Marine Plankton: A practical guide to ecology, methodology, and taxonomy 9780199233267, 0199233268

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MARINE PLANKTON

To all those who love plankton as passionately as we do.

MARINE PLANKTON A Practical Guide to Ecology, Methodology, and Taxonomy

Edited by Claudia Castellani, Plymouth Marine Laboratory, UK and Martin Edwards, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK and University of Plymouth, UK

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3 Great Clarendon Street, Oxford, OX2 6DP, United Kingdom Oxford University Press is a department of the University of Oxford. It furthers the University’s objective of excellence in research, scholarship, and education by publishing worldwide. Oxford is a registered trade mark of Oxford University Press in the UK and in certain other countries © Oxford University Press 2017 The moral rights of the authors have been asserted First Edition published in 2017 Impression: 2 All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without the prior permission in writing of Oxford University Press, or as expressly permitted by law, by licence or under terms agreed with the appropriate reprographics rights organization. Enquiries concerning reproduction outside the scope of the above should be sent to the Rights Department, Oxford University Press, at the address above You must not circulate this work in any other form and you must impose this same condition on any acquirer Published in the United States of America by Oxford University Press 198 Madison Avenue, New York, NY 10016, United States of America British Library Cataloguing in Publication Data Data available Library of Congress Control Number: 2016946644 ISBN 978–0–19–923326–7 DOI: 10.1093/acprof:oso/9780199233267.001.0001 Printed and bound by CPI Group (UK) Ltd, Croydon, CR0 4YY Links to third party websites are provided by Oxford in good faith and for information only. Oxford disclaims any responsibility for the materials contained in any third party website referenced in this work.

CONTENTS ix xi xiii xvii

Foreword Acknowledgements List of Contributors General Introduction

SECTION I  ECOLOGY 1  The Marine Environment N. Penny Holliday and Stephanie Henson 2  Plankton Biodiversity and Biogeography Gregory Beaugrand 3  Phytoplankton Productivity John Raven 4  Zooplankton Productivity Andrew G. Hirst 5  Phytoplankton Biogeochemical Cycles Carol Robinson 6  Zooplankton Biogeochemical Cycles Deborah Steinberg 7  Plankton and Global Change Martin Edwards 8  Plankton and Fisheries Keith Brander

3 12 24 34 42 52 67 81

SECTION II  METHODOLOGY 9  Sampling, Preservation and Counting of Samples I: Phytoplankton Alexandra Kraberg, Katja Metfies, and Rowena Stern 10  Sampling, Preservation and Counting of Samples II: Zooplankton Peter H. Wiebe, Ann Bucklin, and Mark Benfield

91 104

SECTION III  TA XONOM Y Introduction to Taxonomy Rowena Stern, Marianne Wootton, and Claudia Castellani

139

Part 1  Ph y topla nkton Phytoplankton: Diatoms Alexandra Kraberg and Rowena Stern Phytoplankton: Dinoflagellates Alexandra Kraberg and Rowena Stern Phytoplankton: Flagellates Rowena Stern, Heather Esson, and Cecilia Balestreri

151 159 167

vi contents Part 2  Zoopla nkton Protozooplankton: Ciliates 183 Alexandra Kraberg, Rowena Stern, and Michaela Strüder-Kypke Protozooplankton: Radiolaria 189 Rowena Stern, Claire Taylor, Fabrice Not, and Johan Decelle Protozooplankton: Foraminifera 194 Rowena Stern, Claire Taylor, and Saeed Sadri Cnidaria: Scyphozoa and Non-colonial Hydrozoa 198 Priscilla Licandro, Astrid Fischer, and Dhugal J. Lindsay Cnidaria: Colonial Hydrozoa (Siphonophorae) 232 Priscilla Licandro, Claude Carré, and Dhugal J. Lindsay Ctenophora251 Priscilla Licandro and Dhugal J. Lindsay Crustacea: Introduction 264 Claudia Castellani and Marianne Wootton Crustacea: Copepoda 267 Marianne Wootton and Claudia Castellani Crustacea: Branchiopoda 381 Claudia Castellani Crustacea: Cirripedia and Facetotecta 390 Eve C. Southward Crustacea: Ostracoda 410 Martin V. Angel and Anthony W.G. John Crustacea: Decapoda 420 Clare Buckland, Claudia Castellani, Alistair J. Lindley, and Antonina Dos Santos Crustacea: Stomatopoda 465 Claudia Castellani, Clare Buckland, Alistair J. Lindley, David V.P. Conway, and Antonina dos Santos Crustacea: Lophogastrida and Mysida 471 Claudia Castellani, Maiju Lehtiniemi, and Kenneth Meland Crustacea: Amphipoda 490 Tanya Jonas Crustacea: Euphausiacea 505 Alistair J. Lindley Anellida: Holoplanktonic Polychaeta 530 Claudia Castellani and Robert Camp Mollusca: Holoplanktonic Molluscs 538 Silke Lischka and Holger Ossenbrügger Chaetognatha551 Annelies Pierrot-Bults Echinodermata562 David V.P. Conway, Claudia Castellani, and Eve C. Southward Bryozoa568 Judith Fuchs and John Bishop Brachiopoda573 Judith Fuchs and Andreas Altenburger Phoronida577 Judith Fuchs

contents  vii Rotifera581 David V.P. Conway, Robert Camp, and Claudia Castellani Chordata: Thaliacea 584 Priscilla Licandro and Martina Brunetta Chordata: Appendicularia 599 Gabriel Gorsky and Claudia Castellani Chordata: Fish Eggs and Larvae 607 Peter Munk and Jørgen G. Nielsen General Index Species Index

641 667

FOR EWOR D

The ocean pelagic realm is the largest habitat on Earth and this vast three-dimensional space, beneath 71% of the planet’s surface, is home to the plankton. Knowing what goes on in the global plankton community is essential if we are to understand the fundamental role of the pelagic realm in modulating the global environment via its regulatory effects on Earth’s climate and its major contribution to global biogeochemical cycles. With the current focus on global change it is more important than ever to gain some insight into the ecological responses of the plankton to such change. In marine ecosystems, ecological responses to climate warming include biogeographical shifts where species adjust their spatial distributions in response to global warming, phenological shifts where changes in timing of life cycle events have the potential to disrupt trophic pathways, and regime shifts where whole ecosystems undergo abrupt, non-linear change. Finer scale ecological responses also occur at the community level, such as the reduction in body size in ectotherms or increases in species richness. The use of highly sensitive planktonic species as indicators has facilitated ocean monitoring and is especially valuable in managing the marine environment by providing rapid information on issues such as ocean acidification, eutrophication and invasive species. The first section of this book contains informative reviews on plankton-related topics ranging from the marine environment and global change, through to biogeography, productivity and biogeochemical cycles, and the relationship between plankton and fisheries. These authoritative accounts highlight the importance of plankton and help place recent advances in knowledge into their societal and scientific contexts. The second section of this volume is devoted to methodology and comprises two chapters, the first of which outlines sampling, preservation and counting techniques for phytoplankton. Zooplankton is the theme of the other and, after introducing key terminology, this provides a beautifully illustrated critical review the myriad of different sampling systems that have been used for catching or observing plankton over the decades. It also outlines preservation and counting techniques, and both chapters consider the latest advances in genetic sampling methods. The third and by far the largest section is devoted to taxonomy: it serves as a practical guide to the identification and systematics of the most common planktonic organisms found in the North Atlantic and the North Sea. It covers everything from diatoms and dinoflagellates, through the various invertebrate groups, up to the chordates, including fish eggs and larvae. Here in this volume, we benefit from the accumulated knowledge that the contributors captured in the profusely illustrated taxonomic chapters. For each group of organisms, the morphological terminology is introduced and illustrated to ease the non-specialist into the tabular or dichotomous identification keys, and there are thousands of individual line drawings selected to highlight important diagnostic features. In addition, they have devoted considerable effort into providing body lengths for most species, which can often be used as a character to discriminate between similar species. I have no doubt this volume will help to raise identification standards because, in major groups such as the ubiquitous copepods, the only alternative identification guide is very dated—published back in 1933. We are fortunate indeed that the contributing authors have so concisely encapsulated their knowledge and expertise in such a unique and timely volume. Geoffrey A. Boxshall Natural History Museum, London

ACK NOW LEDGEMENTS

First of all this book represents a monumental team effort and its preparation would have not been possible without the contributions, help and continuous support of numerous individuals and organisations around the world. We thank you all. Besides the contributing authors we have many other friends and colleagues to thank who have helped in numerous other ways. For their reviews and comments particularly of the taxonomic section we thank Gerald Boalch, Johan Decelle, and Grazyna Durak for their help with the phytoplankton and protozooplankton chapters, Jean-Claude Braconnot, Aino Hosia, and Peter Schuchert for their advice on the taxonomy of Thaliacea and Hydrozoa; Wolfgang Zeidler and Tammy Horton for their review of the Amphipoda chapter; Karen Osborne for reviewing the holoplanktonic Polychaeta chapter; Masaaki Murano for reviewing the Lophogastrida and Mysida chapter; Maria Grazia Mazzocchi and Iole di Capua for reviewing the Oithona section; Ruth BöttgerSchnac for reviewing the Oncaeidae section; Kate Feller for her help with the revision of the Stomatopoda chapter; and Peter Wiebe for his help with the revision of the Euphausiacea chapter. Particular thanks goes to Antony John and Geoff Boxshall for proofreading most if not all of the chapters in the taxonomy section and for their most insightful comments and suggestions. We would like to thank Martina Brunetta, Astrid Fisher, Kate Brailsford, and Yener Altunbaş for their help with additional proofreading of chapters, the scanning of the line drawings and sourcing of the taxonomic literature and particularly Martina for her meticulous reading of the proofs and the indices. Thank you also to the book project manager, Cheryl Brant, for her effort, support, and patience with what felt like ‘the never-ending task’ of correcting the final proofs of the taxonomic section. We have many colleagues to thank, particularly Russell Hopcroft and Otto Larink, for generously providing us with numerous beautiful images of oceanic and coastal marine plankton. Eric Goberville prepared the maps illustrating the distribution of the copepod species in the North Atlantic. A special thanks to the staff of the National Marine Biological Library (NMBL) at the Marine Biological Association UK (MBA) Sandra Robinson, Barbara Bultman, Emily Miles, and Linda Nobel in particular for their tireless assistance with supplying and sourcing even the most obscure but most helpful taxonomic literature on plankton. The Sir Alister Hardy Foundation for Ocean Science (SAHFOS) and the Royal Society are also duly acknowledged for their part in supporting some of the authors throughout the preparation of this book. We are grateful to all the publishers, institutes and authors for their permission to reproduce in this book previously published images and line drawings. Last but not least we thank the Continuous Plankton Recorder survey taxonomic analysis team for their enthusiastic input and help in the preparation of this work. Claudia Castellani would like to extend her sincere thanks to Linden Hardisty and Stefan Lupu at The Devonshire Nuffield Health and Racquets Club for vitally de-stressing and enjoyable tennis coaching and ‘ball-hitting sessions’ particularly during the final editing of the taxonomy section. Finally, thanks to our families and friends for their unending support, patience and encouragement in bringing this book to life. Claudia Castellani and Martin Edwards

LIST OF CONTR IBUTOR S

Andreas Altenburger, Natural History Museum of Denmark, University of Copenhagen, Denmark Email: [email protected] Martin V. Angel, National Oceanography Centre University of Southampton, UK Email: [email protected] Cecilia Balestreri, The Marine Biological Association, Plymouth, UK Email: [email protected] Gregory Beaugrand, University of Lille, France Email: [email protected] Mark Benfield, Louisiana State University, USA Email: [email protected] John Bishop, The Marine Biological Association, Plymouth, UK Email: [email protected] Keith Brander, DTU-Aqua, Copenhagen, DK Email: [email protected] Martina Brunetta, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Ann Bucklin, University of Connecticut, USA Email: [email protected] Clare Buckland, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Robert Camp, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Claude Carré, Observatoire Océanologique de Villefranche-sur-mer, France Email: [email protected] Claudia Castellani, Plymouth Marine Laboratory, UK Email: [email protected] David V.P. Conway, The Marine Biological Association, Plymouth, UK Email: [email protected] Johan Decelle, Sorbonne Université, France Email: [email protected] Martin Edwards, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Heather Esson, Institute of Parasitology, Academy of Sciences of the Czech Republic Email: [email protected] Astrid Fischer, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Judith Fuchs, The Marine Biological Association, Plymouth, UK Email: [email protected] Gabriel Gorsky, Sorbonne Université, CNRS, Laboratoire d’Océanographie de Villefranche, France Email: [email protected]

xiv  list of contr ibutor s Stephanie Henson, National Oceanography Centre University of Southampton, UK Email: [email protected] Andrew Hirst, University of Liverpool, UK Email: [email protected] Penny Holliday, National Oceanography Centre University of Southampton, UK Email: [email protected] Tanya Jonas, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Anthony W.G. John, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Alexandra Kraberg, Alfred Wegener Institute, Germany Email: [email protected] Maiju Lehtiniemi, Finnish Environment Institute (SYKE), Finland Email: [email protected] Priscilla Licandro, Stazione Zoologica Anton Dohrn (SZN), Napoli, Italy Email: [email protected] Alistair J. Lindley, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Dhugal Lindsay, Japan Agency for Marine-Earth Science and Technology ( JAMSTEC), Japan Email: [email protected] Silke Lischka, Helmholtz-Zentrum für Ozeanforschung Kiel (GEOMAR), Germany Email: [email protected] Kenneth Meland, University of Bergen, Norway Email: [email protected] Katja Metfies, Alfred Wegener Institute, Germany Email: [email protected] Peter Munk, Technical University of Denmark, Denmark Email: [email protected] Jørgen G. Nielsen, Zoological Museum, Natural History Museum of Denmark, University of Copenhagen, Denmark Email: [email protected] Fabrice Not, CNRS, France Email: [email protected] Holger Ossenbrügger, Helmholtz-Zentrum für Ozeanforschung Kiel (GEOMAR), Germany Email: [email protected] Annelies Pierrot-Bults, University of Amsterdam, Institute for Biodiversity and Ecosystem Dynamics, The Netherlands Email: [email protected] John Raven, University of Dundee, UK Email: [email protected] Carol Robinson, University of East Anglia, UK Email: [email protected] Saeed Sadri, SAHFOS and Meteorological Office, UK Email: [email protected] Antonina Dos Santos, Departamento do Mar e Recursos Marinhos Instituto Português do Mar e da Atmosfera, Portugal Email: [email protected]

list of contr ibutor s  xv Eve C. Southward, The Marine Biological Association, UK Email: [email protected] Deborah Steinberg, Virginia Institute of Marine Science, USA Email: [email protected] Rowena Stern, Sir Alister Hardy Foundation for Ocean Science, Plymouth UK Email: [email protected] Michaela Strüder-Kypke, University of Guelph, Canada Email: [email protected] Claire Taylor, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected] Peter H. Wiebe, Woods Hole Oceanographic Institution, USA Email: [email protected] Marianne Wootton, Sir Alister Hardy Foundation for Ocean Science, Plymouth, UK Email: [email protected]

GENER A L INTRODUCTION M a rtin Edwa r ds, Er ic Goberv ille, a nd Claudi a Castella ni The beginnings of the systematic study of the oceans and the planktonic organisms that inhabit them has a long history but really has its modern roots in the intrepid naturalists that were onboard early 18th and 19th century voyages of scientific discovery and exploration. Naturalists such as Edward Forbes who charted the distributions of marine organisms and Joseph Hooker who realised the importance of diatoms in marine food chains were early contributors to this emerging field of marine ecology. By the time of the HMS Challenger expeditions of the 1870s, which set the foundations of modern marine science, it was widely understood that the surface layers of the oceans contained a plethora of small and microscopic organisms. While zoological classifications of residents of the plankton particularly for crustacean copepods had existed since the early 18th century, it was the early pioneering work conducted by Victor Hensen in the mid to late 19th century that really set the stage for plankton research. Hensen established many new techniques on the study of biological oceanography using his own designs of plankton nets and coined the term ‘plankton’ based on the Greek word to wander or drift. He is credited with  being the first scientist to carry out proper quantitative and large scale plankton studies. From this period on oceanographic science took off in the early 20th century with early scientific expeditions by the ships Meteor and Discovery leading the way and vastly increasing our understanding of the oceans. Today oceanographic science still continues to be an important area of research with many maritime nations operating dedicated oceanographic research vessels and shore-based laboratories. With the advancement of technology in the 21st century, from remotely operated vehicles to molecular probes and satellite observations, marine biological research still continues in many forms and the study of plankton still remains highly topical due to the need to sustainably manage our ocean resources and because of the rapid global changes the oceans are experiencing and are set to experience over the next 100 years. The plankton themselves include the free floating photosynthesising life of the oceans (algal phytoplankton, bacteria and other photosynthesising protists), at the base of the marine food-web which provides food for the animal plankton (zooplankton) which in turn provide food for many other marine organisms ranging from the microscopic to whales. The carrying capacity of pelagic ecosystems, explored in the ecological section of this book, in terms of the size of fish resources and recruitment to individual stocks as well as the abundance of marine wildlife (e.g. seabirds and marine mammals) is highly dependent on variations in the abundance, seasonal timing and

composition of this plankton. Marine plankton constitute a huge range of organisms (their biodiversity is described in Chapter 2) and size ranges. They can range from 10 nm (e.g. marine viruses and include femtoplankton and picoplankton) to ~2m for the largest metazooplanktonic species. Zooplankton for example can consist of tiny heterotrophic flagellates (nanoplankton) which range from a few µm in size all the way up to giant jellyfish which can be measured in metres. The main size ranges used for classification and described in this book range from the nanoplankton (2.0–20 µm) which include nanoflagellates which can be either heterotrophic or autotrophic or in many cases mixotrophic but for simplicity are described in the phytoplankton section of this book. The next size class is known as microplankton (20–200 µm) which consists of many diatoms and dinoflagellates and protozoans such as ciliates and foraminiferans. The mesoplankton (0.2–2 mm) are dominated by crustacean plankton and in particular copepods with the larger size range macroplankton (2–20 cm) consisting of large crustaceans such as euphausids, amphipods and larger ­specimens of hydromedusae and ctenophores as well as fish ­larvae. The final size range megaplankton (20–200 cm) is only represented by a few organisms such as chain-forming salps, siphonophores and large jellyfish. Recent reviews have highlighted a worrying decline in taxonomic skills in marine science that has accelerated over the last few decades mainly due to shifting research priorities. However, there has never been a more timely need to increase taxonomic skills due to the threat of future species extinctions. There is a real concern that the oceans will lose a large part of its biodiversity in the coming decades and there will be a need to document and assess these changes. Taxonomy can be considered a foundation of biodiversity science and as such it will be important to pass on taxonomic skills to the next generation of marine scientists. This book will contribute to the training and development of this important but neglected area. This book taxonomically covers all the main organisms of plankton seen between these size ranges mentioned above but focuses more on the mesozooplankton and in particular the crustacean plankton. In standard methods and in the most common form of plankton sampling (detailed in section 2 of this book) these organisms typically dominate the plankton and include up to eight dominant groups: Amphipoda, Cirripedia, Branchiopoda, Copepoda, Decapoda, Euphausiacea, Mysida and Ostracoda. As a consequence the zooplankton crustacea dominate the taxonomic section. Collectively these taxa constitute what is termed the crustacean plankton as opposed to what is collectively known as gelatinous plankton. The gelatinous plankton comprises of

xviii  gener a l introduction highly abundant organisms including the Cnidaria Hydrozoa, and Scyphozoa (common jellyfish), the Ctenophora (comb ­jellies) and the Thaliacea (i.e. doliolids and salps). Other very common zooplankton groups includes the fish larvae, the Chaetognatha (arrow worms) and the Thecosomata and Gymnosomata (pelagic sea snails). The core of the book ­provides a comprehensive, highly informative and referenced identification guide to the marine planktonic organisms found in the North Atlantic basin and adjacent seas. While the main focus is on the mesozooplankton there are sections dealing succinctly with the most important representatives of the ­ ­phytoplankton and the microzooplankton. The book covers predominately boreal and temperate species but also covers some of the most abundant warm water species which have recently appeared in northern latitudes. Additional sections on plankton ecology and methodology provides students and ­professionals with an authoritative overview of the ecological importance of plankton, their collection in the field and ­handling in the laboratory. Ecologically speaking and on a planetary scale, plankton and pelagic ecosystems as a metaphorical collective entity, are perhaps some of the most sensitive organisms to environmental change and one of the most important biological communities on the planet. They inhabit the pelagic zone, which comprises 71% of the planetary surface and a huge proportion of the earth’s total biosphere. They are responsible for the overwhelming majority of marine biological production that fuel marine food-webs and nutrient cycling as well as contributing to approximately half of the world’s oxygen production and carbon sequestration. It is well documented, and explored in this book, that global environmental changes caused by human-activities have had large consequences for the Earth’s oceans. In marine environments the main drivers of change include climate warming, point-source eutrophication, deoxygenation and unsustainable fishing. Furthermore and unique to 75

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the marine environment, anthropogenic CO2 is also associated with ocean acidification. Ocean acidification has the potential to affect the process of calcification and therefore certain planktonic organisms dependant on calcium carbonate for shells and skeletons formation may be particularly vulnerable to increasing CO2 emissions. It is also worth noting that while pelagic systems are undergoing large changes caused by climate change they have also been identified as a form of mitigation of climate change through possible human manipulation of these systems using geoengineering schemes. It has been shown at smallscales that the addition of iron to certain oceanic environments (ocean fertilization) can increase productivity and net export of carbon to the deep ocean. However, this approach is still controversial with largely unknown long-term ramifications for marine ecosystems at the large-scale. Plankton have also been identified as potential biofuels and have more recently been harvested for the food supplement and cosmetic industry (e.g. omega 3 products). All these changes and marine management issues show a clear need to interpret these trends we are observing in the oceans, explored in Section 1 of this book, and effectively monitor these change through various sampling methodologies which are documented in Section 2. Section 3 of the book, while focused on traditional taxonomy and the identification of the plankton, also shows the current distributions of copepod species in the North Atlantic. This baseline biogeographical information will be crucial in documenting any further northward shifts in the plankton over the next coming decades due to climate warming (see for example the changes in the biogeography of Calanus finmarchicus which has occurred over the past 25 years in Fig. I.1). Plankton are tightly coupled to fluctuations in the marine environment and highly sensitive indicators of environmental change such as nutrient availability, ocean ­current changes and climate variability. Some of the most rapid northward movement of organisms caused by climate warming 75

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FIGURE I.1:  Biogeography of Calanus finmarchicus in the North Atlantic showing the northward shift in abundance (as log10(X+1)) from a) 1958 and 1989 to b) 1990 and 2014. Data from the Continuous Plankton Recorder Survey.

gener a l introduction  xix over the whole globe throughout the last few decades have been planktonic. Temperature is a key driver of marine ecosystems and in particular its effects on pelagic populations are manifested very rapidly. This is mainly because 99% of pelagic and planktonic organisms are ectothermic making them highly sensitive to fluctuations in temperature. The rapidity of the planktonic response is predominantly due to their short life-­ cycles and mainly in their passive response to advective changes. For example, phytoplankton fix as much CO2 per year as all terrestrial plants but due to being unicellular they represent at any one time only 1% of the Earth’s biomass and have life cycles measured in days to weeks rather than years to decades like their terrestrial counterparts. It was with this in mind, these rapid changes in the marine environment and the need to monitor and assess these changes this book developed into something that could encompass not only the ecology of plankton but also a guide to their identification and collection. This ambitious book covers many aspects of marine plankton from their taxonomy and identification to their global ecological significance. Highly topical chapters on the ecology of these organisms encompass key areas covering their physical habitat and physiology to their biodiversity and biogeochemical processes. Further chapters explore their crucial roles in fisheries and understanding the impacts of global climate change. In the first ecological section of the book, ­chapter 1 provides an introduction to the marine environment describing the physics and chemistry that set the scene for life in the pelagic realm. Furthermore the climate and circulation of the North Atlantic is explored and how key physical features and processes affect ecosystems via the availability of light and nutrients. In the second chapter plankton biodiversity and biogeography are further described in terms of their habitat and ecological scales in space and time. The chapter concludes on the biogeography and biodiversity of plankton at the global ocean scale. Chapters 3 and 4 describe the productivity of phytoplankton and zooplankton respectively from the initial energy

and chemical requirements for photosynthesis to the rate of production of heterotrophic organisms. Chapters 5 and 6 explores the processes and importance of phytoplankton and zooplankton biogeochemical cycles operating in our oceans describing the biological carbon pump and the impact of increasing atmospheric CO2 on plankton mediated biogeochemical cycles. Chapter 7 focuses on global changes caused by human-activities and their effects on marine plankton and introduces some key concepts of plankton ecology such as the ecological niche concept, plankton succession and the use of planktonic indicators to monitor these changes. The final chapter of the ecological section explores the dependence of fish on plankton and the relationship between plankton productivity and fisheries production. In the second section of the book covers the more practical aspects of plankton research covering the methods in which plankton are sampled, preserved and counted from traditional methodologies to new molecular and automated processes. The extensive section includes information on the collection of plankton samples using a multitude of methods from traditional plankton nets to optical systems and the inter-comparison between the various methods. The section also describes ­preservation and the analysis of plankton samples from basic biomass measures all the way to molecular methodologies and probes and to next generation sequencing. The final section on  taxonomic identification is a comprehensive guide to the  ­myriad of phyla and forms that make up the plankton from  ­single celled protists and algae to the megaplankton. Authoritative taxonomic and informative keys throughout the book are designed to help individuals quickly identify the plankton taxa under investigation. Each chapter describes a ­specific Phylum or Class, depending on the number of species considered. Collectively, these three sections will have a wide appeal to undergraduates and graduates as well as professional researchers and provide an invaluable guide to marine plankton for now and into the future.

PLATE 2:  True colour image from the MODIS-Aqua satellite showing

mesoscale stirring of a phytoplankton bloom in the Barents Sea (August 14th 2011). Bright, greenish areas have high chlorophyll concentration indicated by yellow to orange shading, plus the thin blue lines of the fresh and dark areas have low chlorophyll concentration. Image from the East Greenland and Labrador Currents; deep circulation indicated by the NASA Earth Observatory http://earthobservatory.nasa.gov. (Figure 1.3) wide blue lines that represent the dense northern overflows and export of North Atlantic Deep Water to the southern hemisphere. (Figure 1.2) A PLATE 1:  Schematic of North Atlantic circulation. Shallow circulation

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PLATE 3:  Mixed layer depth in winter (February, top panel, logarithmic colour scale in metres) and in summer (August, lower panel, contour intervals 5m). Base of mixed layer defined as the depth where potential density is 0.125 kg/m3 greater than the surface value. Climatological data from World Ocean Atlas 2009. Image created by Chongyuan Mao, NOC. (Figure 1.5)

PLATE 4:  Spatial and temporal variability in calanoid copepod biodiversity in the North Atlantic Ocean. A, Quantification of diel, seasonal and year-to-year variability in the Bay of Biscay, the northern part of NADR and in the North Sea. B, Spatial changes in the scale at which biodiversity varies in the North Atlantic Ocean. A blue colour suggests that variability varies at a small scale, whereas the red colour indicates that biodiversity fluctuates at a large scale. The main surface currents are indicated. From Beaugrand et al. (2003). (Figure 2.3)

PLATE 6:  Concentration in A, nitrate; B, phosphate; and C, silicate in

sea water. Data, from the World Ocean Atlas. From Beaugrand (2015). (Figure 2.6)

PLATE 5:  Concentration in A, dissolved dioxygen; B, mixed layer

depth (MLD; density); C, bathymetry; and D, sea surface salinity. Dissolved dioxygen and sea surface salinity are from the World Ocean Atlas. Bathymetry data is from the General Bathymetric Chart of the Oceans (GEBCO). MDL (density) data originated from de Boyer Montégut et al. (2004). (Figure 2.5)

PLATE 7:  High Nutrient Low Chlorophyll areas as revealed by the

ratio of standardised nitrates (between 0 and 1) on (standardized chlorophyll (between 0 and 1) +1) in January (top) and July (bottom). Values of chlorophyll greater or equal to 2 mg.m−3 and of nitrates greater or equal to 20µmoles.L−1 were fixed to 1 in the standardized values of nitrates and chlorophyll, respectively. Black areas are regions where the ratio is 0. Data from SeaWIFS and the World Ocean Database. The location of some ocean iron experiments is indicated on the map. IRONEX I and II: IRON fertilization EXperiment (1993 and 1995); SOIREE: Southern Ocean Iron RElease Experiment (1999); SEEDS: The Subarctic Pacific Iron Experiment for Ecosystem Dynamics Study (2001); SERIES: Subarctic Ecosystem Response to Iron Enrichment (2002); EIFEX: European Iron Fertilization Experiment (2004). From Beaugrand (Beaugrand 2015). (Figure 2.7)

PLATE 8:  Biogeography of the global ocean (i.e. biogeochemical provinces) proposed by (A) Longhurst (2007) and (B) recalculated by a numerical procedure from January 1998 to December 2007. From Reygondeau and co-workers (Reygondeau et al. 2013). (Figure 2.9)

PLATE 9:  Mean spatial distribution of some key species or taxa in the North Atlantic Ocean. Modified, from Beaugrand et al. (Beaugrand et al.

2003). (Figure 2.10)

PLATE 10:  Reconstruction of the present-day pelagic biodiversity from the Macro-Ecological Theory on the Arrangement of Life (METAL). From Beaugrand and co-workers (Beaugrand et al. 2013). (Figure 2.12)

A Log10(Specific respiration rate, μl O2 mg C–1h–1)

4 Protozoa Non-calanoid copepods Calanoid copepods Amphipods Euphausiids Tunicates Cnidaria + Ctenophores Fish Mammals (Kleiber)

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Ciliates Chaetognaths R/W~W0 R/W~W–0.25

Log10(Specific clearance rate, ml mg C–1h–1)

6 5 4 3 2 1 0 –10 –9 –8 –7 –6 –5 –4 –3 –2 –1 0 1 2 3 4 5

C Log10(Specific ingestion rate, μg C mg C–1h–1)

4 3 2 1 0 –1 –10–9 –8 –7 –6 –5 –4 –3 –2 –1 0 1 2 3 4 5

D Log10(Specific growth rate, mg C mg C–1h–1)

0 –1 –2 –3 –4 –5 –10–9 –8 –7 –6 –5 –4 –3 –2 –1 0 1 2 3 4 5 6 Log10 (Body mass, mg C)

PLATE 11:  Regressions through mass-specific rates versus body mass for various taxa, 95% confidence intervals for the regressions also shown. All rates are corrected to a common temperature of 15°C. A. respiration rates, mammals are also included for comparison (Kleiber, 1975), B. maximum clearance rates, C. maximum ingestion rates, D. maximum growth rates. The black lines are regressions through all data assuming proportionality between mass-specific respiration rate and body mass raised to power 0 or -0.25. In panel A protozoans include both ciliates and flagellates, in lower panels the ciliates are shown as a separate group and the pink line denotes flagellates alone. Figure adapted from Kiørboe and Hirst (2014). Reproduced with permission from the University of Chicago Press. (Figure 4.4)

CO2

N

2

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+

PO

3–

4

H+ HCO3–

NO3–

– NO 2

CO32–

4

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(Green:Red:Blue)

Phytoplankton Diazotrophs Microzooplankton NO3–

N2O PO

NO2

–

3–

CO2

Mesozooplankton

4

Bacteria/Archaea NH4+

RDOC N2O N2

NO

Faecal pellets

–

2

NO3– NH4+

POM DOM Viruses

PLATE 12:  Infographic of the inter-relationships between the organic and microbial carbon pumps and the microbial cycling of nitrogen and phosphorus. RDOC: recalcitrant dissolved organic carbon, POM: particulate organic material, DOM: dissolved organic material. C:N:P ratios are indicative only. Figure by Robinson et al. shared under a creative commons license at Figshare http://dx.doi.org/10.6084/m9.figshare.1585741. (Figure 5.1)

Temperate species

Warm-temperate species

Subarctic species

Cold-temperate species

1958–1981

1958–1981

1958–1981

1958–1981

1982–1999

1982–1999

1982–1999

1982–1999

2000–2002

2000–2002

2000–2002

2000–2002

60°N 50°N

60°N 50°N

60°N 50°N 2003–2005

2003–2005

2003–2005

2003–2005

60°N 50°N 0.00 0.04 0.08 Mean number of species per CPR sample 1000 km shift northward

0.0

0.4

0.8

0.0

0.4

0.8

0.0

0.4

0.8

Echinoderms 47 days Other meroplankton 27 days Dinoflagellates 23 days

PLATE 13:  Examples of biogeographical and phenological shifts seen over a number of decades recorded by the Continuous Plankton Recorder in the North East Atlantic. Biogeographical shifts: biogeographical changes in plankton assemblages spanning five decades. Warmwater plankton (e.g. warm-temperate species) are moving north and cold-water plankton (e.g. subarctic species) are moving out of the North Sea. Particular rapid movement is seen along the European Continental Shelf up to 1000 km over 50 years. Based on Beaugrand et al. (2003). (Figure 7.1a)

PLATE 14:  Examples from the diatom genera featured above: a. Chaetoceros teres, b. Chaetoceros teres (resting stage), c. Chaetoceros decipiens (arrows are indicating the fused setae), d. Coscinodiscus radiatus, e. Odontella aurita, f. Thalassiosira nordenskioeldii, g. Thalassiosira punctigera (girdle view), arrows are indicating the occluded processes, h. Pseudo-nitzschia cf. pungens (see arrow), i. Asterionellopsis glacialis, j. Rhizosolenia spp. (All images by A. Kraberg, Alfred Wegener Institute, Source: Plankton*net, http://planktonnet.Alfred Wegener Institute.de)

PLATE 15:  Examples of important dinoflagllate taxa in the North Sea, North Atlantic: a. Protoperidinium steinii, b. Ceratium lineatum, c. Dinophysis

acuta, d. Gonyaulax digitale, e. Gyrodinium sp. (source: http://planktonnet.Alfred Wegener Institute.de, image author: Franz Neidl), f. Gyrodinium spirale with ingested prey (a chain of the diatom Paralia sulcata). (Images a-d and f by A. Kraberg, Alfred Wegener Institute, source Plankton*net, http://planktonnet.Alfred Wegener Institute.de).

PLATE 16:  Selected phototrophic flagellates from the North Atlantic: Prymesiophyceae: a. Chrysochromulina rotalis (image by F. Jouenne, CNRS, Station Biologique de Roscoff, source Plankton*net), b. Coccolithus sp. (image by G. Durak, Marine Biological Association, personal communication), c. Prymnesium patilifferum (image by F. Jouenne, CNRS, Station Biologique de Roscoff, source Plankton*net), d. Phaeocystis globosa, fixed in formalin (image by A. Kraberg, Alfred Wegener Institute, source Plankton*net), Pavlovophyceae: e. Pavlova pinguis, mobile stage dominant (image by K. Fresnel, source Plankton*net), Crytophyceae: f. Rhodomonas sp. (image by D. Vaulot, CNRS, Station Biologique de Roscoff, source Plankton*net), g. Chroomonas sp. palmella stage (image by Hoef-Emden, K., Tree of Life Project, licensed under Creation Commons Attribution-NonCommercial-ShareAlike Licence v. 2.5). Euglenophyceae: h. Euglena sp. (image by G. Mathews, Friends of Warnham Nature Reserve), Raphidophyceae: i. Chattonella sp. (image by M.A. Sampayo, source Plankton*net), Pelagophyceae: j. Sarcinochrysis marina var filamentosa. Warm water species with potential pseudofilaments, palemelloid form (image by C. Billard, source Plankton*net), Dictyochophyceae: k. Meringosphaera mediterranea (image by F. Jouenne, CNRS, Station Biologique de Roscoff, source Plankton*net), l. Dictyocha sp. (image by A. Kraberg, Alfred Wegener Institute, source Plankton*net), Bolidophyceae:  m. Bolidomonas mediterranea (image by F. Jouenne, CNRS, Station Biologique de Roscoff, source Plankton*net), Eustigmatophyceae: n. Nanochloropsis gaditana (image by F. Jouenne, CNRS, Station Biologique de Roscoff, source Plankton*net), Prasinophyceae: o. Halosphaera sp. (image by A. Kraberg, Alfred Wegener Institute, source Plankton*net), p. Pyramimonas sp. (image by C. Billard, source Plankton*net), q.  Micromonas pusilla (image by E. Foulon, source Plankton*net).

a

b

c

d

PLATE 17:  Micrographs of selected ciliate species: a. Laboea strobila (image F. Neidl, source Plankton*net), b. Tiarina fusus (image A. Kraberg, Alfred Wegener Institute, source Plankton*net), c. Mesodinium rubrum (image C. Widdicombe, Plymouth Marine Laboratory, source Plankton*net), d. Tintinnopsis cylindrica (image A. Kraberg, Alfred Wegener Institute, source Plankton*net).

A

B

E

F

C

G

D

H

PLATE 18:  Examples of Acantharia: A: Gigartacon Muelleri (Chaunacanthida), B: Phyllostaurus echinoides (Arthracanthida), C: Diploconus fasces (Arthracanthida), D: Lychnaspis giltschii (Arthracanthida), E: Amphilonche elongate, F: Lithoptera fenestra ((Arthracanthida), G. Acanthochiasma spp. (Holocanthida), H: Sticholonche zanclea (Taxopodia). Images A–F were reprinted from Molecular Phylogeny and Morphological Evolution of the Acantharia (Radiolaria), 163/3, Decelle, J., Suzuki, N., Mahé, F., de Vargas, C., Not, F., Fig.3, 435–450., Copyright (2012), with permission from Elsevier. Panel G. from M. Webber, Biosearch). Panel H. from Shimoda marine station, (Tsukuba.ac.uk). Regionally Integrated Marine database. Copyright 2015 Japanese Association for Marine Biology ( JAMBIO).

PLATE 19:  Polycystinea montage showing examples of A: Collodarian, B: Spumellarian, C: Nassellarian, D: Nassellarian. (Images by F. Not,

Station Biologique Roscoff, France, personal communication).

PLATE 20:  The schyphozoan jellyfish Pelagia noctiluca (Cnidaria: Schyphozoa) (Credit F. Lombard, Observatoire Océanographique de Villefranche).

PLATE 21:  The non-colonial hydrozoan jellyfish Pandea rubra

(Cnidaria: Anthoathecata) (credit D.Lindsay, JAMSTEC).

PLATE 22:  The colonial hydrozoan jellyfish Abylopsis tetragona

PLATE 23:  The comb-jelly Beroe cucumis (Ctenophora) (credit R.

(Cnidaria: Siphonophorae) (credit R. Hopcroft, University of Alaska Fairbanks).

Hopcroft, University of Alaska Fairbanks).

PLATE 24:  The calanoid copepod Calanus finmarchicus (Crustacea:

PLATE 25:  The cyclopoid copepod Oithona similis (Crustacea:

Copepoda) (credit R. Hopcroft, University of Alaska Fairbanks).

Copepoda) (credit O. Larink, Techn. Univ. Braunschweig).

PLATE 26:  Cladoceran Evadne nordmanni female carrying embryos

PLATE 27:  Acorn barnacle larvae, nauplius stage (Crustacea:

(Crustacea: Branchiopoda) (credit O. Larink, Techn. Univ. Braunschweig).

Cirripedia) (credit R. Hopcroft, University of Alaska Fairbanks).

PLATE 28:  The crustacean Themisto libellula (Amphipoda: Hyperiidae)

(credit R. Hopcroft, University of Alaska Fairbanks).

PLATE 29:  Crustacean larvae, crab zoea stage (Decapoda: Brachyura) (credit R. Hopcroft, University of Alaska Fairbanks).

PLATE 30:  Meganyctiphanes norvegica (Crustacea: Euphausiacea)

PLATE 31:  Conchoecissa imbricata (Ostracoda: Halocyprididae) (credit

(credit R. Hopcroft, University of Alaska Fairbanks).

R. Hopcroft, University of Alaska Fairbanks).

PLATE 32:  The holoplanktonic polychaetes Tomopteris spp (Anellida:

PLATE 33:  The holoplanktonic polychaetes Travisiopsis spp. (Anellida:

Tomopteridae), (credit R. Hopcroft, University of Alaska Fairbanks).

Typhloscolecidae) (credit O. Larink, Techn. Univ. Braunschweig).

PLATE 34:  Adult stage of the holoplanktonic mollusc Limacina retroversa (Mollusca: Thecosomata) (credit R. Hopcroft, University of Alaska Fairbanks).

PLATE 35:  Adult stage of the holoplanktonic mollusc Clione Limacina

PLATE 36:  Adult stage of the holoplanktonic mollusc Atlanta peroni

PLATE 37:  Adult stage of the holoplanktonic mollusc Carinaria

(Mollusca: Pterotracheoidea) (credit R. Hopcroft, University of Alaska Fairbanks).

lamarckii (Mollusca: Pterotracheoidea) (credit R. Hopcroft, University of Alaska Fairbanks).

PLATE 38:  Adult arroworm Parasagitta elegans (Chaetognata), (credit

PLATE 39:  Brittle star larva, Ophiopluteus stage (Echinodermata),

R. Hopcroft, University of Alaska Fairbanks).

(credit R. Hopcroft, University of Alaska Fairbanks).

(Mollusca: Gymnosomata) (credit R. Hopcroft, University of Alaska Fairbanks).

PLATE 40:  Cyphonautes larva (Bryozoa) (credit O. Larink, Techn. Univ. Braunschweig).

PLATE 41:  Oikopleura gorskyi (Chordata: Appendicularia) (credit R.

Hopcroft, University of Alaska Fairbanks).

PLATE 42:  Chain forming salp Salpa fusiformis (Chordata: Thaliacea)

PLATE 43:  Fish larva of Anarhichas spp. (Cordata: vertebrata) (credit

(Credit F. Lombard, Observatoire Océanographique de Villefranche).

R. Hopcroft, University of Alaska Fairbanks).

PLATE 44:  Fish scale (example of non-planktonic objects often found in plankton samples) (credit O. Larink, Techn. Univ. Braunschweig).

PLATE 45:  Moult (exuviae) of an adult acorn barnacle (example of  non-planktonic objects often found in plankton samples) (credit O. Larink, Techn. Univ. Braunschweig).

PLATE 46:  Scale of a butterfly wing (insect) (example of non-

PLATE 47:  Pinus pollen (plant) (example of non-planktonic objects

planktonic objects often found in plankton samples) (credit O. Larink, Techn. Univ. Braunschweig).

often found in plankton samples) (credit O. Larink, Techn. Univ. Braunschweig).

Sec t ion I

Ecology

Ch a pter 1

The M ar ine En v ironment N. Penn y Holliday a nd Steph a nie Henson

1.1 Introduction The physical environment is the primary control on the growth, distribution and variability of phytoplankton populations in the North Atlantic. Phytoplankton growth is fundamentally reliant on the sufficient supply of two factors: nutrients and light. The availability of both of these factors is determined by the physical environment via changes in mixing and circulation. As phytoplankton have fast growth rates, typically ~ one doubling per day (Lalli and Parsons, 1997), short-term changes in the physical forcing which affect the availability of light or nutrients can have a rapid and pronounced effect on the phytoplankton ­population. Additionally, phytoplankton generally are either not motile or swim slowly, so that their distribution is determined by the physical circulation. Understanding why phytoplankton grow when and where they do therefore relies on an understanding of the physical environment controlling their abundance. Generally speaking, the availability of nutrients and light drive both the seasonal cycle and large-scale spatial variability in phytoplankton populations. Light is required for photosynthesis, with different phytoplankton functional types thriving in different light regimes. Light is attenuated rapidly in the ocean, with irradiance being reduced to 1% of its surface value within the top ~  20–120  m, depending on the clarity of the water. Phytoplankton also require multiple macronutrients (nitrate, silicate and phosphate) and micronutrients (most notably iron) to grow. Nutrient concentrations are greatest below the mixed layer and once surface concentrations have been depleted via phytoplankton uptake, significant quantities of new nutrients can only be supplied by vertical mixing. This may take the form of a deepening mixed layer associated with the onset of autumn (and hence cooler, windier conditions) or a transient event such as a storm or passage of an eddy. This reliance on the supply of both nutrients and light combines to drive the seasonal cycle of phytoplankton growth. In the subpolar North Atlantic, where the ‘textbook’ spring phytoplankton bloom occurs (Sverdrup,  1953), phytoplankton growth is light limited in winter due to deep mixed layers and

short, dim days. The deep winter mixing ensures an abundant supply of nutrients to the upper ocean, but phytoplankton are unable to take advantage due to poor light conditions. With the arrival of warmer conditions and reduced winds in spring, the upper ocean warms creating a stratified layer. Now, with plenty of nutrients available and good light conditions, phytoplankton can bloom. As nutrients become depleted, and grazing by zooplankton becomes important, the phytoplankton abundance is reduced to a summer minimum before increased mixing in autumn introduces new nutrients to the upper ocean. If there is sufficient light, a secondary autumn bloom may occur before strong winds and a cooling ocean increase the mixed layer depth so that light is again limiting to phytoplankton growth. In the subtropics, warm temperatures and weak wind mixing result in relatively shallow mixed layers year-round. Nutrient availability, rather than light, is the principal limiting factor in these regions. The deepening of the mixed layer that occurs in winter (although far less dramatic than in subpolar regions) supplies new nutrients, while light conditions remain suitable for growth, so that increased phytoplankton abundance occurs from late autumn to early spring. In these nutrient limited areas, transient mixing events, such as a passing eddy, may also supply nutrients over a short timescale and thus stimulate phytoplankton growth. In oligotrophic regions where stratification is strong and continual, or in temperate regions in summer which experience strong stratification, a subsurface chlorophyll maximum is often found around the nutricline, provided there is sufficient light at those depths for growth to occur. These considerations of light versus nutrient availability also drive the observed spatial variability in phytoplankton populations, whether on the basin, local or mesoscale. At the largest scales, ocean circulation and basin-wide atmospheric patterns drive the distribution of plankton and are ultimately responsible for the changes in wind stress and net heat flux that control seasonal and spatial variability in mixing. Local-scale variability in mixing can also arise that also affects the growth conditions of phytoplankton, for example from freshwater input due to rivers or ice melt. Persistent oceanic fronts or upwelling f­ eatures that supply nutrients to the upper ocean are often associated

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

4 ecology TIME Century Bi

og

M ) idth

De

ts (w

Decade

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hi ca

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gr

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tio ns (

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e.g

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)

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Upwelling events Diel vertical migration

10 km

100 km

1000 km

10 000 km SPACE

Figure 1.1:  Oceanographic and climatic processes in the North Atlantic controlling where and when plankton grow, acting on multiple time and space scales ranging from mm to biogeographical provinces and from seconds to centuries. AMO, Atlantic Multidecadal Oscillation; NAO, North Atlantic Ocillation. Based on Edwards et al. (2010).

with strong gradients in phytoplankton populations. Finally, at the mesoscale and smaller, eddies and meanders result in transient changes to light or nutrient availability that alter phytoplankton abundance. Clearly, the processes controlling where and when phytoplankton grow act on multiple time and space scales (Fig. 1.1). At the smaller scales, wind-induced Langmuir circulation might be set up in calm seas, or the passing of storms may generate locally-enhanced turbulent mixing, both of which can affect phytoplankton growth and zooplankton accumulation. In this chapter we provide an overview of the marine environment from basin-wide to mesoscale, and give a brief overview of climate-scale temporal variability.

1.2  The Circulation of the North Atlantic The North Atlantic between the equator and the Arctic Circle is  dominated by two major wind-driven features: the great ­circulating pool of warm water that is the subtropical gyre ­(10–50°  N), and the smaller, cooler, anticlockwise subpolar gyre to the north (Fig. 1.2). The gyres are mainly upper ocean, horizontal features transporting huge volumes of water within the North Atlantic. In the subtropical gyre the western side is dominated by the Gulf Stream system which starts in the south as water from the eastern Atlantic and Equatorial Current systems are drawn into the narrow Florida Current. As the Gulf Stream travels northwards it increases in volume as it entrains more recirculating water, until it reaches a maximum of 150 Sv south of Nova Scotia. As the Gulf Stream is forced away from

Figure 1.2: Schematic of North Atlantic circulation. Shallow circulation indicated by yellow to orange shading, plus the thin blue lines of the fresh East Greenland and Labrador Currents; deep circulation indicated by the wide blue lines that represent the dense northern overflows and export of North Atlantic Deep Water to the southern hemisphere. (See Plate 1.)

the coast of the USA and turns eastwards, it loses its coherence, forming meanders and eddies and has considerable variability in the position of the fronts associated with the current. Around 30 Sv of the Gulf Stream water turns northwards and becomes the North Atlantic Current (NAC), while the rest recirculates within the subtropical gyre (Hansen and Osterhus, 2000). The warm waters of the NAC release heat gained in the subtropics to the atmosphere, which contributes to the mild climate of maritime north-west Europe. The current consists of a number of dynamically active fronts and eddies and is very ­variable in location as a result (Brambilla and Talley,  2008). There is much vertical motion associated with the NAC; surface waters are subducted below it while nutrient-rich deep waters are returned to the surface. Most of the NAC water becomes entrained into the ~  30  Sv transported within the subpolar gyre. This is a region of intense water mass transformation, where the influence of the atmosphere and surrounding water masses changes warm saline NAC water into denser, cooler and fresher varieties. The heat loss from the ocean to the atmosphere is one of the most significant features of the region, partly because of the influence on regional climate, but also because it leads to deep convective mixing (600–800 m in the eastern subpolar region, and up to 2000 m in the western subpolar region). The deep water generated by this process draws carbon into the deep layers of the ocean, spreads throughout the intermediate layers of the subpolar region, and forms part of the southward-flowing limb of the overturning circulation. Very saline Mediterranean water is another water type found at intermediate depths in the North Atlantic (Potter and Lozier, 2004). Extensive evaporation in Atlantic water that flows to the eastern Mediterranean generates a return flow of very saline

the m a r ine en v ironment  5 and dense water. The volume of Mediterranean water that enters the North Atlantic is small, but it has an important influence on the stratification of the eastern subpolar gyre. Around 7 Sv of NAC water escapes from the subpolar gyre between Greenland and Scotland, flowing into the Nordic Seas and the surrounding shelf seas (Hansen et  al.,  2008). In the Norwegian Sea, the Atlantic water flows northwards in two main currents, with some entering the Arctic Ocean through the Fram Strait. Atlantic water also gets diverted into the North Sea and the Barents Sea; in the latter it becomes heavily modified by the atmosphere and through coastal run-off before entering the deep layers of the Arctic Ocean (Gammelsrod et al., 2009). Most of the Atlantic water is recirculated within the Nordic Sea gyres though, becoming denser and pooling in the deep basins. The western boundaries of the Greenland and Iceland Seas are dominated by the southward-flowing shallow Arctic water, some of which mixes into the interior of the basin. The dense water formed in the Nordic Seas spills over the shallow ridges that stretch from Greenland to Scotland via Iceland (Eldevik et  al.,  2009). This ‘overflow’ water has the equivalent total transport to the portion of NAC water that flows north through the same region (~ 7 Sv), thereby maintaining across-basin mass balance (Hansen and Osterhus, 2000). The overflow plumes sink on entering the subpolar North Atlantic because of their density; guided by topography they circumnavigate the deep Iceland and Irminger basins before entering the Labrador Sea (Dickson and Brown, 1994). Along with the cooled and freshened NAC water they are exported southwards below the western subtropical gyre as part of the Atlantic Meridional Overturning Circulation (AMOC) (Haine et al., 2008). The continental shelves around Greenland, Canada and the north USA are dominated by a different variety of northern water; these Arctic outflows are cold, fresh and nutrient-rich, which makes them buoyant and productive (Bacon et al., 2008). Across-shelf exchange gradually mixes the Arctic-origin water into the interior of the subpolar gyre, but eventually the Arcticorigin shelf currents meet the Gulf Stream off the eastern coast of the USA and turn into the interior (Rossby, 1999).

filaments from the boundaries of eddies can significantly enhance vertical velocity around the margins. The major ocean frontal systems such as the Gulf Stream and the North Atlantic Subpolar Front continually generate eddies, and a significant portion of transport of the NAC takes the form of eddy transport. Although the Gulf Stream is a nexus of eddy variability, eddies are ubiquitous in the world’s oceans and are a significant factor in the patchiness of primary productivity. In the core of a cyclonic eddy, the surface divergence raises the nutricline, potentially bringing new nutrients into the euphotic zone and thus stimulating phytoplankton growth (McGillicuddy et  al.,  1998). Anticyclonic eddies, with a convergence at their centre, should in theory elicit no phytoplankton response; however, there is evidence that some anticyclonic eddies have elevated phytoplankton abundance around the edges of the core (Martin et al., 1998; Crawford et al., 2007), possibly stimulated by nutrients supplied through submesoscale mixing (Levy et al., 2001; Mahadevan et al., 2008). Eddying flows also alter phytoplankton distribution by advecting populations along meanders and fronts. Satellite ocean colour images (such as in Fig.  1.3) demonstrate the strong influence that mesoscale variability can have on phytoplankton distribution. It can be useful to map the geography of the oceans and seas as a combination of physical and biological features, with fronts or regions of high eddy activity forming natural boundaries to  ecosystems. ‘Biogeographical provinces’ are regions within which biological properties are similar but distinct from other provinces. Provinces are typically defined on the basis of  seasonal variability in the mixed layer depth, sea surface ­temperature (SST) etc., and can be based on fixed locations (Longhurst, 2007) or else be more flexible as physical features change their location as a response to atmospheric conditions or climate variability (e.g. Sarmiento et al., 2004; Holliday et al., 2006). In the most widely used biogeographical province definition (Longhurst,

1.3  Fronts and Eddies Ocean fronts are the regions of rapid changes in density where different water masses meet and geostrophic currents develop. Fronts are characterized by significant vertical changes in velocity (shear), which generates waves or ‘baroclinic instabilities’ that form eddies. Eddies are the oceanic analogue of atmospheric weather systems, but are smaller (50–200 km), last longer (one to a few months), and move more slowly (a few kilometres per day) than atmospheric features. Eddies may circulate cyclonically or anticyclonically and therefore cause the thermocline either to shoal (surface divergence and upwelling) or to deepen (surface convergence and downwelling). Eddies can keep a parcel of cold or warm water isolated from the ambient water for some considerable time, although the stretching of

Figure 1.3:  True colour image from the MODIS-Aqua satellite showing mesoscale stirring of a phytoplankton bloom in the Barents Sea (August 14th 2011). Bright, greenish areas have high chlorophyll concentration and dark areas have low chlorophyll concentration. Image from the NASA Earth Observatory http://earthobservatory. nasa.gov. (See Plate 2.)

