, 527–528, 531, 534, 539, 541, 548, 685–690, 694, 696, 702, 716–717, 719–724, 726–734, 741–744, 746–747, 749–751, 759–760, 768, 770 Heterokont (unequal), 1, 229–231, 334, 388, 408, 413, 426430, 434, 439, 492–493, 495, 498–499, 501–503, 512, 515, 548, 719, 721, 724, 726, 728–731, 757 Isokont (equal), 1, 226, 229–244, 246, 248, 256, 322, 332, 388, 410, 432, 493, 513, 515, 519 Subapical, 7, 392, 401, 410–414, 428–429, 435, 439–440, 733–734, 751 Tinsel, 392, 402, 423, 472–473, 491, 495, 515, 528, 686–687, 717, 734, 742, 744, 746 Whiplash (smooth), 225–226, 254, 423–424, 427, 432, 472, 495, 500, 512, 528, 686, 690 Florida, 15–16, 23, 41, 45, 66, 72, 86, 88, 99, 102, 104, 110, 128, 131, 134, 139–141, 151, 154, 156, 158, 162, 173–174, 184, 187, 216, 218, 221, 371, 377, 433–434, 459, 463, 514, 526, 535–536, 541, 552, 794, 806–807, 809, 811–812 Floridean starch – see Storage product, Floridean starch Food web, 31–32, 34–35, 39, 488, 595, 810, 826

889

Fossil Algae, 387, 687, 706, 793 Cyst, 687, 698 Diatom, 559, 561, 572, 582–587, 661 Pigment, 43, 781 Pollen, 781 Record, 570, 623, 687, 697, 781 Frustule, 5, 7–9, 559–560, 562, 565–569, 571–575, 584, 587, 605, 608–609, 612–613, 616, 623, 627, 630, 638–639, 646–647, 650, 656, 658, 663, 665, 670–671, 675–676, 680–682, 775 Fucosan vesicle (also Physode), 759, 767–771 Fucoxanthin – see Pigment, Fucoxanthin Fungus (-i), 24, 44–45, 180, 236, 258, 315, 364, 514, 702, 819, 821

Gametangium (-ia) – see Reproduction, gametangium Gametophyte, 201–202, 204–205, 212–216, 218, 759 Garibaldi Lake, 13–14 Gas vacuole – see Aerotope Gas vesicle, 23, 61, 64–65, 70, 79–82, 87, 89, 92, 95–96, 101–102, 120, 122, 125, 129, 135, 141, 147, 166, 169, 177 Gelatinous envelope (or matrix; see also Mucilage), 1–3, 60–61, 69–71, 79–82, 89, 92, 95–96, 101, 124–125, 141, 151, 166, 177, 180, 184, 187, 189, 199–200, 206, 208, 210, 212, 215, 226, 228–229, 232–236, 239–244, 246, 248, 259, 262–267, 270, 284–285, 288, 291, 295, 321, 332–333, 344, 363, 365, 367, 369, 371, 375–376, 429, 437, 453–454, 458, 474, 476, 488, 493, 495–502 Gemmae – see Reproduction, gemmae Geology (-ical), 28–29, 34, 40–42, 565, 610, 764 Basalt, 29, 763–764 Granite (see also Canadian Shield), 29, 184, 764 Limestone, 14–16, 29, 36, 44, 63–65, 72, 89, 92, 102, 104, 107, 121, 164, 169, 208, 314, 316, 319, 334, 765, 811 Sandstone, 36, 65, 701, 764 Georgia, 41, 203, 463 Georgian Bay, 206, 513 Germany, 37, 488–489, 514, 526, 536, 584, 759, 768 Girdle band, 361, 372–373, 571, 573, 575–576, 584, 612, 616, 619, 625, 638, 647 Girdle lamella (-ae), 423–424, 432, 474, 528 Glass microscope slide, 25, 316, 410, 490, 514, 539–541, 562, 604–605, 681, 702, 763, 771, 780, 784 Gloeocapsin – see Pigment, Gloeocapsin Golgi body (apparatus), 385, 387, 404, 427, 472, 511–512, 516–517, 528, 531, 533, 565, 567, 734–736, 751 Gone cell, 226, 240, 242–244, 248 Gonidium – see Reproduction, gonidium Grass carp, 819–821, 826

890

Subject Index

Grazer, 23, 25–27, 31, 34–35, 38–39, 43, 205, 819 Great Lakes (Laurentian), 12–14, 18, 25–26, 40, 95, 101, 118, 150, 177, 180–181, 184, 205, 208, 212, 231, 235, 239, 267, 270, 276, 284, 292, 295, 298, 301–302, 306–307, 314, 338–340, 347, 415, 461, 499, 513, 561, 584, 587, 598, 604–605, 607, 628, 638, 659, 661, 663, 807–810, 812 Great Salt Lake, 12, 650 Great Slave Lake, 12, 14, 301 Greater Antilles, 88, 102, 151, 184, 189 Greenland, 18, 139, 144, 184, 369, 472, 486, 526, 537, 552 Guadeloupe, 75, 101, 102, 184, 235–236, 242–243, 412, 415, 446, 459 Guatemala, 14–15 Guild, 605 Gullet (also Flagellar canal, Furrow), 384, 401, 428, 716–717, 721, 724, 726, 734, 736, 741, 743–748

Haematochrome, see Pigment, Astaxanthin Hair cells (see also Seta, Trichoblast), 70, 101, 118, 122–124, 137, 151, 157–158, 164–169, 204, 208, 212–213, 320–322, 324, 330, 333–334, 337–338, 334, 758, 761, 764, 767–768, 770–771 Halophilic species, 66–657, 89, 95, 101, 585 Haplobiontic, 312, 585 Haptonema, 5, 7, 9, 498–499, 511–512, 514–519 Haptophyte scale, 511–519 Hard water (see also calcification; marl), 16, 101, 204–205, 208, 210, 338–339, 369, 376, 535, 585, 676, 681, 701, 764, 763, 811, 825 Harvest (algae or weeds), 757, 811, 817–818, 821 Hawaii, 17, 89, 96, 158, 184, 583 Heavy metal – see Pollution, heavy metal Hepatotoxin (see also Toxin), 64, 806 Herbarium, 68, 81, 206, 316, 465, 562, 588, 630, 767, 769, 772 Herbivory, 19, 22, 25, 32, 35, 64, 205, 408, 765, 783 Heterocyst – see Heterocyte Heterocyte, 20, 116, 119–125, 139, 154–155, 157–184 Heterokont flagella – see Flagellum, heterokont Heteromorphic life history, 201, 512, 696 Heteropolar morphology, 60, 62, 68, 96–102, 106–107, 121–125, 134–136, 151, 158–169, 174–175, 474, 600, 606, 613, 615–617, 676, 682 Heterotrichous morphology, 3, 313 Heterotrophic nutrition (photoheterotrophic), 27, 37, 41, 226, 254, 383, 385, 387, 401, 404, 406, 441, 473, 486, 690, 697, 714, 706–708, 740 Holdfast, 24, 26, 35, 212, 320, 393 Homologous character, 530, 569

Homonym, 89, 246, 248 Hormogonium (-ia), 78, 80, 118–120, 123, 126–127, 129, 132, 135–139, 141, 144, 147, 150–151, 153, 155–158, 160–161, 163–164, 166, 169, 174, 177, 180–181, 183–184, 187, 189 Hot spring (thermal water), 6, 40–41, 59, 66–69, 72–73, 75, 77–78–79, 82, 88, 92, 100, 102, 104, 107, 121, 125–126, 129–130, 132, 135, 139, 141, 144, 147, 158, 184, 187, 189, 197–199, 206–208, 363, 605 Hudson River, 29, 32–33, 37–39, 218 Humic material (substances), 7, 40, 485, 536, 617, 661, 665, 763, 807 Hypocone, 687, 706 Hypolimnion, 18–19, 699–701, 738, 786, 813–816 Hypotheca, 687, 689–690, 692–693, 695, 704, 706 Hyrax, 540, 562, 605

Ice, 11–14, 18, 23, 26, 43–44, 226–227, 258, 346, 363, 367, 369, 409, 433, 463, 486–487, 604, 613, 702, 715, 738, 810 Ichthyotoxin, 512, 739 Illinois, 36, 39, 118, 164, 169, 174, 463, 561 Indeterminant growth, 208, 214–215 Indiana, 15, 41, 80, 243, 246, 340, 408, 809, 815, 820, 824 Indicator species, 7, 364, 405–406, 489–490, 513, 535, 538, 560, 579, 583, 604, 655, 763, 775, 777–782, 786, 788–793, 796, 805–806 Inorganic carbon, 26, 204, 738–739 Intaglio, 571 Intercalary growth (development), 119, 124–125, 187, 210, 321, 333, 339–340, 343, 572, 587, 622, 771 Internal valve, 569, 580, 626, 643–646 Internode, 338 Invasive species (or Invader), 26, 34, 199, 205, 563, 570, 587, 765–766, 795, 811, 816 Inversion – see Eversion Iowa, 80, 337, 404, 412, 449, 454, 458, 461, 463, 536, 552 Irrigation, 180, 805, 808, 810, 817, 826 Isokont flagella – see Flagellum, isokont Isomorphic (stage, life history), 61, 72, 79, 81, 125, 759 Isthmus, 354, 361–362, 368, 373–379, 647

Jamaica, 139, 147, 221, 434, 447, 449, 459 Japan, 68, 82, 428, 513, 551, 696, 699, 759, 763, 765

Kansas, 83, 86, 339, 345, 347, 461, 463, 497, 552, 702, 807 Keel, 7, 9, 413, 529–531, 533, 551, 562, 597, 650, 655, 669–677, 681–692 Kentucky, 561, 775, 782

Keritomy (-ized), 79, 102, 122, 147 Kleptoplastidy, 690, 739

Labiate process, 565–566, 571–572, 574–575, 578–579, 581–582, 584–587, 599–601, 606–607, 611, 613, 616, 619, 623, 656 Labrador, 411, 451, 454 Lake Baikal, 12, 14, 568, 578, 613 Lake Catemaco, 82, 174, 181 Lake Erie, 20, 84, 180, 206, 210, 513, 572, 585, 806, 809, 813 Lake Huron, 26, 206, 210, 212, 314 Lake Mendota, 19, 814 Lake Michigan, 22, 27, 206, 210, 314, 586, 605, 627, 759, 765, 767, 769, 771 Lake Okeechobee, 14–15, 806, 813 Lake Ontario, 206, 210, 340, 813, 817 Lake Simcoe, 206, 210 Lake Superior, 39, 206, 454, 585, 613, 627 Lake Tahoe, 14–15, 18–19, 26, 561 Lake Washington, 23, 813, 825 Lake Arctic, 12–14, 18–19, 26–27, 33, 82, 98, 118, 134, 137, 155, 158, 166, 169, 171, 180, 184, 231, 276, 280, 284, 288, 299, 303, 306, 347, 407, 424, 433, 446, 453–454, 458–459, 461, 490, 535, 552, 561, 584, 604–605, 613, 704 Caldera, 13–16 Cirque, 13 Deep, 12–16, 18–19, 27–28, 42, 87, 107, 573, 583, 587, 670, 701–702, 765, 768, 814–815 Dimictic, 18, 605 Dystrophic, 40, 79, 376, 424–425, 427–429, 433, 439, 441, 443, 446–447, 449, 451, 453–454, 458–459, 461, 489, 616, 646, 699, 709 Eutrophic, 18–21, 23, 25–28, 31, 64–66, 81, 87, 89, 92, 120, 129, 166, 171, 174, 226, 248, 258, 274, 322, 337, 340, 364, 372, 405, 409, 428, 433, 446–447, 449, 451, 453–454, 458, 461, 473, 487, 489, 513, 535–538, 552, 568, 572, 579, 583, 585, 613, 616, 638, 647–648, 662, 699–701, 738, 763, 765, 807, 813, 816, 819, 822 Graben, 14–15 Kettle, 13–14, 702 Landslide, 14–15 Maar, 14–15 Meromictic, 16, 18–19, 42, Mesotrophic, 20, 25, 65–66, 72, 79–82, 87–89, 120, 126, 129, 135, 166, 171, 258, 274, 363, 424, 427, 433, 439, 441, 443, 446–447, 449, 451, 453–454, 458–459, 461, 499, 536–538, 583, 616, 699, 791, 813 Monomictic, 18, 42 Morainal, 14 Oligotrophic, 18–20, 22–23, 25–28, 31, 39, 64–65, 79–81, 126, 129, 139, 155, 258, 363–364, 369–370, 372–379, 451,

Subject Index 453–454, 458, 461, 473, 487, 489, 499, 513, 535–538, 567–568, 585–586, 603, 616, 627, 638–639, 643, 646, 648, 661–662, 699, 791, 818–819 Polymictic, 18 Shallow, 13, 15–17, 19, 23–24, 27, 42, 126, 150, 155, 258, 314, 335, 337, 413, 433, 495, 512, 536, 572–573, 579, 587, 595, 604, 616, 699–700, 766, 771, 808–809, 811, 814–815, 817, 822 Sinkhole, 15–16 Stratification, 17–22, 28, 258, 699–700, 738, 815 Subtropical, 15, 18, 23, 72, 79–80, 82, 87, 95, 135, 139, 161, 166, 169, 171, 174, 184, 187, 215, 315, 332, 340, 343, 377, 535, 537, 694 Tectonic, 14–15, 568, 574 Thermokarst, 13 Tropical, 13, 15, 18, 26, 34, 40, 60, 64–65, 67, 72, 79, 81–82, 129, 135, 139, 164, 171, 174, 189, 340, 365, 377, 485, 535, 552, 581, 647 Volcanic, 16 Laminarin – see Storage product, laminarin Latitude, 32, 42, 204, 486, 765 Laurentian Great Lakes – see Great Lakes, Laurentian Lentic environment – see Lake Leucoplast, 735, 742–743, 746 Leucosin – see Pigment, chrysolaminarin Lichen, 206, 306, 315, 623, 646, 758 Light High light, 407, 513 Irradiance, 17, 23, 26, 35, 38, 64, 489, 700, 766, 811 Low light, 23, 27, 44, 65, 407, 489, 567, 738 Shading, 26, 31–32, 35, 45, 203, 206, 215, 258, 258, 333, 343, 412, 702, 763, 765, 784–785, 819–812, 815–816, 821 Ultraviolet (UV), 44, 66 Limpet, 25–26 Line-encrusted, see CaCO3 Linking spine, 565, 572 Lipid (see also Fatty acid), 35, 312, 474, 574, 738 Littoral (habitat or zone), 6, 16–17, 19, 24–28, 38–40, 66–67, 82, 104, 120, 137, 150, 164, 166, 169, 177, 180–181, 189, 315, 340, 370, 372, 409, 427, 495, 603–604, 623, 757–758, 763–764, 766, 771, 808, 811 Liverwort, 17, 184, 314, 332, 341, 369 Longitudinal canal, 641–642, 647 Lorica, 1–2, 3, 5, 224, 228–229, 236–239, 388, 397–400, 406, 410–411, 414–415, 430, 434–434, 441–442, 458, 473–478, 480, 484–485, 487–488, 491–493, 495–499, 501–502 Lotic habitat – see River Louisiana, 80, 101, 181, 198, 203, 218, 244, 343, 463, 526, 535, 541, 552, 811 Lugol’s iodine – see Fixative, Lugol’s iodine Lutein – see Pigment, Lutein

Macroalga (-ae), 17, 27, 33–34, 205–206, 313, 332, 336, 339, 605, 764–767, 770, 780, 783, 809–811, 814, 819–820 Macrophyte, 6, 24, 31, 39–40, 139, 428, 656, 674, 702, 771, 780, 808, 814, 816, 820 Emergent, 17, 24, 38–39, 315, 810 Submersed, 24–28, 39, 126, 137, 141, 158, 161, 166, 205, 210, 298, 314, 332, 335, 443, 807, 809–811, 816–817, 819, 821 Magnesium (Mg2+), 364 Maine, 203, 463, 749 Malaysia, 202 Management (environmental), 38, 314, 350, 537, 777, 786, 789, 793–796, 805–806, 809, 812–814, 816, 817, 819, 824 Manitoba, 38, 85, 139, 163, 822 Marine (habitats or ecosystems), 5–6, 8, 11–12, 15, 20, 34, 43, 59–60, 64, 66–67, 72, 79, 82, 87–88, 93–94, 96, 101–102, 104–105, 107–109, 129, 132, 135, 137, 144, 147, 150–151, 155, 164, 166, 169, 197, 199–200, 205, 218, 221, 232, 253, 258, 311, 313, 333–337, 339, 342, 346–347, 387–388, 401, 406, 408, 413, 427–428, 485, 511–512, 514, 519, 562–563, 646–648, 676, 698, 701, 704, 716, 738, 748, 758–759, 764–765, 770, 789 Marl, 27–28, 141, 150, 314 Maryland, 497, 702, 814 Massachusetts, 497, 41, 80, 131–132, 144, 169, 181, 237, 243, 340, 425, 428, 451, 461, 463, 748, 766, 771, 825 Mastigoneme, 427 Mayfly (Ephemeroptera), 35, 205, 411, 819 Medulla, 217–219 Mesocosm, 43, 786, 813, 825 Mesokaryotic, 686 Mesospore, 362 Mesotrophic environment –see Lake, mesotrophic Metaboly (-ic), 388, 390, 393–394, 397, 400–401, 410–414, 428–429, 435, 439–440, 485, 495, 499, Metalimnion, 17, 535 Metaphyton (-ic), 38, 59, 64, 70, 72, 79–82, 87–89, 95–96, 119, 122, 124–126, 129, 132, 135, 139, 141, 147, 151, 155, 158, 171, 174, 177, 181, 189, 267, 270, 276, 280–281, 284, 288, 291, 295, 298, 301–303, 306, 424–425, 427, 432–433, 439, 441, 443, 446–447, 4498, 451, 453–454, 459, 461, 463, 780 Mexico, 14–15, 42, 66–67, 72, 80, 82–83, 87, 92–94, 96–104, 106–107, 127, 129, 134, 137, 139, 141, 144, 150–151, 153, 174, 180–182, 184, 203, 206, 208, 215–216, 227, 233, 235–236, 242–244, 411–415, 486, 561, 698, 701, 706–707, 765 Michigan, 337, 363, 370, 411, 424–425, 459, 463, 497, 537, 541, 552, 586, 605, 616, 618, 627, 630, 702, 759, 765, 767, 770, 772, 816,

891

Microplankton, 20 Microscopy, 33, 384 Brightfield – see Microscopy, light Differential interference contrast (DIC), 257, 259, 431, 491, 409, 514, 540, 738–740 Dissection, 206 Electron, 61, 67, 70, 219, 234–236, 242, 246, 253, 259, 335, 365, 388, 392, 396, 400, 402–406, 411, 414–415, 432, 463, 465, 472, 475, 485–486, 491, 500–501, 511–512, 523, 533, 539–541, 561, 672, 691, 740 Inverted, 409 Light, 60–61, 68, 132, 206, 225, 229, 235, 242, 253–254, 258–259, 312–313, 316, 365, 385, 388, 394, 399–400, 402–404, 410–411, 413–414, 427, 471, 484, 491–492, 497, 499–500, 514, 518, 523–524, 528, 533, 539, 561, 571, 605–606, 639, 647, 702, 741, 767 Normarski – see Microscopy, differential interference contrast Oil immersion, 259 Phase contrast, 70, 81, 257, 259, 394, 431, 497, 500–501, 514 Scanning electron (SEM), 234–235, 365, 402, 406, 414, 511, 527, 539–541, 549, 551–552, 566, 574–580, 582, 584–585, 608, 612–615, 617–620, 622, 624–625, 630, 639, 649, 674, 692–693, 695, 703, 708, 734, 741 Transmission electron TEM, 236, 253, 257, 405, 411, 499, 514, 540–541, 551–553, 573, 620, 624, 629, 664 Midge (Diptera), 36, 180 Midwestern United States, 118, 171, 267, 270, 274, 363, 526, 552, 561, 605, 638, 646, 649, 665, 809 Mississippi, 101, 288, 377, 770 Mississippi River, 28–29, 32, 36, 38, 65, 765 Missouri, 152, 451, 463 Missouri River, 28 Mitochondrion (-ia), 387, 459, 690, 731, 735–736, 749–750 Mixotrophy (-ic), 20, 23, 489–490, 514, 534, 701, 739–740, 747 Moniliform morphology – see Colony, moniliform Monitoring, 7, 23, 487, 775, 785–786, 792, 816 Mono Lake (California), 42–43 Monophyletic taxa, 218, 226, 235–236, 239, 244, 245, 387, 534, 570, 604, 659, 697, 737 Monoraphid diatom, 7, 9, 595–598, 601, 604–605, 609, 613, 623, 627 Monosporangium (-ia), 199, 201–202, 207, 210–213, 215, 217–219 Monostromatic (morphology), 207, 210, 218, 220, 317, 329, 347, 494, 503, 770 Montana, 411, 425, 427–428, 443, 447, 449, 451, 453–454, 458–459, 461, 463, 561, 766 Montezuma Well (Arizona), 14, 16, 26

892

Subject Index

Morphotype – see Ecomorphotype Moss, 17, 23–25, 44–45, 63–65, 144, 180, 184, 187, 258, 260, 274, 298, 313, 330, 335–336, 341, 363, 369–370, 378, 407, 490, 703, 706 Mountain (montane) regions, 12–13, 29, 44, 64–65, 68, 81–82, 89, 92, 96, 101, 107, 137, 151, 153, 158, 181, 204, 212, 342, 346, 363, 369, 488, 561, 631, 738, 747, 749, 751 Mucilage – see Algae, gelatinous Mucocyst, 403, 412, 427 Mud, 141, 144, 147, 150–151, 155, 180, 298, 314, 405–407, 429, 433, 463 Multiaxial (filament or thallus), 3–4, 8, 107, 118, 125, 181, 184, 200, 207–210, 217, 219, 295, 317–318, 335, 337–338, 340, 342, 344–345, 434, 459, 758, 771 Multiseriate: See Multiaxial Myxophycean starch – see Cyanophycean starch Myzocytosis, 701

N:P ratio, 22–23, 674, 776, 782, 812–813, 819 Nanoplankton, 20, 64–65, 67, 80–82, 87–89 Naphrax, 540, 562, 605 Nebraska, 16, 407, 561, 814 Necridia (necridic cell), 80, 118–120, 132, 135, 139, 141, 147, 150–151, 155, 158, 164, 166, 168, 189 Neustonic – see Algae, neustonic Nevada, 15, 42, 177, 451, 459, 463, 561 New Brunswick, 332, 454 New England, 19, 29, 40, 181, 434, 526 New Hampshire, 184, 210 New Jersey, 16, 181, 463, 498, 702 New Mexico, 212, 512, 514–515 New York, 13, 16, 19, 42, 79, 89, 216, 218, 340, 347, 463, 526, 537, 552, 770, 792, 817, 819 Newfoundland, 141, 205, 215–216, 339, 489 Nicaragua, 15–16, 18 Nitrate (NO3–) – see Nutrient, nitrate Nitrogen (N2) fixation, 23, 36, 63, 119, 121, 681, 822 North Carolina, 65, 79, 88, 92, 95, 96, 118, 132, 137, 144, 147, 150, 158, 164, 169, 174, 180–181, 184, 216, 218, 288, 295, 303, 338, 363, 370, 413, 415, 425, 427–429, 434, 443, 446–447, 449, 451, 453–454, 458–459, 814, 822 North Dakota, 82, 181 Northwest Territories, 14, 42, 95, 144, 169, 203, 270, 302, 412, 427, 449, 552, 702, 765–766 Nova Scotia, 363, 369, 441, 454, 561 Nunavut, 231, 233, 270, 274, 412, 414–415 Nutrient Ammonium (NH4+), 23, 119, 406, 513, 674, 764 Competition, 19–20, 22, 65, 406, 485, 487, 489, 568, 607, 821

Iron (Fe), 23, 37, 41, 76, 235–237, 301, 406–407, 412, 414–415, 484–485, 495, 497, 501–502, 807, 813–816 Nitrate (NO3–), 7, 23, 119, 208, 257, 364, 406, 513, 674, 699, 700, 764, 823 Nitrogen (N), 45, 117, 119, 121, 124, 177, 254–255, 257, 342, 365, 406, 487, 489, 513, 647, 670, 673–674, 681, 694, 700, 766, 782, 785, 794, 812–813, 819, 822 Nutrient limited (deficiency), 7, 26–27, 37, 39, 118, 164, 204, 254, 257, 512–513, 567–569, 586, 607, 694, 758, 776, 812 Nutrient rich (surplus), 17, 19–20, 39, 43, 45, 280, 295, 299, 342, 406, 572, 579 Phosphorus (P; also phosphate, PO43–), 7, 22–23, 25–27, 32, 36–39, 42–44, 61, 65, 118, 203–204, 256, 258, 406, 485–487, 489, 512–513, 520, 536, 584, 607, 613, 623, 682, 674, 686, 699–700, 764, 776–777, 785, 788, 790, 792, 794–795, 806, 811–814, 823 Requirement, 6, 22–23, 26–27, 44–45, 257, 406, 433, 513, 567 Selenium (Se), 23, 513 Silica (Si), 8, 20–23, 38, 41, 414, 432, 474, 524, 530–531, 533, 539–540, 559–561, 565, 569, 573, 584, 586, 598, 607, 613, 627, 642–647, 656, 662, 664, 812 Trace element, 15, 23, 825

Odor (water quality problem), 38, 64, 338, 473, 487–488, 499, 503, 513, 535, 685, 776–777, 786, 806–807, 809 Ohio, 36, 41, 246, 411, 426, 428, 459, 461, 463, 487, 586, 648, 701–702 Ohio River, 28, 28–29, 32, 37, 572 Oklahoma, 314, 454, 458–459, 461, 463 Oligotrophic environment (see Lake, oligotrophic) Ontario, 81, 95, 129, 132, 139, 169, 171, 180, 208, 210, 231, 233, 239, 242, 248, 267, 270, 377, 411–413, 415, 425, 428, 434, 447, 449, 451, 453–454, 486, 488–489, 513–514, 526, 536, 541, 552, 561, 702, 708, 793, 807, 813, 817, 822 Oogamy – see Reproduction, oogamy Oogonium, 312, 330–332, 338–342, 344, 432, 464–465 Oregon, 12–13, 42, 189, 201, 212, 235, 243, 463, 614, 762, 765, 766, 770 Organic carbon, 23, 31, 204, 406, 604, 647, 793 Organic scale, 480, 484, 493, 499, 512, 514–516, 518–519 Organic-rich (environment), 204, 236, 243–244, 248, 342, 405, 447, 449, 662 Oxygen (O2), 15, 17–19, 27–28, 38–39, 41, 45, 59, 63, 66, 155, 189, 204, 409, 433, 623, 662, 674, 699, 776, 785, 791, 805–806, 810, 816–817, 822, 824–825 Oxygen, depletion, 18–19, 30, 189, 409, 806, 815, 817, 822, 824–825

Pacific Northwest, 40, 561, Paleolimnology (paleoecology), 43, 538, 560, 570, 585, 604, 607, 610, 655, 776, 780–782, 788–789, 792–793 Pallium, 701 Palmelloid (morphology; also Palmella), 257, 260–261, 271, 294–295, 336, 346, 388, 394, 403–404, 406, 409–412, 427, 429–430, 432, 434, 473, 484, 492, 497–498, 500, 502, 736, 739, 744 Panama, 181, 210, 240, 242, 702 Papilla, 231, 235, 248, 266, 301, 319–320, 343, 370, 398, 414, 495, 530, 548–551 Parallelism, 429, 659 Paramylon – see Storage product, paramylon Parasitic (species or stage), 45, 206, 298, 311, 315, 319, 442, 685, 696–697, 701, 706, 708–709, 739, 751 Parenchymatous (morphology), 4–5, 110, 311, 313, 317, 320, 322, 332, 337, 342, 347–348, 494, 500, 758, 771 Parthenospore – see Spore, parthenospore Peat, 38, 40, 66, 88–89, 210, 335, 363, 372, 617 Peduncle, 701 Pelagic (habitat), 17, 23, 39, 66, 147 Pellicle, 5–6, 8, 383–384, 390–392, 395, 399–401, 403–404, 410–413, 690, 704 Pennsylvania, 26, 42, 147, 150, 218, 463 Peridinin – see Pigment, Peridinin Periphyton, 24, 66, 126, 135, 139, 144, 147, 151, 158, 166, 205, 314, 372, 374–376, 378, 595, 598, 604, 672, 674, 779, 785, 789, 810 pH, 11, 15, 23, 26, 43–45, 61, 215–216, 363–364, 485, 487, 490 pH effect, 26, 42, 44, 92, 204, 363–364, 407, 488–489, 513 pH, Inferred pH, 789, 793 Phagotrophy, 383, 385, 387, 401, 403, 408, 473, 489–490, 512, 514, 701, 740 Phenology, 205 Phosphatase (enzyme), 25, 776, 782, 785 Phosphorus – see Nutrient, phosphorus Photoacclimation (also Chromatic adaptation), 45, 61, 65, 738 Photosynthesis (–thetic), 17–18, 20, 23, 38, 41, 43–44, 59, 61, 120–121, 200, 202–204, 217–218, 226, 257, 311, 383–385, 387, 401–402, 406, 415, 423, 473, 485, 489, 686, 690, 696–697, 704, 706–709, 735, 737–740, 751, 776, 782, 785, 816, 823 Phototaxis (-tactic), 119, 465 Phycobiliprotein – see Pigment, Phycobiliprotein Phycobilisomes, 61, 200, 208 Phycocyanin – see Pigment, Phycocyanin Phycoerythrin – see Pigment, Phycoerythrin Phylogenetic (relationships), 1, 68, 225, 227, 229, 235–236, 239–240, 242, 244, 254, 257, 312, 424, 524, 533–534, 569–570, 582, 655, 659, 661, 686, 697, 716, 737, 741–742

Subject Index Phylogeny, 225, 231, 388, 473, 534, 570, 665, 737 Physode – see Fucosan vesicles Phytoplankton – see Algae, planktonic Phytoplankton bloom – see Bloom Picoplankton, 20, 22, 60, 64–65, 67–68, 89 Pigment Accessory, 20, 23, 200, 254, 423–423, 429, 474, 560, 567, 670, 737, 741 Allophycocyanin, 5, 61, 737 Astaxanthin, 43, 312, 342, 392 Carotenoid, 43–45, 61, 126, 141–147, 254, 257, 312, 315, 343, 392, 474, 560, 670, 686, 690, 733, 757 Chlorophyll-a, 6–8, 19, 23, 36–37, 43, 59, 61, 254, 311, 423, 560, 812–814 Chlorophyll-b, 6, 8, 225–226, 254, 311, 384, 423, 451 Chlorophyll-c, 6–8, 423–424, 474, 512–513, 560 Diadinoxanthin, 427, 429, 474, 690, 735 Diatoxanthin, 429, 474, 757 Fucoxanthin, 5, 7–8, 424, 443, 473–474, 528, 560, 690, 757 Gloeocapsin, 61 Heteroxanthin, 427, 429, 443 Loroxanthin, 245 Lutein, 254 Oscillaxanthin, 141 Peridinin, 5, 7–8, 686, 690, 706 Phaeophytin (or Pheophytin), 776, 782–783 Phycobiliprotein (also Phycobilin), 6, 59, 61, 147, 197, 384, 735, 737, 742–743 Phycocyanin, 5, 7–8, 61, 126, 200, 203, 208 Phycoerythrin, 5, 7–8, 20, 61, 126, 147, 200, 203 Scytonemin, 61, 158 Tracer, 39 Vaucheriaxanthin, 427, 429 Xanthophyll, 141, 245, 474 Plankton net, 67, 227, 258, 365, 408, 463, 490, 539, 702 Planozygote – see Cyst, planozygote Plastid (see also Chloroplast), 208, 254–256, 261, 270, 276, 280, 291, 295, 298, 301, 423–425, 427, 432, 435, 438–439, 441, 443, 451, 461, 465, 474, 485, 541, 599–602, 613, 661, 623, 638, 643–650, 655, 675–676, 681–682, 690, 704, 708, 735–737, 742, 744, 767 Plurilocular sporangium, 761 Pollution, 11, 28, 34, 42, 126, 132, 150, 333, 336, 405–406, 409 Acid rain (acidic precipitation), 39, 41, 44, 363–364, 372, 513 Fertilizer, 19, 487, 513, 814, 818 Heavy metals, 41, 205, 340, 345, 414, 785, 781, 825 Nutrient enrichment– see Eutrophication Sewage, 19, 23, 44, 336–337, 406, 514, 778, 794, 813 Polyphosphate (body [-ies]), 61, 623, 823 Polyphyletic taxa, 225, 242, 257, 563, 570, 604

Polysaccharide, 61, 118, 200, 304, 474, 609, 628, 659, 739, 757 Pond, 6–8, 12–16, 19, 24, 26–27, 40, 42, 44–45, 66,72, 80, 82, 87–88, 129, 132, 141, 155, 164, 166, 169, 171, 174, 180–181, 197–199, 205, 226, 257, 267, 270, 272, 274, 276, 280–281, 284, 288–289, 291, 295, 298–299, 301–303, 306, 314–315, 330, 333, 335, 337–341, 344, 346–347, 363, 365, 369–370, 372–379, 385, 405–409, 412–413, 424–425, 427–429, 433, 439, 441, 443, 446–447, 449, 451, 453–454, 458–459, 461, 463, 485–490, 493, 495–496, 499, 501, 512–515, 534–538, 573, 583–587, 649, 668, 700–702, 708–709, 739, 747, 751, 763–765, 771, 805–806, 808, 810–813, 815–819, 821–822, 824–825 Population dynamics, 20–21, 27, 32, 37, 258, 739 Potomac River, 65 Precambrian Shield – see Canadian Shield Preservative – see Fixative Primary production, 19–20, 31–32, 34, 38–39, 41, 43, 63, 782 Protist, 20, 383–385, 388, 403, 405, 423, 473, 490, 511–512, 685–686, 697, 710, 715, 749 Protolichen, 315 Pseudocilia – see Pseudoflagella Pseudocyst, 480, 484, 495–496 Pseudofilament, 1, 3, 62, 69–72, 80–81, 96, 102, 107, 110, 129, 199–201, 208–210, 276, 317, 336, 344, 346, 434, 459–460, 515 Pseudoflagella, 254, 259–261, 263, 267, 270, 284, 295, 298 Pseudoparenchya, 4–5, 96, 181, 199–200, 203, 207, 210, 216, 260, 264, 270, 298, 320, 322, 332, 334–338, 340, 342–343, 434–435, 459–461, 473 Pseudopyrenoid, 274 Puerto Rico, 75, 78, 80, 82, 92–93, 95–97, 99, 101, 104, 106–108, 110, 118, 126–127, 132, 134, 138–139, 145–146, 148, 150–151, 154, 156–157, 160, 162, 176, 178, 183, 212, 220, 363, 370, 433–433, 459, 463 Puncta (-ae, -ate), 333, 438, 567, 571–572, 575, 582, 584–586, 599, 606, 639–641, 643–650, 659–660, 665, 670–671 Pusule, 686, 690 Pyrenoid, 199–200, 207–208, 210, 215, 226–227, 231–233, 235–237, 239–244, 246, 248, 254, 256–257, 260, 263–267, 270–272, 274, 276–277, 280–282, 284–285, 288–289, 291–292, 295, 298–299, 301–303, 306, 312, 317–322, 330, 332–347, 354, 362, 366–367, 369–379, 384, 389–390, 394–395, 397, 402–403, 410–415, 424, 427, 432, 437, 439, 441, 443, 446–447, 449, 451, 453–454, 458–459, 461, 463, 496–498, 500, 515–516, 518, 720–724, 726–727,

893

731–732, 735–736, 742–748, 759, 768, 770–771

Quebec, 14, 16, 26, 35, 38, 203, 215, 377, 379, 434, 458, 526, 541, 561, 709

Raphe, 9, 24, 36, 599, 601–603, 605, 607, 613, 623, 627, 638–650, 656, 658–659, 661–666, 669–670, 672, 675–676, 680–682 Canal (-ed) raphe, 7, 9, 669–670, 672, 675, 681 Keeled raphe, 669–670, 676 Raphe branch, 648, 659, 662, 664–666 Raphe ending, 602, 623, 627, 639–642, 644, 646–650, 656, 658–659, 662–664 Raphe sternum, 603, 623, 647 Raphid valve, 598–599, 601–603, 605, 607, 623–625, 627–630, 641 Reproduction Asexual, 201, 210, 226, 231–237, 239, 241–243, 245–246, 254–255, 257, 288, 312, 322, 332–333, 340, 342–344, 362, 424, 432, 485, 531, 533, 539, 564, 638 Conjugation, 6, 8, 226, 235, 311, 353–356, 362–376 Daughter cell, 61–62, 70–71, 79, 81, 87, 89, 92, 102, 126, 132, 226, 346, 354, 356, 374, 485, 496, 565, 693, 736 Exocyte, 62, 70–71, 101–102, 98–99 Gametangium (-ia), 201–202, 212, 319–320, 322, 338, 343–344, 355–356, 362, 364–366, 369–376, 564 Gemmae, 199, 202, 218–219, 771, 765 Gonimoblast, 207–208, 211–215 Isogamous, 226, 229, 231–233, 235–237, 239–240, 243–244, 246, 248, 254–255, 312, 330, 333–225, 337, 339, 341, 343–344, 347, 365–366, 432, 461. 463, 485, 533, 602, 694 Oogamous, 226, 233, 235–236, 240, 312–313, 318, 322, 330, 332, 338, 340–342, 344, 346, 432, 463, 559, 563 Sexual, 8, 62, 201, 207–208, 210, 212, 218, 221, 226–227, 229, 231–237, 239–246, 248, 254–255, 257, 312, 322, 330–335, 337, 340–347, 353, 362–365, 369–377, 388, 405, 424, 432, 461–463, 485, 531, 539, 559–560, 563–564, 568, 622, 624, 627, 637–638, 649, 655, 659, 664–665, 694, 736 Reservoir (water body), 14, 16–17, 28, 64–65, 72, 81–82, 129, 135, 139, 171, 174, 177, 227, 405, 454, 458, 461, 583, 699–700, 739, 746, 751, 807, 811 Restoration, 788, 974, 806, 811 Rhine River, 37–38 Rhizoid, 24, 27, 35, 200, 207, 209–212, 215, 219–221, 260, 298, 317–320, 322, 332–334, 337–339, 340–342, 344, 347, 363, 370–371, 463–464, 768 Rhizoplast, 528 Rhizostyle, 427, 717, 734, 741, 743

894

Subject Index

Rhode Island, 201, 203, 216, 364, 463 Rice field (also crop, paddy), 45, 66, 79, 121, 174, 177, 180–181, 226 Riffle, 19–30, 35–36, 778–779 Risk assessment, 776–777, 796–797 River / stream, 6, 8, 11–12, 14–17, 28–41, 44–45, 65, 68, 82, 89, 96, 101, 104, 107, 110, 126, 137, 139, 141, 144, 147, 150–151, 155, 158, 161, 164, 166, 177, 180–181, 184, 187, 197–210, 212, 215–218, 220–221, 226, 257, 280, 284, 288–289, 295, 298–299, 303, 314, 330, 332–333, 335–342, 344–347, 363–364, 366, 370, 372, 376, 405, 407, 412, 424, 427–429, 433, 446, 449, 453–454, 459, 461, 463, 487–488, 490, 498, 500, 512–514, 534, 561, 572–573, 575, 579, 584–587, 596, 598, 601, 605, 616, 628, 661, 665, 670, 672, 682, 702, 709, 757, 759, 763, 764–765, 767, 775 Discharge (see also Current Velocity), 28, 32, 37–38 Flood, 16, 28–29, 32–36, 40, 205, 342, 810 Headwater stream (see also Spring–stream), 28, 31, 203, 514 Large river, 28, 32, 37–38, 65, 586, 661 River continuum concept (RCC), 30–31 River order, 28–29, 31–32, 38, 206 Spring-stream, 16, 28, 40–42, 80, 104, 141, 150–151, 155, 161, 181, 198–199, 203, 208, 210, 218, 303, 330, 333, 335, 337–338, 433 River Thames (U.K.), 37, 81, 206 Rocky Mountains, 81 Rotifer, 16, 32, 38, 41, 43, 129, 383, 411, 463, 739, 825 Rubisco, 34, 61, 200, 206, 686, 737–738

Saccoderm desmid, 353 Saline (environments), 6, 11, 23, 42–43, 59, 64, 66, 81–82, 87–88, 96, 107, 123, 132, 135, 137, 135, 137, 139, 150–151, 169, 171, 177, 258, 321, 333, 512, 514–515, 537, 560–561, 572, 574–575, 584–585, 604, 606–607, 638, 646, 650, 707, 766 Salinity, 11–12, 34, 42, 65–67, 80, 177, 189, 514, 565, 569, 604, 648, 738, 766, 782, 787, 789, 791 Sampling method, 25, 206, 315, 365, 486, 490–491, 514, 778 Saskatchewan, 490–491, 514 Scandinavia, 129, 485–486, 513, 537, 661, 759 SCUBA, 315, 767 Scytonemin – see Pigment, Scytonemin Seasonal succession, 19–22, 25–27, 44–45, 203–205, 258, 312, 345, 364, 433, 487–488, 538, 567 Secchi (depth or disk), 15, 19, 513, 779, 783, 806, 812–813, 816 Sediment core, 537, 539, 610, 780–781 Semicell, 354, 356–357, 361–362, 365–369, 372–379

Seta (see also Hair cell), 262, 284, 294, 373 Sheath, 1–3, 20, 60–62, 67, 70–72, 74–80, 82–86, 89–97, 101–102, 104, 107, 110, 118–127, 132, 135–138, 141, 143–151, 154–169, 171, 174, 177, 181–187, 189–190, 208, 210, 239, 242, 244–248, 254, 257–258, 260–263, 265, 267, 274, 276–277, 280, 282, 288–289, 291, 301, 312, 318322, 332, 335, 340, 344–346, 354, 371, 376, 378, 409, 725, 435, 437, 453–455, 482–483, 501, 703, 709 Si:P ratio, 20, 22, 812 Siberia, 12, 585 Sierra Nevada Mountains, 44, 561 Silica (Si) – see Nutrient, silica (Si) Silica scale (also Siliceous scale), 5, 7, 9, 493, 497, 499–502, 512–513, 474–475, 486, 491 Silicon dioxide (SiO2), 559 Sinus, 627, 629 Siphon (also Siphonous morphology), 4, 297–298, 312–313, 344, 462, 465, 810 Sloth, 6, 45, 197–199, 207, 210, 315, 319, 338 Snail, 25–26, 39, 205, 210, 215, 339, 819, 825 Snow algae – see Algae, snow Sodium (Na+), 41–42, 333 Softwater (habitat), 26, 39, 126, 205, 210, 215, 340, 344, 364, 372–373, 412, 485–486, 489, 495, 499, 513, 598, 665, 763, 807, 826 Soil, 6, 11, 38, 40, 44–45, 60, 63, 72, 79, 82, 89, 95, 121–121, 126, 129, 135, 137, 141, 141, 150, 158, 166, 177, 180–181, 184, 197–199, 206, 208, 210, 253, 255, 258, 270, 272, 274, 276, 280, 284, 288, 291, 295, 298, 301, 303, 311, 315–322, 332–337, 341–342, 344–347, 363, 369, 372, 393, 404–409, 424–425, 429, 433, 451, 453, 463, 490, 605, 648, 672, 676 South America, 202, 338, 513, 526, 568, 605, 628, 649, 765, 771 South Carolina, 425, 427, 434, 441, 449, 454, 459, 461, 463, 495, 583, 702 South Dakota, 610, 820 Southeastern United States, 203, 216, 235, 263, 267, 270–271, 274, 276, 280–281, 284, 288, 291–292, 295, 298, 301–303, 306–307, 348, 363, 387, 407, 413–415, 428, 561, 664, 709 Southwestern United States, 42, 104, 203, 216, 334, 407 Species richness, 40, 782, 785, 787, 789 Specific conductance (also Conductivity), 6–7, 40, 88, 171, 204, 206, 208, 210, 212, 215–216, 218, 220–221, 363, 428, 485, 487–488, 513–514, 535–537, 540, 585, 587, 646, 647, 649–650, 662, 676, 681, 738, 740, 742, 764, 766–767, 778, 789, 795, 810 Spermatangium (-ia), 40, 88, 171, 204, 206, 210, 212, 215–216, 218, 220–221, 363, 433, 485, 487–488, 513–514 Spermatium (-ia), 201–202, 210–213