6 ecology is sometimes split into two areas east and west of the MidAtlantic Ridge (NAST-E and NAST-W). Despite winter mixed layers that are similar to the NADR, productivity in the NAST is  lower, although with a similar seasonal cycle of increasing production from January onwards to a peak in March and a secondary autumn maximum. Finally, the North Atlantic Tropical Gyral Province (NATR) encompasses the oligotrophic gyre extending south of the NAST to the North Equatorial Current. The mixed layer depth is shallow and seasonally invariant so that persistent nutrient limitation characterizes this region. In response, phytoplankton growth is curtailed and productivity is uniformly low without a clear seasonal signal. Although the provinces described here are a useful tool for analysis of biological data, it should be borne in mind that the ubiquitous presence of eddies and other small-scale features can introduce large intra-province variability. Figure 1.4:  Biogeochemical provinces as described by Longhurst (2007). Locations are approximate and may change from year to year. ARCT, Atlantic Arctic Province; SARC, Atlantic Subarctic Province; NADR, North Atlantic Drift Province; GFST, Gulf Stream Province; NAST, North Atlantic Subtropical Gyral Province (split into east and west of the Mid-Atlantic Ridge); NATR, North Atlantic Tropical Gyral Province.

2007), the North Atlantic is divided into ~ 14 regions, for example North Sea, Canary Current, Gulf Stream etc. reflecting major oceanographic boundaries. Here we provide a brief description of the seven Longhurst provinces occupying the deep water North Atlantic (Fig. 1.4) (Longhurst, 2007). The Atlantic Arctic Province (ARCT) lies between the coastal currents of Greenland and the northern boundary of the NAC and extends northward into the Greenland Sea. Deep mixing in winter ensures that phytoplankton growth is light limited and the relatively short growing season results in a single peak in production in summer. The Atlantic Subarctic Province (SARC) stretches from south-east of Iceland north-west into the Norwegian Sea and is bounded in the east by the Barents Sea and to the south by an arbitrary line at 60 °N. Winter mixing is not as deep as in the ARCT but is nevertheless sufficiently deep to result in light limitation of phytoplankton growth. Again, there is a single productivity peak in summer, but it exceeds the ARCT production. The ARCT and SARC regions form the northern boundary of the North Atlantic Drift Province (NADR), which separates at its southern edge the NAC from the Gulf Stream. Winter mixing can be almost as deep as in the ARCT, but the single summer productivity peak is greater than in either the ARCT or SARC regions. The Gulf Stream Province (GFST) encompasses a region of strong mesoscale variability from Florida to the Newfoundland Basin. Relatively shallow winter mixed layers allow productivity to increase from February onwards, reaching a peak in early spring. A secondary productivity peak occurs in autumn as nutrient limitation is alleviated by the deepening mixed layer. The North Atlantic Subtropical Gyral Province (NAST) lies south of the NADR, is bounded to the south by the subtropical convergence zone and

1.4  Mixed Layer Depth and Stratification The annual evolution of the surface mixed layer, and the regional variations in the timing and depth of maximum mixed layer depth, is one of the key processes in physical and biological upper ocean process studies. The mixed layer is simply defined as the part of the water column above a shallow pycnocline, characterized by near-homogenous distribution of tracers such as temperature, salinity and nutrients. It is the exchange of turbulent energy and heat with the atmosphere that maintains the mixed layer, while restratification is typically induced by surface warming, the addition of low density freshwater, or through eddy processes. The simple description of what a mixed layer is belies the fact that there is no similarly simple way to globally define the mixed layer depth that is important to individual organisms in terms of light and nutrient supply (Montegut et al., 2004). The problem is partly a result of stratification being a product of temperature and salinity differences (especially relevant in the high latitudes), but also because the base of the mixed layer is a gradient in properties, not a sudden cut-off. However, the main processes responsible for mixed layer development and its variations are generally understood. In the subpolar gyre and Nordic Seas in winter, low pre-existing stratification and large heat loss to the atmosphere lead to convective mixing to depths of around 600–1000 m (historically up to 2400 m has been observed in the Labrador Sea) (Fig. 1.5). During deep winter mixing, water with high nutrient concentration is brought to the surface while phytoplankton populations are exposed to limited light. Thermal stratification develops with warming in the spring, although freshwater spreading over the sea surface from ice melt or rivers can initiate stratification earlier. During the summer months in the high latitudes, wind stress stirs the shallow mixed layer (50–100 m). In the subtropical gyre it is the seasonal cycle of turbulent energy exchange that controls the depths of the mixed layer: 20–40 m in the summer and 100–150 m in the winter. At midlatitudes the winter mixing may increase to 300 m if there is some heat loss associated with the wind stirring.

the m a r ine en v ironment  7 c­onditions. In some cases, single factors such as wind stress (Ueyama and Monger, 2005) or heat flux (Follows and Dutkiewicz, 2002) have been identified as driving variability, but more often a multitude of processes interact to affect phytoplankton, principally mediated through changes in mixing and stratification.

1.5  Long-term Variability of Ocean and Atmosphere Circulation

The maximum depth of winter mixing is a critical factor in setting the nutrients available to phytoplankton. The very deep winter mixed layer in the subpolar gyre ensures plentiful nutrients, although poor light conditions, while the relatively shallow winter mixed layer in the subtropics results in weak nutrient supply. These differences in maximum winter mixing determine that phytoplankton abundance is highest in spring at high latitudes, but in winter at low latitudes (Fig. 1.6). Because of the dependence of phytoplankton success on the relative amounts of stratification and mixing, large interannual variability in both the timing and magnitude of phytoplankton growth arises as a result of variability in meteorological and oceanographic 0

Chlorophyll

1

MLD

A

0.5

500

B

0

0.11 Chlorophyll

logarithmic colour scale in metres) and in summer (August, lower panel, contour intervals 5 m). Base of mixed layer defined as the depth where potential density is 0.125 kg m−3 greater than the surface value. (See Plate 3.) Climatological data from World Ocean Atlas 2009. Image created by Chongyuan Mao, NOC.

MLD

Figure 1.5: Mixed layer depth in winter (February, top panel,

It is important to recognize that the circulation of the North Atlantic as described in this chapter approximately represents the mean state, and that there is considerable variability on a multitude of time and space scales (Hughes et al., 2012). The North Atlantic Ocean and atmosphere act as a partnership, both influencing the behaviour of the other in ways that are not always understood. First we consider atmospheric variability that is important to the North Atlantic marine environment, and later in this section we examine evidence for long-term changes in ocean circulation. Variations in the strength and direction of the wind stress field can sometimes be simply expressed in terms of an index such as the North Atlantic Oscillation (NAO) (Hurrell and Deser, 2010). The NAO is a dominant pattern of variability that reflects changes in the strength and position of the gradient of atmospheric pressure between the subpolar low (typically found over Iceland) and the subtropical high (situated around the Azores). The characteristics of the pressure field determine the location of the storm track and how strong the winds are; both of these parameters considerably impact on the circulation of the ocean through momentum and heat transfer. The NAO index is derived from a statistical analysis of sea-level pressure, is most significant for marine conditions during the winter, and best indicates contrasting conditions where the strength of the high and low pressure centres change. Periods of high NAO index (deeper low pressure, stronger high pressure) are characterized by strong westerlies, a more northern storm track, warm and wet conditions in the eastern subpolar North Atlantic and the eastern subtropics, and cold and dry conditions in the western subpolar North Atlantic. Periods of low NAO index are characterized by fewer and weaker winter storms located further south, cold dry conditions in the eastern s­ ubpolar gyre, but

100 0

100

200 Day of year

300

0

100

200 Day of year

300

0.06

Figure 1.6:  Typical seasonal cycle of mixed layer depth (m; grey) and chlorophyll concentration (mg m−3; black) in A subpolar North Atlantic

and B subtropical North Atlantic. Note the different scales on the y-axes in both plots. Mixed layer depth (MLD) data are from optimally interpolated Argo float profiles (Gaillard et al., 2009) and chlorophyll data are from SeaWiFS satellite.

8 ecology mild conditions around Greenland, and cooler conditions in the subtropics. When the pressure centres shift location, the simple index becomes less useful; these alternative conditions are known as ‘blocking’ (high pressure over Europe and Scandinavia, low pressure over the Labrador Sea) and ‘Atlantic Ridge’ (strong high pressure over the subpolar region, also known as the East Atlantic Pattern). The blocking pattern effectively interrupts and shifts the Atlantic storm track and consequently winters with high incidence of blocking lead to a warmer and more saline subpolar gyre (Hakkinen et al., 2011). The phase of the NAO has an indirect effect on plankton populations through its influence on wind patterns. In positive NAO years, the subpolar gyre strengthens (Marshall et al., 2001) and eastward wind stress increases, resulting in deeper mixed layers. This leads to a delayed spring bloom and an overall reduction in phytoplankton (Henson et al., 2009). In subtropical regions, the winter mixed layer is shallower in positive NAO conditions, resulting in an earlier start to the bloom but lower phytoplankton abundance. In negative NAO years, conditions are approximately reversed and the subtropical gyre expands. A

large ‘transition zone’ that in positive NAO years experiences subpolar type conditions and has the corresponding phytoplankton response becomes more influenced by subtropical conditions. The transition zone at ~ 45–50 °N thus undergoes large interannual to decadal changes in phytoplankton abundance, which is reflected at higher trophic levels (Beaugrand et al., 2000; Greene and Pershing, 2000). Observations and model studies have shown that the physical environment changes significantly on timescales of years to decades, and these changes can profoundly impact ecosystems. The North Atlantic Ocean has a marked warming trend over many decades (a major component of anthropogenic climate change), but superimposed on that trend is multidecadal variability known as the Atlantic Multidecadal Oscillation (AMO, after Kerr,  2000) that includes periods of significant cooling. The AMO can be expressed as an index derived from mean SST with the long-term trend removed. The resulting pattern of variability has been shown to be coincident with other parameters including salinity (Reverdin,  2010), sea level, and sea-ice transport (Frankcombe et al., 2010), and ecological indicators

Box 1.1  North Atlantic climatic time series A 4.00

B

NAO

4.00

2.00

2.00

0.00

0.00

–2.00

NAO

–2.00 0.40

–4.00

0.40

–4.00

0.20

0.20

0.00 –0.20

AMO –0.40 1880 1900 1920 1940 1960 1980 2000 Year

0.00 10.00 7.50 5.00 2.50 0.00 –2.50 –5.00 –7.50 –10.00

AMO –0.20 –0.40

SPGI 1950 1960 1970 1980 1990 2000 2010 Year

Selected decadal indicator time series for the North Atlantic relevant to biological productivity (adapted from Hughes et al., 2012). A, Data period 1880–2009, with anomalies referenced to the long-term mean: the NAO and the AMO (v. 1 in black and v. 2 in grey), and the NAO. Thin black lines show the annual values, and thick lines are the 5-year running means. B, Shorter time series over the period 1950–2009, with anomalies referenced to the 1971–2000 mean: the AMO (v. 1 in black and v. 2 in grey), the NAO, and the SPGI. Data sources are: AMO v. 1, annual mean, NOAA using the Kaplan SST data set and removing the global climate signal as a linear trend; AMO v. 2, annual mean, from the Hadley Centre Sea Ice and Sea Surface Temperature (HadlSST) data set and removing the global climate signal as the mean global SST; NAO, December–March, SPGI with the black line representing the gyre index calculated from modelled data (1960–2005) and the grey line showing the gyre index obtained from altimetry observations (1992–2008).

the m a r ine en v ironment  9 such as salmon recruitment (Friedland et al., 2009), cod populations (Drinkwater, 2009), and coastal phytoplankton distribution (Dixon et al., 2009). The primary mechanism by which the AMO affects phytoplankton populations is via changes in SST (Martinez et  al.,  2009), which are proposed to trigger habitat switching in populations at the thermal limits of their geographic range (Edwards et al., 2013). The AMOC is predicted to slow in the long term as carbon dioxide concentrations rise (IPCC, 2013), and variations in the strength of the AMOC could dramatically alter the heat content and salinity of the North Atlantic. Measurements have not yet shown clear indications of long-term changes in the transport of overflow waters in the subpolar region (Olsen et al., 2008; Jochumsen et al., 2012), but recently a slow decline of the AMOC in the subtropical gyre has been detected (Smeed et al., 2014). Most ocean measurement time series are simply too short to detect multidecadal-scale changes. The subtropical AMOC measurements have, however, shown a wide range of total transport, and during periods of low overturning (e.g. as a response to reduced wind stress, or a changed distribution of wind stress curl) there is a reduction in heat transport from the subtropics to the subpolar region. Changes in the flow of water from the subtropical gyre affect

Time (years)

24 20 16 12 8 4 0

B

400

800

1200

1600

2000 0.3 0.2 0.1 0.0 –0.1 –0.2 –0.3

110 90

Average nitrate concentration (0–1 km), North Atlantic (mmol N m–3)

C

70 50

Freshwater forcing (Sv)

0

Plankton biomass North Atlantic (%)

A Atlantic overturning (Sv)

the circulation of the subpolar gyre; periods of high transport bring more subtropical water into the eastern subpolar regions making it warmer, more saline, and changing the nutrient concentrations (Hatun et al., 2005; Johnson et al., 2013; Desbruyeres et  al.,  2013). This change in heat and salt content propagates into the central subpolar gyre, and along the pathway of Atlantic waters into the deep Nordic Seas and Arctic Ocean (Holliday et al., 2008; Eldevik et al., 2009). The strength of circulation in the subpolar gyre is also influenced by the heat content and dynamic height of the centre of the gyre in the Labrador Sea; this varies on decadal scales and can be represented by a  subpolar gyre index (SPGI) (Hatun et  al.,  2005) (Box  1.1). During periods of high heat loss and deep convection in the Labrador Sea (e.g. as a response to the NAO or low incidence of atmospheric blocking) the sea surface height gradient across the gyre and the strength of circulation (and the index) increase (Hakkinen and Rhines, 2004). The possible interactions between the AMOC and plankton have so far only been explored in model simulations, as data records are insufficiently long to investigate the consequences of AMOC variability. Model studies suggest that if the AMOC slows, primary ­production will decrease strongly, as weaker mixing results in an increase in upper ocean nutrient limitation (Schmittner, 2005) (Fig. 1.7).

26 22 18 14 10

Figure 1.7:  Time series of Atlantic overturning, plankton biomass and nitrate concentration from model simulations (adapted from Schmittner, 2005). A, Freshwater forcing in the North Atlantic (dashed line, right scale) and Atlantic Multidecadal Oscillation (AMO) circulation (solid line, left scale); B, integrated biomass given as percentage changes with respect to year 0; C, average nitrate concentrations. The grey bars enclosing the ensemble mean (solid lines) denote the standard deviation of the different ecosystem models used and represent the uncertainties associated with the formulation and parameterizations of the ecosystem models. The evolution of the physical variables does not vary between the different experiments; hence panel A does not contain shadings.

10 ecology

1.6 Summary The main aim of this chapter has been to provide an overview of the regional circulation of the North Atlantic, and an introduction to the key physical features and processes that affect ecosystems, and especially plankton, via the availability of light and nutrients. There is a natural seasonal cycle in primary production driven by physical processes that determine the light and nutrient levels, but the pattern has strong regional variations. The variations are determined by persistent features on the basin scale (e.g. the main currents and mixed layer regimes of the subtropical and subpolar gyres), as well as transient mesoscale features such as eddies and meanders of fronts. The regional pattern of winter and summer mixed layer is a direct response to atmospheric circulation; a classic example is the contrast between the deep winter mixing in the subpolar gyre

driven by extreme heat loss from ocean to air, and the shallow winter mixing in the subtropical gyre supported by wind stirring only. However, over time there are significant variations in the location of currents and the extent of their eddy activity, or in the depth of winter mixing or the loci of mixing events. All of these examples are the response of the ocean to changes in atmospheric circulation and heat and momentum fluxes, and all significantly impact plankton abundance and distribution and subsequently higher trophic levels in the ecosystem. As well as stochastic variability in ocean and atmosphere circulation, there are sustained patterns of circulation that persist over years or decades, including the NAO and AMO. The physical processes within the coupled ocean-atmosphere system that force changes in the NAO and AMO are still under debate, but it is clear that there are significant biological responses to the changes in environmental conditions that they generate.

R efer ences Bacon, S., Myers, P., Rudels, B., and Sutherland, D.A. (2008). Accessing the inaccessible: buoyancy-driven coastal currents on the shelves of Greenland and eastern Canada. In: Dickson, R.R., Meincke, J., Rhines, P. (Eds.), Arctic-Subarctic ocean fluxes. Springer, Berlin, pp. 703–22. Beaugrand, G., Reid, P.C., Ibanez, F., and Planque, B. (2000). Biodiversity of North Atlantic and North Sea calanoid copepods. Marine Ecology Progress Series 204, 299–303. Brambilla, E., and Talley, L.D. (2008). Subpolar Mode Water in the northeastern Atlantic: 1. Averaged properties and mean circulation. Journal of Geophysical Research 113 (C4), doi:10.1029/2006JC004062. Crawford, W.R., Brickley, P.J., and Thomas, A.C. (2007). Mesoscale eddies dominate surface phytoplankton in northern Gulf of Alaska. Progress in Oceanography 75 (2), 287–303. Desbruyeres, D., Thierry, V., and Mercier, H. (2013). Simulated decadal variability of the meridional overturning circulation across the A25-Ovide section. Journal of Geophysical Research 118, 462–75. Dickson, R.R., and Brown, J. (1994). The production of North Atlantic Deep Water: sources, rates and pathways. Journal of Geophysical Research 99 (C6), 12319–41. Dixon, L.K., Vargo, G.A., Johansson, J.O.R., Montgomery, R., and Neely, M.B. (2009). Trends and explanatory variables for the major phytoplankton groups of two southwestern Florida estuaries, USA. Journal of Sea Research 61 (1–2), 95–102. Drinkwater, K. (2009). Comparison of the response of Atlantic cod (Gadus morhua) in the high-latitude regions of the North Atlantic during the warm periods of the 1920s–1960s and the 1990s–2000s. Deep-Sea Research Part II: Topical Studies in Oceanography 56 (21–22), 2087–96.

Edwards, M., Beaugrand, G., Hays, G.C., Koslow, J.A., and Richardson, A.J. (2010). Multi-decadal oceanic ecological datasets and their application in marine policy and management. Trends in Ecology and Evolution, 25: 602–10. Edwards, M., Beaugrand, G., Helaouet, P., Alheit, J., and Coombs, S. (2013). Marine Ecosystem Response to the Atlantic Multidecadal Oscillation. PloS One 8 (2), e57212. Eldevik, T., Nilsen, J.E.O., Iovino, D., et  al. (2009). Observed sources and variability of Nordic seas overflow. Nature Geoscience 2 (6), 405–9. Follows, M., and Dutkiewicz, S. (2002). Meteorological modulation of the North Atlantic spring bloom. Deep-Sea Research Part Ii-Topical Studies in Oceanography 49 (1–3), 321–44. Frankcombe, L.M., von der Heydt, A., and Dijkstra, H.A. (2010). North Atlantic Multidecadal Climate Variability: An Investigation of Dominant Time Scales and Processes. Journal of Climate 23 (13), 3626–38. Friedland, K.D., MacLean, J.C., Hansen, L.P., Peyronnet, A.J., Karlsson, L., et  al. (2009). The recruitment of Atlantic salmon in Europe. ICES Journal of Marine Science 66 (2), 289–304. Gaillard, F., Autret, E., Thierry, V., Galaup, P., et  al. (2009). Quality control of large Argo datasets. Journal of Atmospheric and Oceanic Technology 26, 337–351. Gammelsrod, T., Leikvin, O., Lien, V., et al. (2009). Mass and heat transports in the NE Barents Sea: Observations and models. Journal of Marine Systems 75 (1–2), 56–69. Greene, C.H., and Pershing, A.J. (2000). The response of Calanus finmarchicus populations to climate variability in the Northwest Atlantic: basin-scale forcing associated with the North Atlantic Oscillation. ICES Journal of Marine Science 57 (6), 1536–44.

the m a r ine en v ironment  11 Haine, T.W.N., Boning, C.W., Brandt, P., et  al. (2008). North Atlantic Deep Water formation in the Labrador Sea, recirculation through the subpolar gyre, and discharge to the subtropics. In: Dickson, R.R., Meincke, J., Rhines, P. (Eds.), Arctic-Subarctic ocean fluxes. Springer, Berlin. Hakkinen, S. and Rhines, P.B. (2004). Decline of subpolar North Atlantic circulation during the 1990s. Science 304 (5670), 555–9. Hakkinen, S., Rhines, P., and Worthen, D. (2011). Atmospheric blocking and Atlantic multidecadal ocean variability. Science 334, 655–9. Hansen, B., and Østerhus, S. (2000). North Atlantic—Nordic Seas Exchanges. Progress in Oceanography 45, 109–208. Hansen, B., Osterhus, S., Turrell, B., et al. (2008). The inflow of Atlantic water, heat and salt to the Nordic Seas across the Greenland-Scotland Ridge. In: Dickson, R.R., Meincke, J., Rhines, P. (Eds.), Arctic-Subarctic ocean fluxes. Springer, Berlin, pp. 15–44. Hátún, H., Sando, A.B., Drange, H., Hansen, B., and Valdimarsson, H. (2005). Influence of the Atlantic subpolar gyre on the thermohaline circulation. Science 309, 1841–4. Henson, S.A., Dunne, J.P., and Sarmiento, J.L. (2009). Decadal variability in North Atlantic phytoplankton blooms. Journal of Geophysical Research-Oceans 114, C04013. Holliday, N.P., Hughes, S.L., Bacon, S., et al. (2008). Reversal of the 1960s to 1990s freshening trend in the northeast North Atlantic and Nordic Seas. Geophysical Research Letters 35 (3), L03614. Holliday, N.P., Waniek, J.J., Davidson, R., et al. (2006). Largescale physical controls on phytoplankton growth in the Irminger Sea Part I: Hydrographic zones, mixing and stratification. Journal of Marine Systems 59, 201–18. Hughes, S.L., Holliday, N.P., Gaillard, F., and Hydrography, I.W.G.o.O. (2012). Variability in the ICES/NAFO region between 1950 and 2009: observations from the ICES Report on Ocean Climate. ICES Journal of Marine Science 69 (5), 706–19. Hurrell, J.W., and Deser, C. (2010). North Atlantic climate variability: The role of the North Atlantic Oscillation. Journal of Marine Systems 79 (3–4), 230–230. IPCC (2013). Climate Change 2013: The Physical Science Basis. Contribution of Working Group I to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change. Jochumsen, K., Quadfasel, D., Valdimarsson, H., and Jonsson, S. (2012). Variability of the Denmark Strait overflow: moored time series from 1996–2011. Journal of Geophysical Research 117, C12003. Johnson, C., Inall, M.E., and Hakkinen, S. (2013). Declining nutrient concentrations in the northeast Atlantic as a result of a weakening Subpolar Gyre. Deep Sea Research I 82, 95–107. Kerr, R.A. (2000). A North Atlantic climate pacemaker for the centuries. Science 288 (5473), 1984–6.

Lalli, C., Parsons, T. (1997). Biological Oceanography: An Introduction. Open University, Elsevier. Levy, M., Klein, P., and Treguier, A.M. (2001). Impact of submesoscale physics on production and subduction of phytoplankton in an oligotrophic regime. Journal of Marine Research 59 (4), 535–65. Longhurst, A. (2007). Ecological Geography of the sea. Academic Press. Mahadevan, A., Thomas, L.N., and Tandon, A. (2008). Comment on ‘Eddy/wind interactions stimulate extraordinary mid-ocean plankton blooms’. Science 320 (5875), 448b. Marshall, J., Johnson, H., and Goodman, J. (2001). A study of the interaction of the North Atlantic oscillation with ocean circulation. Journal of Climate 14 (7), 1399–1421. Martin, A.P., Wade, I.P., Richards, K.J., and Heywood, K.J. (1998). The PRIME eddy. Journal of Marine Research 56 (2), 439–62. Martinez, E., Antoine, D., D’Ortenzio, F., and Gentili, B. (2009). Climate-Driven Basin-Scale Decadal Oscillations of Oceanic Phytoplankton. Science 326 (5957), 1253–6. McGillicuddy, D.J., Robinson, A.R., Siegel, D.A., et  al. (1998). Influence of mesoscale eddies on new production in the Sargasso Sea. Nature 394 (6690), 263–6. Montegut, C.D., Madec, G., Fischer, A.S., Lazar, A., and Iudicone, D. (2004). Mixed layer depth over the global ocean: An examination of profile data and a profile-based climatology. Journal of Geophysical Research-Oceans 109 (C12), C12003. Olsen, S.M., Hansen, B., Quadfasel, D., and Osterhus, S. (2008). Observed and modelled stability of overflow across the Greenland-Scotland ridge. Nature 455 (7212), 519–22. Potter, R.A., and Lozier, M.S. (2004). On the warming and salinification of the Mediterranean outflow waters in the North Atlantic. Geophysical Research Letters 31, L01202. Reverdin, G. (2010). North Atlantic Subpolar Gyre Surface Variability (1895–2009). Journal of Climate 23 (17), 4571–84. Rossby, T. (1999). On gyre interactions. Deep Sea Research II 46 (1–2), 139–64. Sarmiento, J.L., Slater, R., Barber, R., et  al. (2004). Response of  ocean ecosystems to climate warming. Global Biogeochemical Cycles 18 (3), Gb3003. Schmittner, A. (2005). Decline of the marine ecosystem caused by a reduction in the Atlantic overturning circulation. Nature 434, 628–33. Smeed, D.A., McCarthy, G., Cunningham, S.A., et  al. (2014). Observed decline of the Atlantic Meridional Overturning Circulation 2004–2012. Ocean Science, 10 (1), 29–38. Sverdrup, H.V. (1953). On conditions for the vernal blooming of phytoplankton. Journal of Council 18, 287–95. Ueyama, R., and Monger, B.C. (2005). Wind-induced modulation of seasonal phytoplankton blooms in the North Atlantic derived from satellite observations. Limnology and Oceanography 50 (6), 1820–9.

CH A PTER 2

PLA NKTON BIODIVER SITY A ND BIOGEOGR A PH Y Gr egory Beaugr a nd

2.1 Introduction

Holopelagic (oceanic)

Neritopelagic (neritic) Euphotic zone

Dysphotic zone

Mesopelagic zone

1000 m

Depth

Bathypelagic zone

4000 m Ab

5000 m

yss

yss

al

ob

en

pla

in

thi

c Abyssopelagic zone

ench

11000 m

Hadobenthic

nic tr

Ocea

6000 m

Ab

Aphotic zone

ic

l slope

nenta

3000 m

obenth

Conti

2000 m

Epipelagic zone

Bathy

The marine realm covers an enormous area (~ 360 million km ), which represents ~ 70% of the Earth’s surface (Lasserre, 1994; Pauly and Christensen, 1995). The oceanic part of the hydrosphere has an average depth of 3800 m, and ~ 88% of the oceans are deeper than 1000  m (Dinter,  2001). Some areas, called trenches, can be as deep as 11 km (e.g. the Mariana Trench with a depth of 11 033 m). In contrast to the land, the marine environment is three-dimensional, constituted of ~ 1.26 billion km3 of seawater, and this environment is highly structured in all dimensions (Angel,  1993; Haury and McGowan,  1998). Hydrodynamics influence light penetration, the distribution of all forms of energy (exosomatic and endosomatic), nutrients and oxygen. Most oceanic regions are aphotic, which means that only a weak amount of energy exists to sustain biodiversity. With the exception of hydrothermal vents and cold seeps (also called cold vents), where the food web is mainly based on chemosynthesis, deep areas depend on carbon exportation ­taking place from the euphotic zone. Plankton have virtually colonized all biotopes in both neritic and oceanic regions and from the epipelagic (0–200 m) to the hadopelagic zone (Fig.  2.1). Relatively little is known about deeper pelagic realms such as the mesopelagic and the bathypelagic zones. Scientists have only sparse information on these domains. In these deeper waters, the energy on which the food web is based generally comes from organic materials (particulate organic carbon) exported from the epipelagic zone. Haury et al. (1978), by the use of a Stommel (1963) three-­ dimensional time-space plot estimated the spatial and temporal structure of zooplankton biomass variability (Fig.  2.2). At all spatial scales, physical features often represented important ­factors involved in the control of spatial and temporal patterns. They were, however, aware of the limitations of their diagram. The statistical and spectral properties of Stommel-type ­diagrams do not depend on position or time. Consequently, variance forced by different processes can be confused in the diagram and important geographical information may be lost.

Continental shelf

2

Hadopelagic zone

FIGURE 2.1: A schematic cross section of the ocean from the shoreline to the oceanic trench, indicating the main pelagic and benthic habitats. Seamounts and oceanic ridges are not indicated. These topographical features can, however, be inhabited by a rich biodiversity. The depth of both the euphotic and the dysphotic zones vary latitudinally and as a function of plankton concentration and suspended sediments. From Beaugrand (2015).

Figure 2.3A shows the respective influence of diel, seasonal and year-to-year variability in biodiversity in the Bay of Biscay, the northern part of the North Atlantic Drift Province (NADR) and the North Sea. The figure suggests that the Stommel-type diagram of biomass variability may indeed greatly vary spatially. For example, diel variability in calanoid copepod biodiversity is extremely high in the southern part of the NADR, while it is reduced in its north-eastern extremity and over the continental shelf. Year-to-year variability in biodiversity is also high to the north of the NADR and in the North Sea. It is not so in the Bay of Biscay. Figure  2.3B shows how the spatial variability in

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

pla nkton biodi v er sit y a nd biogeogr a ph y   13 Glacial/ interglacial cycles Diel Annual vertical cycles migration

10 000 km 1000 km I100 km H J D F G C 10 km K E B 100 m 1m

A

Space

1 cm

Time

y 000 10 0 y 100 tury Cen

Space

ute

Min

r Yea th n Mo ek Wey Da ur Ho

Biomass variability

A B C D E F G H I J K

: micropatches : swarms : upwelling : eddies and rings : island effects : ‘EI Niño’ events : small oceanic basins : Biogeographic provinces : currents and fronts (length) : currents (width) : oceanic fronts (width)

Time

FIGURE 2.2:  Stommel-type diagram showing the variability in biomass on both spatial and temporal scales. Redrawn from Haury et al. (1978).

A

30 25 20 15 10 5 0

Diel variability Seasonal variability Interannual variability

a Se th N ra l nt

Ce

N or th

er

n

pa

rt

or

of

Ba yo

N

fB

AD

isc

R

ay

Temporal variability

% 35

B

50° N

1200

1000

800 40° N

600

distances (in km)

60° N

400 FIGURE 2.3:  Spatial and temporal variability in calanoid copepod biodiversity in the North Atlantic Ocean. A, Quantification of diel, seasonal and year-to-year variability in the Bay of Biscay, the northern part of the North Atlantic Drift Province (NADR) and in the North Sea. B, Spatial changes in the scale at which biodiversity varies in the North Atlantic Ocean. Blue colour suggests that variability varies at a small scale, whereas red colour indicates that biodiversity fluctuates at a large scale. The main surface currents are indicated. From Beaugrand et al. (2003). (See Plate 4.)

14 ecology

virioplankton bacterioplankton phytoplankton

Abundance

calanoid copepod biodiversity varies in space. In the path of the North Atlantic Current, biodiversity varies at small scales, whereas in the subarctic gyre biodiversity changes at the large scale. Planktonic variability is therefore highly modulated in space and time. This chapter briefly presents the biodiversity and main biogeographic patterns of marine plankton, the causes of such patterns, as well as factors that influence spatial and temporal plankton distribution.

zooplankton metazooplankton

2.2  Marine Biodiversity The oceans hold between ~ 200 000 and ~ 250 000 already inventoried species (Mora et  al.,  2011; Boeuf,  2014). Recent studies have estimated that the oceans may contain between 0.5 and 2.2 million species (Mora et al., 2011; Appeltans et al., 2012), which suggests that we have described between 9.1% and 50% of all marine species so far. Although the oceans contain less species than on land (Mora et al., 2011; Boeuf, 2014), they have a much greater biodiversity at a higher taxonomic level, probably because life rapidly arose in the marine realm after Earth’s formation 4.56 billion years ago (Allègre et  al.,  1995). Traces of organic carbon (biogenic carbon detritus), possibly originating from plankton organisms, have been detected in sedimentary rock of an age of at least 3.7 billion years (Rosing, 1999). Marine metazoan biodiversity appeared ~  700  million years ago and radiated during the Ediacaran (578–542  Ma) and Cambrian (542–485  Ma) periods (Shu et  al.,  2014; Beaugrand,  2015). Terrestrial life appeared only 440 million years ago after the constitution of the ozone layer. It is therefore not surprising that at the phylum level, marine metazoan biodiversity is much greater than terrestrial biodiversity. Out of the total number of recorded metazoan phyla (34), 33 are present in the sea, whereas only 17 occur in the terrestrial realm, with only one endemic. Massive differences exist between the pelagic and the benthic realms (Beaugrand, 2015). All phyla are present in the benthos, whereas only 15 are detected in the pelagic realm. However, classification is changing, and those estimations vary from one author to another (Herring, 2002; Boeuf, 2014). Plankton contains many taxonomic groups that range from 10 nm (e.g. marine viruses) to ~ 2 m for the largest metazooplanktonic species (Fig. 2.4). For example, the Nomura’s jellyfish Nemopilema nomurai can reach 2 m and weigh up to 200 kg (Kirby, 2010). Despite its relatively low biodiversity compared to insects, plankton play an important role in some biogeochemical cycles and in the trophodynamics of marine ecosystems. Phytoplankton creates ~ 50% of the oxygen produced annually by photosynthesis at a global scale and plays a key role in the regulation of the Earth’s climate through its influence on the global carbon cycle, a process termed the biological carbon pump. Plankton are at the base of the ocean’s food chain. The ocean’s total primary production is estimated to be ~ 48.5 Pg (1 Pg = 1015 g) carbon each year (Field et al., 1998). The carbon

nekton

10–8

10–7

10–6

10–5

10–4

10–3

10–2

10–1

100

101

Linear dimension (m) FIGURE 2.4:  Abundance and size of plankton. Size and abundance of the nekton is also indicated for comparison. Adapted from Sieburth et al., (1978).

produced by phytoplankton is transferred to the zooplankton, which are the basis of a global production of 240 million tonnes of fish. Of this, ~ 80 million tonnes of fish are harvested annually by humans. Eukaryotic plankton includes 11 200 catalogued species, of which ~ 4350 are phytoplankton, ~ 1350 are protozooplankton (heterotrophic protists) and ~ 5500 are metazooplankton (holoplanktonic animals) (Sournia et  al.,  1991; Boltovskoy et al., 2005; Wiebe et al., 2010). Therefore, about 7000 species of holozooplankton distributed in 15 phyla have been described (Bucklin et  al.,  2010). Sournia (1994) estimated that between 3500 and 4500 phytoplankton species are described in the sea, which can be compared to the 215 000 plant species described in the terrestrial realm (Mora et al., 2011). A total of 280 species of coccolithophores has been described. The phylum Chaetognatha, exclusively marine, contains 100 species, 20 of which are benthic coastal species (Pierrot-Bults, 1997). PierrotBults further estimated that 250 pteropod molluscs, 400 pelagic cephalopods, 400 pelagic mysids (mostly neritic) and 86 euphausiids (mainly oceanic) have been described so far. Copepoda is the most diverse taxonomic group, and ~ 10 000 marine species have been catalogued (Appeltans et al., 2012), of which 2200 are found in the pelagic zone (Beaugrand, 2015). By analysing 18S ribosomal DNA sequences from 334 plankton size-fractionated samples collected at 47 sampling sites during the Tara Oceans expedition, eukaryotic ribosomal diversity was assessed as ~ 150 000 operational taxonomic units (OTUs) (de Vargas et  al.,  2015). This estimate suggests that the 11 200 inventoried eukaryotic plankton species may only represent less than 7.5% of the total eukaryotic plankton biodiversity. Protists accounted for more than 85% of all OTUs. Most of the unknown species may be found in the pico-nano-plankton fraction. This group (between 0.8  µm and 5  µm; OTU Shannon diversity ≈  4.5) dominated all fractions and was followed by

pla nkton biodi v er sit y a nd biogeogr a ph y   15 the nanoplankton (5–20 µm; OTU Shannon diversity ≈ 2.5), the microplankton (20–180 µm; OTU Shannon diversity ≈ 2.2) and the mesoplankton (180–2000 µm; OTU Shannon diversity ≈ 1.9) fractions.

2.3  Main Characteristics of the Marine Pelagic Realm Although some studies suggest that the interactome (i.e. the whole set of species interactions in a particular community) may explain more variance than environmental factors at both local and large scales (Lima-Mendez et  al.,  2015), large-scale patterns in species distribution and biodiversity suggest this is unlikely to be true. As suggested by the Stommel-type diagram showing biomass variability as a function of space and time scales (Fig. 2.2), plankton are influenced by a large number of environmental factors and as a result are not distributed randomly in the oceans and seas. Plankton biodiversity is constrained by hydroclimatic parameters such as temperature, bathymetry and oceanic surface currents or large-scale ­hydrodynamic features such as the subarctic gyre (Ruddiman, 1969; Rutherford et  al., 1999; Rombouts et  al., 2010; Sunagawa et  al., 2015). Wolfgang Dinter (2001) made a list of the environmental factors that control the geographical range of marine species. The above environmental factors were thought to be the most important factors in explaining the geographical distribution of species. The author also added water quality (e.g. nutrients, salinity and turbidity) that can influence pelagic ecosystems over large areas (e.g. high nutrient low chlorophyll (HNLC) areas). At more local scales, he listed factors such as tidal currents, types of substratum, the harbour effect and freshwater inputs that can be relatively important.

2.3.1  Climatic factors Many climatic parameters influence biodiversity. Atmospheric circulation has a strong effect on surface currents (e.g. Atlantic Meridional Overturning Circulation), upwelling and the structure of the water column (thermocline, halocline and the resulting pycnocline). Wind intensity also affects prey-predator encounter rates (Rothschild and Osborn, 1988) by its effects on oceanic turbulence, and nutrient supply rates by its effects on vertical mixing (Longhurst, 2007). Wind direction, by its control on the dispersal of meroplanktonic species, might strongly affect recruitment of some benthic organisms ( Jolly et al., 2009). Incident solar radiation is also important because it provides the energy that influences sea surface temperature and the structure of the water column. Photosynthetically active radiation (PAR), solar radiation of wavelength 400–700 nm, is also relevant because it represents the level of energy that can be assimilated by photosynthetic organisms (Asrar et  al.,  1989). PAR regulates both the composition and evolution of marine ecosystems, influencing the growth of phytoplankton and in turn the development of zooplankton and fish. This factor is also strongly

influenced by cloudiness. Temperature is a master parameter that influences the spatial distribution of species (Rutherford et  al.,  1999; Rombouts et  al.,  2010). The parameter is largely influenced by atmospheric circulation, incident solar radiation, oceanic circulation and cloudiness in the sky (Beaugrand 2015). In contrast to the terrestrial realm, precipitation or any measure of water availability has in general a small influence in the marine environment, although the parameter may become important in some coastal regions (Goberville et al., 2011).

2.3.2  Marine environmental factors Many environmental factors influence the arrangement of life in the oceans and seas (Fig. 2.5). Dissolved oxygen must remain high enough to support respiration (Pörtner, 2001; Goberville et al., 2010) (Fig. 2.5A). Mixed layer depth is an important parameter for phytoplankton production and controls the spatial distribution of many plankton species (Sverdrup,  1953; Longhurst,  2007) (Fig.  2.5B). Both bathymetry (Fig. 2.5C) and local spatial variability in bathymetry are key determinants of the marine pelagic biodiversity (Helaouët and Beaugrand, 2007). At the bottom, the variation in the bathymetry is also responsible for the creation of very distinct habitats (Helaouët and Beaugrand, 2007). Bathymetry influences light penetration, all parameters enunciated above, including pressure that directly influences plant and animal anatomy and their life history traits. Salinity is also an important factor. Species dispersal can be influenced by salinity through the ability of marine species to osmoregulate (SchmidtNielsen, 1990). Salinity is high over subtropical gyres and in the Mediterranean Sea (Fig. 2.5D). Sediment type (e.g. mud, sand, gravel) is also an important parameter for benthic biodiversity. Oceanic pH influences calcifying organisms such as coccolithophores, foraminifers, corals and pteropods (Orr et al., 2005; Kroeker et al., 2010; Rombouts et al., 2012). Although apparently uniform, the oceans are therefore highly heterogeneous at all spatial scales, creating a variety of habitats that also explains why, locally, marine biodiversity is so diverse.

2.3.3 Macronutrients Macronutrients are often the limiting factor for primary production (Behrenfeld, 2010). Figure 2.6 shows the spatial distribution of some macronutrients (nitrates, phosphates and silicates). Large concentrations of macronutrients are found in extratropical regions (especially in polar regions) and to a lesser extent along the equator. Subtropical gyres are poor in macronutrients.

2.3.4  High nutrient low chlorophyll areas HNLC areas are oceanic regions where phytoplankton standing stock is low despite high concentrations of macronutrients (e.g.  nitrate, phosphate, silicate). Spring blooms are never

16 ecology

FIGURE 2.6:  Concentration in A, nitrate; B, phosphate; and C, silicate

in seawater. Data from the World Ocean Atlas. From Beaugrand (2015). (See Plate 6.)

FIGURE 2.5:  Concentration in A, dissolved dioxygen; B, mixed layer depth (MLD) (density); C, bathymetry; and D, sea surface salinity. Dissolved dioxygen and sea surface salinity are from the World Ocean Atlas. Bathymetry data are from the General Bathymetric Chart of the Oceans (GEBCO). MLD (density) data originated from de Boyer Montégut et al., (2004). (See Plate 5.)

observed. These regions, which cover about 20% of the world’s oceans, are mainly the equatorial Pacific Ocean, the subarctic Pacific Ocean and the Southern Ocean (Fig. 2.7). Small phytoplankton species such as pico- and nano-phytoplankton are more common than diatoms in HNLC areas. Two hypotheses have been proposed to explain this pattern. The first, the iron hypothesis, proposes that macronutrients are never depleted because other micronutrients (e.g. iron) control phytoplankton proliferation (Martin, 1991). Concentrations of

bioavailable iron in these regions are significantly smaller than in other oceanic provinces. The second hypothesis, the grazing control hypothesis, stipulates that phytoplankton cannot reach high standing stocks because they are limited by grazing (Frost, 1987). Although grazing may be important in some regions, iron plays a very important role. Iron is mainly abundant in coastal regions. In contrast, open-ocean waters have much less iron and are said to be infertile in this respect (Martin, 1991). Iron is delivered to the pelagic oceans by dust storms from arid lands, which contain between 3% and 5% iron. Although ocean iron experiments have given contrasting results, some have shown that the addition of iron in some HNLC regions stimulated phytoplankton and rapidly diminished nitrate concentrations. During IronEx I, an area of 64 km² in the Eastern Equatorial Pacific was fertilized with an iron ­sulphate solution (Martin et  al.,  1994). The amount of iron increased from 0.06 nM to 4 nM and was at the origin of a doubling in chlorophyll concentration at the surface. However, the biological response was short lived and after 5  days, phytoplankton growth returned to the values observed before the beginning of the experiment because iron subduction took place at a depth where light limits phytoplankton proliferation.

pla nkton biodi v er sit y a nd biogeogr a ph y   17 January

1.0 0.8 0.6

0.2 0.0

July

1.0 0.8

Ratio standardized nitrate / (normalized chlorophyll +1)

0.4

0.6 0.4 0.2 0.0 FIGURE 2.7:  High nutrient low chlorophyll areas as revealed by the

ratio of standardized nitrates (between 0 and 1) to (standardized chlorophyll (between 0 and 1) +1) in January (top) and July (bottom). Values of chlorophyll greater or equal to 2  mg  m−3 and of nitrates greater or equal to 20 µmoles L−1 were fixed to 1 in the standardized values of nitrates and chlorophyll, respectively. Black areas are regions where the ratio is 0. Data from SeaWIFS and the World Ocean Database. The location of some ocean iron experiments is indicated on the map. IRONEX I and II, IRON fertilization EXperiment (1993 and 1995); SOIREE, Southern Ocean Iron RElease Experiment (1999); SEEDS, The Subarctic Pacific Iron Experiment for Ecosystem Dynamics Study (2001); SERIES, Subarctic Ecosystem Response to Iron Enrichment (2002); EIFEX, European Iron Fertilization Experiment (2004). From Beaugrand (2015). (See Plate 7.)

This experiment showed that iron is therefore an important micronutrient for phytoplankton growth and photosynthesis.

2.4  Plankton Distribution and Partition of the Pelagic Realm Plankton follow most of the main divisions of the pelagic realm. Many partitions of the pelagic realm have been proposed and it is not possible to enumerate all of them. Mark Spalding and colleagues listed the work of Forbes (1856), Ekman (1953), Hedgpeth (1957), Briggs (1974) and Bailey (1998) (Spalding et al., 2007 and references therein). Temperature variability over large timescales explained well the partition of Briggs (1974). Briggs’ partition also considered endemism, each province being based on 10% endemism. An improvement of this partition has been recently proposed (Fig. 2.8). The marine ecosphere was divided into three main ecomes: (1) cold regions (Arctic and Antarctic), (2) cold-temperate regions, and (3) warm-temperate regions. Provinces were also identified.

The main difficulty in partitioning the marine plankton biosphere is related to the dynamic movement of water masses and the locations of surface features, which are influenced by atmospheric conditions. This difficulty led the biogeographer van der Spoel (1994) to separate the biotope of pelagic ecosystems into two components: (1) a stable-biotope component (geographically stable) in which a primary related community lives, and (2) a substrate-biotope component (depending on water mass) characterized by a secondary related community (mixed primary community (Beklemishev,  1961)). An ecosystem is mainly characterized by a primary related community linked to a stable-biotope component, whereas an ecotone is more distinguished by a secondary related community depending on water masses. It is also known that an ecotone can also be characterized by its own biological composition (Ramade, 1994; Beaugrand et al., 2002a; Frontier et al., 2004). The distinction van der Spoel made is fundamental to the correct understanding of how plankton biodiversity is spatially organized in the oceans and seas. Biological partition is rarely achievable with great precision at a global scale because spatial plankton distribution is poorly known. This is perhaps why some authors have proposed partitions based on biogeochemical parameters. The development of satellite technology and the globalization of environmental data sets have enabled the establishment of global biogeography. Figure  2.9A shows the division of the marine ecosphere into biomes and provinces by Alan Longhurst (2007). Four primary biomes (Polar, Westerlies, Trades, and Coastal) and 56 secondary provinces were identified. This partition of the marine ecosphere by Longhurst was mainly based on the characterization of the seasonal cycle of primary production (Longhurst, 2007). Variables used to establish the partition were chlorophyll concentration, mixed layer depth, nutrients, the Brunt-Vaisala frequency, the Rossby radius of internal deformation, photic depth, algal biomass and primary production. These variables allowed the identification of a number of ecological situations: (1) polar irradiance-limited production peak, (2) nutrient-limited spring bloom, (3) winter-spring production with nutrient limitation, (4) small amplitude response to trade wind seasonality, (5) large amplitude response to monsoon reversal, and (6) various responses to topography and wind stress on continental shelves, including coastal upwelling (Reygondeau et al., 2013). Using four parameters (bathymetry, chlorophyll-a concentration, surface temperature and salinity), Reygondeau and co-workers (2013) applied a procedure based on the Non-Parametric Probabilistic Ecological Niche model (Beaugrand et al., 2011) to propose a more dynamical partition of the biogeochemical provinces of Longhurst. The average contour of the provinces was in general in good agreement with those originally proposed by Longhurst (Figure 2.9B). Basing pelagic biogeography on only a few biogeochemical parameters may lead to an overly simplistic scheme because most species are very sensitive to many environmental factors. Biogeographical partitions based on species distribution have been proposed by many authors. Mary Somerville (1780–1872)

18 ecology cold-temperate provinces warm-temperate provinces warm oceans

120°

60°

0°

60°

120°

60°

60°

30°

30° WA EP

0°

0° EA IWP 30°

30°

60°

60° 0 km 120°

60°

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60°

3500

7000

120°

FIGURE 2.8:  The biogeographic division of the marine ecosphere proposed by Briggs and Bowen and based mainly on fish. Cold regions (Arctic and Antarctic) are in white, cold-temperate regions are in dark blue and warm-temperate provinces are in medium blue. WA, Western Atlantic; EA, Lusitania, Black Sea, Caspian, Aral and Benguela provinces in the East Atlantic; IWP, Mediterranean; Sino-Japanese, Auckland, Kermadec, South-eastern Australian and South-western Australian provinces in the Indo-Pacific; EP, California and Peru-Chilean provinces in the East Pacific. Photo courtesy of Briggs and Bowen (2012).

in her book about physical geography divided the marine ecosphere into homozoic zones. Based on Mollusca, Edward Forbes (1815–1854) established nine homozoic zones and related them mainly to marine isotherms. It is unfortunate, however, that the spatial distribution of most plankton species (especially tropical species) remains poorly identified, because such identification would allow a better partition of the biosphere. Pelagic species can be stenograph (i.e. local spatial distribution; ecotone species) or eurygraph (i.e. large spatial distribution) due to the large clines in environmental conditions and the absence of geographical barriers. Because their spatial distribution integrates many environmental parameters, they may be more powerful in partitioning the ecosphere (Fig. 2.10). The figure shows different types of spatial distribution of copepods in the North Atlantic Ocean. Unfortunately, the area under investigation cut the spatial distribution of some species. While we can only see the southern edge of the spatial distribution of Arctic and subarctic species, we can only visualize the northern edge of warm-temperate and subtropical ­species in the figure. Despite this limitation, the figure shows that some species have large (the group Para-Pseudocalanus spp.) or more restricted (Candacia armata) distributional range. Some are cold-water indicators (e.g. the subarctic species Paraeuchaeta norvegica), some are FIGURE 2.9:  Biogeography of the global ocean (i.e. biogeochemical warm-water indicators (e.g. the genus Clausocalanus and provinces) A, proposed by Longhurst (2007) and B, recalculated by a Euchaeta marina), while others are oceanic or pseudo-oceanic numerical procedure from January 1998 to December 2007. From (i.e. occur in both neritic and oceanic regions but preferentially along shelf edges). Metridia lucens has a higher abundance in the Reygondeau et al. (2013). (See Plate 8.)

pla nkton biodi v er sit y a nd biogeogr a ph y   19 Para-Pseudocalanus

Pareuchaeta norvegica 0.2

2

0.15

1.5

0.1

1

0.05

0.5

0 Neocalanus gracilis

0 Metridia lucens

0.04 0.03

0.15

0.02

0.1

0.01

0.05

0 Euchaeta marina

0.2

0 –3

x 10 3

Clausocalanus spp. 1

2

0 Calanus glacialis

0.2 0.15 0.1 0.05 0

Abundance (log10(x+1))

1

0.5 0 Candacia armata

0.1

0.05 0

FIGURE 2.10:  Mean spatial distribution of some key species or taxa in the North Atlantic Ocean. Modified from Beaugrand et al. (2003). (See Plate 9.)

ecotone located in the extension of the Gulf Stream and the North Atlantic Current to the west of the British Isles. This copepod is qualified of mixed-water species because it is found at the boundary between temperate and subarctic waters. The spatial distribution of those species shows that the NADR (sensu Longhurst; see Figure 2.9A) remains highly heterogeneous. Many marine partitions have been proposed. Developments of remote sensing and large-scale ship-based surveys have allowed a better demarcation of the biomes occupied by various taxonomic groups such as coccolithophores (Merico et al., 2003), nitrogen fixers (Westberry and Siegel, 2006) and picocyanobacteria ( Johnson et al., 2006).