Sponge, 45, 258, 272 Spore Spore, aplanospore, 212, 235, 245, 255, 265, 288, 291, 312, 320, 322, 332–336, 340–342, 344–345, 347, 355, 362, 370–372, 375–376, 432, 447, 451, 454, 457–459, 461, 463, 496 Spore, autospore, 288,–289, 298, 322, 341, 424–427, 432, 443–447, 449, 451–456, 458–459, 461, 463, 485, 497, 500, 502. 698, 708 Spore, auxospore, 560, 564, 602, 624, 627, 638 Spore, baeocyte, 62, 70–72, 101–104, 107, 110 Spore, endospore, 199, 201, 207–208, 210, 362 Spore, exospore (see also Exocyte), 62, 70–71, 101–102 Spore, hypnospore, 43–44, 245–246, 463, 708 Spore, parthenospore, 362, 371–372 Spore, planospore, 212, 255–256 Spore, statospore, 432, 439–440, 474, 480, 485 Spore, zoospore, 226, 231–232, 235–237, 239, 254–256, 265–267, 270, 274, 276–277, 280, 282, 284–285, 288, 291, 295, 298, 301–303, 306, 311–312, 320, 322, 326, 330, 332–347, 423–428, 432, 434, 439, 441–451, 453–456, 458–464, 473, 483, 485, 495–502, 696, 709, 739, 759–760, 767–768, 770 Spore, zygospore, 19, 330, 355–356, 362–363, 369–379 Sporophyte, 201–202, 207–208, 211–218, 759 Sporopollenin, 245–255, 312 Spring-fed streams – see River, spring-stream St. Lawrence River, 34, 37, 210, 793 Stagnant water, 6, 89, 204, 322, 336, 340, 347, 363, 371, 413 Stain (microscopy), 70, 259, 403, 411, 491, 498, 500 Alcian Blue, 198, 410 Iodine (test for starch), 5, 226, 259, 403, 409, 454, 463, 491, 514, 702–703, 736, 740 Jensen’s stain, 491, 497, 500 Methylene blue, 295, 403, 476, 491–492, 498, 502 Toluidine Blue O (TBO), 211, 217, 219 Starch – see Storage product, starch Statistical methods, 569, 777, 788–792 Statospore – see Spore, statospore Stauros, 603, 607, 623, 627, 640, 642–643 Stenotherm, 80, 141, 147, 488 Sternum, 599–606, 612–613, 623, 627, 643, 645–648, 656, 665 Stichidia, 199, 202, 208, 219–220 Stickleback (fish), 697, 708 Stigma (-ata) – see Eyespot Stomatocyst, 414, 476, 480, 485–486, 496, 499–503, 531 Stoneflies (Plecoptera), 205

Subject Index Stonewort, 311, 314, 332 Storage product Chrysolaminarin, 5–7, 423, 472, 474, 528, 541, 548 Cyanophycean starch, 6 Floridean starch, 6, 197 Laminarin, 5–6, 8, 757 Paramylon, 5–6, 384, 388–390, 395–397, 402, 407, 409, 411–415 Polyphosphate, 61, 623, 823 Starch (true starch), 5–8, 225–226, 231, 254, 259, 265, 270, 274, 276, 280, 282, 291, 298, 301, 303, 311–312, 318, 335–336, 340–341, 384, 423, 427, 686, 690, 702, 706, 717, 722, 726–727, 735–737, 739–740, 745–748 Stratification – see Lake, stratification Stream – see River / stream (habitats) Stria (-ae), 357, 365–368, 372–373, 403, 437–438, 518, 546, 572, 578, 599–607, 612–613, 616–618, 623–624, 627, 638–640 Stromatolite, 59, 63, 67, 121 Strutted process, 565–567, 571–572, 583–585, 587 Subaerial – see Algae, subaerial Subarctic (habitat or biome), 12, 26, 38, 40, 60, 80, 88, 101, 141, 158, 171, 289, 513, 535, 700–701 Substratum (-a) (surface), 4, 7, 24–30, 32–37, 44, 63–68, 70, 72, 80, 87, 89, 92, 95–96, 101–102, 104, 107, 110, 119, 121–122, 125–126, 132, 135, 137, 139, 141, 144, 147, 150–151, 155, 158, 161, 164, 166, 169, 177, 180–181, 184, 187, 189, 205–206, 220, 260, 270, 272, 280, 288, 291, 295, 303, 316, 354, 363, 371, 376, 378, 388, 393, 405, 408. 410–411, 436, 441, 458, 463, 474, 488, 492, 494–497, 499–503, 565, 583–584, 586, 595, 598, 601–602, 605, 607, 609, 613, 616–617, 623, 638, 656, 659, 661, 663, 669–670, 673, 677, 687, 690–691, 740, 747, 758, 760, 763–764, 766–768, 771, 779, 780, 783, 786, 808–810 Succession – see Seasonal succession Sulcus, 388, 395–396, 410–411, 686–687, 690–691, 706–707, 788 Sulfate (SO42–), 23, 41–42, 66, 372, 377, 406, 514, 662, 807, 813, 816, 821–822, 824–826 Surface bloom – see Bloom Suture, 357, 361, 372–373, 695, 704 Sweden, 204, 513, 588, 758, 763, 794 Synonym (-ous), 96, 158, 164, 212, 229, 232, 267, 344, 376, 379, 388, 425, 427, 458–459, 512, 524, 563, 629, 646, 669–670, 697–698, 706, 708, 742–743, 746, 758, 770–771

Tadpole, 26, 410–412 Taste (water quality problem), 38, 64, 467, 535, 552, 685, 776–777, 786, 806, 809, 816

Temperate environment (region), 8, 12, 17–20, 26, 31, 33, 37, 64–65, 80–82, 87–89, 95, 101–102, 126, 129, 132, 135, 139, 147, 158, 161, 164, 169, 171, 174, 177, 187–189, 202–204, 216, 227, 246, 258, 284–289, 291, 295, 298, 301, 303, 306, 334, 339–340, 345, 364, 485–486, 535, 537, 568, 581, 583, 616, 694, 696, 704, 715, 738, 812–813 Temperature Cold, 12, 17, 40, 59, 66, 80, 88, 95, 101, 107, 129, 141, 147, 288, 303, 342, 488, 499, 535–536, 538, 567, 605, 649, 694, 696, 702, 740, 815, 820 Hot – see Hot spring Warm, 18, 23, 33, 37, 43–44, 65, 80, 88, 101–102, 107, 150, 174, 203–204, 212, 218, 220–221, 314, 407, 409, 434, 463, 488, 513, 535–537, 605, 700, 763, 783 Temporary pond (pool), 19, 44, 66, 257, 364–365, 371, 433, 501, 512 Tennessee, 15, 428, 434, 459, 461, 463 Terrestrial (habitat), 11, 25, 45, 59, 63, 68, 120–121, 144, 150–151, 174, 180, 187, 260, 267, 276–277, 284–285, 288–289, 306, 312, 315, 317–319, 321–322, 334, 341–343, 424, 433, 463, 610, 637, 778, 781, 821 Tetrasporangium (-ia), 201–202, 212–213, 220 Tetrasporophyte, 201–202 Texas, 16–17, 40, 150, 208, 212, 218, 243, 270, 291, 332, 345–346, 425, 427, 512, 702, 739, 749 Theca, 2, 5, 7, 229, 231–232, 690–691, 694, 698, 701, 703–704, 707 Thermal spring – see Hot spring Thermocline, 17, 23 Thylakoid, 5–8, 44, 61, 69, 73, 75–77, 79, 81, 87–88, 101–102, 104, 118–119, 121–122, 126–127, 129–130, 132, 135, 137, 139, 141–142, 144, 147, 149–150, 153, 177, 187, 197, 200, 208, 257, 397, 402, 423, 474, 690, 742, 748 Tierra del Fuego, 486 Total nitrogen (TN), 489, 785 Total phosphorus (TP), 36, 43, 486, 489, 513, 536, 699, 777, 787, 785, 792–793, 794–795, 812–813 Toxin (also Toxic), 6, 63–64, 92, 139, 141, 147, 169, 512, 514, 604, 607, 685–686, 739, 776–777, 782–783, 785–786, 806–808 Travertine, 16, 59, 67, 69, 81, 119, 121, 141, 158, 169 Tree (also Tree bark), 45, 64, 89, 121, 144, 161, 181, 184, 203, 206, 258, 267, 315, 330, 344, 346, 780 Trichoblast, 208, 220–221, 689, 691, 695 Trichocyst, 390, 403, 409, 412, 426–429 Trichogyne, 201–202, 211–216, 218–219 Trichome, 118–132, 134–144, 147–152, 155–158, 161, 164–166, 168–177, 179–181, 184, 187, 189 Trichoptera (see Caddisfly)

895

Trinidad, 463 Trophont stage, 696, 706 True branch, 3–4, 119, 121, 123, 125, 181–190, 200–202, 207–208, 210–221, 259–260, 263–264, 267, 271, 284–285, 298, 313–315, 317–325, 327, 330–344, 347, 354, 364–366, 376, 434–436 Tundra (habitat), 19, 33–34, 121, 158, 180, 184, 197, 203, 215, 422, 446 Turbid (-ity), 23, 27, 31–21, 37–38, 202, 204, 206, 258, 407, 670, 763, 766, 811, 820 Turtle, 12, 313–314, 316, 319, 321, 334, 339–340 Tychoplankton (-ic), 88, 146–147, 408, 424–425, 427, 432–433, 453–454, 458–459, 461, 463, 565, 572, 672, 676, 681

Ultraviolet radiation (UVR), 44–45, 66 Uniaxial (also Uniseriate), 3, 69, 80, 102, 118, 125, 135, 158, 161, 169, 171, 177, 180–181, 184, 187, 189, 200, 207–208, 210–216, 218–219, 264, 317, 322, 326, 330, 332–342, 344–345, 354, 365, 369–377, 434, 459, 461, 600–602, 606, 613, 616–617, 623, 646, 758–760, 768, 771 Unilocular sporangium, 759–760, 762, 765, 767–771 Uniseriate – see Uniaxial United Kingdom (U.K.), 42, 206, 338, 700, 759, 763, 765, 771 Urban (-ization), 19, 39, 45, 342, 794, 813, 816, 826 Utah, 12, 72, 191, 454, 459, 461, 463, 628, 647, 650, 699–700, 763, 766 Utermöhl technique (sedimentation), 67, 514, 539

Valve face, 565–566, 569, 571–572, 583, 585, 587, 600, 613, 616–617, 623, 627, 640–641, 643, 645, 650, 656, 660, 662, 665, 670, 675–676, Valve mantle, 567, 571–573, 578–579, 582–583, 585, 607, 644, 646, 650, 656, 662–663, 665 Valvocopula, 600–601, 623, 644, 648 Van Dorn sampler, 539, 702, 779 Vascular plant, 40, 65, 180, 315, 3198, 321, 363, 365, 377, 405, 424–425, 432, 439, 441, 443, 454, 458–459, 461, 463, 490, 662, 807–808, 810–811, 815–817, 820–821, 824 Venezuela, 93, 95, 202 Vermont, 189 Virginia, 41, 212, 434, 461, 463 Virus (-es), 384, 739, 819 Vitamin, 387, 406, 409, 490, 699–701

Washington (state), 13, 15, 19, 23, 42, 44, 203, 235, 346, 526, 541, 649, 766

896

Subject Index

Water bird (waterfowl), 333, 407, 486, 538, 811, 820–821 Water quality, 13, 39, 59, 406, 489, 560, 598, 655, 782 Watershed (also Catchment), 28–29, 32, 34, 36, 204, 536, 560, 788, 789, 805, 813–814, 816, 826 Wave action (also Scour), 24–27, 137, 347, 453, 808 Weed (or Weedy species), 174, 821, 817, 809–812, 820–821 West Virginia, 42, 459, 463 Western Canada (see also specific provinces), 33, 126, 267, 274, 276, 282, 284, 288–289, 292, 295, 302–303, 306, 552, 646 Western United States (see also specific states), 42, 274, 282, 284, 288–289,

291–292, 295, 298, 301–303, 306, 334, 459, 461, 526, 552, 561, 648, 808–809, 811, 817 Wetland (also Swamp, Marsh), 11, 28–29, 37–40, 66–67, 72, 79–83, 86–89, 95, 102, 104, 137, 147, 151, 155, 158, 151, 164, 166, 174, 177, 180–181, 184, 189, 257, 341, 345, 372, 375–378, 413, 425, 428–429, 433, 458–459, 463, 536, 604, 661–662, 669, 672, 682, 702, 776, 778–780, 785, 788–789, 791, 793, 796 Whorl (branching pattern), 3–4, 207–208, 213–215, 217, 317, 321, 333–334, 337–339, 367 Wisconsin, 12, 19, 22, 74, 76, 78, 83–87, 90, 94, 118, 143–144, 174, 242, 339, 363, 370, 425, 453, 459, 461, 487, 702, 814, 825

Yellowstone National Park, 15, 41, 66–67, 72–73, 75–82, 94–95, 104–105, 107, 118, 126, 128–129, 130–137, 139–140, 144, 147–148, 151–152, 155, 157–158, 186–187, 189–190, 206, 363 Yucatan Peninsula, 15–16, 66 Zebra mussel, 28, 37–38, 813, 816 Zooplankton (see also specific groups), 6, 20, 38, 43, 64, 271, 303, 488, 537, 700, 702 Zoospore, 226, 231, 232–233, 235–237, 239, 254–255, 265–267, 270, 274, 276–277, 280, 284–285, 288, 291, 298, 301–303, 306, 311–312, 320, 322, 330, 332–347, 423–428, 432, 434, 439, 441–451, 453–456, 458–464, 473, 483–485, 495–502, 698, 709, 739, 759

Taxonomic Index

Acanthameba, 737 Acanthoceras, 563, 571–572, 587 magdeburgense, 575 Acanthoica, 512, 515 schilleri, 515 Acanthosphaera, 262, 267 zachariasii, 268 Achnanthaceae, 597–598, 604, 623 Achnanthales, 595–598, 623 Achnantheiopsis, 598, 627 Achnanthes, 34, 41, 45, 597, 601, 604, 607 affine, 623 clevei, 627 clevei var. rostrata, 596 coarctata, 623–624 kolbei, 627 lanceolata, 598, 627 lancolata var. abbreviata, 629 levanderi, 627 linearis, 628 longipes, 623 marginulata, 627

microcephalum, 623 minutissima, 623 ploenensis, 627 pusilla, 43 Achnanthidiaceae, 597, 604–605, 623, 627 Achnanthidium, 597–598, 602, 604–605, 607, 623, 627 affine, 609, 625 deflexum, 625 exiguum, 625 exiguum var. heterovallvum, 625 lanceolata, 36 minutissima, 36 minutissimum, 25–26, 33, 623, 625, 628 Achnanthoideae, 597 Acrochaetiales, 198, 201, 202, 204, 206, 211–212, 213 Actidesmium, 265, 267 hookeria, 268 Actinastrum, 254, 262, 267 hantzchii, 268 Actinella, 655–656, 662

punctata, 657, 661, 664 Actiniscaceae, 697, 706 Actiniscus, 697, 698, 704, 706 canadensis, 702 pentasterias v. arcticus, 688 Actinochloris, 266–267 sphaerica, 268 Actinocyclus, 563, 571, 575 normanii, 572 normanii f. subsalsa, 574 subsalsa, 572 Actinotaenium, 368, 373 diplosporum, 373 perminutum, 373 rufescens, 357 Adlafia, 638, 641, 643 bryophila, 643 muscora, 643 Aeronemum, 431, 435, 459 polymorphum, 460 Agmenellum quadruplicatum, 88

897

898

Taxonomic Index

Akanthochloris, 430, 434, 438, 443, 445 bacillifera, 445 brevispinosa, 445 scherffelii, 445 Albrightia, 125, 184 tortuosa, 186, 187 Alterasynedra, 596 Amblystoma, 45, 291 Amblystomatis, 45 Ambrosia, 298 Ammatoidea, 123, 151–152 normannii, 151–152 yellowstonensis, 151–152 Ammatoideoideae, 151 Amphicampa, 656, 662 eruca, 657, 664 mirabilis, 664 Amphichrysis, 493, 495 compressa, 481 Amphidiniopsis, 691, 697–698, 704, 708 sibbaldii, 693, 702 Amphidinium, 687, 691, 697–698, 704, 706 cryophilum, 702 klebsii, 688 Amphipleura, 638, 640, 643 pellucida, 645 Amphithrix, 137 Amphora, 39, 43, 656, 663, 676 calumetica, 661, 663 coffeiformis, 43 inflata, 662 ovalis, 663 pediculus, 658 perpusilla, 658, 672 Anabaena, 20, 23, 37, 43, 45, 64–65, 68, 79, 119–120, 125, 169–171, 176, 180, 303, 768, 805, 807, 808, 812, 816 azollae, 45, 180 circinalis, 22 cycadearum, 180 fertilissima, 176 flos-aquae, 20 lutea, 176 mendotae, 169 oblonga, 170 perturbata, 169–170 subtropica, 176 variabilis, 176 viguieri, 170 Anabaenoideae, 120, 169 Anabaenopsis, 119–120, 124, 171–172 elenkina, 172 Anabaenopsis (Cylindrospermopsis) raciborskii, 174 Anacanthoica, 512, 515 ornata, 515 Anacystis, 74, 104 nidulans, 81 nigroviolacea, 93 rupestris, 74 Anathece (subgenus of Aphanothece), 72 Ancylonema, 363, 367, 369 nordenskioeldii, 355, 364, 369 Ancylus fluviatilis, 26

Aneumastus, 640, 644 tuscula, 643 Animalcula, 471 Ankistrodesmus, 254–255, 257, 262, 267, 289, 822 falcatus, 269 Ankylonoton, 430, 432, 435, 439 pyreniger, 439–440 salinis, 439 Anomoeoneis, 638, 640, 645–646 brachysira, 40 sphaerophora, 43, 642 Anthophysa, 492, 495 vegetans, 471, 477, 495 Apatococcus, 45, 258, 264, 267, 322, 333, 346 lobatus, 269, 323 Aphanizomenon, 20, 23, 37, 43, 64–65, 119–120, 124, 171–172, 805, 807, 812, 816 aphanizomenoides, 171 flos-aquae, 21, 43, 171–172, 808, 822 gracile, 171 issatschenkoi, 171 schindleri, 171–172 skujae, 171 Aphanocapsa, 40–41, 44, 64–65, 70, 82 arctica, 82 botryoides, 82 conferta, 82 delicatissima, 82 farlowiana, 78, 82 grevillei, 78 holsatica, 82 incerta, 78, 82 intertexta, 82 marina, 82 montana, 89 muscicola, 82 planctonica, 82 protea, 82 pulchra, 92 saxicola, 37 thermalis, 82 tolliana, 82 Aphanochaete, 314, 320, 322 repens, 324 Aphanothece, 62–66, 68–69, 72, 74 bachmannii, 72 bacilloidea, 67, 72 bullosa, 72 castagnei, 72, 74 clathrata, 72, 74 conglomerata, 72 cylindracea, 72 halophytica, 43 karukerae, 72 microscopica, 72 minutissima, 72 nidulans, 72 opalescens, 72 pallida, 72 saxicola, 72 smithii, 72, 74 stagnina, 64, 72, 74 thermalis, 72

uliginosa, 72 utahensis, 72 Apiocystis, 258, 260, 267 brauniana, 269 Apistonema expansum, 771 pyrenigerum, 771 Araceae, 319, 344 Arachnochloris, 430, 433–434, 438, 443, 446–447 major, 445 Arisaema, 315 triphyllum, 328, 344 Aristata, 215 Aristichthys nobilis, 820 Artemia, 43 salina, 512 Arthrodesmus, 376, 378–379 Arthronema africanum, 67 Arthrospira, 66, 122, 139–141 fusiformis, 139 gomontiana, 141 jenneri, 139–141 khanmnae, 140 maxima, 139–140 platensis, 139–140 skujae, 140 Ascoglena, 388, 410, 414–415 vaginicola, 411 Askenasyella, 265, 270 chlamydopus, 271 Astasia, 385 Asterionella, 3, 20–22, 595–597, 600, 606, 613, 631 formosa, 20, 21–23, 605, 607–608, 611, 613 ralfsii var. americana, 607, 611 Asterionellopsis, 596, 607 Asterocapsa, 62, 64, 71, 92–93 divina, 92–93 magnifica, 92 pulchra, 92 Asterococcus, 266, 270 superbus, 271 Asterogloea, 431, 437, 453 gelatinosa, 455 Asteroplanus, 607 Astrephomene, 226, 238, 242, 248 eugenea, 212 gubernaculifera, 243 hermannii, 212 perforata, 243 pygmaea, 212 Audouinella, 34, 202, 205, 207, 211–212 eugenea, 198 helminthosum, 203 hermannii, 36, 198, 201, 204–205, 211, 213, 770 macrospora, 198, 201 pygmaea, 198 tenella, 198, 201, 204 Aulacoseira, 20, 563, 565, 567, 569–572, 583–584 granulata, 573, 576

Taxonomic Index islandica, 576 italica, 576 Aulacoseiraceae, 563 Aulacoseirales, 563, 570 Aulakochloris, 430, 438, 443 striata, 445 Aulosira, 125, 173–174 fertilissima, 174 implexa, 173 laxa, 173 Autosira, 171 Azolla, 45, 180, 338