2.5  Spatial Patterns in Plankton Biodiversity In addition to the main global pelagic divisions, some biodiversity patterns have been observed for a long time. The latitudinal gradient in species diversity is the tendency for biodiversity to be higher towards the equator. Turner mentioned that this gradient is among the oldest observed ecogeographic patterns, already pointed out by Forster in 1778 and von Humbold in 1808 (Turner and Hawkins, 2004). In the marine realm, the pattern has been documented for many plankton species such as ostracods, euphausiids, foraminifers, decapods and copepods, and for

fish (Stehli et al., 1969; Reid et al., 1976; Angel, 1993; PierrotBults, 1997; Rombouts et al., 2009; Beaugrand et al., 2013). This pattern does not appear to be a perfect latitudinal cline, however. For some taxonomic groups, rapid and pronounced spatial changes occur at the middle latitudes (Beaugrand and Ibañez, 2002; Rombouts et al., 2009). For example, at a global scale, Rutherford and colleagues (1999) revealed that foraminifera biodiversity showed a maximum in middle latitudes, a minimum in high latitudes and was intermediate at the equator. This pattern is clearly observed when foraminifera biodiversity is meridionally averaged (Fig. 2.11). Such a hump-shaped relationship has also been observed in the terrestrial realm for birds. A pronounced discontinuity in the diversity of birds was found between the Hadley and the Ferrel cell (Turner and Hawkins, 2004). Factors that contribute to the biodiversity in the marine realm are numerous and belong to a large range of temporal and spatial scales. Geological events that have involved modification to the distribution of continents, open or closure of seaways and changes in the general thermohaline circulation have led to speciation. Climatic oscillations have involved modifications to the geographic distribution of species and to sea levels, which has been shown to contribute to evolution in certain ­circumstances. At a smaller scale, ecological factors act on the ranges of species tolerance, contributing to the spatio-temporal regulation of diversity. All these factors synergistically have

20 ecology 90 °N 70 °N

Latitudes

50 °N 30 °N 10 °N

E

10 °S 30 °S 50 °S 70 °S 90 °S

0

0.2 0.4 0.6 0.8 1 Normalized parameters

FIGURE 2.11:  Latitudinal pattern in pelagic foraminifera biodiversity. Data from Prell et al. (1999).

c­ontributed to set up or maintain species and to shape the ­present-day biodiversity. These large-scale events should still be  considered in the explanation of contemporary patterns of biogeography (Rohde 1992, 1997; Rosenzweig and Sandlin, 1997). What causes the latitudinal gradient in species richness, whether on land or in the sea, has been a topic of debate for decades (Rosenzweig and Sandlin,  1997), and more than 25 hypotheses have now been proposed (Gaston,  2000). While some authors have suggested that the species richness gradients are due to the larger area of the tropical belts (Rosenzweig, 1995), others propose null models of biodiversity such as the neutral theory (Hubbell,  2001) and the mid-domain effect (MDE) (Colwell and Hurtt,  1994). The latter model shows that the ­stochastic distribution of species ranges between poles leads to a peak in species richness at the equator, simply because of a geometric effect in the geographical space (Colwell and Lees, 2000). Perhaps the most compelling hypotheses are those that invoke an environmental control of biodiversity such as environmental stability or energy availability (Rutherford et al., 1999; Beaugrand et al., 2002b; Tittensor et al., 2010). For example, temperature is often suggested to explain large-scale patterns in the distributions of marine organisms (Beaugrand et al., 2010; Tittensor et al., 2010). Mechanisms by which temperature may act to create latitudinal gradients in marine biodiversity remain unclear however (Diniz-Filho et al., 2008), and correlative studies have failed to provide a theoretical basis to explain latitudinal gradients in biodiversity (Rangel et al., 2007). Recently, new mechanistic theories have been proposed and models elaborated to test the effect of the environment, MDE, ecological processes and historical biogeographical processes on the geographical gradients in species richness (Brayard et  al.,  2005; Storch et  al.,  2006; Rangel et  al.,  2007; Gotelli et  al.,  2009). These new types of models showed the strong potential influence of the MDE in both geographical and niche spaces (Brayard et  al.,  2005; McCain et  al.,  2007; Colwell et al., 2009), the potential effect of evolutionary niche dynamics (i.e. the balance between niche conservatism and niche evolu-

FIGURE 2.12:  Reconstruction of the present-day pelagic biodiversity from the MacroEcological Theory for the Arrangement of Life (METAL). From Beaugrand et al. (2013). (See Plate 10.)

tion processes) or speciation/extinction dynamics (Brayard et al., 2005; Rangel et al., 2007). A theory, termed the MacroEcological Theory for the Arrangement of Life (or METAL theory) has recently been proposed to explain the arrangement of marine plankton ­ (Beaugrand et al., 2013; Beaugrand, 2015). The theory is based upon the concept of the niche, sensu Hutchinson (1957), the principle of competitive exclusion (Gause,  1934), and the hypothesis of non-saturation of niches (Rohde,  2005), which embrace the effects of the climatic regime including both its intra- and interannual variability. It explains how temperature, modulated by climate variability, can create gradients in species richness in the oceans by interacting with the thermal tolerance of a species without needing to invoke other processes such as evolutionary niche dynamics or speciation/extinction dynamics or ecological processes such as dispersal and species interaction (Fig.  2.12). The theory suggests that the gradient in species richness is mostly determined by the climatic regime (both mean and variability) of a region and by an MDE in the Euclidean space of the (thermal) niche.

2.6 Conclusion Although plankton is not as diverse as some terrestrial groups (e.g. insect), plankton strongly influence the functioning of the whole marine ecosystem and the Earth system. The arrangement of plankton in the surface ocean is not random; these organisms are strongly influenced by climate, and particularly the temperature regime and its variability. Other secondary but locally important environmental parameters influence plankton biodiversity. The global planktonic system is horizontally divided into three main biomes: warm, temperate and cold-­water oceanic biomes. Secondary factors also enable the delineation of other secondary systems, termed provinces or ecoregions. The locations of these systems influence carbon exportation, ecosystem metabolism and the food web. Global climate change is expected to strongly influence the location of these systems, which will have severe consequences for both regulating (e.g. global carbon cycle) and provisioning (e.g. fisheries, aquaculture) services (Sarmiento et al., 2004; Beaugrand et al., 2015).

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CH A PTER 3

PH YTOPLA NKTON PRODUCTIVITY John R av en

3.1 Introduction Phytoplankton are the planktonic organisms which account for most of the primary production in the ocean (Field et al., 1998). Their characteristic trophic mode is the production of organic compounds using energy from light, and chemical elements from inorganic compounds, known as phototrophy, or more strictly photolithotrophy, from photo (light), litho (rocks, i.e. not organic) trophy (mode of nutrition) (Falkowski and Raven, 2007; Kirk, 2010). This process uses water as the electron donor, with oxygen as a by-product, and the reduction of inorganic carbon (carbon dioxide (CO2) or bicarbonate ions) producing sugars, from which all other cell components are made using inorganic forms of nitrogen, phosphorus and all the other chemical elements needed to produce cells. In contrast to phytoplankton, benthic photosynthetic organisms account for less than 5% of marine primary productivity in the present ocean. Marine phytoplankton are unicellular or colonial organisms ranging in size from less than 1 µm to just over 1 mm, corresponding to about a 1010-fold range of volumes (Falkowski and Raven, 2007; Finkel et al., 2010). They encompass a wide taxonomic range, including the cyanobacteria with bacterial cell organization, and eukaryotic taxa which have acquired and genetically integrated the photosynthetic system of a cyanobacterial symbiont derived from a primary endosymbiotic event in which a eukaryotic cell engulfed a cyanobacterial cell  forming a photosynthetic eukaryote. These hybrid cells evolved to form the Glaucocystophyta, Chlorophyta and (non-­ planktonic) Rhodophyta lineages. These primary endosymbioses gave rise to other photosynthetic taxa through s­ econdary endosymbiotic events involving the ingestion of a photosynthetic eukaryote from the Chlorophyta or, more commonly, the Rhodophyta by a non-photosynthetic eukaryote forming the photosynthetic members of the large groups Alveolata, Ochrista, Cryptophyta, Haptophyta, Excavata and Rhizaria, again with genetic integration of the symbionts (Pawlowski, 2013). Cases of tertiary endosymbiosis are known in dinoflagellates which are members of the Alveolata. Genetic integration involves the

loss of many of the genes in the endosymbiont, and transfer of most of the remainder, mainly c­ oncerned with photosynthesis, to the host cell nucleus. A ­residue of genes needed for photosynthesis are retained in the chloroplast. This parallels the process by which cyanobacterial cells became the original chloroplasts. The endosymbiotic origin of chloroplasts that carry out ­crucial photosynthetic components can be regarded as phagotrophy, as practiced today in many protists, with arrested digestion. Similar processes underlie the acquisition of a functional photosynthetic system by non-photosynthetic phagotrophic eukaryotic cells that occurs in present day cells involving ingestion of microalgae and retention of the whole cell (Flynn et al., 2013). Retention of whole cells, without genetic integration, is an example of true endosymbiosis, while kleptoplasty, the retention of only the chloroplasts from the algal food, is not (Flynn et al., 2013). Another significant role of phagotrophy in the context of phytoplankton is by more immediate digestion, supplying organic carbon as well as nitrogen, phosphorus, iron, vitamins and other nutrients. This added mode of nutrition means that the phytoplankton concerned are phagomixotrophs, practicing a mode of nutrition that occurs to varying extents in most of the higher taxa that lack a cell wall.

3.2  Resources Needed by Phytoplankton 3.2.1  Photosynthetically active radiation and inorganic carbon The defining means of obtaining energy in phytoplankton is by  oxygenic photosynthesis. This involves the absorption of electromagnetic radiation from sunlight, in the wavelength range 400–700 nm (photosynthetically active radiation or PAR) (Falkowski and Raven, 2007; Kirk, 2010). This photon energy between 400 nm and 700 nm absorbed by the light-harvesting complexes in (or on the surface of ) the thylakoid (membrane-­ bounded compartments within the chloroplast) membrane, and involving a range of pigments bound to proteins, is transferred to the photochemical reaction centres where photochemistry is

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

ph y topla nkton producti v it y  25 carried out by protein-bound chlorophyll a (Falkowski and Raven, 2007). There are two sorts of reaction centre. One is known as photosystem II and uses light energy to extract electrons from water to produce oxygen and promote the electrons to a higher energy level (Falkowski and Raven, 2007). This reaction centre uses photons at the energy content equivalent to 680 nm, and can use photons of higher energy equivalence from pigments absorbing in the range 400–680 nm. For each photon arriving at photosystem II, 0.8 electrons are moved (Falkowski and Raven, 2007; Raven et al., 2014). The electrons from photosystem II move energetically downhill to the other reacton centre, that of photosystem I (Falkowski and Raven, 2007). This reaction centre uses photons at the energy content equivalent to 700 nm, and can use photons of higher energy equivalence from pigments absorbing in the range 400–700 nm. For each photon arriving at photosystem I, one electron is moved (Falkowski and Raven, 2007; Raven et al., 2014). Again the electron is promoted to a higher energy level, and then reduces the electron acceptor NADP+ to produce NADPH. Part of the light energy that is not used in the oxidation-reduction reactions of extracting electrons from water and using them in reducing NADP+ is used in pumping protons from the chloroplast stroma of eukaryotes or the cytosol of cyanobacteria into the thylakoid lumen (Falkowski and Raven, 2007; Raven and Ralph, 2015). The photon requirement, oxidation-reduction and proton transfer reactions are shown in equation (1) (Falkowski and Raven, 2007; Raven and Ralph, 2015); the significance of the asterisk superscripts on the O of H2O is shown in the context of equation (4). 9 photon 400 – 700 nm + 2H 2 *O ( lumen) + 2 NADP + (stroma/ cytosol) + 12º H + (stroma/cytosol) → 2 [ NADPH + H + ] (stroma/cytosol) + * O2 ( lumen) + 12ºH + ( lumen) (equation 1) The NADPH produced in equation 1 could, if oxidized by O2, yield 110 kJ per mol electron transferred to O2 in vivo; however, oxidation of NADPH is coupled to reductive, i.e. energy-requiring, syntheses. Energized proton transport into the thylakoid lumen results in a trans-thylakoid proton energy gradient, with the lumen having a lower pH and higher electrical potential than the stroma (Raven and Ralph, 2015). Some of this energy difference released when the H+ returns from the lumen to the stroma or cytosol through the ATP synthase in the thylakoid membrane is used to phosphorylate 1 ADP to 1 ATP for each 4 protons moved. This stoichiometry is based on measurements of H+ fluxes and ADP phosphorylation, although higher H+:ATP values are predicted from the structural biology of the ATP synthase (Raven and Ralph, 2015). The ATP synthesized is far from thermodynamic equilibrium with ADP and phosphate, and the in vivo free energy of hydrolysis is about 55 kJ per mol, per mol ATP. A suitable enzyme, mechanochemical motor (e.g. in flagella), or transmembrane transporter can use the energy from ATP hydrolysis to drive cellular energy-requiring processes (Falkowski and Raven, 2007).

With the H+:ATP ratio of 4, and 12 H+ pumped into the thylakoid lumen for each 4 electrons moving from 2 H2O to  2  NADP+, return of the 12 H+ to the stroma or cytosol through ATP synthase produces 3 ATP (equation 2) (Raven and Ralph, 2015): 12º H+ ( lumen) + 3 ADP (stroma/cytosol) + 3 phosphate (stroma/cytosol) → 12º H + (stroma / cytosol) + 3 ATP (equation 2) (stroma/cytosol) + 3 H 2 O (stroma /cytosol) All phytoplankton use the Calvin-Benson cycle to convert CO2 to CH2O; equation (3) shows the stoichiometric requirement for NADPH and ATP of the cycle at CO2 saturation; all the reactants and products are in the stroma or cytosol (Falkowski and Raven, 2007; Raven, 2009; Raven et al., 2014; Raven and Beardall, 2016): 2 [ NADPH + H+ ] + 3 ATP + 3 H 2 O + CO 2 → CH 2 O + H 2 O + 2 NADP+ + 3 ADP + 3 phosphate (equation 3) Summing equations (1)–(3) gives the overall equation (4), where the superscript asterisks and filled square superscripts distinguish the fate of O from H2O from that of O from CO2. 9 photon 400 − 700 nm + 2 H 2 *O ( lumen) + C¡O2 (cytosol) → CH 2¡O (cytosol) + H 2¡O (cytosol) + *O2 ( lumen) (equation 4) It is important to note that the stoichiometry of 9 absorbed photons supplying the 3 ATP and 2 NADPH required for the reduction of 1 C in carbohydrate from 1 CO2 only applies when the external CO2 concentration is high enough to allow the maximum rate of photosynthesis (Raven, 2009; Raven et  al., 2014; Raven and Beardall, 2016). In seawater in equilibrium with the current atmospheric CO2 mol fraction of 400 ppm CO2, CO2 does not saturate the core carboxylase ribulose-1.5-­ bisphosphate carboxylase-oxygenase (commonly abbreviated to Rubisco). The carboxylase activity of Rubisco is shown in equation (5): 1 Ribulose-1,5-bisphosphate +1 CO 2 +1 H 2 O → 2 3-phosphoglycerate (equation 5) The stoichiometry in equation 5 only applies at saturating CO2 if O2 is present, or at lower CO2 in the absence of O2 (Raven and Beardall, 2016). At non-saturating CO2 in the presence of O2 the competitive Rubisco oxygenase activity occurs (equation 6): 1 Ribulose-1,5-bisphosphate +1 O 2 → 1 2-phosphoglycolate +1 3-phosphoglycerate (equation 6) In a small minority of phytoplankton there is diffusive CO2 entry, giving a lower CO2 and a high O2 at the site of Rubisco activity. This involves an energy cost per CH2O production from CO2 in addition to that shown in equation 3. This energy is required for 2-phosphoglycolate synthesis; this is all lost if the

26 ecology glycolate resulting from the dephosphorylation of 2-phosphoglycolate is lost to the medium. Alternatively, the glycolate is metabolized through the photorespiratory carbon oxidation cycle. This reaction sequence converts 2 glycolate to 1 3-phosphoglycerate and then 1 triosephosphate at the (CH2O) redox level, with production of 1 CO2 and the consumption of O2, and ATP and NADPH. The minimum total photon requirement for photosynthesis with diffusive CO2 influx, Rubisco oxygenase and the photorespiratory carbon oxidation cycle is slightly less than 10 photons per CO2 assimilated into carbohydrate (Raven et al., 2014). The majority of marine phytoplankton have CO2 concentrating mechanisms (CCMs) which concentrate external CO2 at the active site of Rubisco, greatly decreasing Rubisco oxygenase activity and nearly saturating the carboxylase activity. This saves on most of the cost of Rubisco oxygenase and the photorespiratory carbon oxidation cycle, but incurs a minimum additional energy cost of accumulating CO2 of almost 1 photon per CO2, i.e. a total of almost 10 photons per CO2 assimilated into carbohydrate (Raven et  al., 2014; Raven and Beardall, 2016). Many phytoplankton cells with CCMs are saturated for photosynthesis at the present air-equilibrium CO2 concentration, and even more are saturated for growth. However, when growth is CO2 saturated but when photosynthesis is not CO2 saturated in today’s atmosphere, there is a higher organic C content per cell when grown at higher CO2 concentrations (Doney et al., 2012). As will be shown below, additional photons are needed to convert the carbohydrate (C, H and O) produced in photosynthesis into cell material. These photons are required for the uptake and assimilation of the other nutrient elements required for growth, i.e. the major nutrients N, P, S, K, Mg and Ca, and the elements only required in trace amounts such as Fe, Mn, Cu, Co, Ni and Cl. Further photons are needed to provide energy for maintenance processes. Some of the processes involved in growth and maintenance, for example respiration, involve loss of previously gained resources such as C and energy. Since these processes are essential for gaining resources overall, they are considered here under resource gain (del Giorgio and Williams, 2005; Falkowski and Raven, 2007; Raven and Ralph, 2015). The availability of light is among the factors that has shaped the evolution of photosynthetic protists. PAR is attenuated by absorption by particles and by water and substances dissolved in it, by scattering by particles and at the molecular level (Raman scattering) (Kirk, 2010). In an (idealized) optically homogeneous surface ocean there is an exponential attenuation with depth. The attenuation coefficient varies greatly with the particulate (including phytoplankton) content and dissolved organic matter. Strictly light-limiting conditions for growth only apply to phytoplankton spending most or all of their time well below the ocean surface. Such light limitation applies to phytoplankton entrained, i.e. carried along with, vertically circulating water masses in a very deep upper mixed layer meaning that most of their time is spent well below the ocean surface

(e.g. at high latitudes in spring where mixing can occur down to 600 m). Light limitation also occurs for cells occurring deep (e.g. in deep chlorophyll maxima) within the surface ocean when there is little vertical mixing. These depths can vary, so ‘deep’ here means, for a static layer, less than about 300 m from the surface in even the clearest water, while for the most opaque, i.e. highly attenuating, ocean waters the maximum depth at which photosynthetic growth can occur may be as ­little as a few metres (Falkowski and Raven, 2007; Kirk, 2010). The maximum specific growth rate of a given genotype (genetically very closely similar organisms) of phytoplankton depends on the availability of resources such as PAR and nutrients, as well as temperature, that supply activation energy (Raven et al., 2013; Raven and Ralph, 2015). Growth rate also depends on the fraction of biomass occupied by catalysts, including enzymes, transmembrane transporters, mechanochemical motors and pigment protein complexes involved in photon absorption and transfer of the resulting excitation energy, needed for growth and maintenance (Raven et al., 2013; Raven and Ralph, 2015). Crucially, the growth rate depends on the allocation of resources (nutrients and energy) among the catalysts. The allocation of resources is subject, within a genotype, to acclimation processes (see Table 3.1) which, for example, increase the allocation to light-harvesting complexes relative to other catalysts when growth is light-limited. However, there are strong effects on specific growth rates depending on the high-level phylogeny; for example, dinoflagellates generally grow less rapidly than diatoms of a similar size. In this case there is a partial explanation of this disparity: ­dinoflagellates have a 40% lower energetic efficiency of ATP production from organic carbon in oxidative phosphorylation compared to most other phytoplankton organisms (Raven and Ralph, 2015). The variation of specific growth rate with cell size within a high taxon is still not well defined; any decrease in specific growth rate with increasing size is relatively small, and in at least some cases there is a decrease in growth rate with decreasing cell size  for the smallest representatives of an algal class (Raven et al., 2013). Photoinhibition is one reason why a phytoplankton organism does not achieve its maximum specific growth rate when growing in what are otherwise apparently optimal conditions. Photosystem II photochemistry generates damaging products in addition to the ‘normal’ products of O2 and reduced plastoquinone especially as the increasing rate of excitation energy transfer into the photosystem II reaction centre exceeds the rate at which the reduced plastoquinone can be processed by redox and proton pumping catalysts in the thylakoid membrane (Raven, 2011). Photoinhibition encompasses not just the decreased rate of photosynthesis resulting from inhibition by the damaging products of photosystem II activity, but also the protective mechanisms which decrease the possibility of photodamage. Examples of these protective mechanisms are the diversion of excitation energy to some other sink than the ­photosystem II reaction centre and the non-photochemical

ph y topla nkton producti v it y  27 TABLE 3.1:  Characteristics of changes to the function of phytoplankton that can occur over different time periods based on Raven and Geider

(2003).

Process

Characteristics

Timescale

Examples

Regulation

Unchanged genome and proteome; post-translational changes in proteome (covalent modification, ligand binding), changed metabolic fluxes.

Half time seconds to minutes.

State transitions. Nonphotochemical quenching.

Acclimation

Unchanged genome, changed proteome. Changed metabolic fluxes.

Half time 1–2 hours. Similar time course for diel changes in protein expression, synthesis of proteins to replace those damaged in photoinhibition.

Photoacclimation in response to changed photosynthetically active radiation (PAR). Changed expression of carbon dioxide (CO2) concentrating mechanism (CCM) with changes in CO2.

Adaptation

Changed genome through changed allele frequency, or through mutation and fixation; natural selection.

Many generations.

Experimental evolution in response to changed CO2 availability for growth.

quenching of excitation energy which has already reached the  reaction centre. All of the processes considered under ­photoinhibition decrease the rate of photosynthesis under both PAR-limiting and, to a smaller extent, PAR-saturating conditions (Falkowski and Raven, 2007; Raven, 2011). An important final point is that the processes considered under the heading of photoinhibition not only show a decreased rate of photosynthesis at a given PAR relative to that of a hypothetical ‘photoinhibition-proof ’ alga, but also have direct energy costs. These costs are the energy (and other resources) required for damage repair, for avoiding excitation energy arrival at photosystem II reaction centres and for non-photochemical dissipation of excitation in photosystem II reaction centres. There is also a time lag in responding to both increases and decreases in PAR, meaning that if PAR changes more frequently than once every 10 minutes or so it cannot be tracked by the algal responses (Raven and Geider, 2003).

recycling means that, for most nutrient elements, the surface ocean is depleted in the inorganic forms of the element relative to the deep sea (Raven and Geider, 2003; Falkowski and Raven, 2007). The means by which the nutrient elements are replenished after being lost from the ocean by sedimentation and subsequent subduction (and, for N, denitrification and loss to the atmosphere as N2 and N2O) are numerous. They include riverine and aeolian input from terrestrial weathering and biological processes in terrestrial ecosystems and, for N, biological N2 fixation in the ocean producing reduced N and lightning producing oxidized N that enters the ocean as dissolved nitric acid. Consideration of nutrient assimilation below is focused on the three nutrient elements generally considered as limiting photosynthetic primary productivity in the ocean, i.e. N, P and Fe. It is important to recall the significance of phagomixotrophy in many marine phytoplankton; this process can supply N, P and Fe as well as organic C (Falkowski and Raven, 2007).

3.2.2  Nutrient elements: introduction

3.2.3 Nitrogen

The major nutrients C, O, H, N, S, P, K, Mg and Ca and trace elements such as Fe, Cu, Mn, Zn, Ni, Co and Cl are taken up by non-phagomixotrophic phytoplankton in the surface ocean in (generally) an inorganic form. After incorporation into particulate organic matter these are either recycled to the inorganic form in the surface ocean after viral lysis or entry into the grazer food web, or are lost as particulate matter to the deep sea where they are mainly recycled into inorganic form (a small fraction is incorporated into sediments) and ultimately return to the surface in upwellings. This sedimentation and partial

The great majority of marine phytoplankton can only use N as biologically available combined N, i.e. N species in which N is covalently linked to (an) element(s) other than N; they cannot carry out biological N2 fixation. Combined N is lost from the ocean by long-term incorporation into sediments, and, mainly, denitrification, and is supplied to the ocean by riverine and aeolian inputs, by lightning and by biological N2 fixation (diazotrophy) (Tyrrell, 1999; Falkowski and Raven, 2007; Glibert et al., 2016; Raven and Giordano, 2016). The very abundant N source N2 is only available to a limited number of cyanobacteria among

28 ecology the marine phytoplankton. These cyanobacteria include the free-living Crocosphaera and Trichodesmium, and Richelia and the UNCY-A group, which are symbiotic with other phytoplankton (Bombar et  al., 2014). The activities of these diazotrophic cyanobacteria, and of chemoorganotrophic diazotrophs, in the surface ocean are not sufficient to prevent phytoplankton growth in much of the surface ocean being limited by the availability of combined N, even in combination with the combined N input by upwellings and by riverine and aeolian inputs (Tyrrell, 1999). Why diazotrophs, including cyanobacteria, do not supply sufficient combined N to prevent this N limitation of primary productivity is not entirely clear. Potentially diazotrophic organisms grow less rapidly when using N2 rather than combined N under otherwise similar conditions, presumably because resources are diverted from core metabolic processes of growth and maintenance to the resource intensive process of diazotrophy. There are predictions of greater energy (PAR, for reduction of N2 to NH4+, paralleled by H2 production), P, Fe and Mo requirements than for growth on NH4+, and there are field observations of limitations of N2 fixation by Trichodesmium in different parts of its range, by PAR, P and Fe; there seems to be an adequate supply of Mo for diazotrophy in the surface ocean. For non-diazotrophic phytoplankton, there is a range of combined N species that can act as N source. Of these, NH4+, NO3− and NO2− are relatively readily measured in surface ­seawater, and are assimilated by all phytoplankton, with the exception, for example, of some strains of the marine ­picocyanobacterium Prochlorococcus that cannot use NO3- or NO2-. In contrast, a very large range of organic N species can act as N sources for phytoplankton in the surface ocean (Falkowski and Raven, 2007; Tyrrell, 2013; Raven and Giordano, 2016). This makes for complication in measuring the full range of available organic N species and even more difficulty in apportioning the assimilation of N by phytoplankton among these organic N species. In upwelling regions the major N source is NO3−, generated from the mineralization of sinking particulate organic N to produce dissolved organic N, then NH4+ and, by nitrification, NO2− and finally NO3−, in the deep ocean. In the oligotrophic surface ocean, recycling of organic N from phytoplankton grazers and carnivores, including the effects of viral lysis, generates dissolved organic N and NH4+ but does not usually proceed through nitrification to NO2− and NO3− before assimilation by phytoplankton (Raven and Giordano, 2016). Globally, it appears that phytoplankton assimilation of reduced (relative to N2) N forms in ‘recycled production’ exceeds the assimilation of N forms more oxidized than N2 in ‘new production’ (Falkowski and Raven, 2007). It is generally held that ‘new production’ can uniquely remove carbon from the atmosphere by sinking of particulate organic matter, but it must be borne in mind that ‘new production’ is based on upwelled water in which the inorganic nutrients C, N and P are each enriched relative to basal seawater in the Redfield ratio of 106C:16N:1P (by atoms rather than mass), i.e. the ratio in which they are required by marine phytoplankton averaged over time and space. Thus,

upwelled waters supply not only the combined N, and P, used in new production, but also the inorganic C (Doney et al., 2012; Raven and Beardall, 2016). NO3− assimilation has been studied in great detail in marine phytoplankton. NO3− is taken up by high affinity active transport (i.e energy dependent, and able to accumulate NO3− to an electrochemical potential higher than that in the medium) systems which can apparently provide a higher steady-state cytosol NO3− concentration than is found in seawater. Production of the NH4+ used in the incorporation of N into organic matter requires the sequential operation of NO3− and NO2− reductases (Glibert et al., 2016). Phytoplankton biomass production with the Redfield ratio of 106C:16N by atoms requires a gross CO2–C assimilation per unit NO3−–N assimilation of 8:1 by atoms, or a reductant (NADPH or equivalent) ratio of 4:1 of reductant use in CO2–C assimilation relative to NO3−–N reduction. This 25% increase in reductant use per unit biomass produced for growth on NO3− relative to that needed for growth on exogenous NH4+ means that the growth of phytoplankton on NO3− is predicted to require significantly more absorbed photons per C assimilated than growth on exogenous NH4+. However, while this increased photon requirement is sometimes found, in many cases the expected energetic benefit of growth on NH4+ rather than NO3− is not obvious. There are also predictions of greater requirements for Fe and Mo for growth on NO3− than for NH4+. The uptake of NH4+, like that of NO3−, involves high affinity transporters and input of energy by the organism, and seems to result in the accumulation of NH4+ in the cytosol relative to the concentration found in ­seawater (Glibert et al., 2016).

3.2.4 Phosphorus Unlike N, P in nature almost always occurs at the redox level of the oxyanion phosphate which, at the pH of seawater, occurs predominantly as HPO42−. P is supplied to the ocean as HPO4− in rivers, and as aeolian aerosols and dust, and leaves it by longterm sedimentation. P is the ultimate geochemical limiting nutrient for primary productivity, although constraints on diazotrophy mean that N is the proximal limiting nutrient for marine phytoplankton using combined N over much of the ocean (Tyrrell, 1999; Falkowski and Raven, 2007; Tyrrell, 2013). However, there are areas of the ocean in which P limits phytoplankton growth on combined N, as well as other areas where biological N2 fixation is limited, or co-limited, by P supply. As with NH4+ and NO3−, HPO42− entry involves high affinity transporters and an energy input which maintains the HPO42− concentration in the cytosol above that in seawater (Dyhrman, 2016). A significant fraction of the P in seawater occurs as organic phosphates, and most phytoplankton can produce extracellular alkaline phosphatase; this releases HPO42− that can be taken up by cells. A minor component of the P in seawater occurs as organic phosphonates with a C-P bond; these ­natural products can be taken up by some cyanobacteria, with internal cleavage releasing HPO42− (Raven, 2013; Dyhrman,

ph y topla nkton producti v it y  29 2016). Another aspect of the response of phytoplankton to P limitation is the replacement of phospholipids in non-photosynthetic membranes with glycolipids (sulfolipids and galactolipids) typical of photosynthetic membranes, by genetic adaptation or by acclimation (Raven, 2013) (Table 3.1).

3.2.5 Iron Fe is needed by phytoplankton at about 0.01 times the P requirement (Falkowski and Raven, 2007; Marchetti and Maldonado, 2016). Although it is globally a very abundant element, the present oxygenated surface ocean means that most Fe occurs as the insoluble Fe(III). Fe(III) can be accessed by siderophores produced by cyanobacteria which chelate Fe(III); the Fe-charged siderophore can be taken up by cyanobacteria (Marchetti and Maldonado, 2016). Fe(III) can be accessed by surface reductases of eukaryotic algae, producing Fe(II) which can be taken up by the cells before it can be oxidized back to Fe(III) (Marchetti and Maldonado, 2016). These mechanisms are able to prevent Fe limitation of phytoplankton growth on combined N over most of the surface ocean, although diazotrophy by cyanobacteria can be Fe-limited in these ocean areas. Three ocean areas, the Southern Ocean, the Eastern Equatorial Pacific, and the NorthEast Subarctic Pacific, are Fe-limited for phytoplankton growth on combined N (Marchetti and Maldonado, 2016). Planktonic diatoms from Fe-limiting environments have significant decreases in their Fe requirements, at the expense of a restricted capacity to acclimate to varying PAR availability (Marchetti and Maldonado, 2016).

3.3  Limiting the Loss of Resources 3.3.1  Loss of dissolved organic matter by ‘healthy’ cells All marine phytoplankton lose organic C as dissolved organic compounds with a range of relative molecular masses (M) from low Mr glycolate to the larger Mr transparent exopolymeric polysaccharides and proteins (Raven and Ralph, 2015). While some of these compounds have known functions (toxins, siderophores, carbonic anhydrases, alkaline phosphatases), most of the organic carbon lost from the cells has no known function. Marine phytoplankton cells under near-natural conditions have rather variable net dissolved organic carbon loss, ranging from 1% to 10% of net primary productivity (Raven and Ralph, 2015). However, cultures of marine phytoplankton can show a net organic C influx when supplied with high concentrations of assimilable organic C, allowing osmomixotrophy (uptake and assimilation of dissolved organic matter combined with photosynthesis) in the light and osmoorganotrophy (uptake and assimilation of dissolved organic matter) in the dark. This may occur in some organically enriched (e.g. by bird urine and ­faeces) high intertidal rock pools, but not in the open ocean (Raven et al., 2013).

3.3.2  Respiratory losses of carbon dioxide Conversion of organic carbon to CO2 in respiratory reactions is a requirement for algal growth and maintenance (del Giorgio and Williams, 2005). Even in the light, where the production of NADPH and ATP used in biosynthesis could be supplied by the thylakoid membrane reactions of photosynthesis rather than in the respiration of stored carbon, respiratory processes are needed to produce carbon skeletons essential for biosynthesis (Raven and Ralph, 2015). Examples are production of the four carbon dicarboxylic acids required for the aspartate family of amino acids and pyrimidines, and the five carbon 2-oxoglutarate required for the glutamate family of amino acids and (with the exception of mitochondria of Euglena) tetrapyrrols such as chlorins in chlorophylls and haems of cytochromes, algal haemoglobin, and catalase (Falkowski and Raven, 2007; Raven and Ralph, 2015). This biosynthetic use of tricarboxylic acid cycle intermediates requires C3 +C1 carboxylations to supply four carbon dicarboxylic acids, maintaining the functionality of the tricarboxylic acid cycle despite the removal of intermediates. For biosynthesis in the dark, respiration not only produces carbon skeletons, but is also the only source of NADPH and ATP. Maintenance processes use ATP for the synthesis of replacements for damaged proteins (mainly using undamaged amino acids from the damaged proteins) and for recouping leaked solutes (Raven and Ralph, 2015). Maximizing the use of respiratory energy in growth and maintenance can be achieved by avoiding various inefficiencies. One inefficiency is the absence of 2-oxoglutarate dehydrogenase from the tricarboxylic acid cycle of Euglena and many cyanobacteria and its replacement by a less energy-efficient bypass. A further example is the absence of the NADH to ubiquinone proton pumping electron transport reaction from dinoflagellate mitochondria which decreases the ATP produced per NADH oxidized by O2 by 40%. Finally, the use of the alternative oxidase activity which oxidizes reduced ubiquinone decreases the ATP produced per NADH oxidized by oxygen by 60% represents an inefficiency unless the alternative oxidase activity is limited to what is essential for balancing the carbon skeleton synthesis and energy transformation functions of ­respiration (Raven and Ralph, 2015).

3.3.3  Virally induced cell lysis Viral lysis is an important pathway transferring phytoplankton particulate organic matter to the surface ocean food web as both soluble and particulate organic carbon. This viral organic carbon cycling is generally greater than that of dissolved organic carbon efflux by non-infected cells (Falkowski and Raven, 2007). Phytoplankton defences against viruses include, for Emiliania huxleyi, reversion from the calcified diploid heterococcolith-bearing phase to the less-calcified flagellate haploid holococcolith-bearing phase, which is much less sensitive to the virus (Frada et al., 2008).

30 ecology 3.3.4 Grazing Grazing is a major loss of phytoplankton biomass in many habitats (Irigien et al., 2005; Landry et al., 2009; Flynn et al., 2013). The smaller phytoplankton cells are grazed by unicellular phagotrophs, including (as indicated in Section 3.1) mixotrophic algae, while the larger phytoplankton cells are grazed by metazoan grazers. The quantitative extent to which grazing is restricted by toxins produced by some phytoplankton, and ­phytoplankton structures such as spines and mineralization (e.g. calcification by coccolithophores, silicification by diatoms), is unclear. A poorly characterized pathway for the loss of marine phytoplankton is the action of eukaryotic parasitoids (Raven and Waite, 2004).

3.3.5 Sinking Some marine phytoplankton are positively buoyant with low-density components overcoming the effects of solid ballast. Examples are cyanobacteria with gas vesicles, and large diatoms, dinoflagellates and the phycomata stages of some species of the Prasinophyceae with a large fraction of the cell occupied by a vacuole containing solution less dense than seawater. The significance of the large size lies in the large fraction of the cell volume occupied by the vacuole, and by the effect of cell size in Stokes’ law (discussed below) (Lavoie et al., 2016). Other marine phytoplankton cells have densities very similar to, or higher than, that of seawater. It has been suggested that loss of neutral or positive buoyancy by cells infected by viruses or parasitoids, leading to sinking, is a means of purging a population of a minority of infected cells; it is not clear where, or even if, this mechanism is possible (Raven and Waite, 2004). Probably the majority of marine phytoplankton cells, for example coccolithophores (calcified) and smaller diatoms (silicified), are negatively buoyant and so ultimately sink out of the upper mixed layer. These considerations of rising or sinking must be put into the context of cell or colony size. Stokes’ law shows that, for a given excess density of the spherical particle relative to that of the medium, and a given viscosity of the medium, the sinking rate is proportional to the square of the radius of the particle. For buoyant cells the same relationship applies, with the excess density of the particles relative to that of the seawater being negative in this case (Lavoie et al., 2016). These considerations apply to cells lacking flagella. Cells or colonies with flagella, and more or less dense than seawater, can, respectively, move up or down relative to the surroundings. This not only helps restrict cell loss from the upper mixed layer, but also permits diel vertical migration in upper mixed layers where the vertical movement of water in the column is less than the speed of flagellar motility. Such diel vertical migrations may allow for a greater acquisition of PAR (near the surface in the photophase) and nutrients solutes (near the thermocline in the scotophase, i.e. night), thus contributing to resource gain as well as limiting biomass loss. The same is true for vertical periodic (often greater than one day) migration of

large vacuolated cells of diatoms, dinoflagellates and the phycomata stages of Prasinophyceae which can alter their vacuolar density to greater than or less than that of seawater (Lavoie et al., 2016).

3.4  Genetic Adaptation and Phenotypic Acclimation to Habitats Marine phytoplankton, in the form of oxygenic photosynthetic cyanobacteria and eukaryotic algae, occur in the entire range of illuminated habitats in the present ocean, and have coped with the various amplitudes and speeds of environmental change since the global oxidation event, signalling the occurrence of photosynthetic O2 production in the ocean in excess of organic C burial and respiration, some 2.3 billion years ago. The possibilities for a phytoplankton genotype to alter its ­f unctioning in response to a changed environment can be considered under three headings: regulation, acclimation and adaptation (Falkowski and Raven, 2007; Tyrrell, 2013). Regulation (Table 3.1) involves changes to the catalytic activity of a pre-existing component of all proteins in the cell (the proteome) (Raven and Geider, 2003). Examples of photosynthetic responses to changes in incident photosynthetic radiation are state transitions that alter the association of peripheral light-harvesting complexes with photosystem II, and variations in non-photochemical quenching, i.e. the dissipation of excitation energy from a photosystem other than through photochemistry. These processes occur over time periods of seconds to tens of minutes and maximize photosynthetic light use at low irradiances while decreasing the potential for photodamage to photosystem II at higher irradiances (Raven and Geider, 2003; Raven, 2011). Acclimation (Table 3.1) involves changes to the proteome by altering the expression of the existing genome (Raven and Geider, 2003). Again using examples from photosynthesis, a decrease in irradiance increases the number of photochemical reaction centres per cell and/or the number of peripheral light-harvesting complexes per reaction centre, while decreasing expression of downstream catalysts such as those of the photosynthetic carbon reduction cycle. A further example is the decreased expression of CO2 concentrating mechanisms with large increases in CO2 availability. Such acclimatory changes take hours to complete, i.e. a significant fraction of the algal generation time. Since environmental change involves more than one factor, acclimation is increasingly being examined in multifactorial experiments (Doney et al., 2012; Zhu et al., 2016). Both regulation and acclimation have resource costs (Raven and Geider, 2003). For regulation there is the cost of producing the relevant machinery in each generation, and (in some cases) in operating it. For acclimation the costs are those involved in the production of a different balance of proteins. It is not clear how natural selection balances the costs of regulation and acclimation relative to their possible benefits in environments which

ph y topla nkton producti v it y  31 change over a wide range and with high frequency, especially since the known costs and potential benefits may be in different resource currencies (e.g. light and nitrogen) with exchange rates that themselves vary with the environment (Raven and Geider, 2003; Raven, 2011; Lavoie et al., 2016). Adaptation (Table 3.1), in contrast to regulation and acclimation, involves changes to the genome of individuals eventually evolving into genetically distinct populations (genotypes) related to the environment in which they evolved (Raven and Geider, 2003). It is generally taken to involve changes to the nucleotide sequence of a gene or genes, and, less frequently the import of genes by horizontal gene transfer, for example by viral transformation. Another alternative is changes in the frequency of pre-existing alleles in populations. Research in this area, termed experimental evolution, has implicitly focused on changes in the nucleotide sequence of DNA over many generations, and excluded horizontal gene transfer, and on comparing such environmental factors as the present atmospheric CO2 and the CO2 level expected in 2100 ad (Reusch and Boyd, 2013). Such experiments result in a range of new genotypes, some of which appear to be advantageous; however, this possibility needs to be tested in the context of natural selection. Most of these exercises in experimental evolution have involved increased CO2 supply; there is a clear need for more work involving other aspects of environmental change and, especially, experimental evolution under more than one component of environmental change. Clearly phytoplankton have a greater chance of adapting to changing environments than do larger marine organisms with longer generation times (Reusch and Boyd, 2013).

3.5 Productivity 3.5.1 Methodology The methodology used is vital in assessing estimates of primary productivity, for example because different methods are appropriate for different productivities (del Giorgio and Williams, 2005; Falkowski and Raven, 2007; Tyrrell, 2013). Estimates of global marine planktonic primary productivity are quite variable, depending on the methods used (Field et  al., 1998). Measurements of metabolic rates at specific places and times are based on 14C- (or 13C-) labelled inorganic carbon assimilation or, in areas of high primary productivity, net or labelled O2 changes. The labelled inorganic carbon method gives estimates of (net) photosynthesis, dark inorganic carbon assimilation and of organic excretion, but not of gross photosynthesis or respiratory CO2 production (del Giorgio and Williams, 2005). The O2 method gives estimates of net photosynthesis and respiration in the dark and hence (assuming respiration in the light is the same as in the dark) of gross photosynthesis. Enrichment in the isotope 18O2 in high productivity environments can give estimates of the combined O2 uptake by ‘dark’ respiration and light-dependent processes, and of gross 16O2, oxygen production,

in the light. However, these techniques only give values for the specific time and site of measurement, and there are large spatial and temporal variations. Remote sensing, on the other hand, measures chlorophyll reflectance spectra from satellites. This method allows an estimation of chlorophyll in the surface few metres, and involves algorithms based on local estimates for conversion of chlorophyll to primary productivity at the surface, and further extrapolation to provide estimates for the entire photic zone, the lower parts of which are not ‘seen’ by remotely sensed reflectance spectroscopy (Field et  al., 1998; Uitz et al., 2010). This method allows estimates for non-cloudy days for the whole oceans over the time period for which the remote sensing platform is operational.

3.5.2  Local variations Primary productivity is higher in areas where nutrient concentrations are higher (Field et al., 1998; del Giorgio and Williams, 2005; Falkowski and Raven, 2007; Uitz et al., 2010). One example is those places with permanent or seasonal upwellings of nutrient-rich deep ocean water. A second is where there has been previous deep mixing (in winters in higher latitudes),which mixes nutrient-rich deep ocean water into the upper mixed layer. Following shoaling of the thermocline in spring, the presence of high nutrient concentrations in an upper mixed layer that is shallow enough to give a high mean flux of PAR in the upper mixed layer leads to the spring bloom. A third example is where enhanced, anthropogenic, inputs of combined nitrogen and of phosphorus occur from rivers, and/or where combined nitrogen is supplied by atmospheric deposition. These places allow considerable net ecosystem productivity and hence net organic carbon sinking into the deep ocean (Falkowski and Raven, 2007). However, such organic carbon burial does not necessarily involve sequestration of atmospheric CO2 since, for upwelled waters, the increment of phosphorus and of combined nitrogen from mineralization of organic particles which had previously sunk is accompanied by an increment of inorganic carbon in the Redfield ratio of 106C:16N:1P (by atoms) consumed in primary productivity (Tyrrell, 2013). Primary productivity is lower in areas where nutrient concentrations are low, for example in most oceanic gyres. Here the strongly stratified water column limits nutrient transfer from the deep ocean to where there is sufficient light for photosynthesis, so primary productivity is strongly nutrient limited. There is minimal net ecosystem productivity and very limited possibilities of export of particulate organic carbon, with associated nitrogen, phosphorus and iron etc. to the deep ocean (Falkowski and Raven, 2007).

3.5.3  Global summation Estimates of global marine primary productivity vary (Table 3.2). Measurements at specific places and times based on 14C inorganic carbon assimilation or, in areas of high primary productivity, O2 changes, allow time integration (Field et al., 1998; del

32 ecology Giorgio and Williams, 2005; Falkowski and Raven, 2007). Remote sensing allows estimation of chlorophyll in the surface few metres, and involves algorithms based on local estimates for conversion to primary productivity at the surface and further extrapolation to provide estimates for the entire photic zone (Falkowski and Raven, 2007; Uitz et  al., 2010). Remote sensing of ocean colour has also been used to allocate regional (and hence global) primary productivity to higher taxa of phytoplankton with specific pigment compositions (Uitz et  al., 2010) (Table 3.2). The extent to which global marine primary productivity has changed over recent decades, and how it will change with continuing environmental change over the rest of this century, are TABLE 3.2:  Effects on global marine primary productivity of season,

organism size and planktonic-benthic life form, and environmental nutrient availability.

Category

Net primary productivity Pg carbon per year

1

Seasonal April-June

10.9

2

Seasonal July-September

13.0

3

Seasonal October-December

12.3

4

Seasonal January-March

11.3

5

Seasonal Total

48.5

6

Oligotrophic (low nutrient availability) plankton

11.0

7

Mesotrophic (moderate nutrient availability) plankton

27.4

8

Eutrophic (high nutrient availability) plankton

9.1

9

Coastal macrophytes

1.0

10

Trophic Status / Life Forms Total

48.5

11

Microplankton (does not pass a 20 μm filter)

15

12

Nanoplankton (passes a 20 μm but not a 2 μm filter)

20

13

Picoplankton (passes a 2 μm filter)

11

14

Macrophytes

15

Size/Life Form Total

1 47

From Field et al. (1998) categories 1–10 and 14, and Uitz et al. (2010) categories 11–13. Note that the seasonal total and the trophic status / life forms total differ slightly from the size / life form total.

incompletely resolved matters. Over much longer time periods there are indications of changed (relative to today) marine primary productivity during Pleistocene glaciations and during hothouse episodes such as the Cretaceous (Falkowski and Raven, 2007; Doney et al., 2012; Tyrrell, 2013).

Ack now ledgements Discussions with John Beardall, Colin Brownlee, Martina Doblin, Sinéad Collins, Kate Crawfurd, Paul Falkowski, Zoe Finkel, Kevin Flynn, Richard Geider, Mario Giordano, Andrew Johnston, Ian Joint, Janet Kūbler, Elena Litchman, Aditee Mitra, Bruce Osborne, Antonietta Quigg, Peter Ralph, Katherine Richardson, Elisa Schaum, Alison Taylor, Anya Waite and Glen Wheeler have been very helpful. The University of Dundee is a registered Scottish charity, No. SC015096.

R efer ences Bombar, D., Heller, P., Sanchez-Báracaldo, P., Carter, B. J., and Zehr, J. P. (2014). Comparative gemomics reveals surprising divergence of two closely related strains of uncultivated UCYN-A cyanobacteria. The ISME Journal 4, 2530–42. del Giorgio, P. A., and Williams, P. J. Le B. (eds) (2005). Respiration in aquatic ecosystems, pp. 315. Oxford University Press, Oxford, UK. Doney, S. C., Ruckelhaus, M., Duffey, J. E., et al. (2012). Climate change impacts on marine ecosystems. Annual Review of Marine Science 4, 11–37. Dyhrman, S. T. (2016). Nutrients and their acquisition: phosphorus physiology in microalgae. In Borowitzka, M. A., Beardall, J., and Raven, J. A. (eds) The Physiology of microalgae, pp. 155–83. Springer Publishing, Heidelberg. Falkowski, P. G., and Raven, J. A. (2007). Aquatic photosynthesis, 2nd Edition, pp. 512. Princeton University Press, Princeton, NJ, USA. Field, C. B., Behrenfeld, M. J., Randerson, J. T., and Falkowski P. (1998). Primary productivity of the biosphere: integrating terrestrial and oceanic components. Science 281, 237–40. Finkel, Z. V., Beardall, J., Flynn, K. J., et al. (2010). Phytoplankton in a changing world: cell size and elemental stoichiometry. Journal of Plankton Research 32, 119–37. Flynn, K. J., Stoecker, D. K., Mitra, A., et al. (2013). A case of mistaken identification: the importance of mixotrophs and the clarification of plankton functional-classification. Journal of Plankton Research 35, 3–11. Frada, M., Probert, I., Allen, M. J., Wilson, W. H., and de Vargasm C. (2008). The “Cheshire Cat” escape strategy of the coccolithophore Emiliania huxleyii in response to viral infection. Proceedings of the National Academy of Science USA 105, 15944–8.

ph y topla nkton producti v it y  33 Glibert, P. M., Wilkerson, F. P., Dugdale, R. C., et  al. (2016). Pluses and minuses of ammonium and nitrate uptake and assimilation by phytoplankton and implications for productivity and community composition, with emphasis on nitrogen-enriched conditions. Limnology and Oceanography 61, 165–97. Irigien, X., Flynn, K. J., and Harris, R. P. (2005). Phytoplankton blooms: a loophole in microzooplankton grazing. Journal of Plankton Research 27, 313–21. Kirk, J. T. O. (2010). Light and photosynthesis in aquatic environments, 3rd Edition. pp. 509. Cambridge University Press, Cambridge, UK. Landry, M. R., Ohman, M. D., Goericke, R., Stakel, M. R., and Tsyrklerch, K. (2009). Lagrangian studies of phytoplankton growth and grazing relationships in a coastal upwelling ecosystem of Southern California. Progress in Oceanography 83, 208–16. Lavoie, M., Raven, J. A., and Levasseur, M. (2016). Energy cost and putative benefits of cellular mechanisms modulating buoyancy in aflagellate marine phytoplankton. Journal of Phycology 52, 239–51. Marchetti, A., and Maldonado, M. T. (2016). Iron. In Borowitzka, M. A., Beardall, J. and Raven, J. A. (eds) The physiology of microalgae, pp. 252–79. Springer Publishing, Heidelberg. Pawlowski, J. (2013). The new micro-kingdoms of eukaryotes. BMC Biology 11 (40), doi:10.1186/1741-7007-11-40. Raven, J. A. (2009). Contributions of anoxygenic and oxygenic photolithotrophy and chemolithotrophy to carbon and oxygen fluxes in aquatic environments. Aquatic Microbial Ecology 56, 177–92. Raven, J. A. (2011). The cost of photoinhibition. Physiologia Plantarum 142, 87–104. Raven, J. A. (2013). The evolution of autotrophy in relation to phosphorus requirement. Journal of Experimental Botany 64, 4023–46. Raven, J. A., and Beardall, J. (2016). The ins and outs of CO2. Journal of Experimental Botany 67, 1–13.

Raven, J. A., and Geider, R. D. (2003). Adaptation, acclimation and regulation in algal photosynthesis. In Larkum, A. W. D., Douglas, S. E., and Raven, J. A. (eds). Photosynthesis in Algae, pp. 385–412. Kluwer, Dordrecht, The Netherlands. Raven, J. A., and Giordano, M. (2016). Combined nitrogen. In Borowitzka, M. A., Beardall, J., and Raven, J. A. (eds) The Physiology of Microalgae, pp. 143–54. Springer Publishing, Heidelberg. Raven, J. A., and Ralph, P. J. (2015). Enhanced biofuel production using optimality, pathway modification and waste minimization. Journal of Applied Phycology 27, 1–31. Raven, J. A., and Waite, A. M. (2004). The evolution of silicification in diatoms: inescapable sinking and sinking to escape. New Phytologist 162, 45–61. Raven, J. A., Beardall, J., Larkum, A. W. D., and SanchezBáracaldo, P. (2013). Interaction of photosynthesis with genome size and function. Philosophical Transactions of the Royal Society B 368 (1622), doi:10.1098/rstb.2012.0264. Raven, J. A., Beardall, J., and Giordano, M. (2014). Energy costs of carbon dioxide concentrating mechanisms, Photosynthesis Research 121, 111–24. Reusch, T. B. H., and Boyd, P. W. (2013). Experimental evolution meets marine phytoplankton. Evolution, 67, 1848–59. Tyrrell, T. (1999). The relative influences of nitrogen and phosphorus on oceanic primary production. Nature 400, 525–31. Tyrrell, T. (2013). On Gaia: A critical investigation of the relationship between life and earth, pp. 320. Princeton University Press, Princeton, NJ, USA. Uitz, J., Claustre, H., Gentili, B., and Stramski, D. (2010). Phytoplankton class-specific primary production in the world’s oceans: seasonal and interannual variability from satellite obxervations. Global Biogeochemical Cycles, 24 (3), doi: 10.1029/2009GB003680. Zhu, Z., Xu, K., Fu, F., et al. (2016). A comparative study of iton and temperature interactive effects on diatoms and Phaeocystis antarctica from the Ross Sea, Antarctica. Marine Ecology Progress Series 550, 39–51.

CH A PTER 4

ZOOPLA NKTON PRODUCTIVITY A ndr ew G. Hir st

4.1 Introduction Zooplankton is a term used to describe the heterotrophic ­plankton, including both metazoans (multicellular animals) and single-celled protozoa such as ciliates and flagellates. Zooplankton encompass a great diversity of phyla, with an array of life history and ecological traits. Their body size spans over more than 15 orders of magnitude, and they include species with a life cycle of less than a day to many years. Holoplankton are those species which spend their entire life cycle in the water column, while meroplankton are those which have some stage, or part of their life, as a benthic organism, living in or on the bottom substrate. For meroplanktonic forms, the planktonic larvae phase is an effective means to disperse and hence reduce competition between conspecific benthic individuals. Plankton are usually thought of as being unable to migrate horizontally as their swimming speeds are too slow compared with tides and currents. Metazoan zooplankton include taxa such as euphausiids, decapods, amphipods, copepods and appendicularians, as well as gelatinous forms with high body water ­content, such as medusae, salps, doliolids and ctenophores, and the semi-gelatinous molluscs and chaetognaths (Larson, 1986; Kiørboe, 2013). The pelagic environment is a distinct environment with many opportunities but also difficulties to be overcome; this is reflected in strong selection pressures and distinct adaptations of the species which reside here. Autotrophic production is largely dominated by microscopic algae, which contrast starkly with many terrestrial systems with large, slow-growing macroscopic plants. The epipelagic zone (top 200 m of the water column) is a habitat where there are few places to hide from predators. Unlike living in sediment, or in a terrestrial environment, in which there are often structures to hide in and among, pelagic environments typically lack such features. Indeed, many behavioural, ecological and morphological features of ­plankton appear to have evolved to allow predator avoidance, or to allow a population to withstand high predation rates (Verity and Smetacek, 1996). Camouflage from visual hunters is ­possible through special colouration and near transparency, and

b­ ioluminescence may also play a role, while a rapid escape response and advanced abilities to detect predators remotely can be common. The near ubiquity of body transparency across many different phyla within the zooplankton (including in many Arthropoda, Ctenophora, Cnidaria, Chaetognatha, and Chordata such as salps and doliolids) is strongly indicative of the evolutionary pressure to avoid being eaten by visual predators. Avoiding the epipelagic zone during daylight through the daily migration to depth (known as diel vertical migration or DVM) is also incredibly common (Ohman, 1990; Hays, 2003). Varying degrees of migration can be found in organisms from microbes to euphausiids, fish and jellyfish. Species migrate to the surface in order to feed at night, and descend to depth to reside in deeper waters during the day, thereby avoiding sunlit surface waters when they can be more easily seen and preyed upon. Visual and non-visual predators may also perform DVM in order to follow their prey’s migration ambit. Another distinct property of pelagic invertebrates is that their growth and development rates are often very fast (Hopcroft and Roff, 1995; Kiørboe and Hirst, 2014), and they have an ability to grow at near exponential rates through much of their juvenile phase (Huntley and Lopez, 1992; Hirst and Forster, 2013). This is not the case for many terrestrial species and vertebrates. Being able to achieve these feats in a dilute food environment makes it all the more impressive, and indicates these traits may be a prerequisite for long-term success. Given that growth and mortality rates must on some level balance one another (or a species would ultimately go extinct or take over the world), plankton may suffer high mortality rates too (Ohman and Hirche, 2001). Gelatinous organisms are common in pelagic systems. The dual advantage of an inflated body includes both enhanced feeding as larger bodies can typically find more prey, and also partial protection from predation (Kiørboe, 2013), a neat trick in the plankton where both aspects are important. In this chapter we discuss the importance of determining life history and vital rates of zooplankton, and then examine the major ways in which growth and secondary production rates are determined. Finally, we explore mechanistic and empirical frameworks to predict what controls these rates and why.

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

zoopla nkton producti v it y  35 4.1.1  What is secondary production?



P = Bt − B0 + Be (1)

where P is the secondary production of biomass over the period t. This is estimated from the biomass at the end of the period (Bt) minus that at the start (B0), plus that biomass which has been eliminated (Be) over the time period (e.g. through mortality and consumption). We need to consider the eliminated biomass, rather than simply change in the standing stock of biomass, because this is an important aspect of productivity. To give an example, the biomass of fish may be nearly constant throughout the year, yet these fish may have been harvested and biomass removed (which has been replaced by recruitment); as such, simply measuring changes in fish biomass in the ocean over time would not reflect their secondary production. We need to account for both the gains and losses. The first law of thermodynamics states that energy cannot be created or destroyed, but can only be changed from one form to another. Energy for the functioning of most ecosystems comes from sunlight being used in photosynthesis to produce chemical energy. The second law of thermodynamics states that whenever energy is transformed, there is a loss of energy through the release of heat; the activity of organisms, their metabolism and respiration all act as loss terms. Productivity declines between each trophic level because of the inefficiencies and various loss processes. The yield of fisheries is ultimately limited by the primary production, in addition to the efficiency of transfer between each trophic level, and the trophic level the fish occupy. A major driver to understanding productivity of zooplankton is to appreciate what controls the productivity of commercially important species. Key measures of the state of marine food webs involve both the pathways and also the magnitude of biomass flow. Of course, production at lower trophic levels sets an absolute limit to production at higher levels, but beyond this, if there are bottom-up effects, then fluctuations in planktivore production (including many fish) would be expected to correlate positively with changes in production by zooplankton and new primary production. Such a pattern has been found to occur in North Sea fish (Heath, 2005), while fluctuations in plankton have been shown to result in long-term changes in cod recruitment and larval survival in the North Sea (Beaugrand

Fish + Squid Production (g wet mass m–2 yr–1)

Secondary production is analogous to primary production; it is a measure of the rate at which biomass is accumulated by an individual, species or trophic level. Some workers have historically defined it as the production of herbivorous organisms (the second trophic level after primary producers), but it is best to think of it as ‘secondary’ in the sense that it is heterotrophic, and ultimately dependent upon primary production. We define secondary production in such a way as to allow for the fact that biomass may either accumulate, or be transitory, passing to higher trophic levels, dying or decaying through time. Secondary production is therefore defined by the equation:

25 9

20

10

8 15 5

10 4 5 12

7

6

3

0 0

50

100

150

200

250

300

Total Phytoplankton Production (gC m–2 yr–1) FIGURE 4.1:  Carnivorous fish plus squid biomass production as a

function of total phytoplankton production for open-ocean and coastal environments: 1, Atlantic Ocean gyre centre; 2, Atlantic Ocean gyre boundaries; 3, waters off Hawaii; 4, Bothnian Sea; 5, Gulf of Riga; 6, Gulf of Finland; 7, Baltic Sea proper; 8, Nova Scotian shelf; 9, Gulf of Maine; 10, Mid-Atlantic Bight. (Redrawn from Iverson, 1990.) Reproduced with permission from John Wiley & Sons.

et al., 2003). Friedland et al. (2012) found chlorophyll concentration and the ratio of secondary to primary production were positively associated with fishery yields across 52 large marine ecosystems across the globe. The ratio of mesozooplankton production to primary production reflects the efficiency of transfer between two critical food web components affecting fishery productivity. The strong relationship found by Iverson (1990) between primary production and carnivorous fish and squid production is clear in Figure 4.1.

4.2  Rates of Biological Processes in the Plankton For over a century, a common thread in ecology has been the description of rates of biological processes and a drive to understanding what controls their speed. From fast reaction rates within cells, to the slow pace of evolution and speciation, the ‘pace of life’ is a core focus for biologists and ecologists (Kleiber, 1961, 1975; Brown et al., 2004); this focus has similarly been an important issue for those working in the field of plankton s­ tudies. Rather than simply measuring the density and biomass of organisms, we wish to understand how fast changes in populations are, and how they transform material, especially since zooplankton play such an important role in controlling ocean production and biogeochemistry of the ocean (Banse, 1995). Much fieldwork on zooplankton has focused on aspects of population dynamics, including their growth (e.g. Peterson et al., 1991), mortality (e.g. Ohman and Hirche, 2001; Ohman et al., 2002) and fecundity (e.g. Uye, 1981; Runge, 1985). The tug of  war between these rates determines how populations

36 ecology A Life-history rate (d–1)

1

Egg hatch rate

0.1

0.01

Sac spawners egg hatch Broadcasters egg hatch In situ development Food saturated development

Development rate

Weight-specific growth (d–1)

B

Weight-specific fecundity (d–1)

change, and ultimately their success or failure. Over the past few decades, the need for quantitative evaluation of marine ecosystems has been emphasized as a necessary component in the understanding of how climate change and fishing impacts upon marine ecosystems (e.g. Walther et al., 2002; Edwards and Richardson, 2004; Heath, 2005). The most studied group of zooplankton are the copepods. These small crustaceans dominate the mesozooplankton in terms of numbers and biomass, and act as a conduit of material from lower trophic levels of algae and microbes, to higher trophic levels including fish and jellyfish (Atkinson, 1996). The globally available data on epipelagic copepod rates of growth and development (Hirst and Bunker, 2003), fecundity (Bunker and Hirst, 2004), mortality (Hirst and Kiørboe, 2002), feeding (Saiz and Calbet, 2007), respiration and excretion (Ikeda et al., 2001) have been synthesized and analysed for patterns; in particular, the link between these rates and temperature, body size and food availability have been described (e.g. Fig. 4.2). Similar descriptions have recently been completed for gelatinous organisms too (Purcell, 2009; Acuña et al., 2011). We return to some of these relationships in Section 4.4. The high water content of gelatinous organisms allows them, through small energetic input, to attain rapid increases in wet biomass, surface area, prey-capture abilities, and predation rates (Larson, 1991). Some non-gelatinous organisms such as larvaceans, which occupy a mucous house, have a total volume that is more similar to a gelatinous organism when we include this external structure. The house is composed of non-living acellular mucous material that is metabolically inert once secreted (although it needs investment for its production and continued replacement). Because of the effective large volume (gelatinous) nature of these feeding structures, they are able to filter volumes considerably greater than species with similar body carbon that do not have watery bodies (e.g. crustaceans). This helps to explain why gelatinous forms and also those organisms which produce external watery but large feeding structures, such as some molluscs and appendicularians, attain very fast growth rates (Kiørboe and Hirst, 2014). Indeed, the pelagic tunicates, which include appendicularians, salps and doliolids, are some of the fastest growing metazoans on Earth (Heron and Benham, 1984; Hopcroft and Roff, 1995; Madin and Deibel, 1998). In warm tropical waters, the appendicularian Oikopleura can have minimum generation times of ~ 1 day, with mass-specific growth rates ranging from 1.38 day−1 to 3.18 day−1, and averaging 10.7-fold increases in biomass over 24 hours. This is an order of magnitude greater than the copepod Paracalanus crassirostris, which in the same set of experiments did not quite double its biomass in a day, and had growth rates that averaged 0.51 day−1 (Hopcroft and Roff, 1995). Because we typically measure growth in plankton assuming they increase their mass exponentially, and we usually reference this mass increase to that of the animal’s mass itself (i.e. mass increase per unit of body mass per day), growth is usually expressed in units of per day (day−1), where a value of 0.69 day−1 represents a doubling in mass in one day.

1

0.1

0.01

In situ juvenile sac spawners In situ juvenile broadcasters Food saturated juvenile sac spawners Food saturated juvenile broadcasters

C 1

0.1

0.01

In situ adult sac spawners In situ adult broadcasters Food saturated adult sac spawners Food saturated adult broadcasters

0

5

10

15

20

25

30

35

Temperature(°C) FIGURE 4.2:  Life history rates of marine planktonic copepods versus

temperature (°C) including both broadcasting and sac spawning species. The former release their eggs freely into the water column, where they suffer high mortality; the latter keep their eggs attached to their body until hatching. Rates are shown for in situ food and saturated laboratory conditions; this comparison allows the degree of food limitation in the natural environments to be assessed. A, Egg development rates (d−1) of broadcast (grey circles) and sac spawners (grey triangles). Food saturated (large solid circles = broadcasters; large solid triangles = sac spawners) and in situ egg to adult development rates (d−1) (large open circles = broadcasters; open triangles = sac spawners). B, Food saturated mass-specific growth (d−1) of broadcast (solid squares) and sac spawner juveniles (black diamonds). C, Food saturated mass-specific fecundity (d−1) of broadcast (solid circles) and sac spawner adults (solid triangles). In B and C in situ relationships given by heavy lines for comparison, broadcasters dashed lines and sac spawners solid lines. All values are in units of per day to facilitate comparisons. Figure adapted from Hirst and Bunker (2003). Reproduced with permission from John Wiley & Sons.

zoopla nkton producti v it y  37

4.3  Measuring Zooplankton Growth and Productivity 4.3.1 Background Production of heterotrophic organisms is much more difficult and time-consuming to assess than primary production. Measurements of secondary production by large heterotrophs is typically done in a labour intensive way, one species at a time. Consequently, across a region there are rather few estimates of productivity that are comprehensive in the species included. Primary production by contrast is measured using a range of classic methods based on radioisotopes such as 14C (Steeman-Nielsen, 1952), or from variation in oxygen concentration in sealed bottles. Although at present we have no widely adopted analogous method for production by heterotrophic zooplankton, progress is being made on such methodologies (see Section 4.3.6).

4.3.2  Growth rates There are three direct methods for measuring the growth rates of zooplankton. First, the rate of production of eggs and young (progeny); second, the use of cohorts (natural or artificial) to track the growth of young as they increase in size over time; finally, the ‘Moult Rate’ methods. These have been altered and adapted somewhat, depending upon the circumstance, but the differences are largely variations on a theme. In mesozooplankton studies, our measurements are dominated by the study of copepods because these are somewhat easier to examine than other groups, and they also typically dominate this size fraction.

4.3.3  Production of progeny The first, and possibly easiest way to estimate growth rates, is to determine the rate of release of eggs, live born young or gametes by adults. In many zooplankton organisms such growth is commonly assessed in adult females as a rate of egg production (Runge and Roff, 2000). There are some difficulties in such an estimate; for example, in many pelagic organisms growth is indeterminate, i.e. body size continues to change after maturation. In species with such a growth type, both body mass and the output of progeny contribute to production. This is the case for chaetognaths, euphausiids, amphipods and ctenophores. We therefore, strictly speaking, need to be able to assess the somatic body mass growth of animals as well as the reproductive output of adults.

4.3.4  Cohort methods The rate of increase in size or mass of organisms in a population can be determined if a cohort (a group of organisms of similar size or age) can be identified and followed through time. The ability to be able to separate the group from older or younger animals is important because we need to be sure that changes

in size are a result of their growth, rather than being impacted by new recruits or by mixing with other size groups. The main assumption underlying this method is that the same population is being sampled and correctly identified at each time interval. The method is most easily applied when a pulse of reproduction can be followed. These requirements are not often met in marine waters, and defining and tracking cohorts is difficult. Advection and mixing in open water can cause violations of the assumption, while sampling variability also introduces error into the estimates. In addition, the need for frequent sampling can limit the practical applications of this technique. Even so, the cohort method has been applied to estimate growth and production of crustaceans, chaetognaths, medusae and salps for example (Landry, 1978; Heron and Benham, 1984; Lucas et al., 1997; Choe and Deibel, 2000). Kimmerer and McKinnon (1987) cleverly reapplied the cohort method, artificially creating cohorts of similarly sized copepods by gently sieving animals through multiple mesh screens. They incubated these organisms in natural seawater, followed the increases in size of the animals, and hence were able to determine growth rates. This ‘artificial cohort’ method has the advantage that it allows growth to be measured in continuously reproducing populations for which natural ­ cohorts are not observed. The technique has been used in many mesozooplankton, including copepods and appendicularians (Hopcroft and Roff, 1998). An extensive review and refinement of this method, with recommendations for best practice, have been undertaken by Kimmerer et al. (2007).

4.3.5  Moult rate methods Crustaceans are particularly amenable to having their growth rates measured because of features of their life cycle. Not only can they be collected and identified more easily, but unlike many soft-bodied plankton such as jellyfish, it is possible to ascribe distinct life stages to them. As these organisms develop they increase in mass and commonly pass through identifiable stages, they moult, and the sequential stages are often very different in their morphology. By knowing the mass of consecutive stages, and the time it takes to pass between these stages, growth rates can be estimated. Methods employing such ideas are commonly grouped as ‘Moult Rate’ methods. Stage durations, necessary in assessing the time interval over which mass accumulation takes place, are determined from the fraction of organisms which moulted from an incubated stage per day (stage duration, D, in days, is equal to 1/MR, where MR is the proportion of animals moulting out of the stage per day). These methods have been described in detail and appropriate equations developed in Hirst et al. (2005) (Fig. 4.3). In order to estimate production we typically assess growth rates. Production of a population can be thought of as the sum total of the growth rates of all the individuals. As growth is typically assessed in mass-specific units, i.e. the amount of body mass added per unit of organism mass, then product of growth rate (g, units of mass per mass per unit time) and total biomass

38 ecology Stage i

Moult

Moult

Stage i+1

Moult

Stage-specific Method (gi_corr): Arithmetic Mean Mass

Wi_entry

Development Time

Wi_exit Di

Modified Moult Rate Method (gi→i+1): Arithmetic Mean Mass

AWi+1

AWi

Development Time

Combination of Di and Di+1

FIGURE 4.3:  Graphical representation of two methods used to describe growth across moulting stages of zooplankton. The Stage-Specific method determined growth from the change in mass at entry (Wi_entry) and exit (Wi_exit) from a life stage, which has a duration Di. While the Modified Moult Rate method measures change in the arithmetic mean mass of consecutive stages, AWi and AWi+1, these are matched to the appropriate time period based on the duration of the two stages over which the change in these mean weights are achieved. A change in mass must be tied to the appropriate time period to which these changes occur.

(B, units of mass) is the production rate (P, units of mass per unit time):

P = B × g (2)

4.3.6  Biochemical methods New approaches for measuring zooplankton production using biochemical methods have been developed in recent years. One such method involves measuring the activity of aminoacyl-tRNA synthetases (aaRS), an enzyme which catalyses the first step of the protein synthesis and which is essential for somatic growth (Yebra and Hernández-León, 2004). Another approach is based upon the enzyme chitobiase, which is involved in degradation and recycling of chitin exoskeletons in arthropods. Arthropods moult and new exoskeleton is produced as they grow; the concentration of the chitobiase enzyme is therefore related to moulting and growth rates (Oosterhuis et al., 2000; Sastri and Roff, 2000). Further, RNA/DNA ratios have also been used as they are associated with cell multiplication rates (e.g. Dagg and Littlepage, 1972; Wagner et al., 2001). These various techniques are often applied in conjunction with directly measuring mass change in order to calibrate them (e.g. Yebra et al., 2011). Having the potential to be automated and rapid (just like the 14C method is for primary production), the hope is that the biochemical method will eventually become the standard protocol.

4.4  Mechanistic and Empirical Frameworks Our interest in production rates of pelagic organisms stems from a wish to understand ocean biogeochemistry, population dynamics and the yields of commercially important species. Understanding the mechanisms behind what we observe in the natural world, and an ability to make predictions, are key drivers too. Zooplankton growth rates strongly correlate with temperature (see Fig. 4.2) and body size (Fig. 4.4) (Hirst and Bunker, 2003; Kiørboe and Hirst, 2014), as well as the amount of food available and its quality (Pond et al., 1996; Klein Breteler et al., 1999; Jónasdóttir et al., 2009). A highly prominent theory in recent years has been the Metabolic Theory of Ecology (MTE) (West et al., 1997; Brown et  al., 2004), which combines temperature and body size dependence of biological rates into a mechanistic theory. The dependence on body size is based around the idea that to fuel metabolism many organisms distribute material through a branched transport network system (e.g. a closed blood circulatory system). The theory suggests this sets the limit to rates of supply of fuel for metabolism. The temperature dependence part of the theory depends upon activation energies of reactions. Widely applied in terrestrial systems, and increasingly so in aquatic environments too, the theory predicts that development times, longevity, respiration, mortality, growth and feeding rates all scale with body mass in a similar linked way (Brown et al., 2004). The premise is that metabolism, defined as the sum total of the chemical processes that occur in living organisms, underpins all these rates, and hence a similar scaling should pervade them all. Fundamental to the theory is an assertion that organismal rates (R) relate to body mass with a power of ¾, i.e.:

R = a M ¾ (3)

where M is the mass of the organism, and a is an intercept (or normalization) term. A scaling power of 1 indicates that larger organisms do things at the same rate per unit of their body mass as small organisms, while ¾ indicates that larger organisms do things at a slower rate per unit mass than smaller organisms. Simple mathematics dictates that when the rate is expressed in mass-specific terms, r (i.e. the rate per unit mass of the organism), the expression becomes:

r = a M ¾−1, (4)

and therefore,

r = a M −¼ (5)

Recently, the MTE has been brought into question, and has been shown to suffer from important problems (O’Connor et  al., 2007), especially with regards to its applicability to

zoopla nkton producti v it y  39 A Log10(Specific respiration rate, μl O2 mg C–1h–1)

4 Protozoa Non-calanoid copepods Calanoid copepods Amphipods Euphausiids Tunicates Cnidaria + Ctenophores Fish Mammals (Kleiber)

3 2 1 0

–1 –2 –10–9–8–7–6–5–4–3–2–1 0 1 2 3 4 5 6 7 8 9 10

B

Ciliates Chaetognaths R/W~W0 R/W~W–0.25

Log10(Specific clearance rate, ml mg C–1h–1)

6 5 4 3 2 1 0 –10 –9 –8 –7 –6 –5 –4 –3 –2 –1 0 1 2 3 4 5

C Log10(Specific ingestion rate, μg C mg C–1h–1)

4 3 2 1 0 –1 –10–9 –8 –7 –6 –5 –4 –3 –2 –1 0 1 2 3 4 5

D Log10(Specific growth rate, mg C mg C–1h–1)

0 –1 –2 –3

with size with a power of around −¼, when comparing patterns across these taxa, they found that respiration and clearance rates scale very differently from growth and ingestion rates (Fig. 4.4). Mass-specific respiration and clearance rates show a rather similar pattern from the smallest to the largest taxa. Although they decline within individual taxa, they are rather constant with size when viewed more broadly (they follow the slope of zero indicated by the solid black line in Fig. 4.4), which is quite different from the continual decline with increasing size predicted by the MTE (the predicted slope from this theory is indicated by the dashed black line in Fig. 4.4). Such an inconsistency was similarly highlighted by Makarieva et al. (2008). Further work has tested the MTE against a competing geometric theory based around dependence of rates upon surface area (based on the ideas of Rubner, 1883). Many products needed for metabolism, or waste products from metabolic processes that must be removed, are exchanged across the general body surface, especially in many plankton invertebrates. This exchange process may ultimately dictate the body mass dependence of rates, rather than the distribution pathways within the body. Indeed, examining the body mass scaling of respiration rates within species, representing a wide variety of pelagic invertebrate forms, has recently demonstrated a strong significant correlation to the degree of surface enlargement during growth, while at the same time these results falsify the predictions of the MTE (Hirst et al., 2014; Glazier et al., 2015). This suggests that within many pelagic species, rates may be very dependent upon the exchange of necessary materials across the surface itself.

–4 –5 –10–9 –8 –7 –6 –5 –4 –3 –2 –1 0 1 2 3 4 5 6

4.5  The Future

Log10 (Body mass, mg C)

FIGURE 4.4:  Regressions through mass-specific rates versus body mass for various taxa; 95% confidence intervals for the regressions also shown. All rates are corrected to a common temperature of 15°C. A, respiration rates, mammals are also included for comparison (Kleiber, 1975); B, maximum clearance rates; C, maximum ingestion rates; D,  maximum growth rates. R, rate; W, mass. The black lines are regressions through all data assuming proportionality between massspecific respiration rate and body mass raised to power 0 or −0.25. In panel A, protozoans include both ciliates and flagellates; in lower panels, the ciliates are shown as a separate group and the pink line denotes flagellates alone. Figure adapted from Kiørboe and Hirst (2014). Reproduced with permission from the University of Chicago Press. (See Plate 11.)

pelagic invertebrates (Glazier, 2006). Kiørboe and Hirst (2014) tested the predictions of the MTE using data that included many zooplankton and fish species. Assembling growth, respiration, and maximum ingestion and clearance rate data, they found that patterns in nature strongly contradict the MTE. Although within individual taxa, for example ciliates, calanoid copepods, salps or fish, mass-specific rates commonly decline

Although our appreciation of zooplankton production and trophodynamics has come a long way over the past 40 years, much work is still needed. New mechanistic theories are necessary to explain the patterns we observe in vital rates of plankton (Fig. 4.4). Better appreciation of the role of bottom-up and top-down effects within food webs will help determine what controls the structure and dynamics of pelagic communities. Size-spectra data, collected using equipment that allows plankton to be counted and sized, are being exploited to estimate growth and mortality using recently developed mathematical equations (Edvardsen et al., 2002). These offer much promise, and allow rates to be determined in the field rather than in experimental incubations. Improved protocols and automated methods, as well as advances in sampling technology will allow us to better track plankton populations through time in the future.

Ack now ledgements We thank those authors who adapted figures and commented upon the text for us.

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CH A PTER 5

PH YTOPLA NKTON BIOGEOCHEMICAL CYCLES Ca rol Robinson

5.1 Introduction Carbon, nitrogen and phosphorus are essential elements required for all life on Earth. In the marine environment, dissolved inorganic carbon, nitrogen and phosphorus are utilized during phytoplankton growth to form organic material, which is respired and remineralized back to inorganic forms by the activity of bacteria, Archaea and zooplankton. The net result of the photosynthesis, calcification and respiration of marine plankton is the uptake of carbon dioxide (CO2) from the atmosphere, its sequestration to the deep ocean as organic and inorganic carbon and its availability to fuel all fish and shellfish production. The cycling of carbon by marine plankton is inextricably linked to that of nitrogen and phosphorus; thus marine plankton mediate climate through influencing the atmospheric concentration of not only CO2, but also nitrous oxide (N2O). Increasing anthropogenically derived atmospheric CO2 concentrations impact plankton mediated biogeochemical cycles through increasing seawater temperature and dissolution of CO2, leading to changes in water column mixing, availability of light and nutrients, decreasing dissolved oxygen and changing carbonate chemistry. This chapter describes how the activity of phytoplankton, bacteria and Archaea drive the marine biogeochemical cycles of carbon, nitrogen and phosphorus, and how climate driven changes in plankton abundance and community composition are influencing these biogeochemical cycles in the North Atlantic Ocean and adjacent seas.

5.1.1  The biological carbon pump The biological carbon pump (BCP, also known as the organic carbon pump) is the term given to the suite of processes by which CO2 that is transformed by phytoplankton photosynthesis into particulate and dissolved organic carbon (POC, DOC) in the sunlit surface ocean is exported to the deep ocean where it is respired back to dissolved inorganic carbon by heterotrophic prokaryotes (bacteria and Archaea) and zooplankton (see Fig. 5.1). These processes include the passive gravitational

flux of sinking organic particles (marine snow = dead plankton cells, zooplankton faecal pellets etc.), the active flux of DOC and POC mediated by zooplankton vertical migration, and the physical flux of DOC through overturning circulation or subduction. Alongside the BCP, the carbonate pump refers to the biogenic formation (calcification) of calcium carbonate shells by some phyto- and zooplankton. These shells act as ballast, enabling faster sinking (and therefore export) of calcite associated organic carbon. As a result of the BCP and the carbonate pump, together with the physical dissolution of CO2 in seawater (solubility pump), concentrations of dissolved inorganic carbon and macronutrients such as nitrogen and phosphorus are low in surface waters and enriched in the deep ocean, while concentrations of POC and DOC, nitrogen and phosphorus are higher in surface waters and decrease with depth. Approximately 65% of the vertical gradient in dissolved inorganic carbon in the ocean is attributed to the BCP and the carbonate pump, with the rest due to the solubility pump. Recently, a fourth ocean carbon pump has been proposed, the microbial carbon pump, which refers to the formation and transformation of long-lived DOC by microbes including viruses, prokaryotes and protists ( Jiao et al., 2014). The BCP is a major contributor to the global carbon cycle: approximately 100 GT C are produced through photosynthesis by phytoplankton each year, 5–12 GT C yr−1 are exported to the deep ocean, and around 1 GT C yr−1 is buried in sediments (Henson et al., 2011, 2012). Atmospheric CO2 concentrations would be approximately 200 ppm (~ 50%) higher than present day (in 2015) concentrations of ~ 400 ppm if the BCP did not exist (Parekh et al., 2006), with potentially hazardous consequences. Phytoplankton cells die through sinking out of the sunlit euphotic layer, viral infection or grazing by zooplankton. For example, the ability of diatoms to maintain neutral buoyancy is impaired under nutrient stress (primarily silica limitation) leading to rapid sinking of these relatively large cells. Nutrient stress can also promote the excretion of carbon-rich organic material, which causes the phytoplankton cells to stick together forming larger aggregates with faster sinking speeds. A pulse of sinking

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

ph y topla nkton biogeochemica l cycles  43

CO2

N2

H+

H2CO3 N2O NH4+

HCO3–

NO2–

NO3– CO32–

PO43–

C:N:P (Black:Grey:White)

Phytoplankton

Diazotrophs

Microzooplankton NO3–

N2O

CO2

PO43-

NO2–

Mesozooplankton

Bacteria/Archaea NH4+

RDOC N2O N2

NO2

Faecal pellets

–

NO3– NH4+

POM

DOM

Viruses

FIGURE 5.1:  Infographic of the inter-relationships between the organic and microbial carbon pumps and the microbial cycling of nitrogen and phosphorus. RDOC, recalcitrant dissolved organic carbon; POM, particulate organic material; DOM, dissolved organic material. C:N:P ratios are indicative only. Figure by Robinson et al. shared under a Creative Commons license at figshare http://dx.doi.org/10.6084/m9.figshare.1585741. (See Plate 12.)

particulate material of predominantly diatomaceous material can be seen in photographs of the seafloor at 2000 m within 2–3 weeks of the spring diatom bloom in the north-east Atlantic, suggesting sinking rates of 100–150 m d−1 (Billett et al., 1983). Viruses, which reach abundances of 108 mL−1 in marine systems, are major pathogens of phytoplankton and so play a role

in the termination and subsequent export of phytoplankton blooms (Suttle, 2007). Viruses also influence the efficiency of the BCP through infection and lysis of plankton cells. Cell lysis converts living particulate organic material (POM) into fragments of dead POM and dissolved organic material (DOM). This ‘viral shunt’ prevents or short-circuits the flow of carbon,

44 ecology nitrogen and phosphorus from phytoplankton and prokaryotes to higher trophic levels. The formation of cell fragments and DOM reduces the rate at which organic carbon sinks out of the euphotic zone, increases the amount of organic carbon respired back to CO2 in surface waters by heterotrophic prokaryotes and so decreases the efficiency of the BCP. However, viruses can increase the efficiency of the BCP if they increase the export of carbon relative to the export of a nutrient such as nitrogen, iron or phosphorus which limits phytoplankton growth. Viral lysis can produce viral particles rich in nitrogen and phosphorus and cell mineral elements such as iron which are selectively retained in the water column, and carbon-rich compounds such as cell wall material which can sink (Suttle, 2007). Recent culture and field experiments have shown that some viral infected phytoplankton produce high levels of carbon-rich transparent exopolymer particles that cause the dying cells to stick together in aggregates. This increases their sinking rate out of surface waters, and increases the export of carbon relative to inorganic nutrients (Lønborg et al., 2013). In addition to sinking particles (size range 102–104 µm), a ­substantial portion of POC exists as fine, suspended, neutrally buoyant particles, colloids and mucus gels (≤ 102 µm), and around 50% of the POC produced via phytoplankton photosynthesis is transformed through microbial food web processes such as excretion, grazing and viral lysis into DOC. This DOC supports a vast population of heterotrophic prokaryotes and most is respired back to CO2 within hours to days. However, a small fraction of DOC is transformed through the microbial carbon pump and photochemical processes into DOC, which is not readily utilized by heterotrophic prokaryotes (so-called recalcitrant DOC or RDOC) and so can accumulate in the water column for months to millennia ( Jiao et al., 2010; Hansell, 2013). The effective storage of RDOC in the ocean is an important mediator of climate, locking away organic carbon which would otherwise be converted into CO2 to escape to the atmosphere. The majority (~ 90%) of the POC and DOC exported from the euphotic zone is respired back to CO2 by heterotrophic prokaryotes in the mesopelagic or ‘twilight’ zone of the world’s oceans, operationally defined as the region between the base of the euphotic zone and 1000 m depth (Robinson et  al., 2010). This depth horizon contains nearly 75% and 50% of the prokaryote biomass and production, respectively, of the global ocean (Aristegui et al., 2009). There is growing evidence that deep-sea microbial activity is concentrated onto particles and gels, thereby giving access to higher concentrations of organic substrate (Herndl and Reinthaler, 2013). Many marine heterotrophic prokaryotes produce polysaccharides, which help them attach to surfaces. Their exoenzymatic decomposition of the sinking and suspended organic material produces trailing plumes of DOC which attract chemotactic, motile heterotrophs (Azam and Malfatti, 2007; Stocker and Seymour, 2012). Prokaryotic abundance in the mesopelagic zone declines logarithmically with depth, and Archaea are as abundant as bacteria (~ 105 cells mL−1) (Aristegui et al., 2009). While neither bacterial nor archaeal diversity decrease significantly with depth, only

30–50% of the detected prokaryotic phylotypes occur throughout the water column. The remaining 50–70% are present only in distinct depth layers or water masses. For example, the contribution of Gammaproteobacteria to the total bacterial community tends to increase with depth, whereas members of the Bacteroidetes group, common in near-surface waters, are largely absent in deep waters. This change in prokaryotic community with depth is thought to partly result from changes with depth in the concentration and composition of DOM, the primary substrate for prokaryotic heterotrophic metabolism. The importance of microzooplankton (< 200 µm, e.g. heterotrophic protists) and mesozooplankton (> 200 µm, e.g. copepods) as efficient grazers of phytoplankton production and mediators of active carbon flux, recycling of CO2 and transformation of sinking particles is detailed in Chapter 6. The efficiency with which organic material exported out of the euphotic zone and into the mesopelagic zone is remineralized to CO2 or remains as POC and DOC to sink below 1000 m, i.e. out of the mesopelagic and into the bathypelagic zone, is known as the transfer efficiency. This is determined by a combination of the size and type of phytoplankton occurring in the euphotic zone (and therefore the size and type of particles produced when these phytoplankton die and sink, and the size and type of faecal pellets produced by the zooplankton which graze on these phytoplankton), and the transformation of these particles in the mesopelagic zone through fragmentation by zooplankton and solubilization by microbes (Herndl and Reinthaler, 2013). Early studies suggested that the decline or attenuation in the flux of POC through the water column can be described by the relationship: Fz = F100 × ( z /100 )

b

where Fz is the POC flux at depth z (m), F100 is the POC flux at 100 m and b is a dimensionless scaling factor which determines the transfer efficiency and the remineralization depth, i.e. the depth at which the POC is converted to CO2. Flux data collected from six North Pacific sites in 1987 yielded a curve with b = −0.86 (Martin et al., 1987). However, we now know that this simple relationship is not globally applicable and that b  ­varies regionally and seasonally (e.g. from −0.27 to −1.29, with more negative values indicative of greater flux attenuation or lower transfer efficiency). A recent study based on satellite data estimated values of b ranging from −0.5 to −0.9 in the North Atlantic (Henson et  al., 2012). In addition, the simple power equation has been criticized as it relates flux attenuation to a fixed depth of 100 m and so does not take into account the regional variability in the depth of the euphotic zone. A reanalysis of global particle flux data (Buesseler and Boyd, 2009) suggests that regional variability in export of particles out of the euphotic zone and transfer efficiency through the mesopelagic zone can be best described by a combination of euphotic zone net primary production (NPP), the ratio between POC flux at the base of the euphotic zone (Ez, commonly defined as the depth of 1% or 0.1% light penetration) and NPP, and the ratio

ph y topla nkton biogeochemica l cycles  45 between the POC flux at Ez and the POC flux at 100 m below Ez, i.e. the transfer efficiency T100. Seasonal and interannual variability in the transfer efficiency is apparent at the Porcupine Abyssal Plain sustained observatory (PAP-SO) in the north-east Atlantic, where the percentage of the annual organic carbon primary production which reaches sediment traps tethered at 3000 m varies from 0.6% to 1.2% (Lampitt et al., 2010).

5.2  Seasonal Variability In the temperate and subpolar regions of the North Atlantic, seasonal variations in light, nutrient availability, water column mixing and zooplankton grazing generate a seasonal cycle in phytoplankton abundance and production, and therefore in the magnitude of the BCP. During winter, low light and deep mixing prevent rapid phytoplankton growth, but ensure maximum concentrations of inorganic nutrients in surface waters. In spring, day length, light intensity and sea surface temperatures increase, leading to the formation of a distinct layer of warmer surface water lying above a layer of deeper cooler water (so-called stratification of the water column). This layering ‘traps’ the phytoplankton in a shallower surface mixed layer so that they experience higher light intensity, thereby allowing enhanced phytoplankton growth. Phytoplankton biomass increases, producing a ‘spring bloom’, which decreases the surface water concentrations of inorganic nutrients. Nutrient exhaustion and loss to zooplankton grazers and viral infection reduce the phytoplankton biomass to low values in summer. Mixing of nutrients into the surface layer by autumn storms and a reduction in the activity of zooplankton grazers as they progress into their metabolic resting phases in preparation for winter, allow a secondary peak in phytoplankton biomass or an ‘autumn bloom’. Prolonged deep mixing and therefore low light, mean phytoplankton biomass and production return to their annual minima in winter. Within this seasonal cycle of phytoplankton biomass exists a seasonal succession of phytoplankton community structure related to the differing inorganic nutrient and mixing regimes preferred by different phytoplankton groups. To identify this succession requires time-series data collected either in Eulerian mode at a single easily accessible sampling station or in Lagrangian mode where the sampling vessel or platform moves to remain within the same water mass. Within the North Atlantic, Eulerian time-series studies are undertaken at the Bermuda Atlantic Time-series Study (BATS; Lomas et  al., 2013), the PAP-SO (Hartman et al., 2012) and the Western English Channel Observatory (WECO; Smyth et al., 2015), while quasi-­Lagrangian studies were undertaken in 1989 and 1990 at 47–60° N 20° W during the field programmes of the international Joint Global Ocean Flux Study ( JGOFS) (Lochte et  al., 1993) and the UK Biogeochemical Ocean Flux Study (BOFS) (Savidge et al., 1992). Barlow et al. (1993) measured the pigment signatures of phytoplankton groups at 47° N 20° W to show the temporal succession from diatom dominance during the 15-day development

and peak of the spring bloom to the dominance of prymnesiophytes in the 25 days following the bloom. Microscopic analysis of water samples collected from WECO at weekly intervals over a 15-year period show the detailed seasonal cycle of phytoplankton genera (Widdicombe et al., 2010). The spring bloom occurs between late March and early May and is dominated by  chain-forming centric diatoms such as Chaetoceros spp., Thalassiosira spp. or Skeletonema costatum. The prymnesiophyte Phaeocystis and the coccolithophore Emiliania huxleyi bloom in May after the diatom peak, and E. huxleyi blooms again in late July or August. An intense bloom of small pennate and centric diatoms such as Pseudo-nitzschia also occurs during the summer months. Finally, dinoflagellates of the potentially harmful genera Karenia, Prorocentrum and Dinophysis bloom in the warm stratified conditions of late summer before the population returns to an over-wintering community of small phytoflagellates (Widdicombe et  al., 2010). Repeatable seasonal patterns are also seen in the bacterial community composition at WECO (Gilbert et  al., 2012). Order Rickettsiales, dominated by the SAR11 clade, tend to peak in winter when light and primary production are low and inorganic nutrient concentrations are high. The Rhodobacterales, dominated by the Roseobacter clade, tend to peak in spring and autumn when nutrient concentrations are low and primary production is high. Similarly, repeatable seasonal cycles of three ecotypes of SAR11 in the sunlit euphotic zone and deeper mesopelagic waters at BATS are likely related to the supply, quantity and quality of DOM associated with seasonal mixing and stratification of the water column (Carlson et al., 2009). Interannual variation in the timing of water column stratification is also associated with high seasonal and interannual variability in mesopelagic particle flux as revealed by more than 20 years of data collected at the PAP-SO (Lampitt et al., 2010).

5.2.1  Regional variability The flux of organic carbon out of the surface ocean is ­influenced by the size and type of phytoplankton which occur there. The size structure of the phytoplankton community ­varies regionally, with larger and therefore faster-sinking diatoms dominating the temperate nutrient-rich North Atlantic, and smaller and therefore slower-sinking phytoplankton such as the cyanobacteria Prochlorococcus and Synechococcus predominant in the nutrient-deplete subtropical North Atlantic. Two major phytoplankton groups produce mineral material: the coccolithophores, which form calcareous coccoliths through calcification, and the diatoms, which form opaline frustules through silicification. These ballast materials can potentially increase POC export through increasing density and therefore the sinking speed of the particles and by inhibiting organic carbon remineralization and therefore increasing the depth at which the particles are remineralized to CO2. In the euphotic zone of the Atlantic Ocean, up to 20% of organic carbon production can be  attributed to mineralizing phytoplankton (Poulton et  al., 2006). Rates of organic carbon production are at least an order

46 ecology of magnitude higher than rates of calcification, and rates of calcification are 2-fold higher than rates of silicification. Opal tends to dominate the biogenic mineral material at high latitudes, whereas calcite may be more abundant at low latitudes.

5.2.2  Balancing the budget Over an appropriate time period, the mesopelagic ocean is expected to be in steady state, with the sources of organic carbon (e.g. inputs from the surface ocean) balanced by the sinks (mesopelagic heterotrophic respiration and export to the bathypelagic). Hence determining the heterotrophic conversion of organic carbon to CO2, i.e. respiration, provides an important constraint on the export and storage of organic carbon in the deep sea. In addition, being able to apportion the respiration to either prokaryotes or zooplankton is relevant to predictions of future marine carbon storage. Unfortunately, due to methodological difficulties, poorly constrained conversion factors and environmental variability, some recent organic carbon budgets have not balanced, with deep-water respiration appearing to exceed the influx of organic substrates. The efficiency with which prokaryotes convert organic substrates into prokaryotic biomass (prokaryotic growth efficiency = net prokaryotic ­production/(net prokaryotic production + prokaryotic respiration)), the carbon content of a prokaryotic cell, the prevalence of chemoautotrophy, the lateral advection of POC, and the concentration of suspended and slowly sinking POC are the least well constrained parameters in the budget (Burd et al., 2010). However, a recent study using estimates rather than direct measurements of prokaryotic and zooplankton respiration balanced the budget of carbon sources and sinks at the PAP-SO in the North Atlantic, and showed that prokaryotes were responsible for up to 92% of mesopelagic respiration despite the fact that much of the organic carbon was exported as large, fast-sinking particles accessible to larger zooplankton (Giering et al., 2014).

5.2.3  Inorganic nutrient cycling The marine biogeochemical cycling of carbon (C) is inextricably linked to that of nitrogen (N) and phosphorus (P) (Figure 5.1), since N and P are required for protein and nucleic acid (DNA and RNA) production, which make up 50% and 5% respectively of a typical marine phytoplankton cell. A.C. Redfield first recognized the similarity between the particulate elemental composition (C:N:P ratio) of plankton and the C:N:P ratio of dissolved inorganic nutrients in the sea, and this global average 106C:16N:1P (mol:mol:mol) ratio became known as the Redfield ratio. However, since phytoplankton CO2 fixation and nutrient acquisition are relatively loosely coupled, this longterm global average obscures the wide range in phytoplankton elemental composition which can occur. Phytoplankton C:N:P can vary with taxonomic group, the ratio of N:P supply compared to the cellular elemental ratio, and the growth rate of the cells. For example, diatoms tend to have lower C:P and N:P ratios than cyanobacteria, dinoflagellates tend to have higher

C:N and C:P ratios than diatoms, C:P ratios tend to be inversely related to the environmental availability of P, and faster growing cells produce extra RNA, which lowers their C:P and N:P ratios. The cyanobacteria Trichodesmium have significantly higher N:P ratios when fixing N2, with C:N:P ratios of colonies collected from the Sargasso Sea averaging around 600:100:1 (Nuester et  al., 2012). Marine heterotrophic bacteria are enriched in N and P relative to C, with typical ratios of 50C:10N:1P. However, these ratios are also flexible, varying 2to 6-fold depending on species, temperature and composition of the organic substrate (DOC:DON:DOP) (DON = dissolved organic nitrogen; DOP = dissolved organic phosphorus). A recent study of Synechococcus, Prochlorococcus, low nucleic acid content bacteria such as SAR11, and picoeukaryotic phytoplankton in the nutrient-deplete North Atlantic found intracellular C:N ratios below but close to the Redfield C:N ratio of 6.6, but intracellular C:P and N:P ratios up to ten times higher than the Redfield C:P and N:P ratios (Grob et al., 2013). A global survey of phytoplankton and POM provided further evidence that marine organic stoichiometry varies systematically with ecosystem. The data showed a clear latitudinal trend in C:N:P, with ratios of 195:28:1 in warm nutrient-deplete low latitude gyres, 137:18:1 in warm nutrient-rich upwelling zones, and 78:13:1 in cold nutrient-rich high latitude regions (Martiny et al., 2013). The C to nutrient ratio of phytoplankton is both a major determinant of their quality as food for zooplankton and thus influences zooplankton biomass and production, with repercussions further up the food web, and of the quality of sinking particles available for prokaryotic remineralization and thus affects the C export and storage capacity of the ocean. The C:N:P ratios of microzooplankton such as heterotrophic dinoflagellates, which form the grazing link between bacteria and small phytoplankton and mesozooplankton, tend to trace the elemental ratios of their prey. These heterotrophs can accumulate N and P when nutrients occur in excess, through luxury consumption, and use these stores later when available prey are N and P deplete. By contrast, microzooplankton such as ciliates and mesozooplankton such as copepods maintain relatively constant elemental ratios despite the variability in stoichiometry of their prey, and tend to produce faecal pellets with relatively low N:P and C:P ratios (see Chapter 6). Due to preferential microbial remineralization of N and P, the C:N and C:P ratios of sinking particles increase with depth. DOM is particularly enriched in C and N compared to Redfield stoichiometry with recent global averages of 317C:39N:1P and 810C:48N:1P for the degradable and total DOM pools respectively (Letscher and Moore, 2015). The greater depletion of N and P in the dissolved compared to the particulate pool implies more efficient C export by DOM than POM.

5.3  The Nitrogen Cycle The marine N cycle is biologically controlled, with the reservoir of bioavailable N (also known as fixed N, i.e. all forms except

ph y topla nkton biogeochemica l cycles  47 nitrogen gas (N2)) determined by the balance between the microbial processes of nitrogen fixation in surface waters and heterotrophic denitrification and ammonium oxidation in low-oxygen deep waters. Nitrogen fixation, or the reduction of dissolved N2 gas to ammonium (NH4+), is undertaken by a diverse set of prokaryotes including the filamentous cyanobacteria Trichodesmium, the free-living unicellular cyanobacteria Crocosphaera, the symbiotic heterocystous Group A cyanobacteria and heterotrophic bacteria. These N2 fixing or diazotrophic plankton, can therefore thrive in waters where other forms of nitrogen are very low. Heterotrophic denitrification is the reduction of nitrate (NO3−) to N2, with nitrite (NO2−), nitric oxide (NO) and N2O as intermediates, alongside the metabolism of organic carbon to CO2, and can be undertaken by more than 60 genera of bacteria and Archaea (Canfield et al., 2010). Anaerobic ammonium oxidizing (anammox) bacteria fix CO2, alongside the oxidation of ammonia as an energy source and the reduction of nitrite as an electron acceptor, thereby producing N2. Heterotrophic denitrification and anammox therefore close the nitrogen cycle by returning nitrogen gas to the atmosphere. In addition to these microbially mediated sources and sinks of marine N, the predominant transformation of N in the ocean is by the nearly balanced rates of uptake by phytoplankton in surface waters and remineralization by prokaryotes in deep waters. Ammonium is the most energetically favourable source of N for phytoplankton, as it can be assimilated with minimal energy expenditure and therefore all phytoplankton can grow on NH4+ as the only N source. However, NO3− is much more abundant in the ocean than NH4+, with an oceanic inventory of 5.8 × 105 TgN compared to 340 TgN (Gruber, 2008). Therefore, most phytoplankton have the enzymes nitrate reductase and nitrite reductase, which are required to assimilate NO3−. Some examples of phytoplankton species which lack the ability to use nitrate as a source of N include particular clades of the cyanobacteria Prochlorococcus and Synechococcus. In the vast open ocean regions dominated by Prochlorococcus and Synechococcus, surface water dissolved inorganic nitrogen concentrations are very low, and so Prochlorococcus and Synechococcus have evolved to take up what may be the most abundant form of bioavailable nitrogen present, DON, in the form of amino acids (Zubkov et al., 2003). The uptake of the amino acids methionine and leucine by these cyanobacteria in the North Atlantic subtropical gyre (NASG) is significantly enhanced by visible light (Mary et al., 2008), and this photoheterotrophic mode of metabolism is now known to be ubiquitous in all low productivity regions of the global ocean (Evans et al., 2015). Most of the fixed organic nitrogen is returned back to nitrate by the remineralization processes of ammonification and nitrification. Ammonification is the transformation of organic N to NH4+ by heterotrophic prokaryotes which use the oxidation of organic C to CO2 as their source of energy. Ammonium can then serve as an energy source for chemoautotrophic prokaryotes, which oxidize it to nitrite (ammonia oxidation e.g. by

Nitrosomonas spp., Nitrospira spp. and Crenarchaea) and then nitrate (nitrite oxidation e.g. by Nitrobacter spp.), with N2O as a by-product. This two-step process of nitrification is an important source of atmospheric N2O, and N2O is more than 200 times more potent as a greenhouse gas than CO2 (Canfield et al., 2010). At the global scale, nitrification is thought to be the source of about half of the nitrate utilized by phytoplankton (Yool et al., 2007).

5.4  The Phosphorus Cycle In contrast to N, the reservoir of oceanic P is geologically ­controlled, being primarily a balance between river input and sediment burial. The dissolved inorganic P reservoir is predominantly orthophosphate (PO43−), while the DOP pool is a diverse mixture of organophosphates, organic phosphoanhydrides and organophosphonates. In most marine systems, the concentration of DOP is much larger than the concentration of phosphate, often by a factor of ten (Karl, 2014). For example, in the  NASG, phosphate concentrations are below 1 nmol L−1, whereas DOP concentrations range from 50 to 100 nmol L−1. Phytoplankton have a strong preference for the assimilation of phosphate over that of DOP due to the extra hydrolytic step required for DOP assimilation. However, when phosphate is  in  short supply, marine plankton such as Trichodesmium, Prochlorococcus, Synechococcus, and some species of dinoflagellates, coccolithophores, diatoms and bacteria, synthesize hydrolytic enzymes such as alkaline phosphatase in order to assimilate DOP (Mahaffey et al., 2014). In the NASG, Prochlorococcus and low nucleic acid content heterotrophic bacteria account for more than 90% of the uptake of the available phosphate, outcompeting the picoeukaryotic phytoplankton, which contribute only 1% to total microbial phosphate uptake (Zubkov et al., 2007). In order to satisfy their cellular requirements for P, therefore, these picoeukaryotes (also known as plastidic protists) acquire P by feeding on P-rich bacterioplankton cells (Hartmann et al., 2011). Recent work has shown that these plastidic protists are not only the main fixers of CO2 in the open ocean oligotrophic gyres, but are also the major bacterivores (Hartmann et  al., 2012). Once inside the plankton cell, P is incorporated into a range of low molecular weight organic compounds including nucleic acids and adenosine diphosphate and adenosine triphosphate (ADP and ATP). The cell quota of P and the distribution of P within the cell vary between taxa and in response to environmental conditions. The phosphate and DOP assimilated by marine plankton into particulate organic phosphorus is remineralized back to phosphate through the action of heterotrophic bacteria.