Bacillaria, 670, 675 paradoxa, 676–677 Bacillariaceae, 670 Bacillariales, 669, 670, 671, 672, 675, 676, 681 Bacillariophyceae, 7, 201, 656, 669 Bacillariophycidae, 670 Bacillosiphon, 77, 79 gracilis, 79 induratus, 77 Bacteria (see Subject Index) Bacularia, 67, 69, 77 gracilis, 77 indurata, 67, 72, 77 Baetis tricaudatus, 35 Balbianiales, 198, 212, 213 Ballia, 202, 207, 218 prieurii, 199, 218–219 Bambusina, 366, 373, 375 brebissonii, 357 Bangia, 3, 26, 34, 200, 204, 206, 210, 763 atropurpurea, 25, 198–201, 205, 209 Bangiales, 198, 210 Bangiophycidae, 197, 198, 200, 209, 211 Basichlamys, 244–245, 248 sacculifera, 239, 244–245 Basicladia, 314, 321, 339–340 chelonum, 12, 327 Batrachospermaceae, 198, 202, 206, 212–213, 215–216 Batrachospermales, 198, 200–202, 204–205, 212, 214–216, 218 Batrachospermum, 4, 17, 35, 197–198, 200–203, 205, 207, 212–216, 322, 662, 763 ambiguum, 198, 213 anatinum, 198, 201, 213 androinvolucrum, 198 arcuatum, 198 atrum, 198 boryanum, 198, 203 carpocontorium, 198, 203 carpoinvolucrum, 198, 203 confusum, 198 elegans, 198 gelatinosum, 198, 200, 203, 204, 213–215 gelatinosum forma spermatoinvolucrum, 198, 203, 213 globosporum, 198, 214 helminthosum, 198, 202–203, 205, 213

heterocorticum, 198 intortum, 198, 201, 213 involutum, 198, 202–203 keratophytum, 40 louisianae, 198, 213 macrosporum, 198 procarpum, 198 pulchrum, 198 skujae, 199 trichocontortum, 199 trichofurcatum, 199 turfosum, 40, 198–199, 203, 205, 215 virgato-decaisneanum, 198, 215 Belonastrum, 631 Bernardinium, 691, 697–698, 704, 706 bernardinense, 688, 705 Bicosoeca, 477, 491, 495 borealis, 477 kenaiensis, 477 Bicosoecaeae, 484–485 Biddulphia, 584 Biddulphiaceae, 563 Biddulphiales, 563 Biddulphiophycidae, 563, 565 Binuclearia, 318, 344 tatrana, 326 Bitrichia, 478, 492, 495 chodati, 478, 495 longispina, 495 ollula, 478, 495 Blastodiniales, 697, 706 Blennothrix, 119, 123, 150–151, 153 cantharidosma, 151, 153 coerulea, 151 comoides, 151 ganeshii, 151, 153 glutinosa, 151 groesbeckiana, 151 heterotricha, 151 majus, 151 mirifica, 151 ravenelii, 151 Bodanella, 758, 760, 768, 771 lauterbornii, 759, 761, 765, 768 Bodo, 387 sultans, 386 Boldia, 4, 203, 207, 210 erythrosiphon, 198, 209, 211 Borzia, 122, 138–139 trilocularis, 138–139 Borziaceae, 122, 139 Borzinemataceae, 125, 184 Bosmina, 38, 825 Bostrychia, 202, 205, 208, 218, 220 moritziana, 199, 200, 202, 219–220 radicans, 199 tenella, 199, 220 Botrydiaceae, 431 Botrydiales 429, 431, 433–434, 463–464 Botrydiopsidaceae, 430, 436, 450–451 Botryochloridaceae, 430, 436, 451–452 Botrydiopsis, 430, 432, 436, 451, 463 arhiza, 450 Botrydium, 429, 431–434, 463 granulatum, 464

899

Botryochloris, 430, 437, 451, 453 cumulata, 452 Botryococcus, 43, 264, 270 braunii, 43, 271 Bourrellia, 494–495 skuja, 483, 495 Bracchiogonium, 430, 438, 446 ophiaster, 445 Brachionus, 411 Brachysira, 638–640, 645–646 serians, 642 styriaca, 642 Brachytrichia, 119 Bracteococcus, 254, 256, 266, 270 minor, 272 Bradypus, 210, 315 Brasenia schreberi, 811 Bulbochaete, 39, 320, 341 minor, 330 Bumilleria, 431–434, 459, 462 klebsiana, 462 sicula, 462 Bumilleriopsis, 431–432, 436, 457–458 biverruca, 457 closterioides, 457

Calliglena, 412 Caloglossa, 4, 208 leprieurii, 199, 202, 208 ogasawaerensis, 199, 202 Caloneis, 638–639, 646 amphisbaena, 644 schumanniana, 644 silicula, 644 Calothrix, 26, 39, 44–45, 65, 118–119, 164–165 ascendens, 164 braunii, 164 contarenii, 164 donnellii, 164 elenkinii, 164 epiphytica, 164 fusca, 164–165 juliana, 164 kawraiskii, 164 parietina, 164 pulvinata, 164 rivularis, 164 scytonemicola, 164 simplex, 165 stagnalis, 164 stellaris, 164 tenella, 165 Campylodiscus, 43, 670, 672, 675, 681 noricus, 679 Campylomonadaceae, 721–722, 743, 745–746 Campylomonas, 716, 723, 741–745, 748 marssoni, 746 platyuris, 724, 745 reflexa, 721, 723, 745–746 rostratiformis, 722–723, 745 Capartogramma, 638, 640 crucicula, 644, 646

900

Taxonomic Index

Capitulariella, 431, 435, 459 radians, 460 Capsosira, 182 brebissonii, 181–182 Capsosiraceae, 181 Carex, 439, 458 Carteria, 227, 232, 234, 247–248 eugametos, 233–234 nivale, 44 Castor canadensis, 16 Catacombus, 596 Catenochrysis, 523 Catenulaceae, 655–656, 661–662, 663 Cavinula, 638–640, 646 cocconeiformis, 645 Celloniella, 494–495, 499 palensis, 483, 499 Centritractaceae, 431, 436, 457–458 Centritractus, 431, 434, 436, 457–459 belenophorus, 457 brunneus, 457 ellipsoideus, 457 globulosus, 457 Centronella, 596, 613 Cephaleuros, 45, 315, 319, 328, 342 virescens, 315, 328 Cephalomonas, 236, 248 granulata, 236–237 Ceramiales, 199, 202–204, 218–219, 220–221 Cerasterias, 254, 262, 270 irregularis, 272 Ceratiaceae, 697, 707 Ceratium, 2, 23, 691, 697–698, 704 brachyceros, 695, 707 carolinianum, 701–702 cornutum, 694 furcoides, 694 hirundinella, 20, 685, 689, 691, 694–695, 699–702, 707 hirundinella f. hirundinella, 695 hirundinella f. piburgense, 695 Ceratophyllum, 17, 314, 459, 810–811 demersum, 24, 314 Chadefaudiothrix, 431, 434, 459 gallica, 460 Chaetoceraceae, 563 Chaetocerophycidae, 563 Chaetoceros, 563, 565, 571, 574 elmorei, 575 muelleri, 43 Chaetocerotales, 563 Chaetomorpha, 318, 339 Chaetonema, 321–322 irregulare, 324 ornatum, 322 Chaetonemopsis, 330 Chaetopedia, 431, 435, 459 stigeoclonioides, 460 Chaetopeltis, 260, 270 orbicularis, 272 Chaetophora, 314, 320, 322, 330, 497 elegans, 325 incrassata, 764 Chaetophoraceae, 314–315, 348

Chaetophorales, 267, 313, 319, 321–322, 330, 332–338, 340, 347–348, 454 Chaetosphaeridium, 322, 330 globosum, 324 Chalarodora, 266, 270 azurea, 272 Chalkopyxis, 495 tetrasporoides, 484, 496 Chamaecalyx, 62, 65, 70, 96, 100–101 calyculatus, 100–101 clavatus, 100 suffultus, 100 swirenkoi, 100–101 Chamaepinnularia, 638, 641, 646 mediocris, 643 Chamaesiphon, 24–25, 33, 62, 64–65, 68, 70–71, 99, 101, 764, 768, 770 amethystinus, 99 britannicus, 101 confervicolus, 101 geitleri, 101 incrustans, 99, 101, 765 polonicus, 99, 101 regularis, 101 rostafinskii, 101 subglobosus, 99 willei, 99 Chamaesiphonaceae, 60, 62, 68, 71, 96 Chamaetrichon, 320, 332 capsulatum, 323, 332 Chara, 17, 24, 27–28, 39, 311–312, 317, 322, 335, 338–339, 809, 811, 815–817, 819–821, 826 aculeolata, 27 canescens, 331 hispida, 764 tomentosa, 27 vulgaris, 811 Characidiopsidaceae, 430 Characidiopsis, 430, 432, 435, 441, 444 acuta, 444 ellipsoidea, 444 elongata, 444 Characiochloris, 260, 270 characiodes, 272 Characiopsis, 260, 270, 429, 431, 433, 436, 454, 456, 465 acuta, 456 minuta, 272, 427 ovalis, 427 pyriformis, 456 Characium, 254, 258, 260, 270, 441, 454 minutum, 427 sieboldii, 272 Chara/Nitella, 805 Charales, 311–313, 317, 338–339 Charophyceae, 229, 313, 353, 384 Charophyta, 12 Chilomonas, 734–735, 740–741, 743, 748 acuta, 721, 746 paramecium, 721–722, 733, 746 striata, 739 Chlainomonas, 227, 232, 234 kollii, 234 rubra, 234

Chlamydobotrys, 247 Chlamydomonadaceae 225, 228, 231, 233, 235–236 Chlamydomonadales, 225 Chlamydomonas, 23, 45, 225–228, 232, 234–235, 238, 245, 248, 253, 257, 291–292, 301, 429 acidophila, 42 bohemica, 245 sonowiae, 233–234 tetragama, 234 Chlamydophyceae, 313 Chlamydomyxa, 430, 434–435, 441 labyrinthuloides, 442 Chlorakys, 446 Chlorallanthus, 446 Chlorallantus, 430, 431, 433, 446 oblongus, 445 Chloramoeba, 430, 432, 435, 439 Chloramoebaceae, 430 Chloramoebales, 429–430, 432, 434, 439–440 Chlorangiella, 260, 271 pygmaea, 273 Chlorangium, 271 Chlorarkys, 430, 438, 447 reticulata, 445 Chlorcorona, 246, 248 bohemica, 246 Chlorella, 11, 44–45, 254, 255, 257–259, 265, 271, 307, 446 miniata, 44 vulgaris, 273 Chlorellidiopsis, 430, 437, 451 separabilis, 452 Chlorellidium, 430, 437, 451 tetrabotrys, 452 Chloremys, 261, 272 sessilis, 273 Chloridella, 429–430, 437, 445–446 ferruginea, 445 neglecta, 445 Chlorobotryaceae, 424 Chlorobotrys, 424–425, 431, 434, 437, 453–454, 463 regularis, 424–426, 453 simplex, 455 stellata, 425, 427, 453 Chlorobrachis gracilima, 247 Chlorochytrium, 260, 274, 316, 321, 332 lemnae, 273, 314, 323 Chlorocloster, 430, 432, 438, 446 angulus, 445 Chlorococcopsis, 256 Chlorococcum, 45, 254–256, 266, 274, 303, 333 regulare, 425 submarinum, 255, 258 wimmeri, 273 Chlorodesmus, 523, 528 hispidus, 524 Chlorogibba, 430, 438, 446 trochisciaeformis, 445

Taxonomic Index Chlorogloea, 62, 67, 71, 96, 97 cuauhtemocii, 96 epiphytica, 96–97 lithogenes, 67, 96, 97 regularis, 96 tuberculosa, 96 Chlorogonium, 226, 232, 233, 247–248 capillatum, 233, 234 elongatum, 233 euchlorum, 233 tetragamum, 234 Chlorokardion, 430, 435, 439 pleurochloron, 439, 440 Chlorokoryne, 436, 454 petrovae, 456 Chlorokybus, 264, 274 atmophyticus, 273 Chloromeson, 423, 430, 432, 435, 439 agile, 439, 440 parvum, 439 viridis, 439 Chloromonas, 227, 232, 234, 235, 248 anglica, 235 brevispina, 44, 235 clathrata, 235 depauperata, 235 granulosa, 235 minima, 233, 235 nivalis, 44, 235 pinchiae, 44, 235 platystigma, 235 polypyera, 235 Chloromonas (previously Chlamydomonas) nivalis, 43 Chloropedia, 431, 436, 458 plana, 457 Chloropediaceae, 431, 436, 457, 458 Chlorophyceae, 12, 35, 225, 231, 313, 384, 427, 429, 431, 447, 454 Chlorophysema, 261, 274 contractum, 275 Chlorophyta, 6, 12, 45, 253, 259, 311, 313, 353, 384, 441, 446–447, 453, 458, 465, 823 Chlorosaccus, 437 fluidus, 453, 455 Chlorosarcina, 265, 274 brevispinosa, 275 Chlorosarcinopsis, 254, 258 Chlorothecium, 431, 436, 456, 458 capitatum, 456 crassiapex, 456 Chlorotylium, 319, 332 cataractum, 325 Chodatella, 288 Choleochaete pulvinata, 324 scutata, 324 Choloepus, 210 Chondrocystis, 70, 89, 91 bracei, 89 dermochroa, 89, 91 schauinslandii, 89, 91 Chorogloea, 65 Chromista, 384

Chromophyton, 493, 496 rosanofii, 480 Chromulina, 473, 485–486, 489, 493, 495–498, 524 chionophila, 44 globulifera, 497 palensis, 495 stellata, 481 Chromulinaceae, 484 Chroococcaceae, 62, 71, 92 Chroococcidiopsis, 64, 66, 71, 104 cubana, 104 cyanosphaera, 103 thermalis, 104 Chroococcidium, 67, 71, 103–104 gelatinosum, 103–104 Chroococcopsis, 72, 106 fluviatilis, 104, 106 Chroococcus, 40, 41, 43–44, 60, 62, 64–66, 71, 94–95, 813 aeruginosus, 95 cubicus, 95 deltoides, 95 distans, 95 endophyticus, 95 heanogloios, 95 limneticus, 95 limneticus var. subsalsus, 94 microscopicus, 95 minimus, 95 minutus, 95 mipitanensis, 95 multicoloratus, 67, 95 pallidus, 95 polyedriformis, 95 prescottii, 95 refractus, 95 rufescens, 95 schizodermaticus, 95 sonorensis, 95 submarinus, 95 tenacoides, 95 thermalis, 95 turgidus, 95 varius, 95 yellowstonensis, 95 Chroococcus (Linmococcus) dispersus, 94 limneticus, 94 sonorensis, 94 Chroodactylon, 3, 204, 207–208 ornatum, 25, 198–199, 201, 205 ramosum, 25 Chroomonadaceae, 743, 747–748 Chroomonas, 736, 740, 742–743, 747–749 africana, 736 americana, 748 coerulea, 726, 727, 729, 747 nordstedtii, 726–727, 747 oblonga, 726, 729, 747 pochmanni, 726–727, 747 salina, 736 Chroothece, 207–210 mobilis, 198, 203

901

Chrysamoeba, 485, 494–496, 501 mikrokonta, 476 radians, 482 Chrysapion, 503 Chrysapsis, 480, 493, 496 agilis, 480 fenestrata, 480 Chrysarachnion, 494, 496 insidians, 482 Chrysidiastrum, 494 catenatum, 476, 496 epiphyticum, 482 Chrysoamphipyxis, 492, 496, 499, 503 canadensis, 478 Chrysoamphitrema, 492, 496 nygaardii, 478 Chrysobotrys, 497 Chrysocapsa, 491, 495–496 planktonica, 484 Chrysocapsella, 496 Chrysocapsopsis, 483, 494, 503 rupicola, 483, 496 Chrysochaete, 494, 496–497, 500 britannica, 482 Chrysochromulina, 23, 484, 499, 512–513, 515, 518, 739 breviturrita, 7, 23, 512–515, 518–519, 807 inornata, 513, 515, 518 laurentiana, 513, 515, 518–519 onornata, 519 parva, 512–515, 517, 519, 740, 749, 751 strobilus, 512 Chrysoclonium, 473, 501, 503 Chrysococcus, 484, 486, 492, 497, 499 minutus, 478 rufescens, 487 Chrysocrinus, 484–485 Chrysodictyon, 503 Chrysodidymus, 523–526, 528, 530–531, 533–534, 541–542, 551–552 gracilis, 534 synuroideus, 534, 536, 538, 542 Chrysoikos, 497 Chrysolepidomonas, 484, 493, 497, 503 dendrolepidota, 480, 497 Chrysolepidomonadaceae, 484 Chrysolykos, 478, 486–487, 492, 497, 501 planktonicus, 478 skujae, 478 Chrysomallus, 443 Chrysomonadaceae, 471 Chrysophaerella, 523 Chrysophyceae, 19, 73, 313, 414, 429, 440, 443, 471–472, 484–486, 488, 491, 497, 500, 503, 511–513, 523–524, 534–535, 771 Chrysophyta, 5, 9, 24, 259, 757 Chrysopyxis, 492, 497 canadensis, 496 stenostoma, 478 Chrysosaccus, 491, 495, 497 incompletus, 484 Chrysosphaera, 495, 497

902

Taxonomic Index

Chrysosphaerella, 473–474, 486–488, 491, 493, 497 brevispina, 475, 479, 486, 487–488, 491 longispina, 475, 486–488, 491 Chrysostephanosphaera, 494, 497–499 globulifera, 476 globulosa, 482 Chrysoxys, 493, 498 maior, 479 Chytridiochloris, 431, 436, 458 acus, 456 viridis, 458 Cladocera, 16, 20, 38–39, 408, 411, 700, 808, 825 Cladonia, 258 Cladophora, 4, 24, 26–27, 34–35, 39, 101, 107, 208, 288, 314, 316, 318, 321–322, 334–337, 339–340, 605, 763, 768, 783, 808–810, 813, 817, 818–820 amethystinus, 101 fallax, 101 glomerata, 25–26, 34, 43, 314, 327, 608–609, 624, 764, 809–810, 822 halophilus, 101 minutus, 101 portoricensis, 101 Cladophorales, 311–313, 317, 319, 339–340, 454 Cladophorophyceae, 313 Clastidium, 70, 98, 101 cylindricum, 98, 101 setigerum, 98, 101 Cloniophora, 321, 332 spicata, 325 Closteriaceae, 372–373 Closteriopsis, 263, 274 longissima, 275 Closterium, 40, 361–362, 364, 367–368, 372, 377 aciculare, 364, 372 actum, 356 acutum, 372 angustatum, 357 archerianum, 357 closteroides, 357 dianae, 357 gracile, 357 navicula, 357 Coccolithales, 511 Coccomonas, 236, 248 orbicularis, 236–237 Coccomyxa, 258, 265, 274 dispar, 275 Cocconeidaceae, 597–598, 605, 623 Cocconeis, 24, 597–598, 601, 604–606 disculus, 624 fluviatilis, 624 globularis, 28 neothumensis, 623–624 pediculus, 34, 609, 624 placentula, 25, 28, 33, 43 placentula var. lineata, 624 placentula var. rouxii, 624 tomentosa, 28

Codioliales, 313, 347 Codiolum, 347 Coelastrum, 257–258, 264, 274, 453 reticulum, 275 Coelomoron, 65, 70, 82, 84 microcystoides, 84 minimum, 82 pusillium, 82 regulare, 67, 82 tropicalis, 84 Coelosphaerium, 2, 23, 64–65, 70, 82, 84, 87, 303 aerugineum, 84, 87 collinsii, 82 confertum, 87 dubium, 87 kuetzingianum, 84, 87 microcystoides, 82 minimum, 82 minutissimum, 87 naegelianum, 85, 89 subarcticum, 84, 87 vestitum, 82 Colacium, 12, 384, 385–386, 388, 404, 405–406, 408, 409–412, 415 calvum, 408, 411 gojdicsae, 408, 411 libellae, 388, 394, 404, 408, 411 oblonga, 404 oblongata, 404 pisciformis, 404 proxima, 404 rubra, 404 sanguinea, 404 schmitzii, 404 sociabilis, 404 splendens, 404 vesiculosum, 393, 403–404, 406, 408, 411 Coleochaetales, 308, 312–313, 319, 330, 332–333 Coleochaete, 24, 25, 320, 332, 353 scutata, 26 Coleodesmium, 119, 124, 158–159 floccosum, 158–159 wrangelii, 158–159 Colteronema, 125 funebre, 186–187 Compsopogon, 4, 200, 205, 207, 210, 212 coeruleus, 198, 200, 204, 211–212 prolificus, 198 Compsopogonales, 198, 201, 210, 212 Compsopogonopsis, 205, 207, 211–212 leptocladus, 198 Conferva, 312 Conjugatophyceae, 353, 744 Conjugatophyta, 353 Conochaete, 322, 332 Conradiella, 523–524 Contorta (section of Batrachospermum), 215 Corona, 245 Coronastrum, 263, 274 aestivale, 275 Corvomeyenia everetti, 45

Coscinodiscales, 563 Coscinodiscophycidae, 563, 565 Coscinodiscus, 570, 575, 578–579 Cosmarium, 354, 356, 364, 368, 374–376 contractum, 359 eloiseanum, 365 margaritatum, 359 montrealense, 359 pseudoconnatum, 359 quadrifarium f. hexastichum, 359 Cosmioneis, 638–639, 641, 645 pusilla, 646 Cosmoastrum, 378 Cosmocladium, 354, 368, 374, 376 constrictum, 374 pulchellum, 374 pusillum, 376 saxonicum, 356, 374 tuberculatum, 376 tumidium, 376 Crateriportula, 563, 578 inconspicuus, 579 Craticula, 638, 640, 646, 647 cuspidata, 643 Crucigenia, 3, 43, 264, 276 quadrata, 275 Cryptista, 715, 742 Cryptoglena, 384, 385, 388, 401–402, 410, 411, 415 pigra, 396, 400, 412 Cryptomonadaceae, 743–745 Cryptomonadales, 715, 742, 744 Cryptomonas, 23–24, 719–720, 723, 736, 740–741, 743–745, 749 erosa, 719–720, 745 obovata, 719–720, 745 ovata, 719–720, 730, 734, 745 ozolini, 719, 745 phaseolus, 720, 745 pyrenoidosa, 744 rostratiformis, 745 tetrapyrenoidosa, 720, 723, 745 Cryptomonas (Campylomonas) marssonii, 721 platyuris, 721–722 rostratiformis, 721–722, 739 Cryptophyceae, 19, 715, 742, 744 Cryptophyta, 7, 24, 686, 715–716, 737, 742 Ctenocladales, 333, 335 Ctenocladus, 43, 321, 333 circinnatus, 43, 327, 333 Ctenopharyngodon idella, 820 Ctenophora, 596, 600, 606–607 pulchella, 612 Cualobacter, 739 Cyanidioschyzon merolae, 41 Cyanidium, 200, 206, 208 caldarium, 41, 198, 201, 206, 207 Cyanobacteria(-um) , 1–3, 5–6, 8, 12, 19–20, 22–25, 27, 33–37, 39–45, 59–68, 117–121, 132, 135, 137, 139, 144, 187, 189, 205, 315, 320, 332, 364,