5.5  Nutrient Limitation Due to the existence of microbial N2 fixation, which can a­ lleviate N limitation, P is thought to be the ultimate limiting

48 ecology nutrient in the oceans over geological timescales. However, since N2 fixation can be limited by iron (Fe), N is often the shortterm limiting nutrient of non-diazotrophic phytoplankton. The short-term limiting nutrient can vary with microbial population, metabolic pathway and environmental conditions as shown by nutrient manipulation experiments where specific nutrients are added to natural microbial communities enclosed in sample bottles or directly to a water mass labelled with a measurable inert tracer such as sulphur hexafluoride (Moore et al., 2013). While phytoplankton photosynthesis in the tropical Atlantic increases in response to N addition, the greatest increase in N2 fixation occurs after addition of both Fe and P. In the subtropical North Atlantic where bioavailable phosphorus is severely depleted, phytoplankton photosynthesis and biomass increase to a greater extent when both N and P are added compared to the addition of either nutrient on its own, indicating nutrient co-limitation. A compilation of data from nutrient addition experiments reveals coherent basin-scale patterns, and a clear relationship between proximal nutrient limitation of phytoplankton activity and the concentrations of nutrients in the environment (Moore et al., 2013).

5.6  Impact of Increasing Atmospheric Carbon Dioxide on Plankton Mediated Biogeochemical Cycles Open ocean marine systems are increasingly subjected to multiple changes in environmental properties arising from human activity. These changing properties (or drivers) which can cause changes to the ecology and physiology of the plankton and thus biogeochemical cycles, include increases in temperature (which can lead to changes in water mixing and therefore light and nutrient availability, decreasing dissolved gas concentrations, and loss of sea-ice habitat) and increasing seawater CO2 (which leads to decreasing pH and the saturation state of ­carbonate).

5.6.1  Increasing sea surface temperature The ocean takes up > 90% of the heat being added to the Earth system through increasing atmospheric concentrations of anthropogenically derived greenhouse gases. Current predictions assume that atmospheric CO2 will reach 700 ppm by 2100, leading to an increase in seawater temperature of up to 6°C. Increasing seawater temperatures influence marine plankton directly through increasing physiological rates such as growth, reproduction and respiration, indirectly through differential physiological effects on different species or trophic groups thereby affecting predation and competition, and indirectly through changing the physical and chemical properties of the marine environment such as decreasing dissolved oxygen solubility, decreasing mixing of nutrient-rich deep waters with nutrient poor surface waters and decreasing salinity due to ice melt.

Time-series studies of plankton abundance and diversity, and short-term incubation experiments using natural plankton communities, have shown a range of direct effects of increasing temperature on phytoplankton and bacteria. These include a shift from diatom dominated to flagellate dominated phytoplankton communities, an increase in dominance of smaller celled phytoplankton species, an earlier onset of the peak in phytoplankton abundance, a greater increase in the activity of heterotrophic bacteria and zooplankton compared to phytoplankton, a reduction in the accumulation of POM, and an increase in the accumulation of DOM. These effects suggest a potential decrease in export and the efficiency of the organic carbon pump, but assuming that increased DOC is accompanied by increased RDOC, an increase in the microbial carbon pump. The indirect effects of increasing sea surface temperature include an increase in the intensity of stratification between warmer surface waters and cooler deep waters, and a shoaling of the depth of this thermocline. This reduction in vertical mixing reduces the resupply of N and P to surface waters from nutrient-rich deep water and inhibits the re-equilibration of low oxygen, high CO2 deep waters with the atmosphere. In high latitude regions where phytoplankton growth is limited by light rather than nutrients, increasing stratification is expected to increase phytoplankton photosynthesis by trapping phytoplankton in a shallower surface layer thereby increasing light availability, whereas in low latitude regions where phytoplankton are limited by nutrients but not light, increasing stratification is expected to exacerbate nutrient limitation leading to a decrease in phytoplankton photosynthesis (Riebesell et  al., 2009). The decrease in oxygen content of naturally occurring deep water oxygen minimum zones (so-called deoxygenation) could increase the extent of microbial denitrification and anammox, thereby reducing the inventory of fixed nitrogen, unless balanced by an appropriate increase in diazotrophic growth. Deoxygenation of bottom waters could also increase the phosphorus inventory by aiding release from sediments. Increasing global temperatures are expected to have broader impacts on, for example, ocean circulation, coastal upwelling, storm intensity, ice melt and dust deposition, which may all affect the stoichiometry of nutrient supply to phytoplankton and bacteria.

5.6.2  Increasing seawater carbon dioxide concentrations In addition to warming, the ocean absorbs ~ 30% of the anthropogenically derived atmospheric CO2, leading to increasing dissolved CO2 (so-called ocean carbonation) and decreasing seawater pH and carbonate ion concentration (so-called ocean acidification). Stimulatory effects of elevated CO2 on phytoplankton photosynthesis have been observed to differing extents in diatoms, coccolithophores, cyanobacteria and dinoflagellates, suggesting that increasing CO2 will alter competitive interactions between phytoplankton groups and result in changes to phytoplankton community composition. The precipitation of

ph y topla nkton biogeochemica l cycles  49 calcium carbonate (CaCO3) by marine phytoplankton such as coccolithophores to produce protective liths is facilitated by high pH and high carbonate ion concentration. Hence most strains of coccolithophore tested so far show a decrease in calcification in response to elevated CO2 (Riebesell and Tortell, 2011). Such a decrease in CaCO3 ‘ballast’ will reduce the sinking rate of dead coccolithophore cells and therefore reduce the depth at which the cellular organic material is respired back to CO2. CO2 produced at shallower depths will be able to mix into the surface ocean and so re-equilibrate with the atmosphere on shorter timescales (seasons to years rather than decades to centuries), thereby reducing the carbon storage capacity of the ocean and constituting a positive feedback to rising atmospheric CO2 concentrations. Manipulation experiments with plankton cultures and naturally occurring plankton populations suggest that increasing CO2 will also have a significant impact on the marine nitrogen cycle, with increases in global N2 fixation and decreases in nitrification (Hutchins et al., 2009). N2 fixation rates of Trichodesmium cultures grown at pCO2 levels of ~ 750 ppm were 35–65% higher than the N2 fixation rates of cultures grown at present day (~ 380 ppm) pCO2 levels. This positive relationship between pCO2 and N2 fixation rate occurred even in P-limited cultures. However, the positive relationship between pCO2 and N2 fixation rates of the unicellular cyanobacteria Crocosphaera only occurred in iron-replete cultures. When iron was limiting, increasing pCO2 had no effect on rates of N2 fixation. Increasing pCO2 is linked to decreasing nitrification rates of naturally occurring bacterial and archaeal nitrifiers, possibly due to decreases in the pH dependent ratio of NH3/NH4+ where NH3 is the preferred substrate of nitrification. At present, the P cycle appears less likely to be directly affected by rising pCO2, but will be indirectly affected by the anticipated changes in the C and N cycles. A survey of culture data suggests that the C:N and N:P ratios of most phytoplankton will either remain near Redfield or increase with increasing pCO2 (Hutchins

et al., 2009). However, since these effects were species specific, extrapolation to ecosystem responses is difficult.

5.6.3  Interactions between increasing temperature and increasing carbon dioxide Environmental drivers such as warming and ocean acidification do not act independently, but rather have additive, synergistic or antagonistic effects on plankton metabolism. Boyd and Brown (2015) identified three types of interaction: physicochemical interactions in the seawater, interactions that directly affect physiological rates of organisms, and interactions that occur through changes in food web dynamics such as predation and competition. Studies are increasingly assessing the interacting effects of multiple drivers, for example combinations of increasing temperature, increasing CO2, increasing ultraviolet light, and decreasing oxygen, but often do so for only one type of interaction, in one ecosystem and on short-term (acclimated) timescales. Understanding how biogeochemical cycles mediated by phytoplankton and bacteria might change in a changing environment requires future research which takes into account the long-term (adapted) response of ecosystems to all three types of interaction, and the interplay between them.

Ack now ledgements Figure 5.1 was developed during an infographics hack day (Infohackit) organized by Martin Johnson (University of East Anglia) and Peter Moore Fuller (MADE Agency) for the Marine Knowledge Exchange Network (http://marineknowledge.org. uk), which is supported by the NERC Impact Accelerator Award NE/L013401/1 to the University of East Anglia. Many thanks to Chris Bennett, Mark Ng, Elena Garcia-Martin, Jessie Gardner and Jon Bliss for their contributions to the design of Figure 5.1.

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Karl, D. M. (2014). Transformations of phosphorus in the sea: New views of an old cycle. Annual Review of Marine Science 6, 279–337. Lampitt, R. S., Salter, I., de Cuevas, B. A., et al. (2010). Longterm variability of downward particle flux in the deep northeast Atlantic: Causes and trends Deep-Sea Research II 57, 1346–61. Letscher, R. T., and Moore, J. K. (2015). Preferential remineralization of dissolved organic phosphorus and nonRedfield DOM dynamics in the global ocean: Impacts on marine productivity, nitrogen fixation, and carbon export. Global Biogeochemical Cycles 29, 325–40. Lochte, K., Ducklow, H. W., Fasham, M. J. R., and Steinen, C. (1993). Plankton succession and carbon cycling at 47oN 20oW during the JGOFS North Atlantic Bloom Experiment DeepSea Research II 40, 91–114. Lomas, M. W., Bates, N. R., Johnson, R. J., et al. (2013). Two decades and counting: 24-years of sustained open ocean biogeochemical measurements in the Sargasso Sea. DeepSea Research II 93, 16–32. Lønborg, C., Middelboe, M., and Brussaard, C. P. D. (2013). Viral lysis of Micromonas pusilla: impacts on dissolved organic matter production and composition. Biogeochemistry 116, 231–40. Mahaffey, C., Reynolds, S., Davis, C. E., and Lohan, M. C. (2014). Alkaline phosphatase activity in the subtropical ocean: insights from nutrient, dust and trace metal addition experiments. Frontiers in Marine Science 1, Article 73, http://dx.doi.org/10.3389/fmars.2014.00073. Martin, J. H., Knauer, G. A., Karl, D. M., and Broenkow, W. W. (1987). VERTEX: carbon cycling in the northeast Pacific. Deep-Sea Research A 34, 267–85. Martiny, A. C., Pham, C. T. A., Primeau, F. W., et  al. (2013). Strong latitudinal patterns in the elemental ratios of marine plankton and organic matter. Nature Geoscience 6, 279–83. Mary, I., Tarran, G. A., Warwick, P. E., et  al. (2008). Light enhanced amino acid uptake by dominant bacterioplankton groups in surface waters of the Atlantic Ocean. FEMS Microbial Ecology 63, 36–45. Moore, C. M., Mills, M. M., Arrigo, K. R., et al. (2013). Processes and patterns of oceanic nutrient limitation. Nature Geoscience 6, 701–10. Nuester, J., Vogt, S., Newville, M., Kustka, A. B., and Twining, B. S. (2012). The unique biogeochemical signature of the marine diazotroph Trichodesmium. Frontiers in Microbiology 3, 150 doi: 10.3389/fmicb.2012.00150. Parekh, P., Dutkiewicz, S., Follows, M. J., and Ito, T. (2006). Atmospheric carbon dioxide in a less dusty world. Geophysical Research Letters 33, L03610, doi:10.1029/ 2005GL025098. Poulton, A. J., Sanders, R., Holligan, P. M., et  al. (2006). Phytoplankton mineralization in the tropical and subtropical Atlantic Ocean. Global Biogeochemical Cycles 20, doi:10.1029/ 2006GB002712.

ph y topla nkton biogeochemica l cycles  51 Riebesell, U., and Tortell, P.D. (2011). Effects of ocean acidification on pelagic organisms and ecosystems. In Ocean acidification, pp. 99–121. Oxford University Press, Oxford. Riebesell, U., Kortzinger, A., and Oschlies, A. (2009). Sensitivities of marine carbon fluxes to ocean change. Proceedings of the National Association of Sciences 106(49), 20602–9. Robinson, C., Steinberg, D. K., Anderson, T. R., et al. (2010). Mesopelagic zone ecology and biogeochemistry—a synthesis. Deep-Sea Research II 57, 1504–18. Savidge, G., Turner, D. R., Burkill, P. H., et al. (1992). The BOFS 1990 Spring Bloom Experiment: Temporal evolution and spatial variability of the hydrographic field. Progress in Oceanography 29, 235–81. Smyth, T., Atkinson, A., Widdicombe, S., et  al. (2015). The Western Channel Observatory. Progress in Oceanography 137, 335–41. Stocker, R., and Seymour, J.R. (2012). Ecology and physics of bacterial chemotaxis in the ocean. Microbiology and Molecular Biology Reviews 76(4), 792–812.

Suttle, C. A. (2007). Marine viruses—major players in the  global  ecosystem. Nature Reviews Microbiology 5, 801–12. Widdicombe, C. E., Eloire, D., Harbour, D., Harris, R. P., and Somerfield, P. J. (2010). Long-term phytoplankton community dynamics in the Western English Channel. Journal of Plankton Research 32, 643–55. Yool, A., Martin, A. P., Fernandez, C., and Clark, D. R. (2007). The significance of nitrification for oceanic new production. Nature 447, 999–1002. Zubkov, M. V., Fuchs, B. M., Tarran, G. A., Burkill, P. H., and Amann, R. (2003). High rate of uptake of organic nitrogen compounds by Prochlorococcus cyanobacteria as a key to their dominance in oligotrophic oceanic waters. Applied and Environmental Microbiology 69, 1299–304. Zubkov, M. V., Mary, I., Woodward, E. M. S., et  al. (2007). Microbial control of phosphate in the nutrient-depleted North Atlantic subtropical gyre. Environmental Microbiology 9(8), 2079–89.

CH A PTER 6

ZOOPLA NKTON BIOGEOCHEMICA L CYCLES Debor a h Steinberg

6.1 Introduction The structure of planktonic communities profoundly affects particle export and sequestration of organic material (the biological pump) and the chemical cycling of nutrients (Michaels and Silver,  1988; Peinert et  al.,  1989; Legendre and Le Fevre, 1995; Wassmann, 1998; Ducklow et al., 2001). This chapter describes the integral and multifaceted role zooplankton (both protozoan and metazoan) play in the export and cycling of elements in the ocean (Box  6.1), with an emphasis on the North Atlantic Ocean and adjacent seas. Zooplankton consume a significant proportion of primary production across the world’s oceans, and through their metabolism play a key role in the recycling of carbon, nitrogen, and other elements. Global syntheses of grazing impact on autotrophic production indicate that microzooplankton ( 200 μm, including the abundant crustaceans, such as copepods) consuming on average 10% and 40% of the daily primary production for high and low primary productivity regions, respectively (Calbet,  2001, 2008). The various metabolic pathways that mediate biogeochemical cycling of organic matter once it is consumed by ­zooplankton include respiration, excretion, and egestion (with ‘sloppy feeding’ a consideration for mesozooplankton as well). Because of their higher growth rates, grazing rates, and weight-specific metabolism compared to mesozooplankton, microzooplankton may also be the primary regenerators of ammonia (NH4+) in marine systems (Steinberg and Saba, 2008). In contrast, mesozooplankton egestion of sinking faecal pellets make them relatively more important in vertical export and the biological pump. The biological pump is comprised of a suite of biologically-­ mediated processes by which CO2 that is fixed by phytoplankton photosynthesis in the euphotic zone is exported to the deep

ocean, regulating (along with physical processes) air-sea CO2 exchange, carbon export, and ultimately carbon sequestration (Ducklow et al., 2001; Sanders et al., 2014). Zooplankton play a key role in the biological pump by feeding in surface waters and producing sinking particles such as faecal pellets (Turner, 2015) and by actively transporting dissolved and particulate matter to depth via diel vertical migration (e.g. Longhurst et  al.,  1990; Steinberg et  al.,  2000) (Box  6.1). Zooplankton faecal pellets, molts, mucous products, and carcasses help support the metabolism of deep-sea pelagic plankton and fish, as well as benthic communities. Zooplankton also affect the attenuation of sinking particle flux with depth through their feeding and metabolism of sinking POC in the mesopelagic zone (Steinberg et al., 2008), affecting the efficiency by which POM is exported and the sequestration of carbon in the deep ocean. Human or climate-influenced changes in zooplankton communities are becoming well documented, and include ­ shifts in zooplankton species abundance, size-structure, distribution, and phenology (timing of important life cycle events) (Beaugrand et al., 2002; Richardson, 2008; Mackas et al., 2012). Thus, this chapter concludes by addressing how human or ­climate-influenced changes in North Atlantic zooplankton populations may in turn drive changes in zooplankton-mediated biogeochemical cycling.

6.2  Grazing, Metabolism, and Nutrient Cycling 6.2.1 Grazing Microzooplankton are important grazers in North Atlantic food webs, consuming a high proportion of the primary production. Most grazing studies have focused on northern temperate waters during the North Atlantic spring bloom, and the waters of the North Atlantic subtropical gyre. During the North Atlantic spring bloom, the microzooplankton community is

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

zoopla nkton biogeochemica l cycles  53 Box 6.1  Zooplankton and biogeochemical cycling

CO2 grazing phytoplankton

zooplankton

respiration excretion

predation

dissolved organic carbon physical mixing

aggregation formation

sinking aggregates dissolved organic carbon

decomposition bacteria

sinking faecal pellets

diel vertical mitigation sinking carcasses

consumption

respiration

zooplankton

excretion

jelly fall seabed

The role of zooplankton in biogeochemical cycling and the ‘biological pump’. Dissolved inorganic carbon (DIC), carbon dioxide (CO2) and inorganic nutrients are fixed during photosynthesis by phytoplankton in the euphotic zone. Phytoplankton are consumed by herbivorous micro- and mesozooplankton, which through their metabolism (respiration and excretion) regenerate dissolved nutrients and CO2, and excrete dissolved organic carbon (DOC) that is used by phytoplankton or bacteria. Zooplankton egest faecal pellets, which can be reingested, sink, or incorporated into larger sinking aggregates along with phytoplankton. Zooplankton diel vertical migrators feed in surface waters at night and metabolize the food they ingested in the mesopelagic zone during the day. Here mesopelagic zooplankton (along with bacteria) affect particle flux by ingesting and metabolizing sinking aggregates and other particles, leading to the attenuation of sinking particulate organic carbon (POC) with increasing depth, such that only a small fraction of the sinking particulate organic matter (POM) leaving the euphotic zone makes it to the seabed. Dead zooplankton are also part of the particle flux. This includes carcasses of larger gelatinous zooplankton that form blooms in surface waters and sink en masse forming ‘jelly falls’ on the deep-sea benthos. DOC produced in the euphotic zone is partially consumed by bacteria and respired, and the remaining is advected and mixed into the deep sea. Vertical, block arrows denote components of the biological pump. Figure adapted from previous representations by the author incorporated into a Joint Global Ocean Flux Study cartoon, and from Figure 1 in Lebrato and Jones (2011). Phytoplankton and zooplankton images are from the Integration & Application Network (http://ian.umces.edu/symbols/).

54 ecology usually dominated by aloricate ciliates (Verity et  al.,  1993; Stoecker et  al.,  1994; Fileman and Leakey,  2005; Rose et  al., 2009) and occasionally by heterotrophic dinoflagellates and ­tintinnids (Verity et al., 1993; Stelfox-Widdicombe et al., 2000). Microzooplankton herbivory accounted for up to 115% of daily phytoplankton production during the North Atlantic spring bloom as well as in summer (Burkill et  al.,  1993; Verity et al., 1993; Gifford et al., 1995; Rose et al., 2009). This efficient grazing of primary production and subsequent recycling in ­surface waters by microzooplankton implies little available material for export from the surface mixed layer, at least during some phases of the bloom (Verity et  al.,  1993). In contrast, ­mesozooplankton (copepod) herbivory removed up to 5% of daily primary production (3% on average), although metabolic demand estimates suggest phytoplankton contributed only about half of the daily C and N required by mesozooplankton, implying microzooplankton or detritus were important food sources as well (Dam et  al.,  1993). Comparison between the temperate versus subtropical North Atlantic in summer indicated higher mean microzooplankton grazing rates on primary production in the more productive temperate region (1.25 d–1) than the ­oligotrophic subtropics (0.43 d–1) (Stelfox-Widdicombe et al., 2000). This is consistent with a pattern observed along onshore-offshore transects measuring microzooplankton grazing in the subtropical north-east Atlantic Ocean, which indicated higher mean grazing rates in the productive north-west African coastal upwelling region (0.68 d–1) versus oceanic waters of the oligotrophic subtropical gyre (0.37 d–1) (GutiérrezRodríguez et  al.,  2011). These patterns have been attributed to differences in phytoplankton community structure resulting in changes in the efficiency of grazing by microzooplankton (Stelfox-Widdicombe et  al., 2000; Gutiérrez-Rodríguez et  al., 2011). Mesozooplankton (copepod) grazing in the oligotrophic subtropical North Atlantic during spring was on average 22% of daily primary production, and 61% of the primary production for the larger (> 2 μm) phytoplankton only (Huskin et al., 2001).

6.2.2  Metabolism and contribution to requirements for phytoplankton growth Organic matter ingested by zooplankton may be absorbed through the gut and taken up into the organism’s tissues, and subsequently respired as CO2, excreted as dissolved inorganic or organic matter, or put into growth and reproduction. That which is not absorbed is egested as faecal pellets (or lost via ‘sloppy feeding’ as dissolved organic matter). Respiration is a major sink for organic matter in the ocean (del Giorgio and Duarte, 2002), and it is estimated that global, full ocean depth integrated mesozooplankton respiration accounts for 13 Gt C y−1, equivalent to ~ 17–32% of global ­primary production (based on 41–77 Gt C y−1 global primary production; del Giorgio and Duarte,  2002) (Hernandez-Leon and Ikeda, 2005). Thus, zooplankton respiration is a major component of the ocean carbon cycle. Respiration rates of mixed

mesozooplankton or copepods in the subtropical and tropical North Atlantic range from 0.4 to 0.75 μmol O2 mgdw−1h−1 (studies compiled in Table 2 of Lehette and HernándezLeón, 2010). Isla et al. (2004) measured size-fractioned mesozooplankton grazing and metabolism in early autumn along a transect from 50° N to 30° S in the North Atlantic. They found weight-specific respiration rates did not differ significantly between provinces despite a wide range in temperature with latitude (14–24°C), and that total zooplankton C respired was  highest in the eastern tropical Atlantic around the equator,  corresponding to the highest zooplankton biomass (Isla et al., 2004). Zooplankton excretion is a major recycling pathway for N, P, and trace elements, and along with bacterial remineralization, determines the amount of regenerated N available for phytoplankton production. Both macro- and microzooplankton excrete N primarily as ammonia (NH4+), but dissolved organic nitrogen (DON) (e.g. urea, amino acids) can also be a substantial component, contributing 7–89% of the total dissolved N excreted (reviewed in Steinberg and Saba,  2008). Thus, zooplankton are not only important regenerators of N for phytoplankton uptake, but also support bacterial metabolism via excretion of DON. Marine protozoa are the principal regenerators of N, excreting ammonia hundreds to thousands of times their body weight per day (compared to macrozooplankton, which generally excrete < 10% body N d−1) (reviewed in Steinberg and Saba, 2008). Many studies have determined the contribution of zooplankton excretion to phytoplankton production in the North Atlantic (reviewed in Bronk and ­ Steinberg,  2008, for N). For example, in the aforementioned study of North Atlantic mesozooplankton grazing and meta­ bolism along a 50° N–30° S transect, mesozooplankton excretion of NH4+ provided 31–121% of the N, and phosphate (PO43−) 35–83% of the P, required for primary production (Isla et  al.,  2004), indicating the important role of zooplankton in nutrient recycling (Fig. 6.1). Zooplankton faecal pellets can be a major source of POM exported from surface waters (e.g. Wassmann et  al.,  2000; Wexels Riser et al., 2010; see Section 6.3.1) and also of dissolved organic matter via leaching of unabsorbed digestive products from pellets ( Jumars et  al.,  1989; Lampitt et  al., 1990). Due to their high organic content, faecal pellets are also ‘hot spots’ of microbial activity (Smith et  al.,  1992; Thor et  al.,  2003; Tang, 2005). Zooplankton community structure has an important effect on the export of faecal pellets, as pellets from different taxa sink at different rates due to differences between taxa in faecal pellet size, shape, and density—the latter also a result of diet differences (Wilson et al., 2008). For example, Yoon et al. (2001) measured sinking rates of faecal pellets produced by a variety of macro- and mesozooplankton taxa in the north-­ eastern tropical Atlantic, which ranged from 27–160 m d−1 for copepod pellets, 16–341 m d−1 for euphausiids, 44–1170 m d−1 for salps, 65–206 m d−1 for a pteropod, and 120–646 m d−1 for a heteropod. This two order of magnitude range in sinking rates, from ten’s m d−1 for small copepod pellets to 1000 m d−1 for salp

zoopla nkton biogeochemica l cycles  55 ETRA

Gd

CNRY

NAST

NADR

Prim. prod. % N excr. % P excr.

1200

150%

900 100% 600 50%

300 0

% excr.

Prim. prod. (mg C m–2 . d–1)

SATL

–30 S

–20

–10

0

10

20

Latitude

30

40

0%

50 N

FIGURE 6.1:  Percentage of phytoplankton nitrogen and phosphorus demand potentially met by mesozooplankton excretion of NH4+ (%N

excr.) and PO43− (%P excr.) along a transect of the Atlantic Ocean. Oceanic provinces or zones traversed in the transect (from left to right) are: South Atlantic Gyral (SATL), Eastern Tropical Atlantic (ETRA), Guinea dome transition zone (Gd), Canary Coastal (CNRY), North Atlantic Subtropical Gyral (NAST), and North Atlantic Drift (NADR). Figure from Isla et al. (2004).

pellets, is representative of the range seen in taxa across a ­variety of environments (reviewed in Turner, 2002, 2015). Microzooplankton such as protozoans and copepod nauplii also produce ‘mini pellets’ generally < 100 μm in diameter that can contribute to export (Gowing and Silver, 1985; Nöthig and Bodungen, 1989; Gowing et al., 2001). While sinking, faecal pellets may be ingested or fragmented by protozoa, copepods, or larger organisms, providing an important food source for these animals but also affecting export and the efficiency of the biological pump (Lampitt et al., 1990; Noji et al., 1991; Poulsen and Kiørboe, 2005; Poulsen and Iversen, 2008; Wilson et al., 2008). The contribution of faecal pellets to vertical flux and the biological pump will be further addressed in Section 6.3.1.

6.2.3  Elemental stoichiometry and nutrient cycling Heterotrophs such as herbivorous zooplankton maintain relatively constant body elemental ratios (‘homeostasis’), in contrast to the more plastic stoichiometry of their phytoplankton prey, which reflects the ambient nutrient environment (Andersen and Hessen, 1991; Sterner and Elser, 2002; Persson et al., 2010). Because of this, zooplankton elemental composition, and that of their prey, regulates the elemental ratio of products released from zooplankton – such as faecal pellets and excreted dissolved inorganic and organic matter, affecting nutrient cycling and export. Thus a change in either zooplankton or their prey can result in changes in the quantity and bioavailability of regenerated nutrients (e.g. Caron and Goldman,  1990; Pertola et al., 2002), as zooplankton may release C, N or P in different ratios than are present in their phytoplankton prey in order to maintain stoichiometric homeostasis (i.e. a steady chemical composition).

In the subarctic North Atlantic, seasonal changes in C:N:P stoichiometry of the high-latitude, lipid-storing copepod Calanus finmarchicus was measured to investigate changes in dietary demands with season, lipid storage, and growth stage (Aubert et al., 2013). Body C:N and C:P ratios within stages, and without considering the C-rich lipid storage pool, remained constant through end of winter, bloom, and post-bloom despite fluctuations in elemental ratios of the seston food. Copepod faecal pellet N:P and C:P ratios, however, were always lower than their food, suggesting copepods preferentially retained N and C over P, potentially making copepods suppliers of P-enriched particles for export (Aubert et  al.,  2013). Excretion and recycling of elements by zooplankton is often non-Redfield (Steinberg and Saba, 2008), as shown by a wide range of elemental ratios of respiration and inorganic excretion by zooplankton in the Barents Sea: C:N (range 4–44), N:P (range 2–45) and C:P (range 59–910) (calculated from Table 3 in Ikeda and Skjoldal, 1989). Excretion ratios of inorganic (PO4– P:NH4–N) and organic matter (DOC:DON) by diel vertically migrating zooplankton in the Sargasso Sea ranged from 6 to 16, and 5 to 15, respectively, indicating non-Redfield recycling of POM (Steinberg et al., 2002). The stoichiometry of export flux via diel vertical migrators (ingesting prey in surface waters at night and metabolizing below the mixed layer in the day) may contribute to nutrient ratio anomalies seen below the euphotic zone in the North Atlantic subtropical gyre and Sargasso Sea (Steinberg et al., 2002; Anderson and Pondaven, 2003), and elsewhere such as the North Pacific subtropical gyre, where diel vertical migrator excretions are enriched in P  relative to the sinking particle flux, suggesting that active transport may contribute to enhanced P-limitation of primary production documented in surface waters of this region (Hannides et al., 2009).

56 ecology

6.3  Zooplankton and the Biological Pump

The sinking of dead zooplankton could also be a substantial component of the biological pump, although it is not considered in most studies using formalin-poisoned sediment traps, 6.3.1  Zooplankton particle production due to the difficulty of distinguishing carcasses from ‘swimZooplankton produce a variety of sinking particles such as fae- mers’ that entered traps live and died there (Buesseler cal pellets, mucous products (e.g. feeding webs), and molts et  al.,  2007; Frangoulis et  al.,  2011; although see Sampei (crustacean exoskeletons); zooplankton carcasses also can be a et al., 2009 for a method using post-mortem position of copesubstantial POC export (Box 6.1). Faecal pellet production is a pod appendages to make this distinction). Copepod carcasses, variable but important component of the sinking particle flux however, can be abundant in the plankton (Tang et al., 2006), in the North Atlantic and adjacent seas, as shown in Table 6.1. and large accumulations of dead gelatinous zooplankton have There seems to be a general trend of a higher contribution of been noted on the benthos (reviewed in Lebrato et al., 2012). In faecal pellet flux to total sinking POC flux across depths 200 m the western Mediterranean Sea, carbon and nitrogen flux of and shallower in Arctic and subarctic regions of the North zooplankton carcasses measured in a shallow bay by a sediment Atlantic versus subtropical and tropical regions, but the results trap (with a swimmer exclusion device) moored at 38 m are variable and there are fewer studies at these depths in the exceeded that of faecal pellet flux at all times, with the exceplatter two regions (Table 6.1). Faecal pellet flux as a component tion of the beginning of the spring bloom during which faecal of the sinking POC flux clearly does not always decline with pellet flux exceeded other sinking particle types (Frangoulis increasing depth, and often increases, indicating differential set- et al., 2011). The contribution of sinking particle types was also tling of different particle types, or reprocessing of sinking mate- measured in the Gulf of St Lawrence in the eastern North rial (Wilson et al., 2008). Atlantic at 50 m and 150 m in short-term (24-hour) deployments Mesoscale physical processes are important in affecting (Romero-Ibarra and Silverberg, 2011). This study reported a zooplankton faecal pellet export in the North Atlantic. smaller contribution of POC from zooplankton and their body ­ Phytoplankton blooms induced by cyclonic and mode-water parts compared to faecal pellets, although carcasses per se were mesoscale eddies in the Sargasso Sea increased zooplankton not considered (individual zooplankton > 1 mm were considbiomass in the interior of the eddies, resulting in faecal pellet ered swimmers and eliminated from flux calculations) and only flux across 150 m that was 1.5-fold higher inside than outside fragments or exoskeletons (molts) were included. Faecal pelthe cyclonic eddy. Faecal pellet flux accounted for 9% and 12% lets, followed by sinking phytoplankton, were the major of the total POC flux in the cyclonic and anticyclonic, mode-­ ­components of the POC flux, with occasional high flux of water eddy, respectively (Goldthwait and Steinberg, 2008). microzooplankton (tintinnids) (Romero-Ibarra and Silverberg, Faecal pellet flux and size distribution were also measured in 2011). sediment trap samples from 500, 1500 and 3200 m depths in the Gelatinous zooplankton such as pelagic tunicates (salps, Sargasso Sea during a 1-year period in which three mesoscale pyrosomes, doliolids), ctenophores, and cnidarians (medusae, eddies transited through the region (Shatova et al., 2012). Faecal siphonophores) can rapidly form large blooms due to fast pellet contribution to total POC flux was 3–16% of total POC growth rates and life histories that include an asexual phase. flux at 1500 m, with changes in pellet flux and size (increasing or Although low in individual carbon and nitrogen content comdecreasing, depending on the eddy)  following passage of the pared to crustacea (Lucas et al., 2011), gelatinous zooplankeddies attributed to changes in zooplankton abundance and ton sinking ‘en masse’ at the end of a bloom—jelly falls—has assemblage structure (Shatova et al., 2012). been recorded around the world, and may be a substantial, Mucous feeding webs produced by some zooplankton, such rapid export of POM to the benthos (Lebrato et  al.,  2012). as appendicularians (larvaceans), pteropods, and foraminifera In  the North Atlantic Ocean and marginal seas, Lebrato contribute to the sinking particle flux as well. The mucous and  Jones (2009) reported a massive jelly fall of pyrosomes ‘houses’ of appendicularians are used to concentrate food parti- (Pyrosoma atlanticum) on the benthos (at depths of 200 m to at cles, and when the house filters become clogged, the larvacean least 1275 m) off the west coast of Africa, and numerous discards the house with its associated particles (Alldredge et al., P. atlanticum jelly falls have occurred in the Mediterranean Sea 2005). Larvaceans may secrete and discard up to 26 houses d-1 (Bertrand et al., 2002; Lebrato et al., 2012). Salp jelly falls are (Sato et al., 2001) that sink at rates of 10–800 m d-1, depending known from the oligotrophic Sargasso Sea (Wiebe et  al., upon the house size, age, and other factors (Robison et al., 2005; 1979), and scyphozoan jelly falls have been reported in a Lombard and Kiørboe,  2010). Discarded appendicularian Norwegian Sea fjord (Sweetman and Chapman, 2011) where houses contribute 12–83% to total POC flux from surface baited trap studies indicate jelly carcasses are rapidly scavwaters as determined for a range of eutrophic coastal to olig- enged by deep-sea benthic consumers (Sweetman et al., 2014). otrophic open ocean environments, and in the Bay of Biscay Estimating the contribution of jelly falls—not sampled by traand Mediterranean (Ligurian Sea) contribute 17% and 32%, ditional methods such as sediment traps—to the biological respectively, of the total POC flux (Alldredge et al., 2005, and pump will require cameras and other technology (Lebrato references therein). et al., 2012).

zoopla nkton biogeochemica l cycles  57 TABLE 6.1:  Faecal pellets as a percentage of total sinking particulate organic carbon flux measured by sediment traps in the North Atlantic Ocean and adjacent seas.

Location

Depth (m)

% Faecal pellets mean (range)

Reference

9 depths, 20–200 9 depths, 20–200 90 90 90 150 7 depths, 20–200 340 340 340 340 20, 50, 100 15 50 100 200 1430 200 250–300 950–1000 180 150 50

10 (1–37) 21 (2–8, March) (17–40, May) (30–40, July) 3 10 (max = 24) (10–30, April, May) > 85, May (40–70, May–June) 10, June, July 12 2.6 28.8 28.4 21 (max = 45/63, July/Aug) 92 (max = 66) 90) (70– > 90) (70– > 90)

42 200 1500 38 50 150 250 200 500 1000 2000 1000

200 μm) biomass recorded using data from BATS (Steinberg et al., 2012). Over a 17-year period (1994–2010), total mesozooplankton biomass increased by 61%. The increase was positively correlated with sea surface temperature, water-column stratification, and primary production, and tied to multi-decadal climate indices such as the North Atlantic Oscillation. The most likely cause for the increase is bottom-up control by smaller phytoplankton, which have also increased in biomass and production at BATS, translating up the microbial food web to mesozooplankton. As a result of this zooplankton biomass increase, changes in the role of zooplankton in biogeochemical cycling at BATS have occurred. For example, night-time biomass increased at a faster rate than daytime biomass, resulting in a 92% increase in diel vertical migrator biomass (defined as night minus day biomass in the epipelagic zone) over the study period. This increase in diel vertical migrator biomass over time resulted in a near doubling of active transport of C over the 17-year period. The contribution of zooplankton faecal pellets to POC export tripled

over the first decade of the time series, then subsequently decreased. Because total POC export as measured by sediment traps did not significantly increase over the same time period, the relative importance of both diel vertical migration and faecal pellet export compared to total passive flux of POC increased over time (Steinberg et al., 2012) (Fig. 6.2). Long-term changes in epipelagic zooplankton abundance, distribution, and community structure in the North Sea have occurred in response to climate warming and changes in hydrography, as indicated by analyses of Continuous Plankton Recorder (CPR) survey data from 1958 to the present (Beaugrand et  al.,  2002; Beaugrand,  2009; Alvarez-Fernandez et al., 2012). While there is a systematic, subdecadal alteration in the abundance of the boreal copepod C. finmarchicus and the warm-water species C. helgolandicus tied to the North Atlantic Oscillation and to sea surface temperature (during the period 1958–2000; Reid et  al., 2003), the long-term, overall trend is increasing replacement of C. finmarchicus by C. helgolandicus that is also significantly positively correlated with sea surface temperature (Beaugrand et al., 2009). These and other changes that could affect carbon export in the North Sea are discussed by Beaugrand (2009). An increase in phytoplankton (derived from the phytoplankton colour index determined with the CPR) has accompanied the decrease in the abundance of the herbivorous species C. finmarchicus. Beaugrand (2009) suggests as a result there has been a decrease in grazing pressure on phytoplankton resulting in increased export of diatom aggregates; thus there should be increasing carbon export in the North Sea as the climate continues to warm. Another long-term trend may have the opposite effect on carbon export; the mean size of calanoid copepods has decreased in the North Sea. As biogenic carbon originating from smaller copepods in warmer regions cycles at a faster rate (e.g. smaller, slower-sinking faecal pellets and increased weight-specific metabolism of small versus large copepods), the now warmer North Sea system may be changing into an increasingly recycling, as opposed to exporting, environment (Beaugrand et  al.,  2009). Other regions in the North Atlantic may also be shifting towards an increasingly recycling, lower export environment. For example, in the tropical North Atlantic, a 10-fold decrease in epipelagic mesozooplankton biomass from the 1950s to 2000 has been attributed to a decrease in primary production due to increased stratification caused by warming (Piontkovski and Castellani,  2009). This decrease in mesozooplankton coupled with reduced mixing would be predicted to decrease export. There are limited studies on long-term changes in microzooplankton in the North Atlantic, but a study of tintinnid (loricate ciliate) populations from 1960 to 2009 shows the presence of tintinnids in CPR samples has increased in the north-east Atlantic but decreased in the North Sea (Hinder et al., 2012). An experiment testing the effects of warming temperatures and increasing pCO2 on microzooplankton abundance and grazing during the North Atlantic spring bloom indicated that increases in each parameter alone, as well as combined, led to changes in  both phytoplankton and microzooplankton community

60 ecology A

16 14 12 10 8 6

POC flux (gC m–2 y)

4 2 0 1994

B

1997

2000

2003

2006

2009

3 2.5 2 1.5 1 0.5 0 1994

C

Active transport flux (gC m–2 y)

1997

2000

2003

2006

2009

7 6 5 4 3 Egestion (gC m–2 y) 2 1 0 1994

1997

2000

2003

2006

2009

Years FIGURE 6.2:  Long-term changes in zooplankton-mediated export at the Bermuda Atlantic Time-series Study (BATS) station in the subtropical North Atlantic. A, Annual sinking particulate organic carbon (POC) flux across 150 m as measured by sediment traps; B, active transport by diel vertical migrators across 150 m (includes carbon dioxide respiration + dissolved organic carbon (DOC) excretion + POC egestion at depth); C, faecal pellet production by zooplankton in the top 150 m. Figure redrawn from Steinberg et al. (2012), with data points plotted at midpoint of each year.

zoopla nkton biogeochemica l cycles  61 structure, as well as microzooplankton grazing, with changes in  microzooplankton mostly a result of bottom-up control by phytoplankton, rather than changes in microzoooplankton physiology (Rose et al., 2009). These shifts in microzooplankton abundance and community structure could lead to changes in grazing and nutrient regeneration by microzooplankton, and thus in retention or export of carbon and nutrients in food webs in surface waters of different regions of the North Atlantic. Much of what has been predicted in terms of changes in biogeochemical cycling in the North Atlantic due to long-term changes in zooplankton are changes in export from the euphotic zone. Changes in flux through the mesopelagic zone, to the benthos, or in bentho-pelagic coupling, may also be occurring. Lomas et al. (2010) show a 2-fold increase in the attenuation of sinking POC in the upper mesopelagic zone (between 150 and 300 m) during the winter-spring bloom period at BATS from 1997 to 2007. This loss of sinking POC is accompanied by an increase in apparent oxygen utilization, which must be attributed to increased metabolism of sinking POC by mesopelagic zooplankton or bacteria. Although the increase in vertical migrator biomass at BATS suggests mesopelagic mesozooplankton have increased (Steinberg et al., 2012), the lack of a long-term time series of mesopelagic zooplankton makes the mechanism for flux attenuation hard to determine. A decrease in POC flux is predicted to have significant effects on deep-sea benthic community biomass, diversity and function (e.g. Billett et al., 2001; Ruhl and Smith, 2004; Smith et al., 2008). A long-

term increase in the relative abundance of meroplankton ­compared to holoplankton in the North Sea may also be leading to increases in bentho-pelagic coupling (Kirby et  al.,  2007; Beaugrand,  2009), which would also affect cycling of energy and nutrients in the deep ocean. Clearly, the North Atlantic Ocean is experiencing substantial changes in food web structure in response to changes due to climate warming and other anthropogenic influences. How the role of zooplankton in biogeochemical cycles is likely to change as a result is still largely unknown, but certainly predictive biogeochemical models that incorporate zooplankton from different major groups (e.g. Buitenhuis et al., 2006), and experimental work designed to investigate mechanisms leading to observed or predicted changes (e.g. Rose et al., 2009), are important ways forward.

Ack now ledgements Many thanks to Jeanna Hudson, Miram Gleiber, and Joshua Stone who helped with reference search and collection, and to JH for compiling the bibliography. The writing of this chapter was supported in part by grants from the United States National Science Foundation Chemical and Biological Oceanography Programs, award OCE-1258622, and the Division of Polar Programs, award PLR-1440435. This is contribution number 3567 from the Virginia Institute of Marine Science.

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CH A PTER 7

PLA NKTON A ND GLOBA L CH A NGE M a rtin Edwa r ds

7.1 Introduction Global change caused by human activities has had large consequences for the Earth’s biosphere through such effects as climate warming, pollution, loss of biodiversity, unsustainable exploitation of resources, loss of habitats, and alterations to nutrient cycles. These changes have accelerated over the last 50 years as human populations have sharply grown, coupled with unsustainable economic practices. In open ocean environments far from coastal zones it could be argued that human influences such as habitat loss and fragmentation, a strong driver of terrestrial environmental change, are fairly minimal. However, climate warming can actually alter the physical structure of the marine habitat through effects on the stability of the water column, which highly influences the overall biological production and the type of biological communities that exist there. Furthermore and unique to the marine environment, anthropogenic carbon dioxide (CO2) is also associated with ocean acidification. Ocean acidification has the potential to affect the process of calcification, and therefore certain planktonic organisms dependent on calcium carbonate for shells and skeletons (e.g. coccolithophores, foraminifera, pelagic molluscs, echinoderms) may be particularly vulnerable to increasing CO2 emissions. The marine pelagic realm, the habitat for planktonic organisms, is the largest ecological habitat on the planet, occupying 71% of the planetary surface, fuelling the vast majority of marine biological production, and driving marine food webs around the planet. It is estimated that plankton make up 98% of the biomass of all ocean life. Occupying such a large part of the Earth’s biosphere, the pelagic realm also plays a fundamental role in modulating the global environment via its regulatory effects on the Earth’s climate and its role in planetary oxygen production, carbon drawdown and other biogeochemical cycling. Changes to marine pelagic communities caused by increased warming are likely to have important consequences for ecological structure and function, thereby leading to changes to marine biological production and significant feedbacks into the Earth’s climate system. Biologically speaking, changes in temperature have direct consequences for many

physiological processes (e.g. oxygen metabolism, adult mortality, reproduction, respiration, reproductive development) and control virtually all life processes from the molecular to the cellular to whole regional ecosystem level and biogeographical provinces. Ecologically speaking, temperature also modulates both directly and indirectly species interactions (e.g. competition, prey-predator interactions and food-web structures). Ultimately, changes in temperatures can lead to impacts on the biodiversity, size structure and functioning of the whole pelagic ecosystem. At the species level, some of the first consequences of climate change are often seen in a species phenology (i.e. timing of annual occurring life-cycle events) and in species geographical distribution responses. This is mainly because temperature continually impacts the life cycle of the species and naturally the population will respond over time, providing it is not biotically restrained or spatially restricted, attaining its optimum position within its bioclimatic envelope. This can occur within a temporal niche as in seasonal succession (observed as a phenological response) or in its overall biogeographical distribution (observed as a geographical movement in a population) (see Boxes  7.1 and  7.2). These biological changes as well as those changes observed in biodiversity and planktonic abundance and productivity are perhaps the key indicators signifying the largescale changes occurring in our world’s oceans as a consequence of climate warming. In this chapter we introduce some ecological concepts (see Boxes 7.1 to 7.3), show some observed evidence and discuss the use of plankton as ecological indicators of global change with a particular focus on the North Atlantic Ocean.

7.1.1  Sensitivity of plankton to global change Plankton are tightly coupled to fluctuations in the marine environment and are highly sensitive indicators of environmental change such as nutrient availability, ocean current changes and climate variability. Temperature is a key driver of marine ecosystems and, in particular, its effects on pelagic populations are manifested very rapidly (Drinkwater et  al.,  2003). This is

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

A Biogeographical Warm-temperate species

Temperate species

Cold-temperate species

Subarctic species

1958–1981

1958–1981

1958–1981

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1982–1999

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2000–2002

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Mean number of species per CPR sample per assemblage 1000 km shift northward

0.8

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Copepods Diatoms Dinoflagellates Meroplankton Otherholozooplankton

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FIGURE 7.1:  Examples of biogeographical and phenological shifts seen over a number of decades recorded by the Continuous Plankton Recorder (CPR) in the north-east Atlantic. A, Biogeographical shifts: biogeographical changes in plankton assemblages spanning five decades. Warm-water plankton (e.g. warm-temperate species) are moving north and cold-water plankton (e.g. subarctic species) are moving out of the North Sea. Particular rapid movement is seen along the European Continental Shelf up to 1000 km over 50 years. Based on Beaugrand et al. (2003). (See Plate 13). B, Phenological shifts: the change in the timing of the seasonal peaks (in months) for the 66 taxa over the 45-year period from 1958 to 2002 plotted against the timing of their seasonal peak in 1958. In the majority of cases most taxa are moving forward in their seasonal cycles but at different rates. Based on Edwards and Richardson (2004).

pla nkton a nd globa l ch a nge  69 mainly because 99% of pelagic and planktonic organisms are ectothermal, making them highly sensitive to fluctuations in temperature (Atkinson and Sibly,  1997). The rapidity of the planktonic response is predominantly due to their short life cycles and in their mainly passive response to advective changes. For example, phytoplankton fix as much CO2 per year as all terrestrial plants but due to being unicellular they represent at any one time only 1% of the Earth’s biomass and have life cycles measured in days to weeks rather than years to decades like their terrestrial counterparts. In the marine environment the effect of short-term climate variability and interannual variability on populations of higher trophic levels such as seabirds and whales can to a degree be somewhat buffered due to their longer life cycles. In the long term, their ability to undergo large geographical migrations may also help them to mitigate some of the effects of global change. In both cases, this is not applicable to planktonic organisms which in a manner of speaking are at the mercy of the prevailing elements. Biologically speaking, changes in temperature have direct consequences on many physiological processes (e.g. oxygen metabolism, adult mortality, reproduction, respiration, reproductive development) and control virtually all life processes from the molecular to the cellular to whole regional ecosystem level and biogeographical provinces. Ecologically speaking, temperature also modulates both directly and indirectly species interactions (e.g. competition, prey-predator interactions and food-web structures). Ultimately, changes in temperatures can lead to impacts on the biodiversity, size structure and functioning of the whole pelagic ecosystem (Beaugrand et  al.,  2002; Edwards and Richardson, 2004; Beaugrand et al., 2010; Mollmann et al., 2015).

7.1.2  Temperature effects on the pelagic habitat Climatic warming of surface waters increases the density contrast between the surface layer and underlying nutrient-rich waters. The availability of one of the principal nutrients (nitrate) that limits phytoplankton growth has been found to be negatively related to temperatures globally (Kamykowski and Zentara, 1986, 2005). While temperature has direct consequences on many biological and ecological traits, it also therefore modifies the actual marine habitat by influencing oceanic circulation and by enhancing the stability of the water column and hence nutrient availability. The amount of nutrients available in surface waters directly dictates phytoplankton growth and is the key determinant of the plankton size, community and food-web structure (see also Box  7.2). Warming of the surface layers increases water-column stability, enhancing stratification and requiring more energy to mix deep, nutrient-rich waters into surface layers. Particularly warm winters will also limit the degree of deep convective mixing and thereby limit nutrient replenishment necessary for the following spring phytoplankton bloom. A global analysis of satellite-derived chlorophyll data has shown a strong inverse relationship between SST  and chlorophyll concentration

(Behrenfeld et al., 2006). Furthermore, other abiotic variables like oxygen concentration (important for organism size and metabolism) (Pörtner & Knust, 2007), nitrate metabolism (Berges et al., 2002) and the viscosity of seawater (important for the maintenance of buoyancy for plankton and their morphological traits) are also directly linked to temperature. Climate warming is expected to contribute to an overall decline in global ocean oxygen levels (deoxygenation) via reduced oxygen solubility from warming and reduced ventilation from stratification and circulation changes (Keeling et  al.,  2010). This, coupled with increased nutrients from human activities, is thought to have led to an increase in hypoxic events and their continuation, which leads to oxygen minimum zones (OMZs), around the world (Rabalais et al., 2010). So unlike terrestrial environments, where precipitation plays a key role, the chemical and upper-ocean temperature regime in open oceans and its consequent biological composition are inexorably entwined.

7.2  Global Change and Plankton Populations 7.2.1  Climate variability and plankton abundance in the North Atlantic Many of the plankton responses observed in the North Atlantic have been associated with rising temperatures; however, any observational plankton change in the marine environment associated with climate warming should be considered against the background of natural variation on a variety of spatial and temporal scales. Approximating the effects of climate change embedded in natural modes of variability, particularly multidecadal oscillations like the Atlantic Multidecadal Oscillation (AMO) (Edwards et al., 2013), is difficult. In the North Atlantic, at the ocean basin scale and over multidecadal periods, changes in plankton species and communities have been associated with Northern Hemisphere temperature (NHT) trends, the AMO, the East Atlantic Pattern (EAP) and variations in the North Atlantic Oscillation (NAO) index (Harris et  al.,  2014). These have included changes in species distributions and abundance, the occurrence of subtropical species in temperate waters, changes in overall plankton biomass and seasonal length, and changes in the ecosystem functioning and productivity of the North Atlantic (Edwards et al., 2001a; Beaugrand et al., 2002; Edwards and Richardson, 2004; Beaugrand et al., 2009). While contemporary observations over a 15-year period of satellite in situ blended ocean chlorophyll are too short to reveal trends over time and their causes, records so far indicate that global ocean net primary production has declined over the last decade, particularly in the oligotrophic gyres of the world’s oceans (Behrenfeld et al., 2006). Over the whole temperate north-east Atlantic there has been an increase in phytoplankton biomass with increasing temperatures but a decrease in phytoplankton

70 ecology Box 7.1  The ecological niche concept A Energy niche An ecosytem’s exploitable energy space associated with potential energy sources.

Dissipated pathways

Energy dissipation, transformation and gradient breakdown

Closed recycling networks

1 Small amount of available energy or highly seasonal environments

Energy (e.g. solar/chemical)

2 Large amount of available energy and stable systems High biodiversity system, e.g. the tropics. Highly specialized energy niches and a move towards closed nutrient and energy cycles.

Low biodiversity system, e.g. the Arctic. Environmental instability and physiological constraints dictate length and complexity of energy pathways and number of energy niches. Highly variable environments lead to succession regression as energy networks collapse.

Space (e.g. range)

Extension of energy niches, reducing energy loss from system

Seasonal extension of network Loss of energetic pathway and Seasonal specialization in semi-closed dormancy (e.g. recycling cysts) as energy flow is seasonally suspended

Survival range

Time (e.g. phenology)

y e bl ia Va r

An ecological niche represented as a three-dimensional hypervolume (ecospace). The hypervolume defines the multidimensional space of resources (e.g. light, nutrients, structure, etc.) available to (and specifically used by) organisms.

Variable z

B Ecological niche

Optimum range

Variable x BOX 7.1. FIGURE 1:  The concepts of the energy and ecological niche. A, The energy niche concept (an ecosystem’s potential exploitable energy space) depicted for high and low diversity systems. B, The ecological niche represented by a simple three-dimensional space where one can locate the species tolerance limits (from its optimum to its maximum survival range) in a continuous geometrical shape called a hypervolume.

The idea that species only exist in a finite range of environmental conditions or that the presence of an organism in space is the sum of the habitat requirements that allow a species to exist and reproduce forms the most basic concept of the ecological niche (Grinnell, 1917; Elton, 1927). With these early concepts, the requirement-based idea of the ecological niche developed, where it becomes a function that links the fitness of individuals to their environment in a multidimensional hypervolume (Hutchinson, 1957). Hutchinson went on to define two further concepts of niche, the realized niche and the fundamental niche; the fundamental niche being a species potential range, whereas the realized niche is the actual set of conditions where the species lives, including both abiotic and biotic factors. A further distinction made here and in the figure is between an ecosystems exploitable energy space/gradient (energy niche) following thermodynamic laws, and an organism’s ecospace within a biological community following natural selection (ecological niche). This is to reflect the often dual nature of the niche concept as part of a facet of the environment and as a facet of species. For example, marine

pla nkton a nd globa l ch a nge  71 communities can have distinct species similarly evolved to exploit plankton (e.g. baleen whales / filter-feeding sharks) occupying the same energy niche (but different ecological niches) within their ecosystem (an example of evolutionary convergence). The niche concept is fundamental to ecological biogeography and to understanding the response of organisms to global change. For example, temperature continually impacts the life cycle of species, and naturally the population will respond over time, providing it is not biotically restrained or spatially restricted, attaining its optimum position within its bioclimatic envelope. This can be within a temporal niche as in seasonal succession (observed as a phenological response, i.e. a movement in time) or in its overall biogeographical distribution (observed as a geographical movement in a population, i.e. a movement in space). Native niche spaces can also be lost due to extinctions and to invasive species that are introduced to new environments, where the invading species outcompetes the indigenous species. The niche concept is also fundamental in predicting the response of organisms to global change. Ecological niche models, also known as bioclimatic envelope models, habitat suitability models and species distribution models are all reliant on the concept of the ‘ecological niche’. These operational applications have become the prominent statistical method in predicting the distribution of organisms for conservation and climate change management. Finally, it is worth mentioning that while the niche is an important conceptual tool in ecology, it is still difficult to clearly define and measure (Godsoe, 2010; Mcinerny and Etienne, 2012).

biomass in warmer regions to the south (Richardson and Schoeman,  2004). Presumably this is a trade-off between increased phytoplankton metabolic rates caused by temperature in cooler regions but a decrease in nutrient supply in warmer regions. For planktonic diatoms there is not really a predominant multidecadal trend for the North Atlantic basin as a whole, but some regions have shown a strong cyclic behaviour over the multidecadal period. The time signal resembles an oscillation of about 50–60 years and a minimum in diatom abundance around 1980 reflecting changes in the AMO signal (Harris et  al.,  2014). Hinder et  al. (2012) attributed a recent decline in North Sea dinoflagellates relative to diatoms to warming and increased summer windiness (i.e. water-column turbulence). Copepod abundances have been more stable in oceanic regions but have shown a decrease particularly in the southern North Sea (Fox et al., 2009). Climate variability has also a geographically heterogeneous impact on plankton in the North Atlantic and not all regional areas are correlated to the same climatic index. For example, trends in the AMO are particularly prevalent in the oceanic regions and in the subpolar gyre of the North Atlantic, and the NAO has a higher impact in the southern North Sea where the atmosphere-ocean interface is most pronounced. This is also apparent with respect to the NHT where the response is also spatially heterogeneous with areas of the north-east Atlantic and shelf areas of the north-west Atlantic warming faster than the North Atlantic average and some areas like the subpolar gyre actually cooling. Similarly, regime shifts or abrupt ecosystem shifts do not always occur in the same region or at the same time. The major regime shift that occurred in plankton in the late 1980s was particularly prevalent in the North Sea and was not seen in oceanic regions of the North Atlantic. However, a similar regime shift occurred in the plankton colour index 10 years later in the Icelandic Basin and in oceanic regions west of the British Isles (Edwards et al., 2013). The different timing and differing regional responses to regime shifts have been associated with the movement of the 10°C thermal boundary as it moves northwards in the North Atlantic (Beaugrand et al., 2008).

Indirectly, the progressive freshening of the Labrador Sea region, attributed to climate warming and the increase in freshwater input to the ocean from melting ice, has resulted in the increasing abundance, blooms and shifts in seasonal cycles of dinoflagellates due to the increased stability of the water column ( Johns et al., 2001). Similarly, increases in coccolithophore blooms in the Barents Sea and harmful algal blooms (HABs) in the North Sea are associated with negative salinity anomalies and warmer temperatures leading to increased stratification (Smyth et al., 2004; Edwards et al., 2006). It seems likely that an important environmental impact caused by climate change is an increase in the presence of haline stratification in regions susceptible to freshwater inputs resulting in an increased potential for bloom formation. There is some evidence that rising temperatures favour an increase in the abundance and range of pathogenic microorganisms with some marine pathogens shifting their distributions poleward (Vezzulli et  al.,  2012; Burge et  al.,  2014). While global outbreak frequencies of jellyfish abundances and aggregations sometimes follow rising SSTs, the evidence to say that jellyfish are increasing in response to climate warming is still inconclusive (Pörtner et al., 2014).

7.2.2  Biogeographical and phenological changes Some of the strongest evidence of large-scale biogeographical changes from climate warming observed in our oceans comes from plankton studies in the North Atlantic. In a study encompassing the whole of the north-east Atlantic Ocean over a 50-year period, Beaugrand et  al. showed the rapid northerly movements of a key zooplankton group (calanoid copepods) (Beaugrand et al., 2002, 2009). During the last 50 years there has been a northerly movement of warmer water plankton by 10° latitude in the north-east Atlantic and a similar retreat of colder water plankton to the north (a mean poleward movement of between 200 km and 250 km per decade) (Figure  7.3A). This geographical movement is much more pronounced than any documented terrestrial study, mainly due to advective processes and in particular the shelf-edge current running north along the northern European continental shelf. The rapid movement of

72 ecology plankton northward is only seen along the continental shelf, where deeper water is warming much more rapidly. Further along the shelf, plankton are upwelled from this deeper water to make an appearance in the surface plankton community. Hence the plankton have moved 10° latitude northward via mainly deep water advective processes not seen in the movement of surface isotherms. In other areas in the north-east Atlantic the plankton shifts were more moderate and varied between 90 km and 200 km per decade, which is faster than documented terrestrial studies, with their meta-analytic average of 6 km per decade (Parmesan and Yohe, 2003). Interestingly, in the north-west Atlantic, pelagic organisms have been moving southward ( Johns et al., 2001). This initially seems to contradict general thinking of homogenous global climate warming throughout the world’s oceans. However, this movement has been linked to the strengthening of the Labrador Current which has spread colder water southward over the last decade carrying pelagic organisms with cold-water affinities as far south as Georges Bank. These large-scale biogeographical shifts observed in the plankton have also seen paralleled latitudinal movements of fish species distribution (Brander,  2010; Montero-Serra et  al.,  2014). Northward range extensions of pelagic fish species have also been reported for the northern Bering Sea region related to regional climate warming (Grebmeier et  al.,  2006). Climate variability and regional climate warming have also been associated with variations in the geographic range of marine diseases and pathogens (Burge et al., 2014). Phenology, or repeated seasonal life-cycle events such as annual migrations or spawning, are highly sensitive indicators of climate warming. In a long-term study of planktonic organisms and their phenological response to climate warming, many plankton taxa were found to be moving forward in their seasonal cycles (Edwards and Richardson, 2004). In some cases a shift in seasonal cycles of over 6 weeks was detected, again a far larger shift than observed for terrestrial-based observations. Summarizing a terrestrial study of phenology using over 172 species of plants, birds, insects and amphibians, Parmesan and Yohe (2003) calculated a mean phenological change of 2.3 days. It is thought that temperate pelagic environments are particularly vulnerable to phenological changes caused by climatic warming because the recruitment success of higher trophic levels is highly dependent on synchronization with pulsed planktonic production (Edwards and Richardson, 2004). Furthermore in the marine environment, and just as important, the response to regional climate warming varied between different functional groups and trophic levels, leading to mismatch in timing between trophic levels (Figure  7.3B). For example, while the spring bloom has remained relatively stable in seasonal timing over five decades (mainly due to light limitation and photoperiod rather than temperature dictating seasonality) many zooplankton organisms as well as fish larvae have moved rapidly forward in their seasonal cycles. Similarly, other pelagic phenology changes in response to climate warming have been

observed in other areas of the North Atlantic and from other oceans around the world (Mackas et al., 2012).

7.2.3  Biodiversity and invasive species At the North Atlantic scale over the last 50 years, overall plankton biodiversity has been shown to have increased. In particular, increases in diversity are seen when a previously low diversity system like Arctic and cold-boreal provinces undergo prolonged warming events. The latitudinal increase in biodiversity for both phytoplankton and zooplankton has been related to the warming North Atlantic over the last 50 years (Beaugrand et al., 2010). Moreover, the increases in copepod diversity have been paralleled by a decrease in the mean size of zooplanktonic copepods. Similarly, phytoplankton (dinoflagellates) show a relationship between temperature and diversity which could be linked to the phytoplankton community having a higher diversity but an overall smaller size fraction and a more complex food-web structure (i.e. microbial-based versus diatom-based production) in warmer, more stratified environments. Reduced body size as an ecological response to climate warming has many diverse explanations including nutrient availability (see Sheridan and Bickford, 2011). Increased warming and a move towards a dominance of smaller organisms may influence the networks in which carbon flows (i.e. increase in surface carbon residence times) and may have negative consequences for the biological carbon pump and North Atlantic fisheries (see also Table 7.1). Climate warming will therefore increase planktonic diversity throughout the cooler regions of the world’s oceans as temperature isotherms shift poleward. In a recent study using a new model for how biodiversity is arranged in the oceans, future projections estimate that there could be a decrease in biodiversity in warm-water regions of the ocean but as much as a 300% increase in polar regions of the world (Beaugrand et al., 2015). Furthermore, the study showed that when global warming rises above the threshold of 2°C, between 50% and 70% of the global ocean may experience a change in marine biodiversity not seen for at least 3 million years. Climate warming will open up new thermally defined habitats for previously denied non-indigenous species (e.g. newly observed warm-water species in the North Sea seen over the last few decades) and invasive species allowing them to establish viable populations in areas that were once environmentally unsuitable. Therefore, species can arrive naturally as a part of shifting thermal boundaries; however, human-caused introductions (e.g. ballast water exchange, aquaculture) have become a significant cause for concern over recent decades. The introduction of non-native species into a new environment can have significant economic and ecological consequences (Vitousek et al., 1997) as well as introducing potentially HAB forming species (Hinder et al., 2011). While it should be noted that not all introductions lead to invasions or the establishment of viable populations, some species can establish themselves (approximately 10% of introductions are successful). Over the last 100 years there have

pla nkton a nd globa l ch a nge  73 Box 7.2  Plankton ecological succession and successional regression Climax and conservation (K selection)

Phytoplankton ecological succession

dinoflagellates

Self-organization and biodiversity

Successional regression

flagellates

8 9 9 7 8787 8 8 77 8 8 Summer community (narrow niches) 97 89 8 8 8 97 88 Renewal and early 89 99898978887 9 8 7 8 8 succession 6 6 9 867 7 99 88779 8 7 997 8 88 10 7 10 777779788 88 7 99998 9 687 6 6 8 7 8 7 6 1 0 6 9 798 9 10 6 10 01 0 1 10 6 6 767881 5 10 0897 509868689 10 11 106 1 0 9 7 6 59 7 991 10 0 10 010 9 8 10 11 1 10 61 6 9 109 09 0 10 9 7 10 1 0 6 10 1 1 11 10 0 7 6 10 10 1 Autumn 7 6 10 60991 50 10 1 07 6756 6691 10 010 10 6 110 Early summer E 70 91 5 7 656 761 10 bloom 6 69 10010 12 5 5 54 8 7 7 6 1 11 community c 8 6 10 5 5 5 10 5 6 11 155 5 10 1 1 11 5575 5 10 1 01 10 1 111 12 10 54 665 65 12 210 11 0 1 211 12 12 91 10 12 0 10 5 5 10 11 1 1 5 1 0 10 6 2 10 1 0 4 11 1 0 10 5 1 2 12 2 4 6 11 1 1 11 1211 5 4 4 2 12 111 5 6 11 111 12 12 4 4 11 10 11 10 11 11 1 2 12 12 11 11 211 12 44104 4 44 4543 55 4 1 1 1 3 11 11 1 12 112 1 31 35 12 1111 4463 4454 45 4 4 4 4 2 3 11 1 11 4 34 5 Sp 1 33 5 4 11 2 12 2 11 1 11 1 11 1 Spring pring blo bloom (broad niches) 12 12 10 3 212 12 11 1 10 1 3 3 5 5 435 4 1 3 1 2 3 3 1 4 4 3 1 4 4 4 2 12 13 1 2 25 12 12 23 22 3 33 33 4 3 33 3 43 2 1 12 212 1 2 1 2 1 4 2 2 2 21 1 11 1 4 4 4 3 11 11 33 31 2 12 12111 312 3 12 12 23 2 3 2 11 12 312 12 1121 2 12 2 1 32 12 1 2 3 122 1 1 1 3 12 2 2 11 11 2 12212 3 12 212 122 2 2 Start 11 12 11 2 11 2 12 2 12 11 3 1 3 1 3 4 1–12 = month (r selection) diatoms

Maturity and stability of ecosystem BOX 7.2. FIGURE 1:  Multidimension scaling (MDS) plot of phytoplankton community (40 taxa) structure per month (n = 552) through time 1958–2003. The pattern repeats annually from an early successional state in early spring to maximum stability in late summer. After this the community undergoes a successional regression in autumn and winter. The renewal process begins the following year as the ecosystem again organizes energy flow. Arbitrary units describe the increase in self-organization, energy flow, biodiversity and ecosystem stability as ecological succession progresses to climax. Phytoplankton community data from the Continuous Plankton Recorder (CPR) survey.

Phytoplankton, at the base of the marine food web, represent the main structural component of pelagic ecosystems. Annually, phytoplankton undergo a consistent pattern of ecological succession that is tightly coupled to the degree of vertical stability of the water column, which in turn is related to temperature, wind mixing and the chemical environment. The amount of nutrients and their ratios available in the surface waters related to water-column stability directly dictates phytoplankton growth and life strategies and is the key determinant of the plankton size, community and food-web structure. The transition from a turbulent to a stable environment is associated with a phytoplankton succession from diatoms (r selection opportunists), through flagellates (competitors) to dinoflagellates (K selection stress-tolerators) (Margalef, 1978). This early transition from well-mixed waters to stratified waters and the resultant shallowing of the thermocline is characterized by a strong burst in diatom production which is the basis of the classic marine food chain, whereby energy flow passes through the zooplankton to fish and to higher trophic organisms. Trophic chain lengths, biodiversity, homeostasis and control over biogeochemical fluxes (nutrient recycling) tend to increase as successional history advances and systems become more selforganized. In terms of nutrient availability, warming of the surface layers increases water-column stability, enhancing stratification and requiring more energy to mix deep, nutrient-rich waters into surface layers. Climatic warming of surface waters will therefore increase the density contrast between the surface layer and the underlying nutrient-rich waters potentially affecting succession cycles and overall plankton (continued )

74 ecology Box 7.2 Continued production. The availability of one of the principal nutrients (nitrate) that limits phytoplankton growth has therefore been found to be negatively related to temperatures globally (Kamykowski and Zentara,  1986, 2005). Similarly, a global analysis of satellite-derived chlorophyll data shows a strong inverse relationship between sea surface temperature (SST) and chlorophyll concentration (Behrenfeld et  al.,  2006). Low nutrient oligotrophic regions such as subtropical gyres where small flagellates and nanoplankton proliferate are associated with low productivity and microbial systems. Temperature has also the potential to modify ecological succession through its effect on influencing life-cycle events and phenology, potentially creating mismatches in timing between various trophic levels (Edwards and Richardson, 2004). Beyond seasonal succession, at the marine ecosystem scale such as productive temperate marine ecosystems for example, climax communities typify mature ecosystems made up of large, long-lived fish communities. Here, intensive fishing activities can in effect reverse the process of succession by successional regression. For example, large commercial species can be overfished and replaced by smaller, more quickly breeding species (Sherman et  al.,  1981); this is sometimes referred to as ‘fishing down the food web’ (Pauly et al., 1998). This process of successional regression or fishing down trophic levels has been implicated in declining large fisheries around the world and a shift in fisheries from large piscivorous fish towards smaller invertebrates and planktivorous fish populations. It has also been implicated in the possible increase in global jellyfish populations as fishing for planktivorous forage fish species removes potential competitors of gelatinous predators (Purcell et al., 2007).

been an estimated 16 exotic phytoplankton taxa introduced into the North Sea (Nehring,  1998). For example, Edwards et  al. (2001b) showed how the Pacific diatom Coscinodiscus wailesii, after being introduced into the North Sea in 1977, managed to geographically spread and establish itself relatively quickly throughout the region, where it remains an important component of the North Sea phytoplankton community. Apart from these thermal boundary limits moving progressively poleward and in some cases expanding, the rapid climate change observed in the Arctic may have even larger consequences for the establishment of invasive species and the biodiversity of the North Atlantic due to the decreasing areal coverage of Arctic ice. Since the late 1990s, large numbers of a Pacific diatom Neodenticula seminae have been found in samples taken in the Labrador Sea in the North Atlantic associated with the decreasing ice coverage. N. seminae is an abundant member of the phytoplankton in the subpolar North Pacific and has a well-defined palaeo-history based on deep-sea cores. According to the palaeo evidence, this was the first record of this species in the North Atlantic for at least 800 000 years. The reappearance of N. seminae in the North Atlantic, and its subsequent spread southwards and eastwards to other areas in the North Atlantic, after such a long gap, could be an indicator of the scale and speed of changes that are taking place in the Arctic and North Atlantic oceans as a consequence of climate warming (Reid et al., 2007). The diatom species may itself be the first evidence of a trans-­ Arctic migration in modern times and be a harbinger of a potential inundation of new organisms into the North Atlantic. The consequences of such a change to the function, climatic feedbacks and biodiversity of Arctic systems are at present unknown.

7.2.4  Other anthropogenic pressures on plankton populations The global increase in atmospheric CO2 concentration is potentially threatening marine biodiversity in two ways. First, CO2

and other greenhouse gases accumulating in the atmosphere are causing changes to the Earth’s climate system. Second, CO2 is altering seawater chemistry, making the oceans more acidic. Although temperature has a cardinal influence on all biological processes from the molecular to the ecosystem level, acidification might impair the process of calcification or exacerbate dissolution of calcifying organisms. Since ocean acidification has the potential to affect the process of calcification, certain planktonic organisms (e.g. coccolithophores, foraminifera, pelagic molluscs, echinoderms) may be particularly vulnerable to future CO2 emissions (Orr et al., 2005). Apart from climate warming, potential chemical changes to the oceans and their effect on the biology of the oceans could reduce the ocean’s ability to absorb additional CO2 from the atmosphere, which in turn could affect the rate and scale of climate warming. Recent experimental studies are providing increasing evidence that ­rising CO2 levels will affect marine biota and interfere with e­ cological and biochemical processes in the oceans (Pörtner et al., 2014). The majority of studies on negative acidification effects on marine organisms have been based on laboratory studies as field studies are rare (Cornwall and Hurd, 2015). A large-scale field study showed that over the last 50 years some planktonic calcareous taxa were actually increasing in terms of abundance, a trend associated with climate shifts in the NHT. Large increases in abundance are particularly seen for echinoderm larvae, foraminiferans and coccolithophores. The research found that the calcifying plankton were primarily responding to climate-induced changes in temperature during the period ­ 1960–2009, perhaps masking the effects of ocean acidification over this time period. Most of the calcareous taxa recorded in this study exhibited an abrupt shift c. 1996 at the time of a substantial increase in global temperature, and taxa exhibited a poleward movement in agreement with expected biogeographical changes under sea temperature warming (Beaugrand et  al., 2012). In the California Current System, which has become increasingly acidified, no changes in the abundance and

pla nkton a nd globa l ch a nge  75 TABLE 7.1:  Five important ecological responses to climate warming in pelagic ecosystems from the initial mechanisms to eventual ecosystem changes. These arise from initial physiological species responses to population/community responses to eventual ecosystem function and structure changes. 1, 2 and 3 are interconnected through niche preferences. 4 and 5 are interrelated as small size structure in communities is often associated with increased biodiversity.

Biological signal

Potential mechanisms

Ecosystem attributes

Anthropocentric concerns

1. Biogeographical shifts (abundance changes)

Geographical (spatial) adjustment to optimum ecological niche/thermal preference (physiological response and phenotypic plasticity)

Food-web reorganization. Changing diversity. Transitional dynamic regime

Community structural changes. In some cases classed as invasive species. Competitive exclusion of native species

2. Phenological shifts (abundance changes)

Seasonal (temporal) adjustment to optimum ecological niche/ thermal preference (physiological response and phenotypic plasticity)

Successional reorganization of community. Transitional dynamic regime

Decreased temporal synchrony of community and trophic levels leading to potential trophic mismatch

3. Whole ecosystem shifts (regime shifts)

Response to shifting thermal boundaries and decoupling of community structure. Differing sensitivity of species responses to increasing temperature

Whole energy network reorganization/regime shift. New system stability or new dynamic regime

Non-linear and abrupt ecosystem shift (difficult to predict and manage). Change in exploitable species and carrying capacity

4. Reduced body size (in ectotherms)

Response to resource/nutrient limitation. Fitness gains from earlier reproduction and adopting smaller size. Earlier maturation. Better energy utilization and transfer

Energetic dominance of smaller organisms. Longer carbon residence times. Specialized niche subdivisions (narrow niches)

Devaluation of fisheries (possible reduction of biomass and move towards smaller species). Increase in carbon residence times in surface waters (possible reduced carbon drawdown)

5. Increase in biodiversity in some ecoregions (assuming no habitat loss/fragmentation in pelagic systems)

Increase in environmental/ water-column stability. Increase in mutation rates. Move towards smaller size structure of community associated with increasing biodiversity (see 4)

Decreased energy loss from system. Increased homeostatic stabilizing processes. Increased nutrient recycling (move towards closed nutrient cycles). Articulated networks

Positive feedback on climate warming with a possible shift from diatoms to small flagellates. Increasing biodiversity not necessary of benefit to large monospecific populations of exploitable fish species (increase in energy partitioning)

composition of calcifying zooplankton have been observed over the last 60 years (Ohman et al., 2009). However, there is field evidence of shell thinning in modern planktonic foraminifera and Southern Ocean pteropods that have been attributed to acidification trends (Moy et al., 2009; Bednarsek et al., 2012). It is not yet known how much of an effect acidification will have on the biology of the oceans in the twenty-first century, whether rapid climate warming will override the acidification problem, and whether or not species can buffer the effects of acidification through plasticity and adaptation. Equally, ocean acidification could become the main driver of ecological change in oceans around the world in the next few decades.

Globally, eutrophication is considered a major threat to the functioning of nearshore ecosystems, as it has been associated with the occurrence and a perceived increase of HABs. HABs in most cases are a completely natural phenomenon having occurred throughout recorded history, and disentangling natural bloom events caused by natural hydroclimatic variability, global climate change and eutrophication is difficult. For example, increasing temperature, nutrient input fluctuations in upwelling areas, eutrophication in coastal areas and enhanced surface stratification all have species specific responses. Gowen et al. (2012) performed a thorough review of the relationship between eutrophication and HABs and could not reach a

76 ecology c­onsensus about the role of anthropogenic nutrient enrichment in stimulating the occurrence of HABs. Prediction of the impact of global climate change is therefore fraught with numerous uncertainties. There is some evidence that biogeographical range extensions caused by regional climate change have increased the presence of certain HABs in some regions (Edwards et  al.,  2006). Regional climate warming and hydrographic variability in the North Sea have also been associated with an increase in certain HABs in some areas of the North Sea particularly in areas affected by the Norwegian Coastal Current (Edwards et al., 2006). The abundance of certain HAB taxa in the north-east Atlantic has been correlated with increasing SST (Hinder et al., 2012). The increased occurrence of hypoxia in many coastal regions around the world has been associated with an increase in human (e.g. agricultural) activities that result in increased fluxes of nutrients to these waters as well as climate warming (Rabalais et al., 2010). The impacts of atmospheric derived anthropogenic nitrogen on the open ocean have only been recently investigated but may also play a significant role in annual new marine biological production (Duce et al., 2008; Kim et al., 2014). Intensive fishing activities can also directly affect plankton populations and in particular they have been associated with the perceived increases in jellyfish populations. For example, large commercial species can be overfished and replaced by smaller, more quickly breeding planktivorous species (Sherman et al., 1981; Pauly et al., 1998). This process has been implicated in the possible increase in global jellyfish populations as fishing for forage fish species removes potential competitors of gelatinous predators (Purcell et al., 2007). Many species of gelatinous zooplankton are able to rise rapidly in abundance and form extensive aggregations when suitable environmental conditions arise (e.g. a thermal niche or a high availability of prey, possibly due to overfishing of planktivores) and as such the group are potentially indicators of ecosystem instability (Lynam et al., 2011). In oceanic waters in the North Atlantic, depth > 200 m, gelatinous zooplankton abundance between 1946 and 2005 was linked significantly and positively to SST and total copepod abundance. Notably, jellyfish in the north-east Atlantic show cyclic changes in population sizes (c. 20-year cycle in oceanic waters and 30-year cycle in shelf seas). Since 1997 they have been increasing in frequency simultaneously in shelf and oceanic waters (Gibbons and Richardson, 2009; Licandro et al., 2010).

7.3  Summary and Monitoring Change The studies highlighted in this chapter collectively indicate that there is substantial observational evidence that many pelagic ecosystems, both physically and biologically, are responding to  changes in regional climate caused predominantly by the warming of SSTs, ocean current changes and changes to nutrient regimes. These biological manifestations of climatic variability have rapidly taken the form of biogeographic, seasonal,

biodiversity, and species abundance changes and whole ecological regime shifts (see Table  7.1). Due to their sensitivity to change, some of the most convincing evidence for the biological response to climate change in our oceans comes from ­phytoplankton and zooplankton communities. Most of this observational evidence derives from long-term surveys and historical data sets in which change can be observed in the context of multidecadal natural variability and shifting baselines. However, such data sets, particularly for open ocean systems, are rare and regionally based (Edwards et al., 2010). For this reason some of the strongest evidence detected for observed changes in open ocean ecosystems comes from the North Atlantic where an extensive spatial and long-term biological survey exists in the form of the CPR survey. The CPR survey has been in operation in the North Sea and North Atlantic since 1931 and has systematically sampled up to 500 planktonic taxa from the major regions of the North Atlantic at a monthly ­resolution (Reid et al., 2003). Important multidecadal evidence from the Pacific is mainly derived from the California Cooperative Oceanic Fisheries Investigations (CalCOFI) survey operating off the coast of California since 1949 (Ohman et al., 2009). Apart from playing a fundamental role in the Earth’s climate system and in marine food webs, plankton are also highly sensitive contemporary and palaeo indicators of environmental change and provide rapid information on the ‘ecological health’ of our oceans. A plankton species, defined by its optimum niche, in effect has the capacity to simultaneously represent an integrated ecological, chemical and physical variable. With the realization that ocean ecosystems are vulnerable to human threats such as overfishing, climate change, eutrophication, habitat destruction, pollution and species introductions there is an increasing imperative to observe our ocean biology in a more integrated fashion in order to provide the long-term baselines needed for management actions and research (see Box 7.3). Further, there is growing evidence that responses to climate change are usually manifest at the species level and that this taxonomic resolution is often needed to interpret many observed changes. For example, some of the most effective ecological indicators used for assessment purposes in European seas are constructed at the species level (Edwards et al., 2010). Also, the mixed success of conventional management of our oceans, in particular fisheries management, and the continuing human threats posed, have necessitated innovative approaches to the management of our oceans. The ecosystem approach to management (EAM) has been proposed as a more effective and holistic approach for managing and maintaining healthy marine ecosystems and the goods and services they produce by addressing some of the consequences of human use. For example, time-series observations and ecological indicators are useful for adaptive management purposes as once ecological or performance indicators cross over a particular threshold, management actions are triggered. Many international research initiatives such as the Global Earth Observation System of Systems (GEOSS) and The Group on Earth Observations

pla nkton a nd globa l ch a nge  77 Box 7.3  Monitoring global changes with the use of plankton indicators

Biological variable

Example

Notes

Bulk status variables (index)

Chorophyll, biomass, total abundance, EOVs, community size

Not particularly sensitive and difficult to interpret change and predict through models. Cheaper to monitor. Monitored using taxonomic methods, satellite observations and various optical technologies

Sentinel species (univariate taxa)

Individual species of high indicative value and/or key structural species. Targeted indicators for policy and management requirements, e.g. climate change, acidification, fisheries, etc.

Individual species chosen to be highly indicative and highly sensitive to ecosystem change and/or key structural species. Can be highly variable but fluctuation more easy to interpret. Models easier to predict if niche requirements known. Monitored using taxonomic and molecular methods. Molecular methods are no yet completely quantitative; they can be designed to be quantitative for specific taxon groups or functional potential

Community structure based on species information (multivariate taxa)

Multivariate community structure measures. Univariate summaries, e.g. diversity indices

Community indicator fluctuations are statistically more robust and may indicate system-wide changes (e.g. regime shifts). More intensive monitoring required to measure. Monitored using taxonomic and genetic methods

Planktonic ‘indicator species’ are highly sensitive to environmental variability and have had a long history in ecological monitoring of the marine environment (Russell, 1936). They are a relatively quick and easy way to monitor different water masses, define marine habitats and observe changes in food-web structure. Over the past decade, these applications have been refined and used as management tools by developing applied ecological indicators to support specific evolving marine management issues and to provide evidence-based information for policy (Edwards et al., 2010). Planktonic indicators are particularly useful in managing the marine environment as they can provide rapid information on a whole multitude of management issues, such as climate change impacts; fisheries and marine wildlife; eutrophication/pollution; ocean acidification; marine biodiversity and invasive species. While the use of the word ‘indicator’ is still somewhat ambiguous (Heink and Kowarik, 2010), generally, two definitions of indicators exist depending on a biological and managerial perspective, and another one interfaces between these two definitions (applied ecological indicator). The first one is an indicator of an environmental property (e.g. an ecological unit or process) that can be statistically measured, for example an indicative value of a variable. For example in biology, an indicator is an organism so entwined with a particular environmental condition that its presence indicates the existence of those conditions. It is also often used to simplify or aggregate a property of a complex ecosystem so that we can measure a component of the state of the system. The second definition is more familiar in a societal sense and is similar to a performance or pressure indicator (e.g. an economic indicator such as the Dow Jones index), in which management actions can be measured (e.g. a significant change in status or stress is expected to trigger a management response). This definition is often used by policymakers and environmental planners. (continued )

78 ecology Box 7.3 Continued The third definition is an ‘applied ecological indicator’. In this sense, planktonic indicators are used to track responses that are particularly important to policy and management, for example changes in marine ecosystem health, climate change and acidification impacts (i.e. it is a biological response indicator but applied to act as an interface between science and policy). Applied ecological indicators may be bulk indicators, individual species or entire communities. Bulk indices are less sensitive to environmental change and will quite often mask the subtleties that individual species will give you; however, it is thought that bulk indices represent the general functional response of plankton to the changing environment. Bulk indicators such as Essential Ocean Variables (EOVs) are defined as having a high impact in responding to scientific and societal needs and importantly have a high feasibility of sustained observation. Generally, the most revealing and interpretable indicators will use high grade ecological information such as sentinel species and communities to monitor change, as these tend to be more sensitive to change (as opposed to bulk indicators such as measures of chlorophyll) (see table).

Biodiversity Observation Network (GEO BON) are being developed to address these issues and monitor these changes using EOVs and Essential Biodiversity Variables (EBVs). Future biological monitoring of open ocean ecosystems, through an integrated and sustained observational approach, will be essential

in understanding the continuing impacts of climate and environmental change on oceanic systems. This in turn may allow us through international collaboration to mitigate and adaptively manage some of the more detrimental impacts (Edwards et al., 2010).

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CH A PTER 8

PLA NKTON A ND FISHER IES K eith Br a nder

8.1 Introduction With very few exceptions, teleost fish species produce planktonic eggs of around 1 mm in diameter that hatch into small larvae living among and feeding on plankton (Fig. 8.1). The larvae undergo very rapid growth and death processes that are a consequence of small body size. In temperate seas most fish species have evolved a seasonal spawning cycle, with regular spawning areas, in order to maximize their net reproduction, by ensuring that first feeding larvae encounter an adequate plankton food supply before the yolk that they hatch with is used up. The proportion of eggs, larvae and juveniles that survive starvation, predation and transport to unsuitable areas during early life determines the number of young fish that enter the fisheries (the annual recruitment). People have been aware of the interannual variability in recruitment for several hundred years, without understanding the causes: Il doit arriver aux poissons, comme aux animaux terrestres, que certaines années soient plus favorables que d'autres à leur multiplication et à leur accroissement, sans qu’on puisse en assigner précisément la cause (Duhamel du Monceau, 1771). [for fish, as for terrestrial animals, some years must be more favourable than others for their population increase and growth, but it is difficult to determine the precise cause of this] the problem is one of unquestionable difficulty, embracing as it does all the conditions which in any way affect the fish from the egg stage to the time when they are caught (Hjort, 1914).

Much of the scientific investigation into fish population dynamics in Europe and North America has been devoted to this ‘recruitment problem’ and in particular to studying the ‘critical period’ (Hjort, 1926) that fish undergo during early life, mainly in the planktonic stages. A second, closely related reason why studies of plankton are vital for fisheries research is the role of planktonic primary and secondary production (phytoplankton and zooplankton) in supporting the fisheries productivity of different sea areas (Richardson, 2002). Fish stocks can fluctuate greatly over time and there are also large, systematic differences

FIGURE 8.1:  The planktonic development of cod from eggs to larvae, which subsequently settle to the seabed when they reach a length of 35–100 mm (Brander, 2005). The development of copepod species (e.g. Calanus finmarchicus), which cod larvae feed on, is also shown. (Illustration by Glynn Gorick)

in fisheries productivity between sea areas; both temporal and spatial variability can be related to differences in plankton production (Nielsen and Richardson, 1996; Houde, 2009). The relationship between the scientific investigation of plankton and of fisheries has therefore always been a very close one and the study of plankton often originated in and was funded by government departments and organizations with an interest in fisheries. Large-scale, long-term plankton sampling programmes have almost without exception had fisheries distribution, productivity and the dynamics of fish recruitment as their principal motivation (Ohman and Venrick, 2003). The aim in setting up the Continuous Plankton Recorder (CPR) survey in the 1930s was to chart the distribution of plankton in the North Sea in a similar way to the charting of meteorological data: ‘When these charts are examined and the results compared with the positions of herring shoals from year to year, we shall know whether or not we can forecast the position of the fish from the distribution of the plankton . . . These varying

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

82 ecology movements of fish must have a definite ascertainable cause, and once ascertained, forecasting cannot be difficult’ (Hardy, 1926). The Hull Fishing Vessel Owners Association and the Fishmongers’ Company provided financial support. The California Cooperative Oceanic Fisheries Investigations (CalCOFI) plankton surveys began in 1949 as a response to the decline in California sardine (Sardinops sagax) landings from c.  718 000 metric tons in 1936–37 to 118 000 metric tons in 1947–48 (Radovich, 1981). In 1947 the California legislature implemented a landing tax on commercial sardine, specifically earmarked to help identify causes of the sardine decline (Scheiber, 1990), and a joint committee of scientists and industry representatives was formed to oversee administration of the funds. Although the original, clear, practical purposes of these two surveys were at best partially achieved, they have made an immense contribution to our understanding of plankton and of physical and biological oceanography in general. They have provided valuable guidance to policies on issues that had not been thought of at the time they were set up, such as impacts of nutrients and climate change, biodiversity and long-term variability (Brander et al., 2003). A major impediment to using such plankton surveys in any operational application (e.g. location of fish shoals) is the speed and cost of processing samples. The types of remote sensing instrumentation used in meteorology and oceanography can now provide some real-time plankton distribution data and the advent of new technologies for imaging flow cytometry, molecular probes and optical counting are beginning to unlock some of the wealth of information in the archived samples. Sampling fish in the sea in order to estimate their population size and variability is difficult, time-consuming and expensive. In some respects the task is more easily accomplished during the egg and larval stages, when a range of species can be c­ ollected in a systematic survey using the same plankton sampling gear. Some of the applications of such surveys in fisheries management and population dynamics are described in Section 8.2, and Section 8.4 returns to consider the ‘recruitment problem’ and the relationship between plankton and fisheries production.

8.2  Estimating Fish Population Size, Variability and Location from Ichthyoplankton The total number of female fish in a stock can be calculated from the annual total of newly released eggs in the plankton by dividing by the number of eggs released by a single female fish into the sea each year. This is the basis of various egg production methods (EPMs) for estimating fish spawning stock biomass (SSB) (Bernal et al., 2012). EPMs have been used routinely for decades to estimate SSB of northern anchovy Engraulis ­mordax in the Pacific (Parker, 1980; Lasker, 1985a), Atlantic

mackerel Scomber scombrus (Lockwood et al., 1981) and many other species. Although simple in principle, EPMs require a great deal of information about biological processes that determine how many eggs are released (specific fecundity, spawning behaviour and frequency, atresia of eggs, skipped spawning) (DickeyCollas et al., 2012). They also require sophisticated survey and sample design and analysis to account for spawning behaviour, scale effects and physical processes that may transport, aggregate or disperse eggs (Pepin and Helbig, 2012). Many of these issues of sampling, survey design and influence of physical processes also affect population estimates of other forms of plankton, but have probably been explored more thoroughly for ichthyoplankton because of the resources devoted to monitoring fish stocks. The cost of carrying out the surveys and analysis for a major fish stock such as northern anchovy or Atlantic mackerel is high. For example, the triennial mackerel and horse mackerel egg survey involves ships from nine nations and covers the area from Cadiz in southern Spain to Iceland using a standardized sampler and a comprehensive and detailed methodology manual (ICES, 2013). The costs can be justified provided that the estimates produced are sufficiently accurate and precise to be useful in stock management. However, in addition to their value in relation to the immediate goal (e.g. estimating SSB), such large-scale surveys have enormous added value. Examples of scientific issues in estimating fish stocks based on CalCOFI research are given in Box 8.1, which is adapted from Ohman and Venrick (2003).

Box 8.1  Scientific issues in estimating fish stocks based on CalCOFI research • Spatial analysis shows that fishing enhances the climatic sensitivity of marine fishes (Hsieh et al., 2006, 2008) • Multidecadal shifts in North Pacific pelagic ecosystems (Brinton and Townsend, 2003; Lavaniegos and Ohman, 2003) • Fishery-independent estimates of historical population variability, reconstructed from collection specimens (e.g. Butler et al., 2003) • Calibration of bio-optical algorithms for remote sensing of oceanic phytoplankton (O’Reilly et  al., 1998; Kahru and Mitchell, 1999) • Continuous underway fish egg sampling (Checkley et  al., 1997) • Landmark treatise describing the morphology, life history, and time series of geographic distribution of 500 species of marine fish eggs and larvae (Moser, 1996) • Interdecadal warming of the California Current, increase in sea level, and concurrent decline in zooplankton biomass (Roemmich, 1992; Roemmich and McGowan, 1995a; Roemmich and McGowan, 1995b) • Basin theory for population expansion of pelagic fishes (MacCall, 1990)

pla nkton a nd fisher ies  83 • El Niño impacts on marine invertebrates and fishes (Chelton et al., 1982; Butler, 1989; Rebstock, 2001) • Speciation mechanisms in oceanic plankton (Fleminger, 1975; Goetze, 2003) • Daily egg production method for estimating SSB of epipelagic fishes (Lasker, 1985b) • Ontogeny of patchiness in pelagic fishes (Hunter and Coyne, 1982) • Feeding ecology of larval anchovy (Hunter, 1981) • Methodology for quantitative sampling of ichthyoplankton (Smith and Richardson, 1977; Smith et al., 1985) • Stable ocean hypothesis (Lasker, 1975) • Cyclical, historical variations in abundance of sardines and anchovies, from fish scales deposited in varved sediments (Soutar and Isaacs, 1974; Baumgartner et al., 1992) • Correspondence between biogeographic distributions of planktonic organisms and large-scale ocean circulation (CalCOFI Atlas Series; Brinton, 1962; Reid et al., 1978) • Objective definition of recurrent assemblages of plankton (Fager and McGowan, 1963) • Relationship between nutrient distributions and zooplankton biomass on the scale of the Pacific basin (Reid, 1962) • Circulation of the California Current System (Reid et al., 1958; Lynn and Simpson, 1987) • Large-scale teleconnections in the atmosphere/ocean system (Sette and Isaacs, 1960) Adapted from Table 1 in Ohman and Venrick (2003). CalCOFI in a Changing Ocean. Oceanography (Special Issue–Scripps Centennial). 16 (3): 76–85.

Hsieh et al. (2006, 2008) use the spatially detailed, long-term, taxonomically rich data set provided by CalCOFI to show that exploited fish species are more sensitive to climate impacts than unexploited species. They examine geographic distributions of larvae of 13 exploited and 16 unexploited fish species surveyed four times a year on a 54 station grid over the period 1951–2002. Their conclusion, that increased variability of exploited populations due to fishery-induced truncation of the age structure reduces the capacity of populations to buffer environmental events, entails that fisheries must be managed not only to sustain SSB but also to prevent loss of population structure. One of the principal aims in setting up the CPR surveys in the 1930s was to provide fishermen with information to locate target species and improve their catches. However, processing the samples resulted in a delay that made the information operationally useless and in any case new technologies, particularly acoustics, proved to be highly effective for real-time location and tracking of herring and other species. The CPR has, however, proved useful in locating major new fishery resources, such as blue whiting (Micromesistius poutassou) west of Ireland and redfish (Sebastes spp.) in the Irminger Sea (Henderson, 1964a, 1964b). At the time, only Spain recorded any landings of blue whiting and these were from areas south of 52°N (average 13 500 tonnes per year from 1958 to 1966).

After 1967 the international fishery for blue whiting increased to over one million tonnes in 1979–1980 (average 465 400 tonnes per year from 1967 to 1998). More recently the spatial and temporal scale of sampling by the CPR has been an essential input to understanding the distribution, abundance and seasonal dynamics of the major copepod species of the North Atlantic, Calanus finmarchicus (Speirs et al., 2006).

8.3  Dependence of Fish on Plankton The dependence of fish production on plankton production is self-evident, since fixation of carbon by photosynthetic phytoplankton forms the base of the marine food chain that leads to fish (Fig. 8.2). Victor Hensen, who was professor of physiology at the University of Kiel from 1871 to 1891 and the originator of the term ‘plankton’, was particularly interested in the relationship between the microscopic life that he found in the sea and the fisheries productivity in his region (Hensen, 1885, 1911). He argued that measurement of available nutrients and biomass of plankton would provide a quantitative basis for estimating fisheries production. Fisheries production is highest in areas of high plankton ­production, including upwelling areas (e.g. eastern boundary currents), fronts and shelf seas with high nutrient supply (Fig. 8.3). Marine mammals, seabirds and fish that are capable of migrating over long distances often congregate to feed in these high productivity areas. In the north-east Pacific, mean annual fisheries yields of resident species for the period 1960– 1998 in 11 areas were very strongly correlated with annual mean chlorophyll a concentration (p < 0.0001, r2 = 0.87) (Ware and Thomson, 2005). Mean annual fisheries yields for nine areas in the north-west Atlantic for the period 1978–1991 were strongly correlated with annual mean chlorophyll a concentration (p 2: Uncertainties compound when simulating climate, fisheries and marine ecosystems. Deep Sea Research Part II: Topical Studies in Oceanography 113, 312–22. Fager, E. W., and McGowan, J. A. (1963). Zooplankton species groups in the North Pacific. Science 140, 453–60. Fleminger, A. (1975). Geographical distribution and morphological divergence in American coastalzone planktonic copepods of the genus Labidocera. In Cronin, L. E. (ed) Estuarine Research, Vol. I, pp. 392–419. Academic Press, New York, NY. Frank, K. T., Petrie, B., Shackell, N. L., and Choi, J. S. (2006). Reconciling differences in trophic control in mid-latitude marine ecosystems. Ecology Letters 9, 1–10. Goetze, E. (2003). Cryptic speciation on the high seas; global phylogenetics of the copepod family Eucalanidae. Proceedings of the Royal Society of London Series B 270, 2321–31. Hardy, A. C. (1926). A new method of plankton research. Appendix 2. In Kemp, S. (ed.), The Discovery Expedition, 118, pp. 630–2. London: Nature. Heath, M. R. (2005a). Changes in the structure and function of the North Sea fish foodweb, 1973–2000, and the impacts of fishing and climate. ICES Journal of Marine Science 62, 847–68. Heath, M. R. (2005b). Regional variability in the trophic requirements of shelf sea fisheries in the Northeast Atlantic, 1973–2000 ICES Journal of Marine Science 62, 1233–44. Henderson, G. T. D. (1964a). Young stages of blue whiting over deep water west of the British Isles. ICES Annales Biologiques 19, 59–61. Henderson, G. T. D. (1964b). Identity of larval redfish populations in the north Atlantic. Nature 201, 419. Hensen, V. (1885). ‘Über quantitative Bestimmungen des “Auftriebs,” In Mitteilungen für den Verein SchleswigHolsteinischer Aerzte 10, no. 7. Hensen, V. (1911). Das Leben im Ozean nach Zählungen seiner Bewohner. Ergebnisse der Plankton-Expedition der Humboldt-Stiftung. Lipsius & Tischer, 406pp. Hjort, J. (1914). Fluctuations in the great fisheries of Northern Europe viewed in the light of biological research. Rapports et Proces-verbaux des Réunions. Conseil International pour l'Éxploration de la Mer 20, 1–228. Hjort, J. (1926). Fluctuations in the year classes of important food fishes. J. Cons. int. Explor. Mer, 1: 1177, 5–38. Hollowed, A. B., Barange, M., Beamish, R. J., et  al. (2013). Projected impacts of climate change on marine fish and fisheries. ICES Journal of Marine Science 70, 1023–37. Holt, J., Icarus Allen, J., Anderson, T. R., et al. (2014). Challenges in integrative approaches to modelling the marine ecosystems

of the North Atlantic: Physics to fish and coasts to ocean. Progress in Oceanography 129(B), 285–313. Houde, E. D. (2009). Emerging from Hjort’s Shadow. Journal of Northwest Atlantic Fisheries Science 41, 53–70. Hsieh, C., Reiss, C. S., Hunter, J. R., et  al. (2006). Fishing elevates variability in the abundance of exploited species. Nature 443: 859–62. Hsieh, C., Reiss, C. S., Hewitt, R. P., and Sugihara, G. (2008). Spatial analysis shows that fishing enhances the climatic sensitivity of marine fishes. Canadian Journal of Fisheries and Aquatic Science 65, 947–61. Hunter, J. R. (1981). Feeding ecology and predation of marine fish larvae. In Marine Fish Larvae: Morphology, ecology and relation to fisheries, pp. 33–77. University of Washington Press, Seattle, WA. Hunter, J. R., and Coyne, K. M. (1982). The onset of schooling in northern anchovy larvae, Engraulis mordax. CalCOFI Reports 23, 246–51. ICES (2013). Report of the Working Group on Mackerel and Horse Mackerel Egg Surveys (WGMEGS) ICES CM 2013/ SSGESST:04. Kahru, M., and Mitchell, B. G. (1999). Empirical chlorophyll algorithm and preliminary SeaWiFS validation for the California Current. International Journal of Remote Sensing 20, 3423–9. Lasker, R. (1975). Field criteria for survival of anchovy larvae: The relation between inshore chlorophyll maximum layers and successful first feeding. Fisheries Bulletin 73, 453–62. Lasker, R. (ed.) (1985a). An egg production method for estimating spawning biomass of pelagic fish: application to the northern anchovy, Engraulis mordax. US Department of Commerce. NOAA Technical Report, NMFS 36, 99 pp. Lasker, R. (ed.) (1985b). An egg production method for estimating spawning biomass of pelagic fish: application to the northern anchovy, Engraulis mordax, eggs and larvae. Fisheries Bulletin 84, 395–407. Lavaniegos, B. E., and Ohman, M. D. (2003). Long term changes in pelagic tunicates of the California Current. Deep-Sea Research II, 50, 2493–518. Lockwood, S. J., Nichols, J. H., and Dawson, W. A. (1981). The estimation of a mackerel (Scomber scomber L.) spawning stock size by plankton survey. Journal of Plankton Research 3, 217–33. Lynn, R. J., and Simpson, J. J. (1987). The California Current System: The seasonal variability of its physical characteristics. Journal of Geophysics Research 92(C12), 12947–66. MacCall, A. D. (1990). Dynamic Geography of Marine Fish  Populations. Washington Sea Grant Program, Seattle, 153 pp. Mitra, A., Castellani, C., Gentleman, W. C., et  al. (2014). Bridging the gap betweenmarine biogeochemical and fisheries sciences; configuring the zooplankton link. Progress in Oceanography 129, 176–99.

88 ecology Moser, H. G. (1996). The early stages of fishes in the California Current region. California Cooperative Oceanic Fisheries Investigations Atlas, 33, 1–1505. Nielsen, E., and Richardson, K. (1996). Can changes in the fisheries yield in the Kattegat (1950–1992) be linked to changes in primary production? ICES Journal of Marine Science 53(6), 988–94. O’Reilly, J. E., Maritorena, S., Mitchell, B. G., et  al. (1998). Ocean color chlorophyll algorithms for SeaWiFS. Journal of Geophysics Research 103, 24937–53. Ohman, M. D., and Venrick, E. L. (2003). CalCOFI in a Changing Ocean. Oceanography (Special Issue–Scripps Centennial) 16(3), 76–85. Parker, K. (1980). A direct method for estimating northern anchovy, Engraulis mordax, spawning biomass. Fisheries Bulletin 78, 541–4. Pepin, P., and Helbig, J. A. (2012). Sampling variability of ichthyoplankton surveys—exploring the roles of scale and resolution on uncertainty. Fisheries Research 117–118, 137–45. Pörtner, H. -O. et al. (2014). Ocean systems. In Field, C. B. et al. (eds) Climate Change 2014: Impacts, Adaptation, and Vulnerability. Part A: Global and Sectoral Aspects. Contribution of Working Group II to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change, pp. 411–84. Cambridge University Press, Cambridge, United Kingdom and New York, NY, USA. Radovich, J. (1981). The collapse of the Californian sardine fishery. What have we learned? In: Glantz, M. H. and Thompson, J. D. (eds), Resource Management and environmental uncertainty: lessons from coastal upwelling fisheries, pp. 107–36. NY: Wiley. Rebstock, G. A. (2001). Long-term stability of species composition in calanoid copepods off southern California. Marine Ecology Progress Series 215, 213–24. Reid, J. L., Jr. (1962). On the circulation, phosphate-phosphorus content and zooplankton volumes in the upper part of the Pacific Ocean. Limnology and Oceanography 7, 287–306. Reid, J. L., Brinton, E., Fleminger, A., Venrick, E. L., and McGowan, J. A. (1978). Ocean circulation and marine life. In Charnock, H. and Deacon, G. (eds), Advances in oceanography, pp. 65–130. Plenum, NY. Reid, J. L., Jr., Roden, G. I., and Wyllie, J. G. (1958). Studies of the California Current System. CalCOFI Reports 6, 28–56. Richardson, K. (2002). Linking plankton and fish production throughout the history of ICES. ICES Marine Science Symposia 215, 156–63.

Roemmich, D. (1992). Ocean warming and sea level rise along the southwest U.S. coast. Science 257, 373–75. Roemmich, D., and McGowan, J. (1995a). Climatic warming and the decline of zooplankton in the California Current. Science 267, 1324–6. Roemmich, D., and McGowan, J. (1995b). Sampling zoo­ plankton: correction. Science 268, 352–3. Røjbek, M. C., Jacobsen, C., Tomkiewicz, J., and Støttrup, J. G. (2012). Linking lipid dynamics with the reproductive cycle in Baltic cod Gadus morhua. Marine Ecology Progress Series 471, 215–34. Scheiber, H. N. (1990). Californian marine research and the founding of modern fisheries oceanography: CalCOFI’s early years, 1947–1964. CalCOFI Reports 31, 63–83. Schlüter, M. H., Kraberg, A., and Wiltshire, K. H. (2012). Longterm changes in the seasonality of selected diatoms related to grazers and environmental conditions. Journal of Sea Research 67, 91–7. Sette, O. E., and Isaacs, J. D. (eds) (1960). Symposium on “The changing Pacific ocean in 1957 and 1958”. CalCOFI Reports 7, 13–217. Smith, P. E., and Richardson, S. L. (1977). Standard techniques for pelagic fish egg and larva surveys. Fisheries Technolgies 175, 1–100. Smith, P. E., Flerx, W., and Hewitt, R. P. (1985). The CalCOFI vertical egg tow (CalVET) net. NOAA Technology Report NMFS 36, 27–32. Soutar, A., and Isaacs, J. D. (1974). Abundance of pelagic fish during the 19th and 20th centuries as recorded in anaerobic sediments off the Californias. Fisheries Bulletin 72, 257–73. Speirs, D. C., Gurney, W. S. C., Heath, M. R., Horbelt, W., Wood, S. N., and de Cuevas, B. A. (2006). Ocean-scale modelling of the distribution, abundance, and seasonal dynamics of the copepod Calanus finmarchicus. Marine Ecology Progress Series 313, 173–92. Taucher, J., and Oschlies, A. (2011). Can we predict the direction of marine primary production change under global warming? Geophysical Research Letters, 38(2), doi:10.1029/2010GL045934. Thompson, A. B., and Harrop, R. T. C. (1991). Feeding dynamics of fish larvae on Copepoda in the Western Irish Sea, with particular reference to cod Gadus morhua. Marine Ecology Progress Series, 68, 213–23. Ware, D. M., and Thomson, R. E. (2005). Bottom-Up Ecosystem Trophic Dynamics Determine Fish Production on the Northeast Pacific. Science 308, 1280–4.