Taxonomic Index 372, 384, 487, 489, 647, 670, 674–675, 681, 701, 735, 738, 759, 763, 765, 805–809, 812–815, 819, 821–824 Cyanobacterium, 66, 69, 79 cedrorum, 75, 79 diachloros, 75 minervae, 66, 75, 79 Cyanobium, 20, 65, 69, 73, 79, 120 amethystinum, 79 eximium, 73, 79 gracile, 73 roseum, 73, 79 Cyanocystis, 65, 71, 100, 102 hemisphaerica, 102 mexicana, 100, 102 olivacea, 102 pacifica, 100, 102 pseudoxenococcoides, 102 sphaeroidea, 102 valiae-allorgei, 100, 102 violacea, 102 Cyanoderma bradypodis, 315 Cyanodictyon, 68–69, 73, 79 filiforme, 79 planctonicum, 73, 79 reticulatum, 79 tubiforme, 79 Cyanokybus, 71, 93, 95 venezuelae, 93, 95 Cyanomonas, 739, 748 Cyanosaccus, 67 Cyanosarcina, 62, 71, 95 Cyanosarcina (Chroococcus) minutus, 94 minutus var. thermalis, 94 mipitanensis, 94 polymorphus, 94 turgidus, 94 yellowstonensis, 94 Cyanotetras, 70, 83, 87 aerotopa, 87 crucigenielloides, 83, 87 Cyanothece, 20, 66, 69, 77, 79 aeruginosa, 77, 80 lineata, 80 major, 77, 80 Cyanothrix, 78, 80 primaria, 78 willei, 78 Cyathobodo, 498 Cyathomonas, 744 Cyclonexis, 493 annularis, 479, 498 Cyclops, 393, 411 Cyclostephanos, 563, 572, 578–579, 584–585 costatilimbus, 581 damasii, 581 dubius, 579 invisitatus, 581 tholiformis, 581 Cyclotella, 37, 43, 563, 565, 567, 572–573, 579, 581–582, 585, 768 bodanica, 566, 583

cryptica, 569 glomerata, 565 melosiroides, 565 meneghiniana, 569, 585 ocellata, 578 pseudostelligera, 566, 578, 583 radiosa, 578 Cyclotella (Stephanocyclus) meneghiniana, 22 Cyclotubicoalitus, 563, 571, 582–583 undatus, 588 Cylindriastrum, 378 Cylindrocapsa, 318, 340 geminella, 328 Cylindrocapsales, 317, 340–341 Cylindrocystis, 363, 367, 369 brebissonii, 358, 369 crassa, 369 Cylindrospermopsis, 64, 120, 124, 174–175, 807 catemaco, 174 raciborskii, 174–175 Cylindrospermum, 65, 119, 124, 174–175 catenatum, 174 longisporum, 175 minutissimum, 175 stagnale, 175 Cylindrotheca, 670, 675 gracilis, 676–677 Cymatopleura, 676 elliptica, 679 solea, 670, 672, 679 Cymbella, 24, 33, 39, 597, 656, 663–664, 676 affinis, 658 amphicephala, 662 aspera, 662 cesatii, 662 cistula, 661 cymbiformis, 661 lata, 661 minuta, 36 proxima, 658, 661 pusilla, 662 sinuata, 664 turgidula, 658, 661 Cymbellaceae, 655–656, 659, 661–664 Cymbellonitzschia, 670, 675 diluviana, 676–677 Cystodinedria, 690–691, 697, 701, 703, 708 inermis, 688, 696, 705 Cystodinium, 690–691, 697–698, 704, 708–709 bataviense, 688, 694, 696, 705

Dactylococcus, 288 Dacytlococcopis, 40 raphidioides, 76 Daphnia, 411, 700, 808, 813–814, 819 hyalina, 739 laevis, 408 pulex, 408 Dasygloea, 123, 147–148 amorpha, 147 brasiliense, 148

903

calcicola, 147 lamyi, 147–148 yellowstonensis, 147–148 Deasonia, 266–276 granata, 277 Debarya, 366, 369–370 glyptosperma, 355 smithii, 355 Dendromonas, 491, 498 cryptostylis, 477 Denticula, 670, 675–676 tenuis, 677 Derepyxis, 492 anomala, 498 dispar, 478 ollula, 475 Dermatochrysis, 2, 494, 498, 503 reticulata, 482 Dermatophyton, 314, 319, 340 radians, 327 Dermocarpa pacifica, 100 Dermocarpella, 71, 100, 102 hemisphaerica, 100 prasina, 102 protea, 102 Dermocarpellaceae, 60, 70–71, 102 Desmatractum, 266, 276 bipyramidatum, 277 Desmidiaceae, 373–379, 441 Desmidiales, 313, 353–354, 361–365, 372–379 Desmidium, 366, 374 baileyi, 357 grevillii, 357 Desmococcus, 258, 264, 276, 315, 322, 333, 346 olivaceus, 277, 323 Desmodesmus, 254, 256–258, 261, 276, 288, 299 armatus, 255 protuberans, 278 Desmonema wrangelii, 158–159 Diacanthos, 261, 276 belenophorus, 278 Diachros, 430–432, 437, 446 simplex, 445 Diacronema, 512, 515 Diadesmis, 637–640, 647 confervacea, 645 contenta, 645 perpusilla, 645 Diaptomus gracilis, 739 Diatoma, 596, 600, 606–608, 616 hiemale, 613 mesodon, 612–613 tenue, 36, 613 tenue var. elongatum, 612 vulgare, 34, 36, 612–613 Diatomella, 638–639 balfouriana, 638, 645, 647 Dicellula, 261, 276 planctonica, 278

904

Taxonomic Index

Dichothrix, 124, 164–166 baueriana, 166 calcarea, 166 compacta, 166 gypsophila, 166 hosfordii, 166 inyoensis, 166 meneghiniana, 166 orsiniana, 165 rupicola, 166 spiralis, 166 willei, 165 Dichotomococcus, 431, 437, 453 elongatus, 452 lunatus, 451 Dichotomosiphon, 319, 344 tubersosus, 328 Dicranochaete, 321, 333 reniformis, 324 Dictyochlorella, 264, 276 reniformis, 278 Dictyochloris, 266, 276 fragrans, 278 Dictyochloropsis, 276 splendida, 279 Dictyococcus, 266, 277 varians, 279 Dictyosphaerium, 254, 257–258, 263, 280 pulchellum, 279 Didymogenes, 262, 280 palatina, 279 Didymosphenia, 656, 659, 663–664 geminata, 33, 658, 661–662 Difflugia, 458, 461 Dilabifilum, 315, 320, 333 printzi, 325 Dimorphococcus, 263, 280 lunatus, 279 Dinamoebaceae, 697, 706 Dinamoebales, 697, 706 Dinamoebidium, 697–698, 701, 703, 706 coloradense, 688, 702 Dinastridium, 697–698, 703, 708 sexangulare, 688, 705 Dinobryaceae, 484 Dinobryon, 3, 23, 471, 473, 477, 480, 484–489, 492, 497–498, 501–503 acuminatum, 489 attenuatum, 498 balticum, 489 bavaricum, 488–489 borgei, 488–489, 498 cylindricum, 485, 487–489 dilatatum, 498 dillonii, 477 divergens, 477, 487, 489 lorica, 480 pediforme, 488–89 sertularia, 22, 487–490 suecicum, 475, 488, 498 tubaeforme, 498 Dinobryopsis, 498 Dinococcales, 696–697, 701, 708 Dinococcus, 688, 696–698, 703, 708

Dinosphaera, 697–698, 705, 708 palustris, 693 Dinophyceae, 697 Dinophyta, 697 Dinosphaeraceae, 697, 708 Dioxys, 431, 436, 456, 458 inermis, 456 tricornuta, 456 Diplocolon heppii, 158 Diploneis, 638–639, 647 elliptica, 642 finnica, 642 oblongella, 642 smithii var. dilatata, 642 Diplonema, 387, 405 Diplopsalis, 708 Diptera, 700 Discoglena, 402, 412 Dispora, 264, 280 crucigenoides, 279 Distigma, 385 Distrionella, 600, 606, 613 Docidium, 367, 375 baculum, 357 Dolichospermum, 169 Draparnaldia, 4, 35, 314, 321–322, 333–334, 483, 498 glomerata, 325 ravenelii, 325 Draparnaldiopsis, 321, 334 alpinis, 325 Dreissena polymorpha, 37 Ducelliera, 431, 437, 453 chodati, 452 Dunaliella salina, 43 viridus, 43 Durinskia, 697–698, 705, 707 baltica, 689, 692, 694 Dysmorphococcus, 236, 248 globosus, 236 variabilis, 236–237

Echinosphaerella, 262, 280 Ectocarpales, 758–759, 768 Ectocarpus, 758, 760, 768 confervoides, 759, 768 siliculosus, 759, 761, 765–766, 768 Ectogeron, 261, 280 Eichhornia crassipes, 821 Eirmodesmus, 494, 498 phaeotilus, 483, 498 Elakatothrix, 263, 280, 317, 344 americana, 345 gelatinosa, 326 viridis, 326 Ellerbeckia, 563, 565, 571, 584 arenaria, 576, 583 Ellipsoidion, 424–425, 430, 438, 446 acuminatum, 425–427, 446 stellatum, 445

Elodea, 314, 771, 810–811 canadensis, 819 Enallagma civile, 408 Encephalartos, 45 Encyonema, 656, 659, 663–664 helvetica, 662 norvegica, 661 Endochloridion, 430, 432, 438, 446 polychloron, 445 Endoclonium rivulare, 338 Endospora rubra, 95 Enteromorpha, 27, 34, 317, 347, 763, 771 flexuosa, 347 intestinalis, 43, 329, 347 prolifera, 347 Entocladia, 321, 334, 338 polymorpha, 323 Entomoneidaceae, 670, 681 Entomoneis, 670, 675, 681–682 ornata, 672, 678 Entophysalidaceae, 62, 71, 96 Entophysalis, 62, 67, 71, 96–97 atrata, 96 cornuana, 96 lemaniae, 96 lithophila, 96, 97 rivularis, 96 willei, 96, 97 Entransia, 363, 366, 369 fimbriata, 355 Entzia, 697–698, 704, 708 acuta, 690, 693, 695 Epigloeosphaera, 69, 74, 80 glebulenta, 74, 80 Epihydra, 41 Epipyxis, 484, 486–487, 491–492, 498, 501 pulchra, 473 ramosa, 476–477 tabellariae, 475 Epithemia, 25, 34, 39, 43, 669–670, 673, 675, 678, 680–681 turgida, 672 Epithemiaceae, 676 Eremosphaera, 265, 280 viridis, 282 Eremotyl, 280 Erkenia, 493, 498–499 subaequiciliata, 480 Errerella, 288 Euastrum, 361, 368, 375–376 boldtii, 359 divaricatum, 359 humerosum, 359 pseudoboldtii, 359 verrucosum, 359 Eucapsis, 66, 70, 86, 89 alpina, 86, 89 alpina var. maior, 89 minor, 89 Eucocconeis, 597, 598, 603, 606, 623 flexella, 626, 627 flexella var. alpestris, 626

Taxonomic Index Eucyonema minuta, 658 muelleri, 658 Eudinobryon, 498 Eudorina, 24, 239, 241, 242, 247–248 conradii, 239 cylindrica, 239 elegans, 23, 239, 240 illinoisensis, 239 interconnexa, 239 unicocca, 239 Euglena, 2, 23, 24, 383, 385–386, 388, 389–390, 401–406, 407, 408–409, 410, 412, 414–415, 806 acus, 390, 406 adhaerens, 390, 403 agilis, 388 caudata, 391 chadefaudii, 390, 402–403 clavata, 390 deses, 399, 404, 406 geniculata, 402, 406 gracilis, 387, 392, 399, 404–407 granulata, 403 jirovecii, 391 mutabilis, 42, 390, 404, 406–407 myxocylindracea, 388, 403–404, 406 oblonga, 390 obtusa, 406, 407–408 orientalis, 404 oxyuris, 390, 403, 406 oxyuris var. charkowiensis, 406 pisciformis, 404, 406–407 polymorpha, 403, 405 proxima, 406 repulsans, 390 rubra, 405, 407, 412 sanguinea, 407, 412 schmitzii, 405 sima, 400 sociabilis, 391, 406 spirogyra, 390, 401, 406, 412 splendens, 403, 405 stellata, 403–404 texta, 390 tripteris, 390, 392, 404, 406 tristella, 403, 406 truncata var. baculifera, 391 tuba, 391, 404–405, 412 velata, 403 viridis, 384, 390, 403–406 walnei, 403 Euglenamorpha, 388, 410, 412, 415 hegneri, 397 Euglenida, 383, 385 Euglenophyceae, 383 Euglenophyta, 6, 383, 686, 806 Euglenoza, 383, 385, 387 Eunophora, 656 Eunotia, 33, 39, 655–656, 664–665 bilunaris var. mucophila, 662 elegans, 40 exigua, 40, 661 faba, 657 formica, 661

microcephala, 662 papilio, 661 pectinalis, 24 pectinalis var. minor, 661 praerupta, 661 ruzickae, 662 septentrionalis, 662 serra, 657 tenella, 42 Eunotiaceae, 655–656, 661, 664–665, 681 Euplotes, 45 Eusphaerella, 493, 499 turfosa, 479, 499 Eustigmatales, 424 Eustigmatophyceae, 6, 8, 423–424, 426–427, 446–447, 449, 465 Eustigmatos, 424–425, 447, 463 magnus, 425–426 vischeri, 425 Eustropsis, 263, 280, 282 Eutreptia, 385, 388, 401, 404, 410, 413, 415 globulifera, 394, 413 pertyi, 413 viridis, 413 Eutreptiella, 388, 402, 413 eupharyngea, 413 gymnastica, 404 Euvolvox (section of Volvox), 242 Exanthemachrysis, 512, 515 noctivaga, 515 Excentrochloris, 430, 436, 451 gigas, 450 Excentrosphaera, 265 viridis, 282 Exuviaella, 697–698, 701, 703–704, 709 compressa, 689

Falcomonas, 743 Fallacia, 638, 640, 647 pygmaea, 645 Fasciculochloris, 265 boldii, 283 Ferrissia fragilis, 25 Filoprotococcus, 315, 318, 345 polymorphum, 329 Fischerella, 118, 120, 125, 184–185 ambigua, 184–185 letestui, 184 major, 184 thermalis, 184–185 Fischerellaceae, 184 Fistulifera, 639–640, 647 saprophila, 643 Flintiella, 208 sanguinaria, 198, 200, 203 Florideophycidae, 197–198, 200–201 Fortiea, 124, 161, 163–164 bossei, 164 monilispora, 163–164 salinicola, 163–164 Fottea, 282 cylindrica, 283

905

Fottiella, 263, 284 quadrangularis, 283 Fragilaria, 20, 33, 39, 43, 595–596, 599, 606, 610, 709 brevistriata, 616 capucina, 613 construens, 616 crotonensis, 20–21, 605, 607–608, 613–614 crotonensis var. oregona, 614 elliptica, 616 intermedia, 614 lapponica, 616 leptostauron, 616 pinnata, 616 vaucheriae, 25, 613–614 virescens, 616 virescens var. exigua, 616 Fragilariaceae, 596, 598, 604, 607, 610, 613 Fragilariales, 598, 607 Fragilariforma, 596, 599, 606, 613, 616, 631 constricta, 614 virescens, 614 Fragilariophyceae, 595–598, 604, 607 Franceia, 255–256, 262, 284 droescheri, 283 Frankophila, 599, 613 Fremya, 431, 435, 461 sphagni, 460 Fremyella, 164 diplosiphon, 164 robusta, 164 tenera, 164 Fridaea, 320, 334 torrenticola, 325 Fritschiella, 319, 334 tuberosa, 315, 325 Frustulia, 24, 39, 639, 640, 647 rhomboides, 40, 645 rhomboides var. saxonica, 645 saxonica, 40 vulgaris, 645

Galdiera sulphuraria, 41 Gamophyta, 353 Geissleria, 637–638, 640, 647–648 ignota var. palustris, 644 Geitleria, 125 calcarea, 186–187 floridana, 187 Geitleribactron, 70, 98, 101–102 crassum, 98, 102 periphyticum, 98 Geitlerinema, 119, 122, 126–127 amphibium, 126 claricentrosa, 127 claricentrosum, 126 earlei, 126 lemmermannii, 127 splendidum, 126–127 unigranulatum, 127 Geminella, 319, 345 interrupta, 326 minor, 326

906

Taxonomic Index

Geminigera, 734, 742–743 Geminigeraceae, 743 Genicularia, 367–368, 373 elegans, 358 Geosiphon, 180 Glaucospira, 122, 129, 131–132 laxissima, 131 yellowstonensis, 132 Glenodiniopsidaceae, 697, 708 Glenodiniopsis, 697–698, 705, 708 steinii, 693–694 Glenodinium, 707 cinctum, 708 palustre, 708 Gleosphaeridium, 431 Gloechloris, 435 Gloeoactinium, 264, 284 limneticum, 285 Gloeobotrydaceae, 431, 436, 453, 455 Gloeobotrys, 431, 433–434, 437, 454 limneticus, 455 Gloeocapsa, 2, 20, 26, 44, 60, 62–64, 70, 89, 91, 95 acervata, 92 alpicola, 92 alpina, 91, 92 arenaria, 92 atrata, 92 calcicola, 92 caldariorum, 92 conglomerata, 91, 92 crepidinum, 93 decorticans, 92 dermochroa, 91 fusco-lutea, 92 gelatinosa, 91, 92 granosa, 92 kuetzingiana, 92 magma, 95 nigrescens, 92 rupestris, 92 sanguinea, 91, 92 sparsa, 92 sphaerica, 92 thermophila, 92 Gloeocapsopsis, 62, 71, 93, 95 crepidinum, 93 magma, 93 Gloeochaete, 263, 284 wittrockiana, 285 Gloeochloris, 430, 443 planctonica, 444 smithiana, 443 Gloeococcus, 257, 259, 284 pyriformis, 285 Gloeocystis, 258, 265, 284 bacillus, 285 Gloeocystopsis, 291 Gloeodendron, 259, 284 catenatum, 286 Gloeodinium, 696, 707 montanum, 688, 694 Gloeomonas, 232, 248 kupfferi, 235 ovalis, 235

Gloeopodiaceae, 431, 436, 454, 457 Gloeopodium, 431, 436, 454 rivulare, 457 Gloeoskene, 431, 437, 454 turfosa, 455 Gloeosphaeridium, 437, 454 firmum, 455 Gloeotaenium, 263, 284 loitlebergerianum, 286 Gloeothece, 64, 69, 75, 80 confluens, 80 distans, 80 endochromatica, 80 fusco-lutea, 80 heufleri, 75 interspersa, 75, 80 linearis, 80 linearis var. composita, 76 membranacea, 80 opalothecata, 80 palea, 80 prototypa, 80 rupestris, 75, 80 transsylvanica, 81 Gloeotila, 319, 345 contorta, 326 Gloeotilopsis, 318, 345 sterile, 326 Gloeotrichia, 25, 39, 119, 124, 166–167, 770 echinulata, 23, 166–167 natans, 166 pilgeri, 166 pisum, 166–167, 764 Godlewskia, 65, 71, 101 Golenkinia, 257, 262, 284 radiata, 286 Golenkiniopsis, 262, 284 solitaria, 286 Gomontia, 319, 334 holdenii, 323 perforans, 314 Gomphoneis, 655–56, 659, 661, 663 elegans, 659 eriense, 665 eriense var. variabilis, 660 herculeana, 33, 35, 659, 665 olivacea, 660, 665 olivaceum, 662 quadripunctata, 665 Gomphonema, 24–25, 39, 604, 655–656, 659, 663, 665 abbreviatum, 662 acuminatum, 660 acuminatum var. brebissonii, 662 angustatum, 661–662 apuncto, 660–661 augur, 662 brasiliense, 661, 666 gracile, 24, 661 grovei, 661, 666 hebridense, 661 lingulataeforme, 666 manubrium, 661 minutum, 662

olivaceum, 36 parvulum, 661–662 sphaerophorum, 660, 662 subtile, 662 truncatum, 660, 662 Gomphonemataceae, 655–656, 659, 661–662, 665 Gomphosphaeria, 60, 65, 70, 86–87 aponina, 64, 86–87 cordiformis, 87 irieuxii, 86 lacustris, 85 multiplex, 64, 87 natans, 64, 67, 86–87 salina, 87 semen-vitis, 67, 86–87 virieuxii, 87 wichurae, 89 Gomphosphenia, 663, 665–666 grovei, 661 lingulataeforme, 661 Gonatozygon, 366–368, 373 brebissonii, 358 monotaenium, 359 Gongrosira, 33, 319–320, 335, 764 debaryana, 314, 325, 335 incrustans, 25 pseudoprostrata, 335 scourfieldia, 335 Goniaceae, 225, 228, 239, 243–246 Goniochloris, 430–431, 433, 438, 446 fallax, 465 sculpta, 445 Goniomonadaceae, 742, 744 Goniomonadales, 742, 744 Goniomonadophyceae, 742 Goniomonas, 719,–20, 734–736, 740–744, 749 truncata, 719–720, 733, 744 Gonium, 242–243, 248 discoideum, 244 formosum, 244 multicoccum, 243–244 octonarium, 244 pectorale, 227, 243–244 sacculifera, 245 sociale, 227, 238 Gonyaulacaceae, 697, 707 Gonyaulax, 687, 697–698, 701, 704, 707 palustris, 708 spinifera, 689, 692 Gonyostomum, 426,–428, 463, 465 depressum, 428 latum, 428 semen, 428 Granulochloris, 236, 248 spinifera, 237–238 Groenbladia, 366, 375 neglecta, 375 undulata Groenlandiella, 443 Guillardia, 743 theta, 734 Gumaga nigricula, 35