SEC T ION I I

Methodology

CH A PTER 9

SA MPLING, PR ESERVATION A ND COUNTING OF SA MPLES I: PH YTOPLA NKTON A lex a ndr a K r a berg, K atja Metfies, a nd Row ena Ster n

9.1 Introduction

9.2  Phytoplankton Sampling Methods

Phytoplankton comprise a very diverse array of organisms, including diatoms, dinoflagellates, smaller flagellates and the  picoplankton (plankton < 3 µm in diameter). Bacterial phytoplankton consist primarily of photosynthetic cyanobacteria. Eukaryotic phytoplankton belong to protists, a loose group of eukaryotic microbes that exclude fungi, animals and plants. Although all of these were originally subsumed under the term ‘phytoplankton’, some groups such as the dinoflagellates are now known to contain many heterotrophic species (commonly termed microzooplankton). We will include the microzooplankton in our description of sampling methods. A multitude of different devices are available for the collection of phytoplankton samples and their enumeration (Booth, 1993; Birk et al., 2012). The exact method to be used depends to some extent on the target organisms of interest but also on whether the study is to be qualitative or quantitative. Although many methods appear to be quite similar, it is vital to carefully think about the sampling protocol prior to starting a sampling campaign. This is particularly true if data are to be collected over a long period of time, for instance as part of a long-term monitoring programme. Different methods have varying levels of precision and accuracy. This means that they affect the ways in which individual data sets can be interpreted, and methods therefore have to be kept consistent within time series to avoid creating artefacts (e.g. community shifts where there are none). Here we describe the most widely used sampling and analysis techniques, both traditional and recently emerging, and discuss them in light of their advantages and disadvantages.

9.2.1  Qualitative and semi-quantitative methods Qualitative methods are designed to assess the species richness but not their abundance in a given area, for example to establish a checklist. For such studies it is important that even rare species are captured, and therefore nets with which the plankton can be concentrated are used. A basic plankton net is a very simple construction with a conical net (usually made of nylon) for filtering the plankton (Horner, 2002; see Fig. 9.1). The narrow end of the net is attached to a collecting vessel. The broader opening of the filtering net is held open by a metal hoop. A bridle system is attached to the hoop so that the net can be pulled behind a ship to provide a horizontally-integrated sample, or lowered through the water column to give a depth-integrated sample. An example of this type of net is the Apstein net. However, many variations exist on the basic set-up, which needs to be taken into account when comparing data from different studies. The mesh sizes used vary, but commonly used are 5 µm, 20 µm and 80 µm nets for phytoplankton and microzooplankton, 125–150 µm for mesozooplankton and 500 µm for larger plankton including fish larvae. This kind of net sample will result in an integrated sample of the community within the entire sampled volume. Flow meters can be attached to either the broad net opening to measure the volume of water passing it, or behind the narrow end of the net to measure how much water was actually filtered (e.g. Gulf III sampler). To sample quantitatively, the net has to be pulled very slowly as otherwise turbulence near the net opening will cause the water to be diverted from the net without passing through

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

92 methodology A

B

C

Sleeve Hoop

D

E

Net

Collecting vessel

F

G

FIGURE 9.1:  A, Rosette sampler; B, single Niskin sampler; C, schematic of a basic plankton net; D, flow meter; E, Sedgewick Rafter chamber; F, schematic of the components of an Utermöhl chamber; G, Utermöhl chamber with all components in place.

the filter mesh. To decrease the filtered volume and to reduce turbulence the net opening can be narrowed, for example by a canvas or plastic sleeve. A more complex version of a quantitative net sampling device is the Continuous Plankton Recorder (CPR), which has been operated by the Sir Alister Hardy Foundation for Ocean Science in the Atlantic since 1931. This sampling device has specifically been designed for plankton trawls over long distances (Warner and Hays, 1994). The CPR is towed behind ships of opportunity, usually merchant ships. The CPR works with a filter band of silk which is wound through the CPR on rollers. The water entering the CPR at the front-end aperture is filtered through the silk on which the plankton gets trapped. As the CPR moves through the water, the band of silk also moves within the internal mechanism of the CPR. The length of silk used during the tow is proportional to the length of the tow (the silk moves by about 10 cm per 10 nautical miles (18.52 km)). In this way the organisms can be quantified.

Advantages/Disadvantages Even if pulled slowly, the netting will often become clogged with time, reducing the filtered volume. Therefore, for accurate counts of the phytoplankton, assemblage net samples are not advisable. As plankton nets are pulled vertically or horizontally through the water, they also only provide an integrated sample of that body of water. On the other hand, they are likely to capture even quite rare organisms as long as these can withstand the pressure occurring when the water passes through the net. More complex set-ups with several nets that can be used in sequence are also available. With these multinet samplers, more than one sample can be taken at different depths. However, mesh sizes used are usually larger and therefore geared towards zooplankton.

9.2.2  Quantitative methods Subsurface water samplers A gentler way of taking a sample is the use of water samplers. These are cylindrical in shape and are closed at both ends with a rubber stopper. The two stoppers are held open with a wire before deployment. When the sampler has been lowered to a chosen depth, a dropping weight (messenger) is sent down to release the wire so that the two ends of the sampler close. The resulting water sample is of a known volume and from a discrete water depth. Again a variety of types and makes are available, but two that are in very common use are the Niskin and Nansen samplers (Fig. 9.1B). Many different variations of this very basic set-up are available depending on what fraction of the plankton/phytoplankton is to be sampled. If discrete samples through the whole water column are required, a so-called rosette sampler can be used (Fig. 9.1A). This combines a number of Niskin samplers in a circular set-up, with the samplers set up around the periphery of a metal frame, with the space in the centre available for the attachment of further devices for measuring parameters such as temperature, salinity or pigments. The distinct advantage of this set-up is that the sampling equipment only has to descend through the water column once, preventing excessive turbulence and mixing of the water column prior to sampling.

Advantages/Disadvantages A clear advantage of using samplers such as Niskin or Nansen bottles is that samples are obtained from a discrete depth and with a defined volume. The sampled water will also contain the whole plankton community, i.e. it is not restricted to a specific size range. In addition, damage to delicate organisms can

methodology: ph y topla nkton  93 ­ sually be avoided. On the down side, as the sample is not conu centrated and only a relatively small volume of water is sampled, there is a greater risk of missing rare species.

9.3  Sample Analysis 9.3.1  Sample fixation While for taxonomic investigations it is often advisable to use live material, studies involving the generation of large numbers of samples requiring long-term storage will need to be chemically preserved. Many fixatives are in use, the most common for phytoplankton being Lugol iodine solution. Raw water samples to be used in routine monitoring will usually be fixed in a weak Lugol solution of 1% or less. Microzooplankton on the other hand require higher concentrations of the fixative (5%+). Other fixatives such as gluteraldehyde are often used to preserve small flagellates in the picoplankton size range. However, it is toxic and should therefore be used under a fume hood while wearing protective gloves. The use of fixatives is not entirely unproblematic, as they can shrink cells and/or distort their shape (Menden-Deuer et al., 2001; Zarauz and Irigoien, 2008). With all methods for quantitative use it is extremely important to set appropriate reference conditions if samples are to be taken repeatedly, for example during the course of routine monitoring. For samples in a given programme to be comparable they need to be taken with standard equipment and protocols. They also need to be fixed in a consistent manner. Therefore large initiatives like the Helsinki Commission (HELCOM) and the International Council for the Exploration of the Sea (ICES) have usually developed standard protocols and manuals to be used uniformly by their members (Sournia, 1978; HELCOM, 2000; ICES, 2005).

9.3.2  Utermöhl method Phytoplankton samples for biodiversity assessments using quantitative counts are usually not concentrated during sampling, i.e. they present the species composition and density of the raw water sample. To make them amenable to counting they need to be concentrated. One of the most widely used methods for quantitative counts is the Utermöhl method using so-called Utermöhl chambers (Lund et al., 1958). These consist of a base plate with a well into which samples are poured to concentrate them, a chamber or tower placed on top of the well, and a cover slip to be placed on top of the tower (Fig. 9.1G,H). Utermöhl towers are available in different predefined volumes so that samples of different initial density can be concentrated to a roughly similar final density in the well. The sample is poured into the tower and left to settle in the well for several hours or days. A rule of thumb is that the samples should be left to settle for 2 hours per cm of height of the Utermöhl tower. A note of caution: it is very important to report the length of time the chambers have settled and the

v­ olume of Utermöhl chamber that was used. Differently sized and shaped taxa will have very different sinking velocities. If the settling process is too short and contains many small cells for instance, the entire community might not be present in the well. For intercomparison of results, metadata on the Utermöhl procedure are therefore vital. Once the sample has settled sufficiently, the liquid in the Utermöhl tower is decanted, leaving only the sample in the well, which can now be counted using an inverted microscope. Here techniques also vary depending on the size and abundance of the organisms to be counted. For large cells occurring in low densities, the whole field should be counted at a low magnification. Smaller organisms should be counted at higher magnifications. If these cells occur at very high densities, it might take a long time to count all cells in the well. In these cases a small number of tracks might be counted (either from top to bottom or left to right edge of the well). The volume of this track can be calculated as a proportion of the volume of the whole well, giving a factor with which the count has to be multiplied to give the count for the entire well. The Utermöhl counting method is designed to assess the entire the phytoplankton community. For cultured material, but conceivably also for very dense raw water samples (e.g. during a bloom), other counting chambers are also available. These have in common that only a very small volume of water is being analysed and that there is no prior sample concentration step. Well-known chamber types are the haemocytometer and the Sedgewick Rafter chamber. The latter is calibrated to hold exactly 1 ml of sample, i.e. this is really only suitable for the densest phytoplankton or cultured material. The finely calibrated grid of the chamber also facilitates counts of even smaller volumes by counting just a subset of grid cells, or by random fields (an indirect method that is used to assess abundance in a sample where cell densities are very high and counting the entire sample in the chamber would be impractical).

9.4  Automated/Semi-Automated Systems Increasingly, counts are not just carried out manually. More and more automatic and semi-automatic devices are used for phytoplankton enumeration at different levels of resolution (see Fig. 9.2). To some extent these devices combine the processes of sample collection and analysis. With equipment such as flow cytometers, size classes and the trophic state of organisms (autotrophic/mixotrophic/heterotrophic) in a sample can be discriminated. A device gaining popularity for species to family specific counts is the FlowCam. This is similar to a flow cytometer in that plankton is passed through a capillary tube via a detector. However, in addition, images are also made of the detected organisms. These can be used to construct a ‘library’ of images that is used to identify cells using image recognition software. However, the size range of ‘countable’ taxa is limited.

94 methodology A

B

C

D

E

FIGURE 9.2:  Examples of devices for semi-automated sampling: A, FlowCam device; B, BBE fluorescence desktop device; C, fluorescence probe for onboard operation to be run via autoplug or data cable (arrow); D, composite FlowCam images of the species Eucampia zodiacus; E, graphic output of the FluoroProbe showing a depth profile of major algae classes as well as total chlorophyll.

Another device that is now increasingly used and is similar to Flowcam is the Imaging FlowCytobot. This has proven itself to be very useful for long-term insitu deployments Tools such as the FlowCam (http://www.fluidimaging.com) or ZooScan (http://www.obs-vlfr.fr/LOV/ZooPart/ZooScan/) will not be able to replace manual counts as only a subset of the total plankton community under investigation can be quantified (i.e. the subset that the machine is trained to identify). Moreover, there are as yet no internationally accepted standards for assuring the quality of the data derived from these methods. However, they are a very good support tool particularly in combination with other methods as they can document the sampling process with image material (Garmndia et al., 2013; Wert et al., 2013). For a broader overview of major phytoplankton groups, ­fluorescence-based probes are also available, for instance the FluoroProbe, which allows the detection of total diatoms, cyanobacteria, chlorophytes and cryptophytes and in addition yellow matter via selective excitation of accessory pigments characteristic for these different groups. This device can be used for high frequency in situ measurements, for instance to obtain depth profiles of pigment distribution prior to the deployment of sampling devices to sample discrete depths (Catherine et al., 2012). This ability of the FluoroProbe to collect high frequency measurements is an advantage, as is its ease of deployment. However, it does not provide species specific information, and there are as yet very few studies that check FluoroProbe measurements against field data. This should be  done in any new environmental setting that this device is deployed in.

Note on data archiving No sampling equipment is completely unbiased, for instance with respect to undersampling particular components of the phytoplankton community (Majaneva et al., 2009). The sample protocols are as important as the data themselves and should be archived alongside the actual data, which themselves require consistent long-term archival in an accepted open access repository.

9.5  Molecular Methodologies 9.5.1  Molecular fingerprinting methods: measuring plankton community changes During the past two decades a variety of molecular fingerprinting methods have been established to profile the diversity of protistan communities. Molecular fingerprinting methods targeting the ribosomal operon-like terminal restriction fragment length polymorphism (T-RFLP), automated ribosomal ­intergenic spacer analysis (ARISA), single-strand conformation polymorphism (SSCP) or denaturing gradient gel electrophoresis (DGGE) are well-established cost-effective molecular genetic tools to elucidate variability in the composition of bacterially-­ derived microbial and protistan communities (Muyzer, 1999; Moeseneder et  al., 2001; Savin et  al., 2004). Molecular fingerprinting is routinely applied to assess the spatio-temporal dynamics of microbial, fungal or protistan

methodology: ph y topla nkton  95 c­ ommunities in a wide variety of different environments. The methods offer an alternative to microscopic counts or molecular assessments of species communities based on clone library analyses or high resolution sequencing of barcoding genes. The general line of action for molecular fingerprinting is based on the extraction of nucleic acids (usually genomic DNA), followed by the amplification of a target gene with universal primers, for example ribosomal genes, and finally the analysis of the polymerase chain reaction (PCR) products depending on the fingerprinting technique. Among the molecular fingerprinting techniques, T-RFLP and ARISA (Fig. 9.3) do not provide information on the community composition in terms of individual protist species because they do not involve identification of organisms in a sample by sequencing target genes. Instead, these two methods quickly provide an overall picture of differences in the composition of different communities. T-RFLP and ARISA are based on analysing the composition of differently sized DNA fragments generated from ribosomal operons present in a protistan community. It is assumed that each differently sized DNA fragment represents a different type of protist. T-RFLP fingerprinting is based on the sequence heterogeneity of a certain target gene

among different protist species, for example within the 18S rDNA. The sequence heterogeneity within the target gene is translated into a set of differently sized DNA fragments by amplification of the target gene with fluorescently labelled ­universal primers and subsequent digestion with restriction enzymes. Universal primer sets are suited to amplify the target gene from different protist types in a sample. Restriction enzymes recognize restriction sites that contain specific DNA sequences. Typically restriction sites are nucleotide motifs of 4–6 bases that cut DNA strands at this site. In regard to this, T-RFLP is based on the assumption that the target genes of different protist types have different sequences that determine the restriction site in the target gene. Furthermore, the restriction reaction results in a set of differently sized fluorescently labelled restriction fragments of the target gene that reflects the community composition in a protist sample. ARISA is independent of the sequence or the restriction profile of the target gene. It is based on directly analysing the size variability of the internal transcribed spacer (ITS) regions (ITS1 and ITS2) of the ribosomal operon. The ITS1 and ITS2 are located between the genes coding for the 18S rRNA (also called small subunit (SSU) rRNA) and the 28S rRNA (also called the large subunit (LSU) rRNA).

T-RFLP

ARISA Restriction

GGCC

GGCC GGCC

Species A

Species A PCR-Fragment 18S rDNA

Species B

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PCR-Fragments ITS

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Community profile deduced by size separation on a capillary sequencer 60

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FIGURE 9.3:  Detection principle of terminal restriction fragment length polymorphism (T-RFLP) and automated ribosomal intergenic spacer

analysis (ARISA) subsequent to the amplification of the target gene via polymerase chain reaction (PCR).

96 methodology They are separated by the gene coding for the 5.8S rRNA. The ITS1 and ITS2 do not code for proteins. Therefore there is a high degree of variability in the sequence and the length of these regions. By analogy with T-RFLP, it is assumed that different types of protists have differently sized ITS regions. In an ARISA analysis, a protistan community is reflected by its composition of differently sized fragments of the ITS region that have been amplified with universal fluorescently labelled primers. The composition of the fluorescently labelled and differently sized DNA fragments is determined for both T-RFLP and ARISA by electrophoretic size separation and laser detection of the fluorescent signal. DNA standards of known size are included in the analyses as size standards. The obtained electropherograms allow quantification of the number and size of different protist types in a sample. Thus, the composite of differently sized DNA fragments in a sample is for both methods a characteristic molecular fingerprint of a protistan community that is very well suited for a comparison of community composition. SSCP or DGGE are molecular fingerprinting methods that provide an overview of differences in the composition of microbial and protistan communities. However, unlike T-RFLP and ARISA it is possible to obtain information on the taxa contributing to the communities. SSCP and DGGE are gel-based fingerprinting methods that allow distinct bands to be cut out for subsequent sequencing of the target gene. Both methods are based on the analyses of sequence heterogeneity of a target gene among different protist species. DGGE involves the separation of different PCR amplified variants of the target gene using an electrophoresis gel that contains a linear gradient of a denaturing agent (a mixture of urea and formamide). It is assumed that variants of the target gene with different sequences differ in their melting behaviour. On the linear denaturing gradient of the DGGE electrophoresis gel, the DNA fragments stop migrating at different positions in the gel. As a result of this, different variants of the target gene are distributed on the gel according to their sequence. SSCP is a molecular fingerprinting method in which a target gene is amplified with universal primers. A single-strand piece of DNA is generated from the PCR product by digestion of the opposite strand with an exonuclease that recognizes a phosphorylated 3´ end at the end of one of the double strands. The resulting single strands are separated via gel electrophoresis on a non-degrading acrylamide gel. For SSCP it is assumed that variants of the target gene differing in their sequences have different single-strand conformations or folding, which impacts the migration through the gel.

Advantages/Disadvantages Molecular fingerprinting methods, in particular T-RFLP and ARISA, are cost-effective and quick. They provide the possibility of analysing and processing large numbers of environmental samples in parallel with relatively low effort. Thus they are very useful techniques for monitoring and evaluating variability

in protistan communities with high spatio-temporal resolution. Compared to other molecular fingerprinting techniques, T-RFLP and ARISA are advantageous because they have a high degree of reproducibility. Comparative studies dedicated to the applicability of T-RFLP and ARISA for the assessment of prokaryotic communities in the soil and the water column revealed that both methodologies provided similar results in respect to differences in the microbial community structure. However, their major drawback is that both methods are largely qualitative. They are not useful methods to assess the true richness or composition of protistan communities. Differences between the fingerprinting techniques were observed with respect to bacterial richness and diversity estimates. T-RFLP and ARISA provide only very limited information on the species that contribute to the communities. Currently there is only very little information on the taxonomic significance of T-RFLP or ARISA fragments of protists. In this respect, SSCP and DGGE are advantageous over T-RFLP and ARISA. They provide the possibility to sequence distinct bands identified on the gel. Furthermore, the process of denaturation on a DGGE gel is a stepwise process, as discrete portions of the DNA fragment suddenly become single stranded within a narrow range of denaturing conditions. Thus it is relatively easy to pick distinct bands for further sequencing. However, DGGE has been criticized for its limited sensitivity. The potential of DGGE for the assessment of the rare microbial and protistan biosphere (< 1% of the microbial biosphere) is limited. Furthermore, it is difficult to establish and reproduce the denaturing gradient on a DGGE gel. Thus, the reproducibility of DGGE is limited.

9.5.2  Identifying protists de novo from environmental DNA using clone library sequencing and next-generation sequencing Phytoplankton, microzooplankton and heterotrophic microbial eukaryotes come under the collective term of protists. It is estimated that there are between 26 000 and 74 400 catalogued protist species (Pawlowski et al., 2012) compared to predicted estimates of between 0.7 and 1 million total protist species in existence (Appeltans et al., 2012). This disparity is mainly due to the difficulty in culturing protists, the main way to characterize these organisms. It is thought that most protistan diversity cannot be cultivated in the lab. For example, in 1997 it was thought that 12% of bacterial diversity had not been cultivated, but by 2007 these estimates had risen to 70% as sequencing revealed more species (Achtman and Wagner, 2008). To circumvent these issues, genetic methods have been developed to differentiate protistan (also called eukaryotic microbial) communities within mixed environmental samples (see Fig. 9.4 for summary). DNA sequencing is the main genetic method to identify taxa conclusively. A DNA sequence can vary in length but has sets of characters that, ideally, are unique to that species for exclusive identification. There are two approaches commonly used. The first is a selective approach to narrow down or even pick out individual cells of interest and

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Flow cytometry

Pipette

Indivudual/population separation

Mixed environmental sample

DNA extraction individual cells or mixed environmental samples PCR amplification and sequencing of barcode marker 1. 2. 3. 4.

ATGGACCTTTAGCAATTAACGAGCTAAGTAGCCGA... ATGGACCTTTAGCAATTTACGAGGTAAGTAGGCGA... ATGGACCTTTAGCAATTTACGAGCTAAGTAGGCGA... ATGGACCTTTAGCAATTAACGAGCTAAGTAGCCGA...

Comparison with barcode database: LD, Genbank Species A Species B

Specimen Vouchers

Species B

Specimen Vouchers

Species C

Barcode assessment

Genus

Genetic distance

Poor barcode

Apply genetic distance boundaries Visualization of diversity: phylogenetic tree 1 4

Species/genus separation

Species

Specimens

Specimen Vouchers

Specimens

Specimen Vouchers

Good barcode

No separation

Species

Genus

Genetic distance

Species A 3 2

Species B

Species C

Picked cells phylogeny

Species A Species B Species C

Environmental phylogeny

FIGURE 9.4:  DNA barcoding process: a mixed population of cells can either be separated by manual isolation with a pipette. Flow cytometry is less selective, separating groups of cells based on size and/or pigment, but it can be useful if the cells have unique pigment characteristics. Alternatively, DNA barcode sequences of a mixed sample can be analysed with taxa-specific primers (e.g. diatoms) or more general primers (e.g. eukaryotes), depending on what is needed. Each barcode sequence can be compared to a database of the same or similar known voucher species to identify the genetic distance range for each species to determine a good versus poor barcode marker. Phylogenetic trees are used to visualize the groups and applying genetic distances to each group can help define species groups – and whether there are subpopulations within a species, e.g. Species B in the picked cell phylogeny. Environmental phylogenies that are performed on a larger group of taxa, e.g. eukaryotes, may not be as accurate as species-specific tests, so individual species will be identified with less accuracy (e.g. Species B and C are merged in the environmental phylogeny) but more taxa will be identified in the sample overall.

98 methodology obtain the DNA sequences of one, several DNA markers or even, in rare cases, an entire genome from the cell, providing both genetic and taxonomic information. Cells are isolated individually by pipette or they can be separated by flow cytometry. After isolation, many cells are identified by a process called DNA barcoding (see Fig. 9.4): a short, DNA signature used as a taxonomic proxy for species identification. A DNA barcode is a short stretch of DNA (around 500–700 bp; bp = base pairs) obtained from a standardized region of a DNA marker, normally the mitochondrial gene, cytochrome oxidase I (COI). The use of a standardized region allows the user to compare the DNA barcode of an unknown taxa with a database of DNA barcodes from similar taxa to determine the closest similar taxon. Normally members of a species are very similar to each other, with only 1–2% difference in their DNA barcode sequences. In contrast, members of different species show a 10-fold difference between each. In a taxon group in which genetic boundaries within and between species have been established, it is easy to determine the species group to which the unknown belongs. Such approaches are recommended when identifying just one or a few unknown individual cells, or characterizing a potential new member of a genus, as these methods are labour intensive. DNA barcoding relies on the fact that the taxonomy has clearly demarked species within the taxon group and that all members have evolved over time at equal rates. Many brown algae, red algae, diatoms, dinoflagellates, some ciliates and foraminifera have established DNA barcodes and genetic boundaries with which to identify a species (see Barcode of Life Database, BOLD). In reality, the species concept for many protists is vague or unknown, has incomplete taxonomic information and/or has evolved at different rates. For example, within a commonly found phytoplankton group, Dinophyceae, which are mostly free-living, one genus, called Symbiodinium, has become symbiotic and has evolved at a much faster rate than other members to the extent that the genetic diversity of most Dinophyceae genera is less than a fifth of that of Symbiodinium’s. To further complicate matters, many protists have multiple genetic histories due to endosymbiosis; for example, cryptophytes are heterotrophic organisms that have acquired a red algal chloroplast and nuclear genome, now reduced to a remnant. In total, cryptophytes have four genomes including the mitochondria that have and are evolving at different rates. As a result, researchers cannot use one marker to compare all protists and have used alternative taxonomic DNA markers as barcodes to distinguish their protist group of interest. Recently it has been agreed that protist DNA barcoding is performed using a primary DNA barcode, the variable V4 subregion (V4) of the small ribosomal gene (SSU), or V4-SSU. For species resolution, an additional secondary taxon specific marker, commonly COI, the ribosomal ITS or D1-D2 region of the large ribosomal gene (LSU) is recommended. In many cases, however, the overall diversity of a community is of interest, in which case an environmental barcoding (also called environmental sequencing) or metagenetic analysis can

be applied to the sample. Environmental barcoding describes the process in which the entire environmental community is characterized using one or a few targeted taxonomic DNA markers, without any prior morphological identification required. A similar term often used is metagenomics, but this method also extends to the characterization of many different parts of the genomes, in a non-targeted manner, of every organism in that sample (the metagenome). The result of an environmental barcoding project is a set of sequences from one or a few parts of the genome that represent the total taxonomic diversity in that sample. The output of a metagenome is a set of sequences representing all the genomes of the organisms present in that sample and can be used to identify the functional potential of a community, for example genes involved in oil metabolism. The standard essential steps in an environmental barcoding project using Sanger sequencing are shown in Figure 9.5: DNA extraction, PCR amplification of one or a few taxonomic DNA markers, cloning to separate PCR products belonging to different taxa to create a clone library and sequencing the cloned PCR products. This results in a few hundred sequences of 500– 900 bp in length that mostly correspond to abundant taxa (Pedros-Alios, 2006), giving a biased view of the community. Rare taxa are often missed. Protistan communities often change by succession rapidly and rare taxa may become abundant. To determine temporal community stability, it is necessary to characterize the entire community. In the last 5 years, next-­ generation sequencing (NGS) technology has allowed scientists to obtain thousands of taxonomic DNA sequences (called reads) per sample, allowing the whole community to be identified (Fig. 9.5). The term ‘next generation’ is somewhat out of date as it is now routinely used. It consists of four different technologies: Illumina HiSeq and MiSeq, Roche 454, Ion Torrent Personal Genome Machine (PGM) and Life Technologies SOLiD (reviewed by Scholz et  al., 2012). A third-generation sequencing technology called single molecule real-time or SMRT sequencing (Chin et al., 2013) also exists but is used less frequently. It is beyond the scope of this chapter to describe these technologies in detail but all essentially add on a labelledDNA sequence tag to the PCR amplified products that enables each different PCR product to bind to a corresponding DNA tag fixed to a support matrix enabling millions of sequencing reactions to occur in a miniaturized reaction chamber. This process negates the requirement for the labour-intensive and expensive cloning step previously used. The price of NGS is rapidly decreasing and is more economical than clone library sequencing. However, the size and quality of sequence reads are lower, typically between 150 and 600 bp, although increased sequence length capability is rapidly developing. DNA sequencing can identify taxa in a mixed sample without the need for morphological information. It can estimate total diversity, both known and unknown, and can be used to determine occurrence or absence of organisms in different samples. For example, the presence of just one harmful algae species is enough to determine shellfish closure. In any sample, there are

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DNA from mixture of cells in an environmental sample

PCR amplification of taxonomic DNA marker

Bacterial cloning NGS sequencing

Sanger sequencing products from a clone-library

NGS sequencing reads from amplified PCR products FIGURE 9.5:  Alternative DNA sequencing approaches. Library cloning involves capturing each polymerase chain reaction (PCR) product representing individual cells into a plasmid cloned into a bacteria that can be grown to amplify the number of products that can then be extracted from the bacteria and sequenced. There are a variety of next-generation sequencing (NGS) technologies, but most commercially available involve labelling the PCR products with a universal label that can be attached to a solid substrate and each PCR product representing a species will be sequenced at the attachment site to form clusters of sequences. Both sequencing approaches will incorporate errors inherent in the PCR amplification process. The scale of NGS is much larger than clone library so diversity can be explored at a greater depth; however, errors in PCR step plus those specific to each NGS technology are massively amplified compared to clone library, which are sequenced using more accurate Sanger sequencing methodology.

100 methodology many dead organisms. To determine the diversity of living components of samples, RNA sequencing can be used. RNA is a nucleic acid produced to synthesize proteins and so can only be produced by living cells. However, often it is required to quantify organisms. The steps involved in DNA sequencing, especially the DNA extraction method, the target DNA marker used, the PCR and the sequencing processes all create different kinds of bias in the type of organisms represented in the resultant DNA sequence set. Just as morphological identification is biased towards larger organisms with more distinguishing features, genetic methods can be biased by chemical and physical properties. Quantification by DNA sequencing is not recommended and instead other methods must be employed.

Advantages/Disadvantages: clone library versus next-generation sequencing DNA sequencing is the gold standard of genetic identification. Clone libraries represent PCR products from species that are encapsulated into plasmids that are then introduced into bacteria through the cloning process. These transformed bacteria can be grown, further amplifying the PCR product. The PCR products are sequenced by accurate Sanger sequencing methodology. However, the efficiency and biases in the cloning process and the laborious individual selection process mean that clone libraries generate a few hundred sequences per sample at most, leading to bias in sample representation. NGS has a far greater sequence output (thousands per sample) to provide better diversity representation, but it can only handle smaller sized PCR products, and each new sequencing technology has its own inherent errors that are magnified to a greater degree because of the scale of output.

9.5.3  Molecular probes (methods to obtain information on the occurrence of selected species) For more than two decades, numerous environmental and phylogenetic studies based on clone library analyses, and more recently NGS of barcoding genes, have been carried out. They provide a wealth of molecular-based information on the structure and variability of aquatic microbial or protistan communities. As a consequence, the number of publicly available

Reference database with large numbers of sequences e.g 18S rRNA

sequences of barcoding genes is continually growing, providing the background information for the design of specific hierarchically organized molecular probes that target the barcoding gene (e.g. the 18S rDNA) at different taxonomic levels (Quast et al., 2013) (see Fig. 9.6). Usually, molecular probes are single stranded and have a length of 18–25 bases. They are very useful tools related to the detection and monitoring of selected types of microbes and protists in environmental samples, especially in the pico-sized ( 1000 m (Fig. 10.3C). A multiple net system was attached to the bottom of the DeepTow system and used for sampling within a few tens of metres of the deep-sea floor in the 1980s (Fig. 10.3D). This net system was adapted for use on DSRV Alvin for near-bottom studies of plankton in the vicinity of hydrothermal vent sites in the 1990s. On other benthic habitats, such as coral reefs, fixed or stationary net systems that orient to the current’s flow and filter out zooplankton drifting by, nets pushed by divers, and traps have been used to capture plankton close to the bottom.

Closing cod-end systems Sophisticated net systems began to be developed in the 1950s to collect animals at multiple specific depth intervals on a single tow. Single nets equipped with closing cod-end devices preceded multiple net systems by only a few years. One, the Mark III Discrete Depth Plankton Sampler (DDPS), was developed by Aron et  al. (1964) and had four catch chambers separated by solenoid-activated damper doors attached to the back of an IKMT, or a net with a 1 m diameter. It was one of the first to carry underwater electronics to sample depth and temperature, and to send the data up a single conductor cable for display at the surface. The LHPR, a modification of the CPR, was developed in the 1960s as a cod-end sampler (Longhurst et  al.,  1966; Wiebe, 1970). The recorder box, attached to the back end of a net, had Nitex nylon gauze strips that were advanced in discrete steps (15–60 seconds) by an electronics package on the tow frame

(Fig.  10.3C). Also recorded were pressure, temperature, and flow into the net. The LHPR underwent a number of revisions to improve its performance (Haury et al., 1976; Williams et al., 1983). A descendant of the LHPR developed in the 1990s by Dunn et al. (1993) is ARIES (Fig. 10.3E). It has a multiple codend system, water sampler, data logger, and an acoustic telemetry system. Although closing cod-end systems are still being used by some investigators, they suffer from several biases that are difficult to eliminate. Smearing (the spreading of a discrete distribution caused by organisms entering the net at the edge having a longer transit time), stalling (the intermittent passage of plankton down the net and into the cod-end sampler), and hang-up (the permanent adherence of zooplankton to the net mesh) all cause significant error in the patterns of distribution being sampled.

Multiple net systems Paralleling the development of closing cod-end samplers was the development of multiple net systems. The design of the Bé Multiple Plankton Sampler (MPS – Bé et al., 1959) was initially, in the late 1950s, messenger operated, and then, in the 1960s, pressure actuated. It provided the basis for the Bedford Institute of Oceanography Net and Environmental Sensing System (BIONESS), with ten nets, developed in the 1980s (Sameoto et al., 1980; Fig. 10.4A). The Multinet, a modified version of the MPS that carried five nets and was opened and closed electronically via conducting cable, was developed in Germany at about the same time (Weikert and John,  1981; Fig.  10.4B). Another variant, developed by Terazaki (1991) at the Ocean Research Institute (Tokyo, Japan) (which is now the Atmosphere and Ocean Research Institute (AORI) in The University of Tokyo), was the Vertical Multiple Plankton Sampler (VMPS). The Tucker trawl system also gave rise to a series of opening and closing net systems. In the mid-1960s, Davies and Barham (1969) used timing clocks to open and close the Tucker trawl mouth. Also in the 1960s, the rectangular mouth opening trawl (RMT), also known as the rectangular midwater trawl, which was opened and closed acoustically, was described by Clarke (1969). The RMT was expanded into the NIO combination net (RMT 1+8) by Roe and Shale (1979); it was equipped with nets with 1 m2 and 8 m2 mouth openings. This was expanded into a multiple net system with three sets of 1 m and 8 m nets controlled acoustically and then later by a microcomputer unit connected by conducting cable to an underwater electronics unit (Dimmler and Klindt, 1990). In the 1970s, 5-net and 9-net Tucker Multiple Net Trawls were developed by Frost and McCrone (1974). The system was powered electrically through conducting wire and controlled from the surface. Soon after, another modified Tucker trawl system, the Multiple Opening/Closing Net and Environmental Sensing System (MOCNESS), with nine nets and a rigid mouth opening was built (Wiebe et al., 1976, 1985). The current versions of the MOCNESS, which range from systems with a

methodology: zoopla nkton  109

FIGURE 10.4:  A, A 1-m2 BIONESS being washed down after a day tow in Storfjorden, Norway, during the gear intercomparison study

aboard the R/V Johan Hjort in June 1993. B, A 0.5-m² Multinet being deployed in the South Atlantic Ocean from the R/V Polarstern (ANTXXIV/1) on 15 November 2007. C, A 1-m2 MOCNESS being launched in the Drake Passage on ARSV Lawrence M Gould (cruise 11-10) on 11  November 2011. D, The 10-m2 MOCNESS being launched in the South Atlantic Ocean from the R/V Polarstern (cruise ANT-XXIV/1) on 8 November 2007.

0.25 m2 mouth opening for sampling microzooplankton to one with a 20 m2 mouth for sampling midwater fish and micronekton, are computer controlled (Fig.  10.4C,D). Sensors include pressure, temperature, conductivity, fluorometer, transmissometer, oxygen, and light.

Moored plankton collection systems There are very few instrument systems that autonomously ­collect samples of plankton from moorings. Most have been patterned after the CPR or LHPR. The O’Hara Automatic Plankton Sampler built in the 1980s had two rolls of Nitex mesh on which plankton were filtered out of water drawn through

the sampler by a battery powered motor (O’Hara, 1984). Lewis and Heckl (1991) built a modified version of the O’Hara system using a series of nets attached to a belt that was moved in steps across a flow tunnel by a battery powered motor. The Moored, Automated, Serial Zooplankton Pump (MASZP) was built in the late 1980s (Doherty and Butman, 1990) again using two strips of plankton gauze on supply spools that cut across an intake tunnel and were wound onto a take-up spool at discrete intervals during sampling by a battery powered computer controlled pumping system. A commercial v­ersion, the ­ Zooplankton Sampler (ZPS) is available from McLane Research Laboratories, Inc. The lack of such systems may be due to the difficulty of powering them for long periods underwater.

110 methodology 10.2.2  Optical systems Light propagation in seawater has limited range compared to that in air due to both absorption by plants, other particles, dissolved organic matter, and the water itself, and scattering off suspended particles. As a result, optical instruments to detect or image zooplankton in situ must do so at close range (cm to m). The development of useful imaging systems for zooplankton has been challenging. Zooplankton encompass a broad size range, vary in opacity from nearly transparent to completely opaque, and exist at a broad range of numerical densities depending on time of year, location, and depth. Identification to genus or species level frequently requires resolving subtle morphological features. Consequently, researchers have been seeking imaging systems capable of collecting high-resolution images of the contents of relatively large volumes of water. Until recently, limitations in digital imaging technology and light propagation generally prevented imaging systems from collecting high-resolution images from large volumes. The rapid development of higher-resolution imaging sensors (either charge-coupled devices (CCDs) or complementary metal-oxide semiconductor (CMOS)), driven largely by the consumer digital camera and machine vision markets, has resulted in increases in the pixel density of these chips from a few megapixels (MP) to 36 MP or more. As a consequence, it is becoming feasible to image the contents of larger volumes of water without sacrificing resolution. Advances in imaging hardware have resulted in a challenge for users and developers of imaging systems. It is now possible to collect hundreds of thousands to millions of images in a short period of time. A system imaging at 30 Hz would collect over 2.5 million images per day. Just as zooplankton ecologists have learned that it is far easier to collect samples of zooplankton with nets than it is to process those samples, users of digital zooplankton imaging systems are potentially faced with a parallel problem, in that it is far easier to collect images of the water than it is to isolate, count, and identify the zooplankton in those images. To overcome this bottleneck, the community has recognized that the development of user-friendly, efficient image processing software is just as important as the imaging hardware itself. While this review is largely focused on imaging hardware, the importance of image processing software cannot be understated. There are two general types of instruments based on whether they produce an image of the zooplankton present in a water volume (e.g. video, photographic (film), and digital camera ­systems) or use the interruption of a light source to detect and estimate the size of particles (e.g. optical plankton counters).

Image-forming systems mounted on non-opening/ closing nets One of the challenges that imaging systems face is to quantify the contents of a volume of water that is sufficiently large to  contain a representative number of individuals as well as

ensuring that the volume is adequate to encompass most targets in their entirety. Early zooplankton imaging systems utilized either photographic 35 mm film or low-resolution video cameras. Both systems were constrained by relatively low sample volumes, the former because of the slow rates at which consecutive frames could be acquired in order to obtain images along the entire length of a tow, while the systems based on low resolution CCD chips were constrained to a small volume in order to achieve adequate image resolution. One solution to the low sample volume problem was to mount the imaging hardware near the cod end of a net in order to pre-concentrate zooplankton and ensure that multiple individuals were present in each image. Examples of such camera-net systems include a 35 mm system (Ortner et al., 1981; Olney and Houde, 1993) and the video-based Ichthyoplankton Recorder (Froese et al., 1990; Lenz et al., 1995). More recently a new imaging system called Lightframe On-sight Keyspecies Investigation (LOKI) was developed to quantify zooplankton spatial heterogeneity on scales of  100 µm and can be deployed to depths of 3000 m. The UVP is a commercially available system (manufactured by Hydroptic in France). The UVP5 was recently used to characterize zooplankton within the California Current frontal region (Ohman et al., 2012). ZOOVIS was another example of a system designed to image a larger volume of water per image than the VPR while using a light sheet to constrain the dimensions of the image volume and thereby avoid imaging organisms that were out of focus (Benfield et al., 2004). ZOOVIS was a towed system consisting of a down-looking 4 MP digital camera aimed at a flat, forward projected light sheet produced by a pulsed xenon strobe (Fig. 10.6). By matching the depth of field of the camera to the light sheet, only in-focus images of organisms were obtained. The system was designed to image either a 340 ml or 3.4 L volume. Data were telemetered to the surface via an electro-optical cable where they were written to disk and visually sorted and their contents enumerated. The system has been successfully deployed in a coastal fjord and was used to groundtruth the composition of acoustic scattering layers (Trevorrow et  al.,  2005) where it obtained clear images of solitary and aggregated euphausiids Euphausia pacifica. A later system designed for use on an ROV was called ZOOVIS-ROV with the specific goal of ground-truthing the composition of thin plankton layers. This system employed four 532 nm lasers projected backwards towards a forward-facing camera in an off-axis configuration. A 4  MP digital camera synchronized to the lasers collected images from a volume of ~40 ml at user-defined rates of up to 8 Hz. Data were sent directly to the surface over an electro-optical tether from the ROV. Two other systems that are capable of imaging the contents of larger volumes of water are SIPPER and ISIIS. Both systems utilize line scan cameras that provide a continuous record of the contents of a narrow slab of water that passes through a light sheet. In the case of SIPPER, the optics are placed within a tunnel, while ISIIS images a volume using a light sheet projected across an unconstrained region of water towards a camera. One of the advantages of these continuous systems is that they provide data on the spatial interrelationships among zooplankton and other particles in the water column and can therefore provide data on micro- to fine-scale patchiness, as well as interspecies associations and relative orientation among organisms. SIPPER has undergone improvements since the initial instrument was described (Samson et  al.,  2001). The current version (SIPPER-3) images the contents of water as it passes through a 96 mm × 96 mm tunnel with sub-100 μm resolution at a tow speed of 3.0  knots (~  1.5  m/s). Data are recorded internally. SIPPER image processing is supported by PICES software that performs segmentation, classification and analysis.

FIGURE 10.6:  Stand-alone plankton imaging systems. A, Underwater Vision Profiler 4 (Image: G. Gorsky, photo); B, Underwater Vision Profiler

5 (Image: D. Luquet, OOV); C) Large Area Plankton Imaging System (Image: E. Horgan, WHOI); D, Zooplankton Visualization System (Image: M. Benfield, LSU/WHOI); E, ZOOVIS-ROV off Oregon. The imaging system is the can and conical frame projecting from the bottom of the ROV (Image: M. Benfield, LSU/WHOI); F, ZOOVIS-Deep (Image: S. Cook, LSU); G) ISIIS (Image: K. Longnecker, WHOI); H, ISIIS-2 (Image: Bellamare, LLC); I, LOKI (Image: H. Lilienthal, AWI); J) SIPPER (Image: D. Remsen, USF); K, LOPC (Image: R. Lopes, USP); L, ZOOPS-O (Image: Y. Lindemann, HUJ); M, OPC on MININESS (Image: M. Benfield, LSU/WHOI); N, Imaging FlowCytobot (Image: H. Sosik, WHOI).

114 methodology SIPPER ­systems have been deployed on the high-resolution sampler (HRS) tow-vehicle, an autonomous underwater vehicle (AUV), and an autonomous surface system. Publications relating to SIPPER include Remsen et al. (2004) and Luo et al. (2004, 2005). Optical systems generally image far smaller volumes than are sampled by traditional nets. Even at imaging rates of 30 Hz, the volume imaged for a given period of time remains well below the volume that would be filtered by even small nets over the same period. With the development of the ISIIS, this is beginning to change. ISIIS (Cowen and Guigand,  2008) was designed to image a much larger volume than previous imaging systems. It accomplishes this by employing a shadowgraph optical set-up that enables both a long depth of field as well as imaging of nearly transparent organisms due to their subtle density differences relative to seawater. Moreover, the image volume is telecentric, which means that objects are the same apparent size no matter where in the depth of field they are located. Like SIPPER, ISIIS utilizes a line scan camera and it produces a continuous record of the undisturbed contents of the water at a maximum rate of 162 L/s while being towed at 5.0 knots (~ 2.6 m/s). ISIIS is a tethered instrument connected to a vessel by an electro-optical cable. The system can be mounted on a v-fin tow-body or on a new vehicle equipped with an active undulation flight control system (Fig. 10.6). The images from ISIIS have a pixel resolution of 70 μm. ISIIS is commercially available from Bellamare (San Diego, USA), and image processing software based on algorithms described in Tsechpenakis et al. (2008) is available, while even more capable software is under development. McClatchie et  al. (2012) and Timmerman et  al. (2014) contain analyses of data collected with ISIIS. The Deep Zooplankton Visualization System (ZOOVISDeep) also employs shadowgraph optics to achieve a telecentric, long depth of field. Unlike ISIIS, ZOOVIS-Deep employs a 5 MP area scan camera to capture images from the contents of a 100 ml volume (2.22 cm tall × 2.65 cm wide × 17 cm deep) at 15  Hz yielding a volumetric sampling rate of 1.5  L/s. Each image has a resolution of 10.8 µm pixel-1. The system is self-contained with battery power sufficient for several hours and on-board recording. A highly collimated red LED with a 5 µs pulsewidth eliminates motion blur and provides the images with holographic information that enables reconstruction of objects located outside of the in-focus depth of field. The system can be deployed to depths of 2000 m in profiling mode, or tow-yo’d in the upper 200 m by paying out, and hauling back cable. It has recently been deployed in a turbid region of Chesapeake Bay where it collected clear images of copepods, ctenophores, cnidarians and other zooplankton (Bi et al., 2013). The LOKI system (Schulz et al., 2010) discussed briefly earlier in this section utilizes a digital imaging system that constrains entrained water containing zooplankton within a narrow depth of field, within which high-magnification and high-resolution images can be obtained. LOKI is self-contained and has been deployed to 491 m (Schulz et al., 2010). The image

volume and the frame rate of the system described in Schulz et al. (2010) were not clearly explained. The description of the dimensions of the imaged volume is not clear, but appears to be 13 mm × 18 mm × 4 mm deep (0.94 ml) and the images have a resolution of 13.4 µm pixel-1. ZOOPS-O is a combined acoustic and optical imaging system designed to quantify zooplankton. It was developed by Jules Jaffe at the Scripps Institution of Oceanography. A 2 MP area scan camera images the contents of a 106 ml volume at 40 µm pixel-1 resolution. ZOOPS-O was deployed in the Red Sea during 2013 where it collected high-resolution images of a variety of zooplankton. The Jaffe Lab has also recently deployed a new 9 MP imaging system called the Scripps Plankton Camera (SPC). The SPC is a moored colour imaging system that employs dark-field, white light illumination to image the contents of a 10 ml volume at high resolution (7.4 µm pixel-1). The images are processed and segmented in real time and served on the Web (http://spc.ucsd.edu/). This system has sufficient resolution to image both zooplankton as well as larger phytoplankton. Phytoplankton and microzooplankton imaging systems have developed in parallel with those described above, albeit at a slower pace. The FlowCam is a commercial instrument developed by Fluid Imaging Technologies that images phytoplankton and other particles in the lab from 1 µm to 2 mm. It is not an in situ instrument, although an in situ prototype was developed and successfully tested. The FlowCam utilizes a pump to draw the contents of a sample through a narrow imaging cell where a digital camera (colour or greyscale depending on the model) images the contents of the water sample. The user can vary the flow rate and the method of triggering an image (regular frame rate or triggered by fluorescence). The Imaging FlowCytobot is an in situ imaging instrument designed by Heidi Sosik and Robert Olson at the Woods Hole Oceanographic Institution. Imaging FlowCytobot is primarily designed to collect high-resolution images (1  µm greyscale images of marine phytoplankton and microzooplankton from 10  µm and larger) using a high-resolution digital camera mounted in series as part of a flow cytometer system (Olson and Sosik,  2007). The system processes water at a rate of 0.25 ml min-1 with data served on the Web from a continuous data stream (http://ifcb-data.whoi.edu/mvco). The resolution is adequate for identification to genus, or in some cases species, level and a MATLAB-based image classification system enables automated classification of taxa with high accuracy.

Holographic systems Digital holography can provide measurements about the three-dimensional spatial interrelationships among particles and plankton within a defined image volume. No other imaging technique yields this information. Essentially, each hologram digitizes the contents of a small volume of the ocean. Once back in the lab, this information can be reconstructed as a series of two-dimensional slices through the volume, which can

methodology: zoopla nkton  115 then be analysed. Herein lies both the potential of, and at present the limitations of, digital holography—while holograms contain a great deal of high-resolution data on the identities and spatial interrelationships of the contents of a volume, the holographic images are large data files and they require substantially more computational time to process than two-dimensional digital images. A number of research holographic systems have been developed and deployed; however, there is currently only one commercially available system. An in situ holographic imaging system called the holocamera (Katz et  al.,  1999; Malkiel et  al.,  1999) was deployed from a manned submersible and other platforms. The holocamera employed a ruby laser to produce an in-line hologram of the contents of a 6.3 cm diameter cylinder whose length could be between 10 cm and 68 cm. This camera produced holograms within the volumetric range 198– 1346 ml, and the self-contained system was capable of acquiring up to 300 holograms per deployment. The group that produced the holocamera also developed a laboratory system designed to study plankton behaviour (Malkiel et  al.,  2003). The original holocamera was quite large, and in 2005 they described a much smaller, free-floating, neutrally-buoyant, holographic imaging system that was capable of performing pre-programmed vertical profiles (Pfitsch et  al.,  2005). This smaller system, called Holosub, was connected to the vessel by an optical fibre that was re-terminated prior to each deployment. One enhancement in this system was the incorporation of two digital cameras each imaging orthogonal volumes of 40.5 ml, and a lens can be added to the system to produce higher-resolution images at the cost of a smaller volume imaged (Pfitsch et  al.,  2005). Talapatra et al. (2013) describe the application of the Holosub in the coastal waters of Washington State. In the UK, another holographic system called the eHoloCam was developed at the University of Aberdeen. This system, described by Watson et al. (2004), was a derivative of an earlier, much larger holographic imager called HOLOMAR (Watson et al., 2003). The eHoloCam is a lightweight, self-contained system in a housing rated to 1500 m. It produces in-line holograms with 5 µm lateral, and 100 µm axial, resolution, from a volume of 36.8 ml at rates of 5–25 Hz (Sun et al., 2008). The system was commercially marketed in 2007 by CDL (now Teledyne CDL) but no longer appears to be part of their product line. More recently, another in-line, self-contained holographic imager called the digital holographic imager (DHI) was described by Loomis et al. (2007). This system images a volume of 1.3  L with 9  µm lateral and 200  µm axial resolutions at sub-1 Hz frame rates. The system is self-contained and can store up to 1600 holograms on flash memory. New software designed to rapidly process the content of the DHI holograms was described by Li et al. (2007). In 2010 the system was deployed on an ROV and used to measure the size distributions of oil droplets at depths of 1500 m in the Gulf of Mexico during the BP oil spill. This demonstrated that instruments designed with a biological oceanographic purpose can provide important data about other particles in the oceans.

Two holographic imaging systems are commercially available. The LISST-HOLO is marketed by Sequoia Scientific Inc. (Bellevue, WA, USA). This system integrates holographic imaging and a particle size analyser and has pressure and temperature sensors. Descriptions of other holographic imaging systems produced since the holocamera, eHoloCam, and DHI include: Graham and Nimmo-Smith (2010), Bochdansky et al. (2013), and Tan and Wang (2013). The digital in-line holographic microscope (DIHM) marketed by 4Deep (Halifax, NS, Canada; formerly Resolution Optics, http://4-deep.com) includes a submersible instrument designed to produce high-resolution holographic images of phytoplankton (Bochdansky et al., 2013). The DIHM employs a 4.2 MP digital camera to image the contents of 1.8 ml at 7 Hz.