Taxonomic Index Gymnodiniaceae, 697, 706 Gymnodiniales, 697, 706 Gymnodinium, 691, 697–698, 704, 708 acidotum, 688 caudatum, 702 cruciatum, 702 fuscum, 705–706 helveticum, 690 marylandicum, 702 paradoxum, 694 triceratium, 706 Gyrodinium, 687, 691, 697–698, 704, 706 pesillium, 688 Gyromitus disomatus, 512 Gyrosigma, 638–639, 648, 650 parkeri, 641 scalproides, 641

Haematococcaceae, 225, 228, 232–233 Haematococcus, 231–232, 248 carocellus, 232 lacustris, 45, 232 pluvialis, 231–232 Haidadinium, 697–698, 701, 703, 708 ichthyophilum, 688, 702 Hammatoidea, 151 Hannaea, 596, 600, 606, 631 arcus, 33, 36, 613, 615 arcus var. amphioxys, 615 Hantzschia, 45, 670, 675, 681 amphioxys, 672, 676–677 Hanusia, 743 Hapalosiphon, 24, 40, 44, 65, 119, 125, 188–189 aureus, 189 brasiliensis, 189 confervaceus, 189 delicatulus, 189 flexuosus, 189 fontinalis, 189 hibernicus, 188–189 intricatus, 189 pumilus, 27, 189 tenuis, 189 welwitschii, 188–189 Haplotaenium, 362, 367, 375, 377, 379 minitum, 357, 375 sceptrum, 375 Haptophyceae, 7, 471, 511 Haptophyta, 1, 5, 7, 9, 807 Harpochytrium, 458 Hassallia, 119, 124, 158, 160 byssoidea, 160 discoidea, 160 granulata, 160 Hazenia, 315, 320, 335 mirabilis, 323 Heimansia, 354, 368, 374–376, 379 pusilla, 360 Heliapsis, 492, 499, 502 mutabilis, 479 Helicodictyon, 315, 320, 335 planktonicum, 323

Helminthogloea, 430, 435, 443 ramosa, 444 Hemidiniaceae, 697, 707 Hemidinium, 688, 696–698, 703–707, 709 nasutum, 693, 695 Hemidiscaceae, 563 Hemiselmidaceae, 743, 748 Hemiselmis, 743, 748–749 amylifera, 748 amylosa, 726–727, 729 virescens, 738 Hemisphaerella, 431, 436, 458 operculata, 456 Heribaudiella, 4, 760, 762, 768, 771 fluviatilis, 25, 33, 757–759, 763–766, 770 Heterochloris, 429, 430, 435, 439 mutabilis, 439 viridis, 439–440 Heterochromonas, 502 Heterococcus, 431, 435, 459, 461, 465 ramosissimum, 460 Heterodendraceae, 431 Heterodendron, 431, 434, 461 pascheri, 460 Heterodesmus, 431, 437, 453 bichloris, 452 Heterogloea, 430, 434, 436, 443–444 endochloris, 444 minor, 444 Heterogloeaceae, 430 Heterogloeales, 429–430, 432, 434, 441, 444 Heterohormogonium, 80 schizodichotomum, 78 Heteroleibleinia, 65, 122, 132, 134–135 kuetzingii, 134 minor, 134 profunda, 134 pusilla, 134–135 versicolor, 135 Heteroleibleinioideae, 135 Heteromastix, 230 angulata, 230 Heteropediaceae, 431 Heteropedia, 431, 435, 459, 461 polychloris, 460 Heterothrix, 431, 434, 461 exilis, 462 Hildenbrandia, 4, 34, 202, 204, 207, 218, 763–764, 768 angolensis, 199, 218–219 rivularis, 25, 33, 204, 218, 765, 770 Hildenbrandiales, 199, 201–203, 218–219 Hippodonta, 640, 648 capitata, 643 hungarica, 643 Homoeothrix, 65, 122, 135–137, 763–764 crustacea, 136–137 janthina, 136–137 stagnalis, 137 varians, 136–137 violacea, 137 Hordeum, 822 Hormathonema, 96

907

Hormidiopsis, 319, 345 crenulatum, 345 ellipsoideum, 326 Hormidium, 313, 346 Hormotila, 285 blennista, 287 Hormotilopsis, 255, 285 Hyalella azteca, 825 Hyalobryon, 498 Hyalosynedra, 596 Hyalotheca, 364, 366, 376, 441, 458 dissiliens, 358, 376 mucosa, 376 Hybrida (section of Batrachospermum), 215 Hydra, 45, 272 viridis, 258 Hydrilla verticillata, 816 Hydrococcaceae, 71, 96 Hydrococcus, 62, 65, 71, 96, 98 cesatii, 96, 98 rivularis, 96 Hydrocoleum, 123, 147, 149–151 groesbeckianum, 149 homoeotrichum, 149–150 Hydrocoleus, 151 Hydrodictyon, 24, 255, 257, 259, 285, 808, 810, 818, 822–823 reticulatum, 287 Hydrosera, 563, 571, 583 whampoensis, 577 Hydrurus, 494, 500 foetidus, 12, 35, 476, 483, 488, 499 Hyella, 60, 62, 67, 72, 107, 109 balani, 67, 107, 109 caespitosa, 107, 109 fontana, 107, 109 gigas, 107 kalligrammos, 107 linearis, 107 littorinae, 107 pyxis, 107 seriata, 109 tenuior, 67, 107 vacans, 107 Hyellaceae, 60, 62, 71, 107 Hymenomonas, 512–513, 515, 519 coccolithophora, 516 danubiensis, 516 prenanti, 512 roseola, 511–512, 514, 516, 519 scherffelli, 516 Hypheothrix (Symplocastrum) parciramosa, 148 Hypnodinium, 690–691, 696–698, 704, 709 sphaericum, 688 Hypophthalmichthys molitrix, 820

Ilsteria, 431, 437, 453 quadrijuncta, 452 Imantonia, 512 Inactis, 119, 137

908

Taxonomic Index

Infusoria, 384–385, 471 Ischnura verticalis, 394, 408 Isochrysidales, 511, 513 Isocystis, 124, 174, 176–177 infusionum, 176 planctonica, 176 Isthmochloron, 430, 439, 445, 447 lobulatum, 445 trispinatum, 445

Jaaginema, 119, 122, 126, 128 filiforme, 126 neglecta, 128 subtilissima, 128 Johannesbaptistia, 69, 78, 80 pellucida, 78, 80 primaria, 78 schizodichotoma, 78, 80

Kansodinium, 697–698, 708 ambiguum, 693, 694, 702 Karayevia, 596–598, 603, 607, 627 clevei, 596, 609 clevei var. rostrata, 596, 626 laterostrata, 626 Kathablepharidaceae, 715–716, 749–751 Kathablepharis, 716, 732, 735, 739–740, 743, 749, 750–751 ovalis, 731–732, 733, 740, 750–751 phoenikoston, 731–732, 733, 740, 749–751 Katodinium, 687, 691, 697–698, 704, 706, 709 auratum, 702 fungiforme, 701 spiroidinoides, 688, 705 Kentrosphaera, 266, 288 facciolae, 287 Kephyrion, 486–487, 492, 497, 499, 503 obliquum, 478 Kephyriopsis, 501 Keratella, 393, 411, 825 Keratococcus, 263, 288 bicaudatus, 287 Keriochlamys, 265, 288 styriaca, 287 Keriosphaera, 430, 438, 447 gemma, 448 Khawkinea, 384–385 Kinetoplastida, 385–387, 402 Kirchneriella, 262, 288 obesa, 289 Klebsiella, 388, 411 Klebsormidiales, 313, 345–347 Klebsormidiophyceae, 313 Klebsormidium, 45, 313, 318, 344–345 flaccidum, 315 klebsii, 326 rivulare, 42 Kobayasia, 639, 648 subtilissima, 645 Kolbesia, 597–598, 603, 607 amoena, 627

kolbei, 626 ploenensis, 626–627 suchlandtii, 627 Koliella, 318, 326, 346 Komma, 741, 743, 749 caudata, 726–728, 747 pochmanni, 728 Komvophoron, 122, 139–140 groenlandicum, 139 jovis, 139–140 minutum, 140 schmidlei, 140 Kybotion, 492, 499 eremita, 479 Kyliniella, 207, 210 latvica, 198, 209

Lagerheimia, 255–256, 262, 288–289 Lagynion, 492, 499 macrotrachelum var. oedotrachelum, 478 Lamprothamnium, 27, 317, 338 buckellii, 338 longifolium, 338 Larix, 429 Lauterborniella, 261, 288 elegantissima, 289 Leibleinia, 122, 132–133, 135 calotrichicola, 133 Lemanea, 33, 203–205, 208, 211, 216–217, 763 borealis, 199, 203 fluviatilis, 33, 35, 199, 203–205, 214, 216–217 fucina var. parva, 199 mamillosa, 202, 204 Lemaneaceae, 199, 203–206, 216–217, 219 Lemmermanniella, 68 Lemna, 274, 314, 316, 332, 429, 443, 459, 627, 708 minor, 628 wollei, 821 Lemnicola, 597–598, 603, 607, 623 hungarica, 627–628 Lepidochrysis, 474, 502 Lepochromulina, 492, 496, 498–499 bursa, 478 Lepocinclis, 388, 394, 401, 406–407, 410, 413, 415 capito, 394 fusiformis, 394 marssonii, 394 ovum, 394, 413 playfairiana, 413 salina, 413 Leptochaete, 137 Leptolyngbya, 63, 65–66, 119, 122, 132, 134–135 cartilaginea, 134 foveolarum, 134 nostocorum, 134 Leptolyngbyoideae, 132 Leptosira, 315, 320, 335–336 mediciana, 323 Leptosiropsis, 335

Leucocryptos, 716, 735, 749 acuta, 746 Leuvenia natans, 451 Limbata, 450 Limnococcus, 95 Limnothrix, 126, 128–129, 155 guttulata, 129 redekei, 128–129 vacuolifera, 129 Linnothrix, 122 Lithoderma arvenensis, 759 fluviatile, 759, 770 fontanum, 759, 770 zonatum, 759, 763, 770 Lithomyxa, 69, 80–81 calcigena, 81 Lobococcus, 289 Lobocystis, 264, 288 dichotoma var. mucosa, 289 Lobomonas, 232, 248 rostrata, 233 Lophodiniaceae, 697, 707 Lophodinium, 691, 697–698, 705, 707 polylophum, 689, 695, 701 Loriellaceae, 125, 184 Lucas, 743 Lutherella, 431, 433, 436, 456, 458 adhaerens, 456 globulosa, 456 Luticola, 637–638, 640, 648 goeppertiana, 644 mutica, 644 Lyngbya, 3, 24–25, 39–41, 43, 45, 66, 123, 132, 135, 151, 154–155, 807–809, 813, 819, 823 aeruginea, 132 aestuarii, 151 angustissima, 135 bijahensis, 135 birgei, 151, 154 calitrichicola, 132 cartilaginea, 135 confervoides, 151 contorta, 135, 155 giuseppei, 155 hahatonkensis, 155 intermedia, 154 lagerheimii, 135 magnifica, 154 maior, 151 martensiana, 151 meneghiniana, 151 patrickiana, 155 rubra, 135 salina, 151 spirulinoides, 151 splendens, 154 subterranea, 135 subtilis, 135 tenuis, 135 vesiculosa, 135 wollei, 808–809 yellowstonensis, 135

Taxonomic Index Lyngbyopsis, 123, 145, 150 willei, 145, 150

Macrozamia, 45 Magnolia, 328, 343 Malleochloris, 260, 288 sessilis, 290 Malleodendraceae, 430 Malleodendron, 430, 435, 443 caespitosum, 443 gloeopus, 443–444 Mallomonadaceae, 523–524 Mallomonas, 7, 523–552, 807 acaroides, 532 acaroides var. acaroides, 537–538, 542, 550 acaroides var. muskokana, 529–530, 535–536, 538, 542, 550 adamas, 524 akrokomos, 531, 542, 549 alpina, 537 annulata, 542, 549 asmundiae, 538, 542, 550 canina, 536, 542, 549 caudata, 525, 529, 535, 542, 549 corymbosa, 525, 537–538, 543, 550 crassisquama, 535, 539, 543, 550 cratis, 543, 550 dickii, 524, 527, 537, 543, 549 doignonii, 537 doignonii var. tenuicostis, 543 duerrschmidtiae, 529, 535, 537–538, 543, 550 elongata, 529, 537–538, 543, 549 fenestrata, 531 galeiformis, 535, 537–538, 543, 550 hamata, 524, 536, 538, 544, 548 heterospina, 530, 537, 538, 544, 549 hindonii, 536, 544, 549 lychenensis, 527, 538, 544, 548 mangofera, 544, 549 mangofera var. foveata, 544 matvienkoae, 538, 544, 549 paludosa, 536, 538 papillosa, 544, 549 portae-ferreae, 537–538, 545, 550 pseudocoronata, 528, 537–538, 545, 550 pugio, 536, 538, 545, 549 punctifera, 532, 537, 545, 548 retorsa, 531 splendens, 528 striata, 545, 550 tonsurata, 524–525, 527, 532, 537, 539, 545, 550 torquata, 524, 537–538, 545 torquata f. simplex, 545, 549 torquata f. torquata, 549 transsylvanica, 524, 537, 546, 548 Mallomonopsis, 523–524 Mantellum, 70, 83, 87 rubrum, 83, 87 Martyana, 596, 600, 606, 613 martyi, 615 Massartia, 706

Mastigocladaceae, 125, 189 Mastigocladus, 41, 66, 125 laminosus, 66, 189–190 Mastigophora, 385 Mastogloia, 39, 43, 569, 604, 638–639, 648 smithii var. lacustris, 644 Medicago, 822 Melastomaceae, 343 Melosira, 20, 33–34, 563, 565, 571–572, 583 undulata, 567, 576, 583 varians, 33, 576, 583 Melosiraceae, 563 Melosirales, 563 Meridion, 596, 600, 606 anceps, 616–617 circulare, 616–617 circulare var. constrictum, 617 Meringosphaera, 430, 438, 447 tenerrima, 448 Merismogloea, 431, 437, 454 polychloris, 455 Merismopedia, 43, 65–66, 69, 83, 87–88 angularis, 83, 88 convoluta, 88 elegans, 83, 88 elegans var. maior, 83 gardner, 88 glauca, 88 major, 88 punctata, 83, 88 smithii, 67, 83, 88 tenuissima, 88 Merismopediaceae, 69, 82 Merotrichia, 427–428 capitata, 426, 428 Mesodictyon, 563, 582, 584 Mesostigma, 228, 248 grande, 228–229 viride, 228 Mesotaeniaceae, 353 Mesotaenium, 354, 363, 367, 369 berggrenii, 369 degreyi, 369 endlicherianum, 358 kramstai, 364 Micractinium, 261, 288 pusillum, 290 Micrasterias, 2, 40, 361, 364, 368, 375–376 foliacea, 359, 366, 376 johnsonii var. ranoides, 360 muricata, 365 pinnatifida, 360 Microchaetaceae, 123, 158, 161 Microchaete, 124, 163–164 robinsonii, 163–164 tenera, 163 Microchaetoideae, 161 Microcoleoideae, 147 Microcoleus, 20, 24, 39, 66, 123, 139, 149–150 chthonoplastes, 149 lacustris, 150 purpureus, 139 vaginatus, 149

909

Microcostatus, 638, 640 krasskei, 645, 648 Microcrocis, 70, 84, 88 gigas, 88 irregulare, 84 irregularis, 88 obvoluta, 84, 88 pulchella, 84, 88 Microcystaceae, 62, 70, 89 Microcystis, 23, 37, 43, 60–61, 63–65, 68, 70, 90, 92, 805, 807, 822 aeruginosa, 20, 43, 90, 92 comperei, 90, 92 flos-aquae, 92 glauca, 92 ichthyoblabe, 92 natans, 92 pulchra, 90 smithii, 92 splendens, 94–95 viridis, 92 wesenbergii, 67, 90, 92 Microglena, 493, 499, 523–524 butcheri, 481, 499 punctifera, 499 Micromonadophyceae, 229 Microneis, 604 Microspora, 318, 341, 431 willeana, 326 Microsporales, 317, 341 Microthamniales, 336 Microthamnion, 321, 335 kuetzingianum, 325, 336 strictissimum, 336 Mimosaceae, 343 Mischococcaceae, 431, 436, 457 Mischococcales, 270, 430, 432, 434, 443, 445, 448, 450–459 Mischococcus, 429, 431, 433, 436, 454 confervicola, 457 Moina, 411 Monadodendron, 498 Monallantus, 430, 432, 438, 447 pyreniger, 448 Monimiaceae, 343 Monocilia, 459 Monochrysis, 487, 493, 499–500 vesiculifera, 481 Monodopsidaceae, 424 Monodopsis, 424–425, 427 subterranea, 426, 447 Monodus, 430, 439, 447 chodatii, 448 ovalis, 427, 447 subterraneus, 447 Monoraphidium, 263, 289 nanum, 447 pusillum, 290 Monostroma, 317, 347 latissimum, 329 Mougeotia, 24–26, 39, 354–355, 363, 366, 369–370, 370, 372, 393, 768 Mougeotiopsis, 363, 366, 370 calospora, 355, 370

910

Taxonomic Index

Muelleria, 638–639, 648 gibbula, 641, 649 terrestris, 649 Myriophyllum, 314–315, 322, 335, 811 brasiliense, 811 spicatum, 24, 816 Myrmecia, 254, 266, 276, 289 pyriformis, 290 Myxochloridaceae, 430 Myxochloris, 430, 435, 441 sphagnicola, 441–442 Myxosarcina, 71, 104 amethystina, 104 gloeocapsoides, 104 rubra, 104

Naegeliella, 494, 500 flagellifera, 483 Najas, 314–315, 322, 335 marina, 24, 810 Nannochloris, 20, 24, 265, 291 bacillaris, 290 occulata, 427 Nannochloropsis, 424–425, 427, 465 occulata, 427 oculata, 426 Nautococcus, 266, 291 piriformis, 292 Navicula, 24, 34, 39, 43, 45, 597, 637, 640–641, 643, 645, 647 avenacea, 35 capitata, 648 cocconeiformis, 646 contenta, 44 cryptocephala, 27 gastrum, 650 gregaria, 36 hasta, 641 hungarica, 27, 648 levanderi, 649 mutica, 648 paludosa, 648 pelliculosa, 647 placenta, 641 pupula, 650 pygmaea, 647 rhychocephala, 641 saprophila, 647 similis, 648 soehrensis, 646 subinflatoides, 43 subtilissima, 40, 648 tantula, 44 tenuicephala, 27 (tuscula group), 644 Naviculaceae, 637, 639, 665 Naviculales, 656 Neidiopsis, 639, 649 levanderi, 642 Neidium, 638–639, 649 densestriatum, 642 hitchcockii, 642 iridis, 642 Nemalionopsis, 207, 218

tortuosa, 199, 217 Neochloris, 256, 266, 291 aquatica, 292 Neomysis mercedis, 813 Neonema, 431, 434, 461 quadratum, 462 Neonemataceae, 431 Neospongiococcum, 266, 291, 303 gelatinosum, 292 Neosynedra, 596 Nephrochloris, 430, 435, 439 incerta, 439–441 salina, 439 Nephrocytium, 262, 291 agardhianum, 292 Nephrodiella, 430, 438, 447 brevis, 447 nana, 447 phaseolus, 448 Nephroselmis, 228, 230, 247 olivacea, 229–230 Netrium, 354, 363, 367, 370, 379 digitus, 358 minus, 370 oblongum, 358 Nitella, 17, 27, 317, 332, 339, 348, 811, 820, 821 flexilis, 331, 339 hookeri, 27 Nitellopsis, 27–28, 338 obtusa, 28 Nitzschia, 22, 24, 39, 43, 669, 672, 675–676, 681 acicularis, 672, 677 amphibia, 671, 677 angustata, 677 communis, 43 denticula, 677 filiformis, 673 frustulum, 43 holsatica, 672 inconspicua, 670 monoensis, 43 palea, 43–44, 673, 677 scalaris, 670 sinuata var. tabellaria, 677 Noctiluca, 388 Nodularia, 64, 66, 120, 125, 177–178, 807 baltica, 178 harveyana, 67, 177–178 litorea, 178 sphaerocarpa, 177 spumigena, 43, 177–178 spumigena var. minor, 178 willei, 177–178 Nostoc, 11, 26–27, 39–40, 45, 63, 68, 119–120, 125, 158, 177, 179–180 aureum, 180 commune, 179–180 edaphicum, 179 linckia, 180 minutum, 180 paludosum, 179–180

parmelioides, 36, 180, 764, 770 pruniforme, 180 sphaericum, 180 sphaeroides, 180 verrucosum, 33, 764, 770 Nostocaceae, 118, 123, 169 Nostocales, 117–120, 123, 155, 158, 161, 164, 169 Nostochopsaceae, 125, 187 Nostochopsis, 125, 187–188 lobata, 188–189 Nostocoideae, 120 Notosolenus, 407 Novarino, 743 Nupela, 637, 641, 643, 649 Nuphar, 40, 427, 459, 461 Nymphaea, 427, 459, 461 odorata, 811

Ochromonas, 2, 429, 472–473, 485, 487, 493, 495–496, 500–502, 523–524 monicis, 490 sphaerocystis, 476, 480 tuberculata, 485 vulcania, 489 Octacanthium, 376, 379 octocorne, 360, 376 Octogoniella, 261, 291 sphagnicola, 293 Odontidium, 612 Oedocladium, 319, 341 hazenii, 330 Oedogoniales, 311–313, 315, 317, 319, 341, 348 Oedogonium, 24, 35, 39, 40, 44, 314, 316, 318, 322, 340, 441, 458, 497, 696, 708, 709, 808, 810, 822–823 croasdaleae, 330 Oestrupia, 639 zachariasii, 644, 650 Oligochaetophora, 322, 336 simplex, 324 Oncobyrsa, 96 Onychonema, 366, 376 filiforme, 360 Oocardium, 354, 364, 366, 376 stratum, 356 Oocystidium, 264, 291 ovale, 293 Oocystis, 43, 255, 264, 291, 301, 822 lacustris, 293 Oodesmus oederleinii, 481 Oodiniaceae, 697, 706 Oodinium, 690, 691, 697–698, 701, 703 limneticum, 688, 702, 706 Oophilia, 260, 291 amblystomalis, 293 Opalozoa, 749 Opephora, 596, 613, 615 ansata, 616 martyi, 616 Ophiocytiaceae, 431, 456, 459

Taxonomic Index Ophiocytium, 429, 431–433, 436, 457, 459 arbusculum, 457 capitatum, 457 longipes, 457 majus, 459 mucronatum, 457 parvulum, 457 Ophrydium, 45 versatile, 662 Oronectes immunis, 819 Orthoseira, 563, 565, 571, 584 dendroteres, 576 Orthoseiraceae, 563 Orthoseriales, 563 Oscillatoria, 20, 23–24, 27, 39–41, 43–45, 65, 123, 126–127, 132, 141, 155–156, 807, 809, 813, 823 amphibia, 126 amphigranulata, 129 anguina, 155 claricentrosa, 127 curviceps, 155 depauperata, 155 earlei, 127 filiforme, 126 funiformis, 155 jenensis, 156 lacustris, 147 limnetica, 129 limosa, 155–156 margaritifera, 155 obtusa, 156 ornata, 155 princeps, 155–156 proboscidea, 155 redekei, 129 refringens, 156 rhamphoidea, 155 rubescens/agardhii, 806 sancta, 155–156 splendida, 126 tenuis, 808 Oscillatoria (Planktothrix) rubescens, 806 Oscillatoriaceae, 118, 120, 122, 150–151 Oscillatoriales, 117–119, 121, 126, 129, 132, 135, 137, 139, 147, 151 Ourococcus, 288 Oxyneis, 596, 601, 606, 616–617, 621 binalis, 621 binalis var. elliptica, 621

Pachycladella, 261, 292 umbrinus, 293 Pachycladon, 292 Palmella, 260, 292, 295 miniata var. aequalis, 294 Palmellochaete, 261, 295 tenerrima, 294 Palmellococcus, 271 Palmellocystis, 301 Palmellopsis, 260, 295 gelatinosa, 294

Palmodactylon, 295 Palmodictyon, 255, 264, 295 varium, 294 Pandorina, 226, 239, 241, 248, 822 charkowiensis, 241 colemaniae, 241 morum, 226–227, 240–241 unicocca, 241–242 Pannus spumosus, 65 Parabasalia, 387 Paracoenia, 41 Paradoxia, 261, 295 multiseta, 294 Paralemanea, 203, 205, 208, 216–217 annulata, 199–200 catenata, 199 mexicana, 199, 217 Paralia, 583 Paraliaceae, 563 Paraliales, 563 Paramecium, 686, 690 bursaria, 258 Paraphysomonadaceae 474, 513, 523 Paraphysomonas, 474, 486, 488, 491, 500, 523 butcheri, 485 campanulata, 486 foraminifera, 486 imperforata, 485–486 sediculosa, 486 sideriophora, 486 sigillifera, 486 vestita, 477, 486–487, 491 Parietochloris, 256 Pascheriella, 247 Pascherina, 245, 248 tetras, 246, 247 Paulschulzia, 263, 295 pseudovolvox, 296 Pavlova, 512, 515 Pavlovales, 511, 513 Pectodictyon, 264, 295 cubicum, 296 Pediastrum, 43, 254–255, 257–258, 263, 295, 301 duplex var. typicum, 296 Pedimellophyceae, 471 Pedinomonas, 228, 230, 247 maior, 230 minor, 229–230 noctilucae, 388 rotunda, 230 Pedinopera, 236, 238, 248 granulosa, 237–238 rugulosa, 238 Pedinophyceae, 225, 229, 231 Pelagodictyon, 563, 584 Pelagophyceae, 471 Peniaceae, 373 Penium, 361–362, 367–368, 373 didymocarpum, 373 exiguum, 357 phymatosporum, 373 silvae-nigrae, 373

911

spinospermum, 373 spirostriolatum, 357 Peranema trichophorum, 403 Percolozoa, 387 Peridiniaceae, 697, 707 Peridiniales, 697, 707 Peridiniopsis, 697–698, 704, 707 berolinensie, 701 cunningtonii, 694, 696, 699 lubieniensiforme, 694 penardiforme, 692 penardii, 694, 699 polonicum, 691–692, 701 quadridens, 689, 691–692 thompsonii, 702 Peridinium, 697–698, 704, 709 balticum, 707 bipes, 694, 696, 699 cinctum, 690, 694 cinctum f. tuberosum, 691, 707 dybowski, 707 gatunense, 690–692, 694, 699–702 inconspicuum, 23, 686, 694, 699, 707 limbatum, 694, 699–702, 707 lomnickii, 699 pusillum, 699 volzii, 694 willei, 691–692, 694, 699, 707 wisconsinense, 702 Perone, 430, 436, 451 dimorpha, 450 Peronia, 656, 663, 665 fibula, 657 Peroniella, 431, 434, 436, 458 hyalothecae, 456, 458 planctonica, 458 Petalomonas, 387 Petalonema, 124, 160–161 alatum, 160–161 byssoidea, 161 involvens, 160 Pfiesteria piscicida, 685, 739, 814 Phacotaceae, 225, 228, 236–237, 239 Phacotus, 236, 238, 248 angustus, 238 glaber, 238 lenticularis, 237–238 subglobosus, 238 Phacus, 388, 395, 399–402, 404, 406–407, 410, 413, 416 agilis, 395 chloroplastes, 413 chloroplastes f. incisa, 413 curvicauda f. anomalus, 395 elegans, 395 longicauda, 395 longicauda var. insecta, 395 longicauda var. tortus, 395 mariana, 395 monilata, 395 orbicularis var. longicauda, 395 pleuronectes, 395 polytrophos, 395

912

Taxonomic Index

Phacus (continued) trimarginatus, 395 triqueter, 395 Phaeaster, 500 Phaeobotrys, 503 Phaeocystales, 511 Phaeocystis, 512 Phaeodermatium, 488, 494, 500 rivulare, 481, 500 Phaeogloea, 491, 495, 500, 503 mucosa, 483, 500 Phaeophyceae, 8, 757, 765 Phaeophyta, 12, 19, 24, 758, 766 Phaeoplaca, 473, 494, 497, 500 thallosa, 481 Phaeoschizochlamys, 473, 495, 500, 503 mucosa, 484 Phaeosphaera, 491, 495, 500 gelatinosa, 481 Phaeothamnion, 473, 494, 500–503 confervicola, 481, 502 Phaeothamniophyceae, 443, 471, 473, 491, 503 Phalansterium, 499 Phillipsiella, 523 Phormidium, 26, 34–35, 41, 63, 65–66, 123, 132, 135, 141–142, 807, 809, 819 autumnale, 141–142 favosum, 141 fonticolum, 141 formosum, 142 geysericola, 128 inundatum, 141 minnesotense, 141 retzii, 141 richardsii, 142 uncinatum, 141 Phormidiaceae, 120, 122, 132, 139, 147, 150–151 Phormidioideae, 139 Phragmites, 39 Phycopeltis, 315, 319, 343 arundinacea, 328 Phyllobium, 260, 295 sphagnicola, 296 Phyllogloea, 259, 295 fimbriatum, 296 Phyllosiphon, 315, 319, 328, 344 arisari, 328 Phymatodocis, 364, 366, 377 alternans, 377 nordstedtiana, 358, 377 Physolinum, 344 Physomonas, 500 Phytodinales, 696–697, 708 Phytodiniaceae, 697, 708 Phytodinium, 697–698, 704, 709 simplex, 688, 705 Pinnularia, 24, 39, 41, 638–639, 646, 649–650 braunii, 42 mesolepta, 644 microstauron, 42 viridis, 40, 644

Piscinoodinium pillulare, 706 Pithophora, 321, 334, 340, 808–810, 815, 818, 820, 823 oedogonia, 327 Placoma, 96 Placoneis, 650 abiskoensis, 641 Plagioselmis, 731–732, 744, 749 nanoplanctica, 730–732, 748 ovalis, 732 prolonga, 748 Plagioselmis (as Rhodomonas) minuta, 43 Plagiotropis, 638–639, 641, 650 Planktolyngbya, 122, 133, 135 bipunctata, 135 capillaris, 135 contorta, 133, 135, 155 limnetica, 133, 135 regularis, 135 tallingii, 133, 135 Planktosphaeria, 265, 270, 295 gelatinosa, 297 Planktothrix, 37, 65, 120, 123, 141, 143, 147, 155 agardhii, 141, 143 cryptovaginata, 141 mougeotii, 141, 143 prolifica, 141 rubescens, 141, 143 Planktothrix (Oscillatoria) rubescens, 23 Planorbis contortus, 26 Planothidium, 597–598, 602, 607, 623, 627 apiculatum, 629 frequentissimum, 629 lanceolatum, 627, 629 oestrupii, 629 peragalli, 629 pseudotanense, 629 Plantae, 384 Platydorina, 227, 239, 241, 248 caudata, 240–241 Platymonas, 231 subcordiformis, 231 Plectonema, 20, 45, 119, 123, 144, 155, 819 batrachospermi, 144, 146 edaphicum, 146 flexuosum, 146 murale, 146 tenue, 144 tomasinianum, 152, 155 wollei, 155 Pleodorina, 239, 241, 248 californica, 240–241 indica, 241 japonica, 241 Pleurastromsarcina, 274 Pleurastrophyceae, 313 Pleurastrum, 313, 322, 335–336 insigne, 323 Pleurocapsales, 60

Pleurocapsa, 62, 72, 107–108, 110 crepidinum, 108 epiphytica, 108 fluviatilis, 104 minor, 44, 108, 110 minuta, 108 varia, 110 Pleuroceridae, 210 Pleurochloridaceae, 430, 443, 445, 448, 450 Pleurochloridella, 430, 435, 443, 473 vacuolata, 444 Pleurochloridellaceae, 430 Pleurochloris, 430, 437, 446–447 commutata, 44, 425, 447 magna, 425, 447 polyphem, 425, 447 pyrenoidosa, 448 Pleurochrysis, 484, 515 carterae, 43 carterae var. dentata, 512, 514–515, 518 Pleurocladia, 757–758, 760, 767–768, 771 lacustris, 759, 761, 763–765, 766, 769–770 Pleurococcus, 267, 276, 333 Pleurodiscus, 363, 366, 370 borniquinae, 355, 370 Pleurogaster, 430, 438, 447 lunaris, 448 Pleurosigma, 638–639, 650 elongatum, 641 Pleurosira, 563, 571, 584 laevis, 577, 584, 586 Pleurotaenium, 361–362, 366–367, 375, 377 coronatum, 358 Pliocaenicus, 563, 566, 572, 581, 584 Polyblepharaceae, 225, 228, 229, 231–232 Polyblepharides, 228, 230, 247 fragariiformis, 230 singularis, 229–230 Polychaetophora, 321–322, 336 lamellosa, 324 Polycystis, 92 firma, 92 incerta, 78 marginata, 92 montana f. minor, 82 pulverea, 92 Polyedriella, 430, 433, 439, 447–448 aculeata, 448 helvetica, 427, 447–448 Polyedriopsis, 262, 298 spinulosa, 297 Polygoniochloris, 430, 431, 433, 437, 448–449 regularis, 448 tetragona, 448 Polykyrtos, 430, 435, 441 vitreus, 440 Polylepidomonas vaculata, 489 vestita, 489 Polysiphonia, 202, 208, 221 subtilissima, 199, 220–221 Polytaenia, 371 trabeculata, 355

Taxonomic Index Polytoma, 226, 232, 235, 248 granuliferum, 235 uvella, 233, 235 Polytomella, 228, 230, 248 agilis, 230 citrii, 229–230 Pontederia cordata, 821 Pontosphaera stagnicola, 516 Porochloris, 261, 298 filamentarum, 297 Porphyridiales, 198, 208–209 Porphyridium, 3, 200, 206–208 aerugineum, 200 purpureum, 198, 209 sordidum, 198 Porphyrosiphon, 123, 144–145 fuscus, 144–145 notarisii, 144–145 robustus, 145 versicolor, 144–145 Porterinema, 758, 760, 767–768, 770 fluviatile, 759, 763, 765–766, 769, 771 marina, 771 Potamogeton, 17, 315, 335, 459, 461, 819 nodosus, 821 pectinatus, 28, 810, 812 Poterioochromonas, 23, 473, 492, 501 malhamensis, 477 Prasinophyceae, 225, 229, 231–232, 313, 384 Prasiola, 11, 34, 45, 317, 342 fluviatilis, 35 mexicana, 33, 329, 342 Prasiolales, 312, 317, 342 Printzina, 319, 343 ampla, 343 Prismatella, 430, 439, 449 hexagona, 448–449 Proales werneckii, 463 Prorocentrum, 697, 709 Proteomonas, 734, 736, 741, 743 Protista, 383–385, 473, 511–512, 685–686, 697, 715 Protococcus, 267, 276 Protococcus-Pleurococcus, 315, 322 Protoctista, 385 Protoderma, 260, 298, 314, 320, 332, 336 involvens, 319 sarcinodeum, 297 viridae, 323 Protoeuglena, 388 Protosiphon, 256, 258, 260, 298, 322, 344 botryoides, 297, 315, 328 Protozoa, 24, 44, 171, 260, 272, 383–385, 391, 404, 686, 737, 749 Prymnesiales, 511, 513 Prymnesiophyceae, 471, 511, 516, 690 Prymnesium, 511, 515 parvum, 512, 514–515 saltans, 514 Psammodictyon, 672 Psammothidium, 597–598, 602, 607, 623, 627 altaicum, 630

lauenburgianum, 630 levanderi, 630 rosenstockii, 630 subatomoides, 630 Pseudanabaena, 65, 122, 129–130, 139, 155, 807 catenata, 44, 129 galeata, 130 limnetica, 129–130 lonchoides, 130 mucicola, 129 thermalis, 129 Pseudanabaenaceae, 118, 120–121, 126, 129, 132 Pseudanabaenoideae, 126 Pseudendoclonium, 320, 336 basiliense, 323 basiliense var. brandii, 337 prostratum, 336 submarinum, 337 Pseudoactiniscus, 697–698, 705 apentasterias, 688, 702, 706 Pseudobodanella, 758 peterfii, 759, 771 Pseudocharaciopsidaceae, 424 Pseudobohlinia, 261, 298 americana, 297 Pseudobumilleriopsis, 459 Pseudochaete, 321, 337 gracilis, 324, 337 Pseudocharaciopsis, 424–425, 441 minuta, 426–427 ovalis, 425, 427, 446–447 texensis, 427 Pseudocharacium, 270, 441 Pseudodendromonas, 498 Pseudokephyrion, 486–487, 492, 498–499, 501 alaskanum, 478 auroreum, 475 Pseudophormidium, 63, 119, 123, 144, 146 batrachospermi, 144 flexuosum, 146 Pseudopolyedriopsis, 430, 438, 449 skujae, 448 Pseudoschizomeris, 315, 317, 346 caudata, 326 Pseudostaurastrum, 424–425, 430, 439, 447–449, 465 enorme, 448, 465 hastatum, 448, 465 limneticum, 426–427, 449, 465 Pseudostaurosira, 596, 599, 606, 608, 610, 613, 616 brevistriata var. elliptica, 618 brevistriata var. inflata, 618 robusta, 618 Pseudotetraedron, 430–431, 439, 447, 449 neglectum, 450 Pseudulvella, 320, 337 americana, 323, 337 Pteromonas, 236, 238 aculeata, 237–238 angulosa, 238 cordiformis, 238

cruciata, 238 sinuosa, 238 Pulchrasphaera, 289 Punctastriata, 596, 599, 606, 613, 616, 618 Pyramimonas, 735 parkeii, 749 Pyrenomonadaceae, 722, 743, 746–747 Pyrenomonadales, 737, 743, 746–748 Pyrenomonas, 2, 742–744, 746, 749 ovalis, 722, 724, 746 Pyrobotrys, 226–227, 245, 247, 248 casinoensis, 246–247 stellata, 247 Pyrrhophyta, 7, 12, 124, 687, 697

Quadricoccus, 263, 298 verrucosus, 299 Quadrigula, 262, 298 closteroides, 299

Raciborskia, 708 bicornis, 688 Radaisia, 72, 108, 110 confluens, 110 epiphytica, 108 gardneri, 108, 110 willei, 110 Radiococcus, 264, 298 nimbatus, 299 Radiocystis, 69, 73, 81 elongata, 73, 81 fernandoi, 81 geminata, 73, 81 Radiofilum, 318, 346 conjunctivum, 326 Radiosphaera, 267 Rana, 45 clamitans, 412 pipiens, 412 Raphidiastrum, 378 Raphidiella, 431, 437, 453 fascicularis, 452 Raphidiopsis, 124, 173, 180 curvata, 173, 180 mediterranea, 174, 180 Raphidium, 280 Raphidomonadales, 427 Raphidonema, 315, 318, 346 nivale, 44, 326 Raphidonemopsis, 318, 346 sessilis, 326 Raphidophyceae, 40, 423, 427, 465 Rayssiella, 263, 298 hemisphaerica, 299 Reimeria, 656, 659, 663–664 sinuata, 658, 662 sinuata f. antiqua, 658 Rhabdoderma, 65–66, 69, 76, 81 compositum, 76, 81 curtum, 81 lineare, 76, 81 zygnemicolum, 76

913

914

Taxonomic Index

Rhabdogloea, 69, 76, 81 hungarica, 81 smithii, 76, 81 subtropica, 81 Rhinomonas, 743 Rhizochloridaceae, 430 Rhizochloridales, 430, 432, 434, 441–442 Rhizochloris, 430, 435, 441 mirabilis, 441–442 stigmatica, 441 Rhizochrysis, 494, 496, 501 scherffelii, 482 Rhizoclonium, 208, 318, 321–322, 334, 340, 763, 771, 808–809 hieroglyphicum, 314, 327, 340 Rhizolekane, 430, 435, 441 sessilis, 441–442 Rhizoochromonas, 485, 493, 501, 503 endoloricata, 480, 501 Rhizophydium fugax, 739 sphaerocarpum, 364 Rhizosoleniaceae, 563, 567 Rhizosoleniales, 563 Rhizosoleniophycidae, 563 Rhodochytrium, 260, 298 spilanthidis, 300 Rhododendron, 343 Rhododraparnaldia, 201–202, 207, 212 oregonica, 198, 201, 212–213 Rhodomonas, 736, 739, 741–743, 746, 749 lacustris, 738 lens, 738 minuta, 748 ovalis, 725 Rhodophyta, 6, 12, 19, 34–35, 41, 197–198, 200, 202–203, 204–205, 735 Rhoicosphenia, 597, 604, 656, 661, 663 abbreviata, 659, 666 curvata, 34, 659 Rhoicospheniaceae, 597, 655–656, 659, 661, 665–666 Rhomboidella, 430, 439, 449 oblique, 450 Rhopalodia, 43, 669, 670, 672–675, 680–681 gibba, 678 Rhopalodiacaeae, 670, 676 Rhopalodiales, 669–672, 674–675, 681 Richteriella, 288 Rigida, 412 Rivularia, 65, 118–119, 124, 166, 168–169, 764 aquatica, 168 biasolettiana, 169 compacta, 169 dura, 168, 169 globiceps, 169 haematites, 169 minutula, 169 planctonica, 166 Rivulariaceae, 118, 124, 164 Romeria, 121, 129–130 alascense, 129 elegans, 129 elegans var. nivicola, 129

heterocellularis, 129 hieroglyphica, 129 leopoliensis, 129–130 mexicana, 129 nivicola, 130 Rosenvingiella, 342 Rossithidium, 597–598, 602, 607, 625, 628 duthiei, 630 linearis, 609, 627, 630 pusillium, 630 Rotifera, 383 Roya, 367, 370 Rufusia, 45, 199, 207, 210 pilicola, 198, 201 Rufusiella, 697–698, 701, 703, 709 insignis, 688, 702

Saccochrysis, 493, 501 piriformis, 481, 501 Sacconema, 124, 167, 169 rupestris, 167, 169 Sagittaria, 322, 332 Salicornia, 333 Sarcomastigophora, 383 Scenedesmus, 44, 254, 256–258, 262, 298, 301, 453, 768 acutus, 300 armatus, 255, 276, 299 f. quadricauda, 256 quadricauda, 44 trainorii, 256 Scherffelia, 228, 230, 248 phacus, 229–230 Schizochlamydella, 259, 263, 301 gelatinosa, 300 Schizochlamys, 301 Schizodictyon, 284 Schizomeris, 317, 337 leibleinii, 329 Schizothrix, 34, 39, 44, 63, 65–66, 119, 122, 136–137, 148, 150, 763 acuminata, 150 aikenensis, 137 californica, 150 chalybea, 150 constricta, 136–137, 150 friesii, 150 giuseppei, 150 hancockii, 150 mexicana, 150 muelleri, 150 parciramosa, 148, 150 penicillatum, 150 purpurascens, 148, 150 richardsii, 150 rivularis, 150 sauterianum, 150 stricklandii, 150 taylori, 150 telephoroides, 150 thermophila, 135 violacea, 136 wollei, 148

Schizotrichaceae, 121, 137, 150 Schmidleinema, 125, 184–185 cubanum, 184–185 indicum, 184 roberti-lamyi, 184 Schroederia, 261, 301 setigera, 300 Scirpus, 17, 439, 458–459, 461 Scoliopleura, 638–639 peisonis, 641, 650 Scotiella, 266, 301 cryophila, 44 turberculata, 300 Scotiellopsis, 266, 301 rubescens, 300 Scourfieldia, 228, 230, 247 cordiformis, 229–230 Scytonema, 3, 39, 66, 119, 123, 155, 157–158, 161 arcangelii, 158 capitatum, 157 cincinnatum, 158 crispum, 158 crustaceum, 158 dubium, 158 endolithicum, 67 longiarticulatum, 157 myochrous, 158 ocellatum, 158 tolypothrichoides, 158 Scytonemataceae, 123, 155 Scytonematopsis, 118, 123, 157–158 fuliginosa, 157–158 hydnoides, 157–158 Secale, 822 Selenastrum, 262, 301, 785 capricornatum, 302 Selenophaea, 491, 495, 501, 503 granulosa, 482, 501 Sellaphora, 637, 640, 650 americana, 643 pupula, 643 Semiorbis, 656, 662 hemicyclus, 657, 665 Setacea (section of Batrachospermum), 215 Siderocelis, 265, 301 minutissimus, 302 Simulium, 35 Siphonales, 312, 319, 321, 344 Sirodotia, 200, 205, 208, 215 delicatula, 203 huillensis, 199, 216 suecica, 199, 214, 215 Sirogonium, 354, 363, 366, 370 illinoiense, 355 Skeletonema, 563, 565, 569–571, 585 potomos, 582 Skeletonemataceae, 563 Sklerochlamys, 430, 438, 449 pachyderma, 450 Smithsonimonas, 227 Snowella, 64–65, 70, 85, 88 fennica, 85, 88 lacustris, 88 litoralis, 85, 88

Taxonomic Index Snowella (continued) rosea, 67 septentrionalis, 88 Sorastrum, 261, 301 spinulosum, 302 Sphacelariales, 758–759, 771 Sphaeropleales, 312, 317, 342 Spermatozopsis, 228, 230–231, 247 exsultans, 229, 231 Sphacelaria, 758, 760, 767–768 fluviatilis, 759, 763, 765, 771 lacustris, 759, 765, 767–769, 771 Sphaerellocystis, 265, 301 aplanospora, 301 ellipsoidea, 302 Sphaerellopsis, 236 Sphaeridiothrix, 473, 494, 501, 503 compressa, 483 Sphaerocystis, 43, 258, 265, 301 schroeteri, 302 Sphaerodinium, 697–698, 705 fimbriatum, 691, 693–694, 702, 708 polonicum, 708 Sphaeroplea, 318, 342 annulina, 328 Sphaerosorus, 431, 437, 453 coelastroides, 452 Sphaerozosma, 366, 376–377, 458 vertebratum var. latus, 358 Sphagnum, 40, 291, 295, 298, 363, 370–371, 373, 375, 378, 427–428, 432, 439, 441–443, 451, 458, 461, 463, 490, 662, 709 Sphinoclosterium, 364–365 Spiniferites, 698 Spiniferomonas, 473, 485–488, 491, 493, 497, 501, 523 abei, 475, 486, 489 bilacunosa, 487, 489 bourrellyi, 474, 480, 487–489 cornuta, 475, 489 coronacircumspina, 489, 491 minuta, 491 serrata, 488, 491 silverensis, 475, 486 takahashii, 491 trioralis, 486–489 Spinoclosterium, 367, 372 cuspidatum, 356, 372–373 Spinocosmarium, 361, 368, 377 laconiense, 377 quadridens, 359, 365, 377 Spirirestis, 191 rafaelensis, 191 Spirogyra, 24, 39, 314, 340, 353–354, 362–365, 370–372, 708, 808–810, 818, 820, 822–823 floridana, 371 juergensii, 355 majuscula, 364 wrightiana, 355 Spirotaenia, 280, 354, 363, 367, 371 condensata, 357 densata, 371 Spirulina, 122, 131–132

caldaria var. magnifica, 131 gigantea, 132 labyrinthiformis, 132 laxa, 132 major, 131–132 meneghiniana, 131 nordstedtii, 132 platensis, 139 princeps, 132 stagnicola, 131–132 subalsa, 131 subtilissima, 132 weissii, 131–132 Spirulinoideae, 129 Splendidae, 533 Spondylomoraceae, 225, 228, 246, 248 Spondylomorum, 245, 248 quaternarium, 246–247 Spondylosium, 366, 377 pulchellum, 377 pulchrum, 358 rectangulare, 358 Spongilla, 272 lacustris, 45, 258 Spongiochloris, 256, 266, 302 spongiosa, 302 Spongiococcum, 291, 303 alabamense, 302 Spongomonas, 499 Spumella, 491, 502 sociabilis, 477 Stanieria, 66, 71, 102–104 cyanosphaera, 67, 103–104 sphaerica, 104 Staurastrum, 354, 361, 363–364, 368–369, 377–379, 447, 456, 458 anatinum f. curtum, 360 bioculatum, 360 claviferum, 360 dejectum, 23 longipes, 364 tribedrale, 360 turgescens, 360 Staurodesmus, 363, 368–368, 376, 378–379 crassus, 364 cuspidatus, 360, 364 extensus var. joshuae, 364 sellatus, 364 subtriangularis, 360 triangularis var. limneticus, 364 Stauroforma, 599, 606 exiguiformis, 616, 619 Stauromatonema, 125, 181 viride, 181–182 Stauroneis, 24, 40, 638–640, 650 acuta, 642 anceps, 27 kriegeri, 642 phoenicenteron, 642 Staurosira, 596, 599, 606, 610, 616 construens var. pumila, 619 construens var. venter, 619 Staurosirella, 596, 599, 606, 608, 610, 613, 615–616 ansata, 620

915

leptostauron, 616, 620 pinnata, 620 spinosa, 620 Stenopterobia, 670, 676 sigmatella, 672, 679 Stephanocostis, 563, 572 chantaicus, 582, 585, 588 Stephanocyclus, 563, 572, 585 caspia, 585 cryptica, 579 gamma, 579 meneghiniana, 37, 569, 579, 585 quillensis, 585 striata, 579 Stephanocyclus-(Cyclotella) meneghiniana, 22 Stephanodiscaceae, 563, 570 Stephanodiscus, 20–22, 37, 563–565, 567–568, 570, 572–573, 578–579, 583–586, 807 binderanus, 565, 580 binderanus var. oestrupi, 580 excentricus, 585–586 hantzschii, 38, 565, 580 hantzschii f. tenuis, 580 lucens, 586 niagarae, 566, 569, 580 reimerii, 569 rhombus, 585–586 superiorensis, 569 yellowstonensis, 569 Stephanoporos, 492, 502 sphagnicola, 478 Stephanosphaera, 231–232, 248 pluvialis, 231–232 Stichochrysis, 493, 502 immobilis, 481, 502 Stichococcus, 44, 318, 345–346 bacillaris, 42 subtilis, 326 Stichogloea, 473, 491, 495, 502–503 doederleinii, 481 olivacea, 487 Stichosiphon, 62, 65, 70, 99, 102 exiguus, 99, 102 filamentosus, 102 gardneri, 102 regularis, 102 sansibaricus, 99, 102 willei, 99, 102 Stigeoclonium, 39, 44, 314, 320–321, 334, 337, 808, 810 farctum, 338 lubricum, 325 tenue, 44 Stigonema, 35, 40, 119, 125, 181, 183–184 congestum, 183 elegans, 183 hormoides, 184 informe, 181, 184 mamillosum, 183–184 mesentericum, 184 minutum, 181, 184 minutum var. saxicola, 184 mirabile, 184

916

Taxonomic Index

Stigonema (continued) ocellatum, 184 panniforme, 184 thermale, 184 turfaceum, 184 Stigonematales, 117–120, 123, 181, 184, 187, 189 Stipitococcaceae, 430 Stipitococcus, 430, 433, 435, 441–442 apiculata, 442 crassistipitatus, 442 vas, 442 vasiformis, 442 Stipitoporos, 430, 435 polychloris, 442 Stomatochroon, 319, 343 lagerheimii, 328 Storeatula, 734, 736, 742–744, 749 rhinosa, 722, 724–725, 746 Streptophyta, 229 Stromatochroon, 315 Strombomonas, 2, 388, 403, 410, 413–414, 416 acuminata, 400 conspersa, 414 costata, 414 deflandrei, 400 fluviatilis, 400 giardinana, 400 gibberosa, 400 lackeyi, 400 longicauda, 400 ovalis, 400, 414 rotunda, 400 schauinslandii, 400 taiwanensis var. bigeonii, 413–414 tambowika, 400 urceolata, 397, 400 verricosa, 400 verricosa var. zmiewika, 400 volgensis, 400 Stylobryon, 491, 502 abbotti, 472, 477, 502 Stylochrysalis, 492, 502 aurea, 478 parasitica, 502 Stylococcus, 502 Stylodinium, 690–691, 696–698, 701, 703, 709 globosum, 688, 705 longipes, 702 Stylonema, 208 Stylosphaeridium, 260, 303 stipitatum, 304 Surirella, 39, 670, 672, 680 angustata, 678 linearis var. constricta, 678 ovata, 678 tenera, 678 Surirellaceae, 670, 681–682 Surirellales, 669–672, 675, 681–682 Symbiodinium, 686–687 Symploca, 119, 123, 142, 144 borealis, 144 cartilaginea, 144

cavernarum, 144 ciliata, 144 dubia, 144 hydnoides, 142 kieneri, 144 muralis, 144 muscorum, 142, 144 nemecii, 144 thermalis, 144 Symplocastrum, 119, 123, 137, 148, 150 Syncrypta, 493 volvox, 479 Synechococcaceae, 62, 68 Synechococcus, 20, 22, 24, 41, 63, 65–66, 69, 73, 76–77, 79, 81–82 aeruginosus, 77, 80 bigranulatus, 82 cedrorum, 79 koidzumii, 82 lividus, 66, 76, 82 maior, 77 minervae, 79 nidulans, 76, 81, 88 sigmoideus, 82 vulcanus, 66, 82 Synechocystis, 20, 65, 69, 78, 88–89 aquatilis, 78, 88 fuscopigmentosa, 88 minuscula, 88 primigenia, 89 salina, 88 thermalis, 78, 88 willei, 78, 89 Synedra, 22, 595–596, 600, 606–608, 613, 616, 620, 631 ostenfeldii, 621 parasitica, 620, 672 radians, 22 rumpens var. fusa, 621 ulna, 25, 36, 621 ulna var. aequalis, 621 ulna var. danica, 621 Synedra-Fragilaria, 25 Synedrella, 631 Synochromonas, 502 Synura, 7, 471, 502, 523–526, 528–531, 533–536, 538, 541, 546–548, 551–552 australiensis, 551 curtispina, 538, 546, 551 echinulata, 528, 529, 536, 538, 546, 551 echinulata f. leptorrhabda, 551 echinulata f. leptorrhabda, 546 lapponica, 524, 530, 533–534, 538, 541, 546, 548, 551 mammillosa, 551 mollispina, 546, 551 petersenii, 525, 527–528, 534–535, 547, 551 petersenii var. praefracta, 529 sphagnicola, 528, 530, 534, 536, 538, 541, 547, 551 spinosa, 524–525, 528, 537, 547, 551 spinosa f. longispina, 538

splendida, 534 uvella, 22, 528, 535, 547, 551 Synuraceae, 523 Synurophyceae, 7, 471, 474, 494, 513, 523–526, 528–530, 533–537, 541, 552 Synuropsis, 493, 502–503 gracilis, 479

Tabellaria, 3, 22, 24, 39, 595–597, 601, 606, 608, 621, 623 binalis, 617 fenestrata, 20, 622 flocculosa, 21, 621, 622 quadriseptata, 622 Tabellariaceae, 596, 598, 616 Tabellariales, 598, 616 Teilingia, 366, 378 granulata, 359 Teleaulax, 734, 742–743 Termemorus laevis, 357 Terpsinoë, 563, 571 musica, 575, 586 Tessellaria, 524, 526, 530, 533, 541, 548, 551, 552 volvocina, 527, 534, 548 Tetmemorus, 367–368, 378 Tetrabaena, 244, 248 socialis, 227, 238, 244–245 Tetrabaenaceae, 225, 228, 239, 244–246 Tetracanthium, 368 Tetrachrysis, 473, 494, 496, 500, 502–503 dendroides, 476, 484, 502 Tetracyclus, 596, 601, 606, 617, 623 glans, 622 Tetracystis, 254, 258, 264, 303 texensis, 304 Tetradesmus, 262, 303 wisconsinensis, 304 Tetradinium, 690, 691, 697–698, 703, 709 javanicum, 688, 705 simplex, 702 Tetraedriella, 430, 438, 449–450 enorme, 465 hastatum, 465 limneticum, 465 regularis, 450, 465 spinigera, 450 trigonum, 465 Tetraedron, 254, 262, 303, 465 regulare, 465 victoriae, 304 Tetragoniella gigas, 449 regularis, 449 Tetraktis, 431, 437, 453 aktinastroides, 452 Tetrallantos, 262, 303 lagerheimii, 304 Tetramastix, 825 Tetrapion, 503 Tetraplektron, 430, 433, 439, 449–450 torsum, 450 tribulus, 450

Taxonomic Index Tetraselmis, 228, 230–231, 248 cordiformis, 229, 231 subcordiformis, 231 Tetraspora, 255, 259, 303, 322, 498 gelatinosa, 304 Tetrasporopsis, 473, 482, 494, 498, 503 fuscescens, 482 perforata, 482 Tetrastrum, 261, 303 heterocanthum, 305 Tetratomococcus, 298 Tetreutreptia, 385, 388, 408 pomquetensis, 401 Thalassiocyclus, 563, 572, 586 lucens, 579, 586 Thalassiosira, 563, 565, 567, 571, 575, 584–587 pseudonana, 43 weissflogii, 579 Thalassiosiraceae, 563 Thalassiosirophycidae, 563 Thalpophila, 125 cossyrensis, 189 imperialis, 189–190 Thamniochaete, 320, 337–338 huberi, 324 Thecadiniaceae, 697, 708 Thompsodinium, 697–698, 704, 707 intermedium, 689, 692, 702 Thoracomonas, 236, 238, 248 feldmanii, 238 phacotoides, 237–238 Thorea, 207, 218–219 hispida, 199, 219 violacea, 33, 199, 218–219 Thoreales, 199–200, 203, 216–217, 219 Tilapia, 820 Tolypella, 27, 317, 339, 345 nidifica, 331 Tolypothrix, 26, 34, 119, 124, 161–162, 184 amoena, 162 bouteillei, 161 distorta, 25, 161 lanata, 161 papyracea, 162 penicillata, 161–62 robusta, 162 setchellii, 161 tenuis, 161–162 tenuis f. minor, 161 willei, 162 Tolypotrichoideae, 158 Tomaculum, 264, 306 catenatum, 305 Torquatae (section of Mallomonas), 550 Tortitaenia, 371, 379 obscura, 371 Torytaemia, 367 Trachelomonas, 23, 384, 388, 403–404, 406, 410, 414–416, 806 abrupta, 398 acanthostoma, 398 aculeata f. brevispinosa, 399 armata, 398 armata f. inevoluta, 398

armata var. longispina, 398 armata var. steinii, 398 bulla, 399 charkowiensis, 399 cylindrica, 397–398 dubia, 397–398 dybowskii, 398 erecta, 397 girardiana, 399 grandis, 397, 407 hexangulata, 399 hispida, 398, 406 hispida var. coronata, 398 hispida var. cremulatocollis f. recta, 398 hispida var. papillata, 398 hispida var. punctata, 398 horrida, 398 intermedia, 397 kelloggii, 397 lacustris, 397 lefevrei, 406 mammillosa, 399 playfairii, 399 pulcherrima, 398, 407 pulcherrima var. minor, 398 robusta, 398 rotunda, 397 scabra var. longicollis, 399 similis, 399 speciosa, 399 spectabilis, 399 spirillifera, 415 superba, 398 superba var. duplex, 398 superba var. spinosa, 398 superba var. swirenkiana, 398 sydneyensis, 398 triangularis, 397 volvocina, 397, 407 volvocina var. compressa, 397 volvocina var. punctata, 397 Trachychloron, 430, 438, 449–450 depauperatum, 450 fusiforme, 450 Trachycystis, 430, 438, 447, 451 subsolitaria, 450–451 Trachydiscus, 430, 439, 450–451 ellipsoideus, 450 lenticularis, 450 sexangulatus, 450 Trapa natans, 24 Trebouxia, 254, 258, 266, 306 parmeliae, 305 Trebouxiophyceae, 313 Trentepohlia, 45, 315, 320, 343–344 aurea, 328 Trentepohliales, 312–313, 342–344 Trentepohliophyceae, 309 Treubaria, 261, 306 triappendiculata, 305 Tribonema, 44, 341, 429, 431–432, 433–434, 461–463 aequale, 462 affine, 461

917

regulare, 462 viride, 461–462 Tribonemataceae, 431 Tribonematales, 429, 431, 433–434, 459–460, 462 Tribophyceae, 270, 423–424, 427, 429–430, 432, 441, 447, 465, 473 Triceratiaceae, 563 Trichocoleus, 122, 137–139, 150 acutissimus, 138–139 erectiusculus, 138 minor, 139 purpureus, 138 sociatus, 139 Trichodesmium, 64, 119, 123, 143–144, 147 iwanoffianum, 143 lacustre, 143, 147 Trichodiscus, 338 elegans, 324, 338 Trichophilus, 319, 338 welcheri, 315, 323 Trichoptera, 35 Trichormus, 65, 120, 125, 171, 176, 180 anomalus, 180 doliolum, 180 fertilissimus, 176, 180 luteus, 176 subtropicus, 176 variabilis, 176, 180 Trichosarcina, 315 polymorphum, 345 Triploceras, 362–363, 365, 367, 378 gracile, 357 Triticum, 822 Trochiscia, 262, 306 hystrix, 305 Tryblionella, 670, 672, 675, 677, 680–681 Tuomeya, 203, 205, 208, 216 americana, 199, 204–205, 214 Turfosa (section of Batrachospermum), 215 Tychonema, 122, 146–147 bornetii, 147 bourrellyi, 146–147 Typha, 17, 332, 427–428, 443, 458–459, 461, 764

Ulnaria, 631 Ulothrix, 34, 313, 318, 345, 347, 370, 808, 810 zonata, 25–26, 35, 314, 326 Ulotrichaceae, 348 Ulotrichales, 40, 313, 317, 337, 340, 344–347 Ulva, 253, 312 Ulvales, 311, 313, 317, 336–337, 347 Ulvella, 336 Ulvophyceae, 313, 384 Uroglena, 23, 485, 487–488, 493, 502–503 americana, 489 volvox, 488 Uroglenopsis, 473, 476, 485, 487–488, 491, 493, 499, 502–503 articulatus, 489

918

Taxonomic Index

Uronema, 313, 318, 344, 347 elongatum, 326 Urosolenia, 563, 571–572, 575, 587 Utricularia, 363

Vacuolaria, 427–428 virescens, 426 Vallisneria, 17 Vampyrella, 696, 708 Vaucheria, 4, 11, 322, 341, 423, 429, 431–434, 463–465 aversa, 464 bursata, 433, 464 dillwynii, 433, 464 fontinalis, 433, 464 geminata, 433, 464 Vaucheriaceae, 431 Vaucheriales, 429, 431, 433–434, 463–464 Virescentia (section of Batrachospermum), 215 Vischeria, 424–425, 427 helvetica, 427, 447 punctata, 427 stellata, 426–427 Vitreochlamys, 232, 248 fluviatilis, 233 Volvocaceae, 225–228, 233, 239–240, 242–243, 246 Volvocales, 37, 225–227, 235–236, 245 Volvochrysis, 502 Volvox, 225–227, 239, 241–242, 248, 312 aureus, 240, 242 carteri, 242 carteri f. weismannia, 242 dissipatrix, 242 globator, 242 perglobator, 242 powersii, 242

prolificus, 242 rousseletii, 242 spermatosphaera, 242 tertius, 242 vegetans, 471, 495 weismannia, 242 Volvulina, 239, 242, 248 pringsheimii, 242 steinii, 240–241, 243 Vorticella, 171

Westella, 264, 306 botryoides, 305 Wislouchiella, 236, 238, 248 planctonica, 237–238 Wolfia, 627 Wollea, 125, 180–81 bharadwajae, 181 saccata, 173, 181 Woloszynskia, 691, 697–698, 703, 705, 707 apiculata, 694 cestocoetes, 702 pseudopalustre, 694 reticulata, 689, 695, 699, 702, 704 Woronichinia, 60, 61, 64, 68, 70, 85, 87, 89 elorantae, 89 fremyi, 89 karelica, 89 klingae, 67, 85, 89 naegeliana, 67, 85, 89 Xanthidium, 363, 368, 376, 378–379 controversum, 379 hastiferum, 360 Xanthophyceae, 429, 440, 473 Xanthophyta, 24

Xenococcaceae, 60, 71, 104 Xenococcus, 65, 72, 106–107 angulatus, 107 bicudoi, 106–107 candelariae, 107 chaetomorphae, 107 cladophorae, 107 deformans, 107 gilkeyae, 107 lamellosus, 107 pallidus, 107 pyriformis, 107 schousboei, 107 willei, 106–107 yellowstonensis, 107 Xenotholos, 72, 106–107 huastecanus, 106 kerneri, 106–107

Yamagishiella, 239, 241, 242, 248 unicocca, 240, 242

Zoochlorella, 271 Zygnema, 3, 24, 33, 36, 39, 354, 363, 366, 371–372, 708 conspicuum, 355 frigidum, 355 Zygnemaphyceae, 353 Zygnemaphyta, 353 Zygnematales, 12, 19, 39–40, 311, 313, 353–354, 362–365, 369–372, 379 Zygnematophyceae, 313, 353 Zygnemopsis, 366, 371–372 decussata, 355 Zygogonium, 39, 45, 363, 366, 372, 497 ericetorum, 355

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