Particle-detection systems Plankton and other particles can be counted as they pass through a light beam. The interruption of the beam provides information on their numbers, while the degree of attenuation of the light signal on a photodetector can provide estimates of the cross-sectional area of a particle. This principle has been used to great effect by the Optical Plankton Counter (OPC)—a commercial instrument that has gained widespread popularity in the zooplankton community and which has provided numerous insights into plankton ecology. The OPC consists of a sampling tunnel (22  cm wide × 2  cm high) within which water carrying plankton and other particles passes through a collimated light beam projected on a photodetector. When light from the incident beam is interrupted by a particle, the output signal by the photodetector is reduced by an amount proportional to the cross-sectional area of the particle. By digitizing the change in the response by the photodetector, the equivalent spherical diameter (ESD) of the particle can be estimated. The initial OPC (Herman, 1988) had a lower particle size detection limit of 550  µm ESD. Improved optics reduced this lower threshold to 220 µm (Herman, 1992). Focal Technologies commercialized the instrument, and the OPC is in widespread use throughout the oceanographic community. The relatively low cost and ease of use of the OPC have made it a popular instrument and there is a large body of literature associated with the OPC. A partial list of publications relating to the OPC includes: Heath et al. (1999), Grant et al. (2000), Labat et al. (2002), and Remsen et al. (2004). While the OPC has gained widespread usage, it can prove a challenge to correctly convert the information it provides on size distributions into numbers of taxa because the ESD frequency information is taxonomically ambiguous on its own. For this reason, most studies that have used the OPC most effectively have relied on independent information (usually from nets) about the relative composition and size distributions of the dominant taxa in the system of interest. Moreover, where gelatinous organisms are prevalent, the partial attenuation of the light beam by translucent organisms can be particularly problematic. At high particle densities, coincident transit of

116 methodology two or more particles through the beam created confusion because the particles were treated as a single particle. In part, as a response to the need to address these issues in the OPC, and also to take advantage of technological advancements, the next generation of the OPC incorporated a laser and a higher-resolution analogue to digital converter (ADC). The Laser Optical Plankton Counter (LOPC: Herman et al., 2004) is the latest version of the OPC and is also marketed as a commercial instrument by Brook Ocean Technologies. The finer resolution of the light detector and use of an ADC with a much faster response time than the module used in the OPC means that particles are digitized multiple times as they pass through the tunnel of the LOPC. This produces a low-resolution approximation of the shape of the particle as it moves through the light sheet. The LOPC has a sampling intake tunnel that is larger than the original OPC allowing a greater volume to be sampled, and the faster ADC allows it to work in much higher plankton densities than the original OPC because it is less ­sensitive to the effects of coincidence (Herman et  al.,  2004). The LOPC has also gained widespread usage and a growing body of literature reflects its popularity. Some recent publications describing LOPC data include: Gaardsted et  al. (2010), Trudnowska et al. (2012), Basedow et al. (2013), Marcolin et al. (2013), and Espinasse et al. (2014).

10.2.3  High-frequency acoustics In addition to optical imaging systems, active acoustic systems are the most capable remote sensing technology currently available and are essential for observing zooplankton in situ. While the transmission of light in seawater is quite limited (as noted above), the transmission of sound at low and moderately high frequencies (1  Hz to 100  kHz) is much greater. Above 100 kHz, sound is more rapidly attenuated largely because of absorption due to the salt (principally magnesium sulphate) in seawater. In spite of this limitation, it is high frequency sound in the 1 kHz to 1 MHz range that is proving exceedingly useful for studies of zooplankton and nekton because it can be used to detect the presence of animals 10s to 100s of metres away from the transducer producing the sound. Thus the use of high to ultra high frequency acoustic scattering techniques has become a common component of many observational programmes in biological oceanography. Stanton (2012) has reviewed the development of a wide range of acoustic systems and their operation over the past 30  years to study zooplankton and fish, including theoretical modelling, laboratory experimentation, instrumentation development, and at-sea experiments. Foote and Stanton (2000) provide a broad overview of current acoustic systems and their use. The acoustic systems have evolved from single-frequency, analogue echo sounders to a plethora of digital systems. The active acoustic systems include single-frequency systems (single-beam, dual-beam, split-beam, multiple beam) and multiple frequency systems. A number of these are described and illustrated in Wiebe and Benfield (2003). Some are hull-mounted

systems, while others have been deployed over the side in towed vehicles (e.g. Wiebe et al., 2002, 2013) (Fig. 10.7A,B); Stanton et  al.,  2010; Knutsen et  al.,  2013 – Figure  10.7C and ROVs (Greene and Wiebe, 1991; Jaffe et al., 1995), used on profiling systems (Holliday and Pieper, 1995; Postel et al., 2007; Lavery et al., 2010 – Figure 10.7D), or used bottom mounted at fixed observatories (Ashjian et al., 1998; Klevjer and Kaartvedt, 2011; Sato et al., 2013). Most narrowband systems typically transmit a gated sine wave, but some narrowband systems have employed a linear frequency modulated signal (‘chirp’) to increase the range resolution and to improve the signal to noise ratio (Ehrenberg and Torkelson, 2000). More recently, broadband systems that span a frequency range useful for studying zooplankton and micronekton have been developed (Stanton,  2009; Stanton et  al.,  2010 (1.7–100 kHz); Lavery et al., 2010 (150–600 kHz)). These broadband systems provide nearly continuous frequency coverage over a broad range of frequencies that enable much better discrimination of backscattering sources (e.g. living organisms, physical microstructure, bubbles, suspended sediments) and improved estimates of size and numerical density of fish and zooplankton. Volume backscattering (integration of the energy return from all sources in a given ensonified volume, i.e. echo integration) and target strength (TS) (echo strength from individual backscattering sources) are the two fundamental acoustic measurements (Lavery et  al.,  2007). Either or both of these measurements can be obtained depending upon the construction of the echo sounder and transducers. Only volume backscattering can be determined directly with a single-beam transducer, whereas a multiple beam acoustic system can also provide individual TS (Stanton, 2012). Interpretation of the backscattering signal in terms of biologically meaningful properties is a complex problem (Lavery et al., 2007; Stanton, 2012) primarily because the backscattering is a function of the acoustic properties of the sources. Even if all of the backscattering is from biological entities, their contribution depends on their size, shape, orientation, and material properties (sound speed contrast and density contrast), which are highly variable among the taxa. No single model can account for the frequency dependent variation that characterizes different taxa in mixed assemblages. Over the past 20 years, three classes of models have been developed to account for backscattering by zooplankton with 1) fluid-like bodies (e.g. copepods, euphausiids, amphipods, salps), 2) elastic shells (pteropods) and 3) gas inclusions (siphonophores) (Stanton et  al., 1994,  1998; see Lavery et  al.,  2007 for details of these models). Traditionally, multiple frequency and broadband acoustics data has been combined with one or more models to interpret the data in terms of animal abundance and size (Holliday and Pieper, 1995; Lawson et al., 2008; Lavery et al, 2010). The quality of these inverse solutions depends strongly on the biological complexity of the water that is ensonified and the number of frequencies or the broadband frequency range used. Lavery

methodology: zoopla nkton  117

FIGURE 10.7:  A, The BIo-Optical Multi-frequency Acoustical and Physical Environmental Recorder (BIOMAPER-II) locked in the docking mechanism and ready to be brought on board the R/V Endeavor in the Gulf of Maine on 13 December 1999 (Wiebe et al., 2002). B, The ‘Greene Bomber’ towed v-fin being launched in the Slope Water south of New England from the R/V Connecticut on 15 July 2010. It carried a Hydroacoustic Technologies Inc. four-frequency acoustic system (43, 120, 200 and 420 kHz). C, The MESSOR towed body being deployed in the Iceland Sea north of Iceland from the R/V GO Sars on 8 May 2013. It carried multiple down-looking Simrad EK 60 transducers as well as a suite of other environmental sensors, a VPR, and a multi-net. D, The WHOI-Edgetech broadband profiling system being deployed in the mid-Atlantic Bight from the R/V Oceanus in July and August 2006. It had 4 channels/transducers ranging from 160 to 600 kHz, a CTD, and a roll-pitch-yaw sensor (Lavery et al., 2010).

et al. (2010) and Stanton (2012) describe the current constraints associated with the inversion problem. Acoustics per se cannot be used to identify species or taxa of plankton. ‘Ground-truthing’ is central to accurate interpretation of acoustics data. Net tow or optical data are needed to confirm the presence of the scattering types inferred from the acoustics data. With collocated acoustics and net tow data, a procedure often referred to as the ‘Forward Problem’ can be carried out that enables comparison of observed volume ­backscattering levels versus those predicted on the basis of net catches (Wiebe et al., 1997, 2013; Lavery et al., 2007). The procedure involves determining the abundance and length of the individuals in each sample and then using the appropriate backscattering model for the individual’s taxonomic group to calculate their backscattering cross section. Estimates of expected backscattering cross sections for each individual are summed over all individuals in each taxon and then over all taxa to yield an estimate of the total expected volume backscattering strength in the volume defined by the depth range sampled by the net. The prediction is then compared with the observed volume backscattering coefficient from the collocated acoustics data. The degree of agreement between the estimated and the

observed backscattering values provides a measure of how well the sources of backscattering are understood.

10.3  Intercomparison of Zooplankton Sampling Systems The diversity of sampling equipment that has developed since the beginning of quantitative sampling in the late eighteenth century has given rise to the need to intercompare the samplers and to evaluate their performance. Sources of error are primarily escapement of individuals through the net mesh, avoidance of the mouth of the net system, and variation caused by the irregular patchy distribution of the organisms in the field (Skjoldal et al., 2013). For quantitative studies, the volume of water filtered by a net needs to be accurately measured (Box  10.2). Most studies were comparisons between pairs of samplers, or comparisons of variants of the same gear or different mesh sizes on the same gear (see Table 2 in Wiebe and Benfield, 2003). A recently published study intercompared and evaluated methods for sampling and determination of zooplankton

118 methodology Box 10.2  Calculation of volume filtered In quantitative sampling of zooplankton, the volume of water filtered by a net or other sampling device (pump) needs to be measured. The normal procedure is to use a calibrated flow meter mounted in the net mouth to measure the distance travelled by the net. For nets with a bridle, the flow meter should be located halfway between the centre and the rim of the mouth (Tranter and Smith, 1968; Working Party No. 2, 1968). The distance travelled multiplied by the mouth area of the net equals the volume filtered assuming 100% filtration efficiency (see Smith et al., 1968). In the case of a vertical tow, the net is lowered to depth and then hauled back to the surface. The flow meter should be designed so that it does not spin while the net is lowered to depth and the net only filters water on the upward portion of the tow. In the case of an oblique tow, the flow meter is operational from the time the net enters the water until it arrives back at the surface. If a net stops accepting all of the water encountered by the net mouth as  a result of the net mesh being clogged, this will cause misrepresentation of the water column sampled. Assessment of clogging can be done by use of one flow meter positioned in the net mouth and another positioned outside the rim of the net (Smith et al., 1968). The second flow meter can provide a means of evaluating the filtration efficiency of the net. Careful calibration of flowmeters (in terms of revolutions per distance travelled) is essential for quantitative studies.

d­ istribution and biomass of a number of sampling systems currently in use, including the WP-2, CalCOFI 1-m Ring Net, MOCNESS, BIONESS, MultiNet, LHPR, Gulf V, OPC, and CPR (Skjoldal et  al.,  2013). Many of these net systems are used at time-series sampling sites around the North Atlantic (Table 10.1). Net mesh size had a major influence on the biomass and species composition of the zooplankton community. There was a consistent relationship between escapement through the mesh and the width of the organisms. In comparisons of nets with 333 μm, 180 μm, and 100 μm mesh, about 50% of the organisms escaped through the mesh at an individual width equal to the mesh size (see Figure 34 in Skjoldal et al., 2013). Active avoidance of the sampler, while not a significant problem for the mesozooplankton, is important for the larger macro- and megazooplankton. Traditional methods to reduce the avoidance have been either to increase the towing speed or to increase the size of the net mouth opening (Clutter and Anraku,  1968). However, increased towing speed to reduce avoidance gives rise to increased extrusion of smaller organisms through the net mesh and adds to the loss due to escapement. Nets large enough to reduce avoidance are difficult to deploy from most research vessels. Recent efforts to reduce avoidance have been to use bright flashing lights to create a blinding effect so that individuals cannot see the net, since strong net avoiders such as krill and fish have good visual acuity (Sameoto et  al.,  1993; Wiebe et  al.,  2004,  2013; Simard and

Sourisseau, 2009). For example, use of a strobe light increased the catch of krill by factors of 5 to 10 or more and with the strobe light on, acoustic and net estimates of their abundance agreed (Wiebe et al., 2013). In the Skjoldal et al. (2013) study, an important conclusion was that different vertical, oblique, and multiple opening/closing net systems produced similar estimates of zooplankton biomass and abundance when operated with comparable mesh-sized nets and a sufficiently high mesh open area to mouth opening ratio. This study also emphasized that no single net system is able to effectively sample the broad size spectrum of zooplankton ranging from the microzooplankton to the megazooplankton.

10.4  Preservation of Samples Much has been written about the preservation, storage, and analysis of plankton samples. The types of fixatives used to preserve zooplankton samples depends upon the objectives of the research and how the samples are to be analysed. A comprehensive review of the methods of zooplankton preservation was a product of the Scientific Committee on Oceanic Research (SCOR) working group 23, which worked from 1968 to 1972 (Steedman, 1976). More recent reviews focused on preservation of protozooplankton or for zooplankton genetic studies can be found in Gifford and Caron (2000) and Bucklin (2000). The following is a brief review of the methods that are widely used.

10.4.1  Preservation for sample enumeration and taxonomic morphological analysis For zooplankton sample enumeration and taxonomic morphological studies, a seawater formalin solution containing about 4% formaldehyde buffered with sodium tetraborate decahydrate is generally used (Steedman,  1976; Postel et  al.,  2000). Buffering the preservative to a pH of about 8.2 is especially important for calcareous shelled zooplankton such as pteropods. With too little buffering, the calcareous shells will dissolve. The pH of a newly formalin-preserved zooplankton sample will often decrease over time, and so monitoring of the sample pH and rebuffering is essential to maintain the sample pH until the preservative stabilizes, especially in the first few days and weeks after collection. Since formalin and its fumes can be carcinogenic, it is best to work with formalin-preserved samples in a fume hood. For microzooplankton (heterotrophic flagellates, dinoflagellates, ciliates, and sarcodines), the complexity of the make-up of the animals requires several different preservation methods in order to assess the entire suite of taxa in a sample (Stoecker et al., 1994). Commonly used are formaldehyde, glutaraldehyde, Bouin’s solution, and Lugal’s iodine solution (Gifford and Caron, 2000). For heterotrophic dinoflagellates, formalin or glutaraldehyde is suitable since they do not interfere with chlorophyll fluorescence. Formalin is commonly used

methodology: zoopla nkton  119 TABLE 10.1:  Sampling gear and net mesh used at time-series sites in the North Atlantic. The information in this table was excerpted from

O’Brien et al. (2013), which has additional information on the sites.

Site

Sampling gear

Sampling mesh

Site

Sampling gear Sampling mesh

USA/NEFSC Ecosystem Monitoring Program

Bongo net (60 cm)

333 µm

Germany/Leibniz Institute for Baltic Sea Research, Warnemünde (IOW)

WP-2 Net (57 cm)

100 µm

Canada/Atlantic Zone Monitoring Program (AZMP)

Ring Net (75 cm)

200 µm

Sweden/Swedish Meteorological and Hydrological Institute (SMHI)/HELCOM Monitoring

WP-2 Net (56 cm)

90 µm

USA/Bermuda Rectangle net (80 Atlantic Timecm × 120 cm) series Study (BATS)

202 µm

Germany

Apstein and CalCOFI

150 µm, 500 µm

Iceland/Icelandic Spring Cruise

WP-2 Net

200 µm

United Kingdom MSS Inshore Ecosystem Monitoring Programme

Bongo net (40 cm)

200 µm

Faroe Islands/ Faroe Marine Research Institute (FAMRI)

WP-2 Net

200 µm

Plymouth Marine Laboratory (PML)

WP-2 Net

200 µm

Norway/IMRBergen

WP-2 Net (56 cm)

180 µm

Spain Zooplankton Ecology Group/University of the Basque Country (ZEG/ UBC)

Ring Net (25 cm)

200 µm

Finland/Finnish Environment Institute SYKE/ HELCOM Monitoring

WP-2 Net (56 cm)

100 µm

Spain/Instituto Español de Oceanografía (IEO)—Spain; seRies temporAles De oceanografIA en eL norte de ESpaña (RADIALES)

Juday Net (50 cm)/WP-2 Net (38 cm)/ Bongo net (40 cm)

250 µm/ 200 µm/ 200 µm

Estonia/HELCOM Juday Net (36 cm) Monitoring

100 µm

Portugal/Instituto Português WP-2 Net do Mare da Atmosfera (50 cm) (IPMA)

200 µm

Portugal/Centre of Marine Sciences

200 µm

Latvia/National monitoring programme of Latvia

WP-2 Net (57 cm)/ 100 µm/160 µm Juday Net (36 cm)

Poland/HELCOM Monitoring

WP-2 Net (57 cm)

100 µm

Note: HELCOM, Baltic Marine Environment Protection Commission—Helsinki Commission.

WP-2 Net (40 cm)

120 methodology for shelled sarcodines, but precaution is needed to maintain the pH of the solution for calcareous forms and, in the case of acantharia, the concentration of strontium in the preservation liquid with the addition of strontium.

10.4.2  Preservation of samples for genetic analysis Zooplankton samples and specimens intended for genetic, genomic, or transcriptomic analysis should be collected so as to ensure that they are alive and healthy (usually indicated by their transparency) at the time of preservation or freezing. Samples intended for genetic analysis should be collected so as to minimize the length of time between net capture and recovery. Damage can be minimized by collection in short-duration near-surface net tows using large, baffled, and solid cod ends (see Fig. 10.1C). Samples can be split immediately upon capture to allow rapid preservation of a fraction to be used for genetic analysis. Specimens can be identified under a microscope using a specimen stage that can be cooled—or located in a cold room – to ensure specimens are alive at the time of preservation or freezing. For specimens intended for gene expression analyses, ‘collection effects’ (i.e. damage and disturbance resulting from net capture) may be significant and generate artefactual results; special care should be taken in designing appropriate collection strategies. The broad range of molecular approaches and protocols may be divided according to whether the focus of analysis is DNA or RNA (Fig. 10.8). This is a critical determinant of the sample preservation methods to be used. Formalin is not a good preservative if the zooplankton are to be subject to genetic studies. In general, DNA can be preserved most easily in undenatured alcohol (95% or 100% ethanol). The sample should be drained of seawater before the ethanol is added. There should be 3 to 4 times more alcohol than plankton volume. Alcohol as a preservative is much less effective than formalin, and spoilage of overcrowded or high-biomass samples is a distinct likelihood. It is important that the alcohol be changed 24 hours after the initial preservation, with continued changes until the alcohol remains clear. Long-term storage of alcohol-preserved zooplankton samples for genetic analysis should be under temperature-­controlled conditions (ideally −20°C to 4°C). DNA is rapidly denatured and destroyed at temperatures above 25°C. Bucklin (2000) provides additional details on collection, alcohol preservation, and storage of zooplankton samples for DNA analysis. Additional methods for preservation of zooplankton samples for analysis of DNA include acetone (Goetze and Jungbluth, 2013), as well as various buffer (salt) solutions, for example DESS (dimethyl sulphoxide, disodium EDTA, and saturated NaCl) (Yoder et al., 2006). For some groups, especially gelatinous zooplankton taxa, buffer solutions may have significant advantages in preserving both morphological structures and DNA for genetic analysis (see Laakmann and Holst, 2013). Genetic analysis of zooplankton has been slowed by preservation and storage of samples in formalin solutions, which are

Target gene sequencing Genomic fingerprinting (SNPs) Fragment analysis (Microsatellites)

Species identification Population genetics Phylogeny

Whole-genome sequencing

Whole transcriptome sequencing

Stress responses

Target gene expression (q-PCR)

Physiological condition

Gene function (ontology)

Metabolism

FIGURE 10.8:  A broad range of molecular approaches and protocols used for analysis of zooplankton samples may be divided according to whether the focus of analysis is DNA or RNA, as shown in the diagram. Requirements for sample collection, preservation, and storage are determined by the molecular properties to be examined (see text for explanation).

not always adequately buffered for pH. Numerous protocols exist to recover DNA from formalin-preserved tissue, including for zooplankton (Bucklin and Allen,  2004; Kirby and Lindley, 2005; Laakmann and Holst,  2013) and fish (Zhang,  2010). However, both the aqueous matrix and the rapid acidification of unbuffered formalin exacerbate the difficulty of recovering DNA from archived zooplankton samples. Long-term storage in buffered formalin does not preclude some types of molecular genetic analysis, but the yield of DNA and the success of molecular analyses depend on the length of time the tissue has been exposed to formalin and the pH of the solution (see Tang, 2006). Quantitative examination of gene expression requires analysis of RNA, which can be preserved by deep freezing in liquid nitrogen or on dry ice. RNA can be preserved for long periods by keeping specimens deep frozen in liquid nitrogen or in an ultracold (−80°C) freezer. Various custom-made and commercially available buffer solutions may be used for zooplankton samples intended for analysis of RNA. An example is RNA ­laterTM Storage Solution (Sigma-Aldrich Co.), which—according to the manufacturer—preserves RNA stably for 1 week at room temperature and indefinitely at −20°C. Regardless of the medium of preservation and storage, the greatest danger to zooplankton samples intended for genetic

methodology: zoopla nkton  121 analysis is heat. RNA is particularly labile and can be maintained only in cold (in buffer solutions) or ultracold conditions. DNA is rapidly consumed by enzymes at temperatures above freezing; DNA is denatured—even if preserved in buffer or alcohol—at temperatures above 37°C.

10.5  Analysis of Samples A detailed review of the methods of analysis of zooplankton samples is provided in Postel et al. (2000). Population genetic analysis of zooplankton is reviewed by Bucklin (2000); also see Hellberg (2009) and Bucklin et  al. (2010a). This section will briefly highlight some of the more widely used methods.

10.5.1  Determination of biomass, taxonomic composition, and size by traditional methods Biomass measures A common starting point is the determination of the total biomass of the zooplankton in the sample. Volumetric (settled volume, displacement volume) and gravimetric (wet mass, dry mass, ash free dry mass, and carbon/nitrogen) methods provide bulk estimates (Postel et  al.,  2000). The volumetric measures are preferred when other measurements of the taxa or species in the samples are needed. Settled volume is the least precise and is not recommended because it is difficult to convert the measurement to more meaningful constituents such as carbon or nitrogen. Direct measurements of carbon and nitrogen are

Box 10.3  Determination of abundance from a plankton sample D = N × S/V where: D = Density of organisms in numbers per cubic metre N = Number of organisms S = Split factor V = Volume of water filtered As an example, a sample is taken with a 1 m diameter ring net on an oblique tow that filtered 150 m3. The sample was split twice and a count of the animals in a 1/4 split was made. For one species (sp1) a count of 100 individuals was obtained and for a second species (sp2) the count was 4. The estimates of the number of individuals per m3 are: sp1= 100 × 4/150 = 2.67/m3 sp2 = 4 × 4/150 = 0.11/m3 Assuming the individuals in the sample were randomly distributed while being split, their distribution in the sample can be assumed to follow a Poisson distribution (Lund et al., 1958; Postel et al., 2000). The confidence limits on the counts can be computed using the fact that for the Poisson distribution, the variance = mean. The 95% limits for sp1 are therefore = ± 1.96 × sqrt(100) = 19.6, so 100 ± 19.6 = 80.4 to 119.6, and for sp2 = ± 1.96 × sqrt(4) = 3.92, so 4 ± 3.92 = 0.08 to 7.92. These values can be used to give the limits for the /m3 estimates: sp1upper limit = 4 × 119.6/150 = 3.19 sp1upper limit = 4 × 80.4/150 = 2.14 sp1upper limit = 4 × 7.92/150 = 0.21 sp1upper limit = 4 × 0.08/150 = 0.002 How many individuals of a species need to be counted? The answer depends on the level of confidence that is required in the study and how abundant the species is. For rare species, the entire sample may need to be scanned to count the few present. For more abundant species, an aliquot containing 100 to 200 may provide the required counting precision (see Postel et al., 2000 Table 4.11). However, it is important to note that if the animals in the sample being subdivided are not randomly distributed, the variance will be larger and these error estimates based on the Poisson distribution will be conservative (Longhurst and Seibert, 1967). Also the counting error is likely to be small relative to the field sampling error that is subject to the patchy non-random distribution of most zooplankton (Wiebe and Holland, 1968). Based on work by Winsor and Clarke (1940), it has been common practice to log-transform (base 10) the abundance estimates to remove the dependence of the variance on the mean and to stabilize the variance. Once the variance, s2, of a single observation is obtained, the square root of the variance (the standard deviation, s) is multiplied by the t-distribution 95 limit value (in most cases, 1.96). The antilog of this value and its reciprocal provide 95% limits of a single observation. Typically, replicate tow variability ranges from 1/2 to 2 times to 1/6 to 6 times a single observation (Wiebe and Holland, 1968), which is usually much larger than the counting error.

122 methodology the most accurate, but their measurement results in the destruction of the sample. There are additional biochemical approaches such as the determination of organic constituents that can be measured on bulk samples, for example protein, lipids, carbohydrates, ATP, and chitin as well as elements in addition to carbon and nitrogen, for example hydrogen and phosphate (Postel et al., 2000). Total biomass depends on the taxon composition of the samples and the partition of the biomass according to taxon is often desired.

Taxa identification and counting A higher level of quantitative analysis involves identification and counting/measuring of taxa, species or stages of species in a sample to determine the abundance per taxon, species, or stage (see Box  10.3). The traditional way of making these measurements is with a binocular microscope often with dark-field illumination and equipped with a calibrated ocular scale to measure individual lengths or widths, together with a multiple mechanical counter to register counts of individual species (Fig. 10.9). A Bogorov style tray (Bogorov, 1927) constructed with a series of connected channels is often used to facilitate the counting of species in a sample (Figure 10.9). A sample is poured into the tray and then under 6× or 12× magnification the tray is moved along from the starting channel to the end while counting the species as they are encountered. A skilled counter can use the mechanical counter to register each individual with little or no recourse to look at the ­counter. The zoom capabilities of modern microscopes allow the counter to look for key taxonomic characters to ensure accurate identification of a species. A more expansive discussion of how microscopic analyses are performed is given by Postel et al. (2000).

Newer techniques can provide some of the same information from analysis of high-resolution image scans of a sample (Benfield et al., 2007). The digital images can be made on a flatbed scanner from a silhouette image of a sample (Ortner et al., 1979; Davis and Wiebe, 1985) or from a direct scan of a sample (Grosjean et  al.,  2004; Gorsky et  al.,  2010), or with a ­digital camera or video system attached to a microscope. Automated and semi-automated methods of analysing digital images of samples have been developed beginning in the 1980s (see review by Benfield et  al.,  2007). A number of software packages (some open-source) are now available to enable rapid analysis of the images and provide enumeration of abundances, size measurements, and biomass. These include Visual Plankton (Davis et  al.,  2005), ZooProcess with Plankton Identifier (Gorsky et al., 2010), ZooImage (Bell and Hopcroft, 2008), the WHOI Silhouette DIGITIZER (Little and Copley, 2003).

Size-frequency measurements The length and width measurements of individuals often made during the counting process can be used for a variety of purposes. They can be combined with allometric methods to ­enable the computation of their wet mass, dry mass, or carbon mass (Postel et  al.,  2000; Kiorboe,  2013). For example, Davis and Wiebe (1985) used silhouette derived taxon abundance and length measurements (Fig.  10.10) to estimate changes in the trophic structure of a warm-core ring. Mauchline (1998) has summarized a long list of conversion equations to interchange between morphometric and biomass measures for calanoid copepods, and Siegel and Nicol (2000) have done the same for euphausiids. Fewer conversion factors are available for other zooplankton taxa. With individual specific size, other computations based on body size can be made of physiological rates, i.e. Banse and Mosher’s (1980) conversion of body size to productivity, Hirst and Bunker’s (2003) conversion of body size to species productivity, and Ikeda (1985) and Ikeda et al.’s (2001, 2007) conversion of body size and temperature to respiration and excretion. With the bulk dry mass measure and the total counts of the zooplankton in the sample, the biomass concentration divided by the abundance provides an average individual specific dry mass, which may be sufficient for the first order estimation of physiological rates. Of particular importance is the need to describe how the length measurements were made and to conform to the ways they were made by previous investigators in order to make valid comparisons. For example, Mauchline (1980) describes the many ways the lengths of euphausiids have been made over the years, and it is important to know which length was used when making comparisons.

Multiple sample use FIGURE 10.9:  A traditional binocular microscope set up for counting

zooplankton, including a camera mounting adapter, top and bottom lighting, a Bogorov style tray ready for counting, and a stacked array of laboratory counters. Note that as part of the process, the samples can be sieved to provide size fractions for further processing (Be´, 1968).

Frequently, zooplankton samples must serve multiple objectives and must be analysed in ways that are incompatible with a single preservation procedure. In addition, for some analyses, the sample may be too large for the analysis procedure and

methodology: zoopla nkton  123 Examples of Zooplankton Standard Lengths

Thaliacea

Amphipoda

Ctenophora

Copepoda

Euphausiacea

Mysidacea

Ostracoda

Cladocera

Crustacean larvae

Siphonophora

Medusae

Decapoda

Chaetognatha

Larvacea

Radiolaria

Heteropoda

Pteropoda

Foraminifera

FIGURE 10.10:  Examples of a variety of zooplankton types with guides for making standard length to be used with length to wet weight

conversion formulas in the Silhouette Digitizer program (Little and Copley, 2003). Redrawn from Isaacs et al. (1971, Figure 1).

must be split into aliquots. The two predominant splitters are the Folsom Splitter (McEwan et al., 1954) and the Motoda Box  Splitter (Motoda,  1959) and subsequent modifications (e.g. Longhurst and Seibert, 1967; Scarola and Novotny, 1968). There are also subsamplers that extract a small volume from the ­sample for analysis (e.g. the Stempel pipette: Hensen, 1887; Frolander, 1968; Postel et  al.,  2000). As with all subsampling devices, errors arise during the subsampling procedure. A number of studies have examined the statistics of subsampling both theoretically (Ricker,  1937; Venrick,  1971) and empirically

(McEwan et  al.,  1954; Longhurst and Seibert,  1967; Sell and Evans,  1982; Van Guelpen et  al.,  1982; Griffiths et  al.,  1984). Generally, counting error arising from sample splitting is smaller than the variation associated with field sampling, but the actual subsampling error can exceed the error expected from a random model primarily because the animals in a ­sample clump together giving rise to a bias in the splitting (Sell and Evans, 1982; Van Guelpen et al., 1982). In cases where more than one preservation procedure is needed, or parts of a sample are needed for an analysis that will

124 methodology Zooplankton Processing

Volumes and Species of Scyphozoa and Ctenophora

Cod end Plankton Splitter Formaldehyde

1:10

Formalin 1

St. 543 “Johan Hjort” 24–12–03 WP2-56 cm - 180 μm 1/ sample 350–0 2

2 Preserved Samples

Seawater

Seawater

2000 μm

Sieving Tray

1000 μm

180 μm

Sieving Tray

Fish larvae Krill Shrimp Freshwater 1000–2000 μm

>2000 μm

180–1000 μm

2.346 g

Drying Oven 70 deg.

Freezer

Scale (mg)

Dry weight samples

FIGURE 10.11:  A diagram illustrating the treatment of zooplankton samples collected during the zooplankton gear intercomparison study in Storfjorden Norway on the R/V Johan Hjort in June 1993. The samples were split with a Motoda splitter. Reprinted from Skjoldal et al. (2013, Figure 4) with permission.

methodology: zoopla nkton  125 CMarZ Sample Handling Protocol On Deck 1) Wash nets carefully 2) Put cod-ends into numbered buckets with ice to keep samples cool. 3) Move buckets expeditiously into the walk-in cold room. 4) Samples logged in.

Stratified and Integrated Samples In Wet Lab (1) Three teams (Picker/Recorder) Remove/Record large specimens of:

In Lab

1) Gelatinous forms 2) Fish 3) Macrozooplankton/nekton

Photo Imaging Species Identification Dissection/Preservation

All specimens removed recorded in CMarZ Specimen Log

Integrated Samples

Stratified Samples

In Wet Lab (2)

In Wet Lab (2) Splitter Team splits and preserves samples 1/

split: (A) removed, sieved, preserved in formalin 1/ split again to: 2 1/ split: (B) for live picking in main lab 4 and then alcohol preserved for later texonomic analysis 1/ split: (C) preserved in alcohol. 4 2

Splitter Team splits and preserves samples 1/ split: 2

(A) used for live picking, then preserved in alcohol

1/ split: 2

(B) to formalin for later taxonomic analysis (preservation under hood)

Ship Laboratory Sample Analysis Split: (A) archived after used for silhouette analysis Split: (B) used for taxonomic analysis on board Split: (C) used for archive

In Lab BARCODE vial with specimens Extraction, PCR, Sequencing

FIGURE 10.12:  A diagram illustrating the treatment of zooplankton samples collected during Census of Marine Zooplankton cruises to  the  north-west Atlantic on the R/V Ronald H Brown (06-03) in April 2006 and the R/V Polarstern (ANT-XXIV/1) in November 2007 (Wiebe et al., 2010).

render the sample useless for other analyses, the sample can be split before preservation and the subsamples can be processed differently. A recent example of this is given by Skjoldal et al. (2013) in which the samples collected for a gear intercomparison study were processed for multiple objectives (Fig. 10.11). In this case, the samples were split and one-half preserved and used for quantitative counts and sizing of the species present, and the other half sieved into three size fractions for dry mass  determinations after first sorting out large individual

euphausiids, shrimp, and fish larvae. A second example involved sampling on two Census of Marine Zooplankton cruises in the Atlantic Ocean where sampling of different depth intervals between the surface and 5000 m took place (Wiebe et al., 2010). In this case, the stratified samples (after most large gelatinous forms, fish, and macrozooplankton/nekton were removed for individual analysis) were split (Fig. 10.12) with half preserved in formalin for laboratory study, including biomass estimates (e.g. displacement volume), species counts, and other quantitative

126 methodology analyses. The other half was split again with ¼ (split B) for live picking of species that were identified by skilled taxonomists and then sent to the shipboard lab for DNA barcoding or were frozen in liquid nitrogen for later genetic study. The other ¼ (split C) was preserved in alcohol for additional genetic study. A similar protocol was applied to the sample that integrated the water column during the net deployment to depth.

10.5.2  Genetic analysis of zooplankton samples Molecular genetic, genomic, and transcriptomic approaches (see Box 10.4) can be used to examine zooplankton taxonomy and systematics, population genetics and demographic history, physiological responses to environmental conditions and change, phylogeny and evolution, among others. Species of any zooplankton group may be discriminated, identified, and detected using DNA or RNA variation (Fig. 10.8). Characterization of population genetic diversity and structure yields new understanding of population connectivity and dispersal pathways of zooplankton in ocean currents. Physiological responses of organisms may be detected by gene expression analysis using whole-transcriptome sequencing and quantitative PCR (qPCR). Environmental sequencing and metagenetic analysis (i.e. the large-scale analysis of taxon richness via the analysis of homologous genes) of unsorted zooplankton samples is allowing rapid analysis of biodiversity at multiple levels of organization: genetic, population, species, guild, and ecosystem. Analysis of genomic diversity may help in the understanding and prediction of whether and how zooplankton populations and species may adapt and evolve in response to climate change.

using ­ morphology and molecules for identification of zooplankton species, has become increasingly widespread ­ (McManus and Katz, 2009; Cornils, 2015). DNA or RNA variation can be used to develop accurate, reliable, and rapid tools for discrimination and identification of species, including morphologically cryptic ones. DNA sequence variation of the mitochondrial cytochrome oxidase subunit I (COI) barcode region has been widely used for species identification of zooplankton (Bucklin et al., 2010a, 2010b, 2011; Jennings et al., 2010a, 2010b; Ortman et al., 2010; Li et al., 2011; Cheng et al., 2013; Laakmann and Holst, 2013; Maas et al., 2013; Blanco-Bercial et al., 2014). Other marker gene regions have also been used, including mitochondrial 16S rRNA (e.g. Goetze,  2010; Lindsay et  al., 2015), nuclear intervening transcribed spacer (ITS-1) (Lehtiniemi et al., 2013), and 28S rDNA (Hirai et al., 2013). Nongenic characters used for species identification are based on genomic fingerprinting, including detection of single nucleotide polymorphisms (SNPs) (De Faveri et al., 2013) and insertion-deletion polymorphisms (Smolina et al., 2014).

Population genetics, phylogeography and phylogeny

Many important ecological and evolutionary processes are reflected in spatial and temporal patterns of genetic variation of species, including genetic structure (i.e. the distribution of genetic variation within and between populations), connectivity (gene flow among populations), and demographic history. Among ocean processes that may drive population genetic structuring of zooplankton are currents, persistent eddies, ocean gyres, and other physical ocean structures at mesoscale to large scales (Peijnenburg and Goetze, 2013). Geological features—continents, islands and other land forms, continental Integrative morphological–molecular taxonomy shelves, seamounts, and ocean ridges—may form natural barTaxonomic analysis of zooplankton samples can be time-­ riers to dispersal. In contrast, cosmopolitan or circumglobal consuming and complicated, requiring expertise in species species may exhibit genetic differentiation of populations at identification spanning the many groups occurring in the plank- large scales (e.g. between ocean basins) due to reproductive isotonic assemblage. Use of integrative taxonomic approaches lation caused by limited dispersal among distant populations (Dayrat, 2005; Goldstein and DeSalle,  2011), especially those (Blanco-Bercial et  al.,  2011a; Peijnenburg and Goetze,  2013). Genetic analysis can provide a window into the evolutionary history of a population or species. Related analytical approaches include phylogeography, the description of the geographical disBox 10.4  Glossary of genetic terms tributions of the genetic lineages within a population or species (e.g. Blanco-Bercial et al., 2011a), and phylogeny, the reconstrucGene expression: Conversion of the information encoded in a gene tion of the evolutionary relationships among the population or (DNA) first into messenger RNA (mRNA) and then to a protein. species and its relatives (e.g. Blanco-Bercial et al., 2011b). Gene ontology: Automated reasoning from a relational database of homologous genes whose functions are known or inferred, usually for a model species. Genomics: Study of the full complement of hereditary information of organisms. Metagenetics: High throughput DNA sequencing of one or more target gene markers in mixed or environmental samples. Transcriptomics: Study of all RNA molecules (mRNA, rRNA, tRNA) in cells.

Environmental sequencing and metagenetic analysis Methodological advances in high throughput DNA sequencing are allowing analysis of unsorted environmental samples (i.e. environmental sequencing) and metagenetics (i.e. the largescale analysis of taxon richness via the analysis of homologous genes). These novel approaches can provide detailed and accurate biodiversity assessments of marine communities (Bourlat et al., 2013; Ji et al., 2013). The taxonomic complexity of marine

methodology: zoopla nkton  127 zooplankton assemblages, potentially including numerous cryptic, novel, rare and introduced species, makes such approaches particularly useful in detecting what has been termed the ‘hidden diversity’ of zooplankton (see Lindeque et al., 2013). Machida et  al. (2009) pioneered the use of environmental barcoding, also called metabarcoding (i.e. DNA sequencing of COI from unsorted samples) for metazoan zooplankton assemblages. A number of metagenetic studies of species biodiversity in marine environments based on COI sequences have now been published (e.g. Bourlat et al., 2013; Ji et al., 2013). A similar approach can be used for different genes to examine diversity at various taxonomic levels. The nuclear small-­ subunit (18S) rRNA shows consistent patterns of divergence across invertebrate and vertebrate taxa and reliably discriminates genera, families, and higher taxonomic groups (Creer et al., 2010). Different portions of this gene sequence have been the basis of high throughput sequencing analyses of metazoan diversity (Creer et al., 2010; Fonseca et al., 2010; Bik et al., 2012), including zooplankton (Lindeque et al., 2013; Pearman et al., 2014). The 18S rRNA gene has significant advantages for ­accurate classification of novel sequences and reliable amplification with consensus primers; it provides group-specific markers that complement the less reliable but more variable COI barcode region. Novel approaches to data analysis and visualization of results may soon facilitate near real-time biodiversity assessments based on metagenetic analysis (Stoeckle and Coffran, 2013). These advances will yield accurate, high-resolution, and rapid characterization of marine zooplankton species diversity at high time- and space-resolution for a variety of basic and applied uses.

Genomics Whole reference genomes are complete or nearing completion for tens of zooplankton species, although most data are not yet publicly available. These invaluable resources will allow full

analysis, interpretation, and annotation (i.e. description of the genes and assignment of function to the gene products) for genomes of ecologically important, non-model species found in pelagic ecosystems. Understanding the adaptation potential of zooplankton to climate change will require knowledge of the evolution of genes and genomes, including rates of mutation, which are controlled by regions of the genome and impacted by many factors, including population size and connectivity (Barton, 2010; Stapley et al., 2010).

Transcriptomics Gene expression is an indicator of physiological condition and response to environmental stress (Hofmann and Todgham, 2010). Analysis of differential gene expression can be used to understand the underlying molecular bases of physiological responses of organisms to environmental conditions, and thereby seek to predict consequences of environmental stress and climate change for individuals, populations, species, and communities. Gene ontology can identify the functions of differentially-expressed genes; interpretation of whole-transcriptome gene expression profiles is allowing discovery and analysis of important biochemical pathways that mediate physiological condition and response of organisms (Wang et  al.,  2009). Recent studies have examined methods for RNA extraction for transcriptomics (Zhang et al., 2013; Asai et al., 2015). A limited number of reference transcriptomes for marine zooplankton species has been published (e.g. Clark et al., 2011; Ning et al., 2013; Toullec et al., 2013; Batta-Lona, 2014; Lenz et al., 2014; Mojib et al., 2014). In addition, examination of transcriptomic (gene expression) responses to environmental stress have been done for a number of species (for review, see Dam,  2014). Publication of research papers in peer-­reviewed journals usually requires submission of the molecular data to a public repository or website. When this is not the case, the DNA and RNA sequence data, assemblies, and annotations are not available for viewing or additional analysis by others, unless obtained directly from the author(s).

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SEC T ION I I I

Taxonomy

INTRODUCTION TO TA XONOM Y Row ena Ster n, M a r i a nne Wootton, a nd Claudi a Castella ni 1.1  General Introduction Taxonomy is the scientific discipline of describing, delimiting and naming organisms. It is the foundation of biodiversity science, and taxonomic identification underpins studies of ecology, physiology, conservation, evolution and more recently environmental policy, as issues and new legislation regarding sustainable management of our oceans have come to the forefront. Morphological characters have always been the primary basis of identification, and the invention of the microscope more than 400 years ago allowed minute organisms to be seen in detail, as first described by van Leeuwenhoek and Hooke (Shannon, 2016). Since then, microscopy has been key to the progression of the biological sciences and to the identification of thousands of planktonic species. Thanks to the development of DNA-based identification, over 20 years ago, species can now additionally be identified using differences in their DNA sequences. Genetic differences can be seen as additional traits which, together with morphological traits, define a species, and this is known as systematic classification. The technology surrounding the collection and identification (both microscopical and molecular) of organisms has now advanced so much that we are able to collect, sort, photograph and identify to broad taxonomic levels, thousands of small plankton cells per minute using automated platforms while underwater, i.e. in situ. Chapters 9 and 10 of this book cover current and emerging methodologies used to collect and identify organisms. These new approaches have accelerated the rate at which we can observe and analyse planktonic organisms. As a result, recent decades have seen both an increase in the number of new marine species discovered and, rather worryingly, an increasing trend in the number of species which have disappeared or have drastically declined as a result of global climate change and human impact on marine ecosystems. Most marine phyla have one or more planktonic stages in their life cycle, usually the larval stages, which often are also the most vulnerable to changes in environmental conditions. Hence, being able to identify and recognize planktonic organisms is key to both the protection and the sustainable management of marine resources. Yet, the link between many planktonic larval stages (e.g. some Decapoda, Stomatopoda, Polychaeta and parasitic Cirripedia) and their adult stages, be

they planktonic, benthic or parasitic, remains unknown even for relatively well-studied oceanic regions of the world such as the North Atlantic. The disparity in the terminology used by different taxonomists to describe the same morphological features within and across taxa has generated a great deal of confusion for taxonomic identification in general. Moreover, additional confusion in species identification arises when one species is given multiple names or synonyms by different investigators because they were not aware it had already been described, or because the taxon name is still debated. There are many synoptic guides describing a wide variety of phytoplankton taxa over broad geographical regions of the North Atlantic in a single manual such as Algae (Graham et al., 2009), Identifying Marine Phytoplankton (Tomas, 1997), Phytoplankton of Norwegian Coastal Waters (Throndsen et al., 2007) and Coastal Phytoplankton Photo Guide for Northern European Seas (Kraberg et al., 2010) to name but a few. By contrast, there is no single guide which covers even the most common zooplankton taxa found in the North Atlantic in a single manual akin to the two volumes of the South Atlantic Zooplankton guide (Boltovskoy, 1999). Many, but by no means all, zooplanktonic taxa have been illustrated in a series of ­taxonomic publications by the International Council for the Exploration of the Sea (ICES) (Fiches d’Identification du Zooplancton / Identification Leaflets for Plankton) and even these require updating and correcting to incorporate the latest taxonomic information and classifications. No single plankton guide for the North Atlantic covers both the phytoplankton and the zooplankton in a single volume. With the current drive to produce global data sets on plankton that will underpin issues such as climate change impacts and biodiversity, this book will help to ensure that the data fed into these global data sets are accurate and up-to-date from a taxonomic perspective. The ­discipline of taxonomy and the community of taxonomists have been in a state of critical decline over the last few decades. Despite new automated identification technologies, the human eye and the humble microscope still remain relevant to the identifying microorganisms to species level. With this in mind, this book represents an important resource, imparting taxonomic expertise and supporting the accurate identification that will generate the data needed to address all issues concerning marine biodiversity.

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

140 ta xonom y

1.2  The ‘Tree of Life’

of feeding (trophic) modes including autotrophy (photosynthetic), heterotrophy (carnivorous), mixotrophy (both autotrophic and heterotrophic), saprotrophy (feeding on dead and decaying matter) or can be parasitic or symbiotic. In the last 20 years there has been a major systematic reorganization of protists based on gene phylogenies, biochemical profiling and electron microscopy imaging analysis that better reflects the complex evolutionary history of protists (Keeling and Palmer, 2008; Cavalier-Smith, 2010; Raven, 2013). There are now seven eukaryotic supergroups (Fig. 1 and Table 1) described in the latest revision of the Tree of Life of Eukaryotes (Adl et al., 2012). However, an old-fashioned and informal terminology is still commonly used to describe higher taxonomic classification in protists, as summarized in Table 1, which in many cases no longer directly corresponds to currently recognized taxa groups. However, lower classification descriptions of microbial eukaryotes remain largely unchanged. In this book, higher-level classification is based on Adl et al. (2012), but this source is not comprehensive for lower-level classification descriptions. Therefore for microbial eukaryotic classification below class level, this book uses the taxonomic databases AlgaeBase

While there are several difficulties in defining species, there are also challenges in defining higher taxa categories where character traits are no longer taxa specific, such as cell size or the possession of flagella, that are common to multiple and non-related organisms. Another difficulty is that character traits may become altered, lost or cryptic (e.g. only present at a genetic level) in a lineage during the course of evolution leading to mis-categorization. As the traits of many microscopic planktonic organisms can only be explored using electron microscopy or at a genetic level, relationships between unicellular organisms have only  recently been tested. The eukaryotic tree of life formerly contained five main kingdoms within the domain Eukarya: Animalia, Fungi, Plantae and Protozoa and Chromista (Cavalier-Smith, 1998, 2010). The ­latter two comprise an ancient and diverse group of mostly ­single-celled, microbial eukaryotes that are neither plant, ­animal nor fungi, estimated to contain 0.7–1.0 million species (Appeltans et al., 2012). These unicellular organisms vastly outnumber multicellular animals and exhibit a variety

Archaeplastida Stramenopiles

Chloroplastida Various

Glaucophyta

Diatomea

Rhodophyta

Phaeophyceae

Discoba

SAR

Excavata

Malawimonas

Metamonada

Ciliophora

Alveolata Dinoflagellata

Dictyostelia

Apicomplexa

Acantheria

Amoebozoa

Tubulinea

Nucletmycea Polycystinea Fungi

Rhizaria

Foraminifera

Opisthokonta

Rigidifilida

Cryptophyceae

Animalia

Cercozoa Haptophyta

Apusomonadida

Collodictyonidae

Choanomonada

Amorphea

Ancrymonadida

Incertae sedis Breviata

Incertae sedis

Bacteria Archaebacteria

FIGURE 1:  Tree of Eukaryotic Life according to Adl et al. (2012), showing the major supergroups of eukaryotes. Bacteria and Archaebacteria are indicated to show their relative placement compared to eukaryotes but are not described in this book. The taxa underlined and in bold type are supergroups, see Table 1. Thicker lines in the tree indicate taxa that are described in this book. SAR are a supergroup containing Stramenopiles, Alveolates and Rhizaria, while Amorphea encompass Opisthokonta, Amoebozoa and Apuzomonadida.

introduction to ta xonom y  141 TABLE 1:   Higher eukaryotic taxa groups described by Adl et al. (2012) and their corresponding older classifications based on Cavalier-Smith

(1998).

Supergroup

Superphyla

Old classification (Kingdoms)

Description

Examples in this book

Amorphea

Opisthokonta

Animalia, Fungi

One flagellum now or ancestrally

Animals

Amoebozoa

Protozoa

Produce pseudopodia, but lack flagella

 

Stramenopiles (AKA Heterokonts)

Chromista

Two unequal flagella now or ancestrally

Diatoms

Alveolata

Chromista

Possess cortical alveoli-flattened sac-like structures now or ancestrally

Dinoflagellates, Ciliates

Rhizaria

Protozoa and Chromista United only by molecular data

Foraminifera, Radiolaria

Archaeplastidia

Plantae

Chloroplasts derived from primary endosymbiosis

Chlorophytes

Excavata

Protozoa

Multiple morphological and genetic similarities

Euglenophyta

SAR

(http://algaebase.org) for phytoplankton (Guiry and Guiry, 2009) or the World Register of Marine Species (WoRMS) (for Protozoa). The reader should note that in Chapter 3, some taxa are classified as incertae sedis, which means their placement is still unknown. By comparison, the Metazoa (Animalia) remains relatively unchanged. The classification of all animals in the rest of the taxonomy section is based on the taxonomic reference database WoRMS, which still uses the five kingdom system.

1.3  Scope and Structure of the Taxonomy Chapters The taxonomic section of this book serves as a practical guide to the identification and systematics of the most common planktonic organisms found in the whole of the North Atlantic and the North Sea. Visual key maps of the major classes of ­phytoplankton (Fig. 2) and heterotrophic flagellates and protozooplankton (Fig. 3) are also provided, which use features ­visible under light microscopy. Please note that there are limits to identifying species using this method because many of the features visible with the light microscope overlap between taxa, and therefore they are not definitive traits for identification. Examples of different types of larger, multicellular planktonic animals are shown in Figure 4. This initial identification should

guide the reader to a specific chapter where the taxa of interest are dealt with. Each chapter describes a particular phylum or class, depending on the number of species considered. Each taxonomic chapter consists of: (1) a general introduction and a photographic illustration of a live representative, information on (2) the life cycle and (3) ecology, plus (4) the generalized morphology of the taxon, summarizing the key identification features used in the text and displaying them on annotated line drawings. There is also a section (5), which indicates the systematic placement of the taxon described within the tree of life, and lists the key marine representative illustrated in the chapter (usually to genus or family level). This section also includes information regarding the taxonomic authorities responsible for the classification adopted in each chapter, and recent changes which might have occurred, and it lists relevant taxonomic sources. Wherever possible, a dichotomous key is provided to facilitate further identification of the taxa using simple external morphological features to, typically, species, genus or family level depending on the complexity of the identification. In those cases where identification to family or genus is too complex, such as for example in the Copepoda and fish larvae, a table listing the genera or families is provided instead of a dichotomous key. Both the key and the table should lead the reader to the final section of each chapter where the genera or families are described and illustrated in more detail by highlighting specific representatives on identification cards. Each identification card

FIGURE 2:  Key to gross morphology of taxa commonly found in marine phytoplankton in the North Atlantic. Star indicates organisms that have acquired plastids through kleptoplastidy. Numbers indicate the sections where they are described.

FIGURE 3:  Key to gross morphology of taxa commonly found in marine microbial eukaryotic heterotrophic plankton in the North Atlantic.

144 ta xonom y Phylum Cnidaria (Page 198 and 232)

Non-Crustacean Zooplankton Phylum Ctenophora

Class Hydrozoa

Class Scyphozoa Medusa stage

Ephyra larval stage

(Page 251) (colonial) Siphonophorae

(non colonial) Medusa stage

Phylum Anellida

Phylum Mollusca (Page 538)

Class Polychaeta (Page 530) Adult worm

Phylum Phoronida (Page 577)

Larval stages of benthic polychaetes

Phylum Rotifera

Gymnosome

Phylum Brachiopoda

(Page 581)

(Page 573)

Thecosome

Phylum Chaetognatha (Page 551)

Actinotroch larva Adult Arrow-worm

Phylum Chordata Class Thaliacea (Page 584)

Sub-phylum Vertebrata (Page 607)

Class Appendicularia (Page 599) Oikopleura

Fritillaria

Fish eggs

Fish Larvae

Doliolida

Ciliated Larvae Phylum Bryozoa (Page 568) Cyphonautes larva

Phylum Echinodermata (Page 562) Echinopluteus

Ophiopluteus

Bipinnaria

Phylum Mollusca (Page 538) planktonic molluscs larvae

Auricularia

FIGURE 4:  Quick Picks. Pictorial key to multicellular zooplankton taxa illustrated in this book.

Throcophore

Veliger

introduction to ta xonom y  145 Crustacean Zooplankton Phylum Arthropoda Sub-class Copepoda (Page 267) Cyclopoida

Calanoida Eggs

Harpacticoida

Monstrilloida

nauplii

Sub-class Thecostraca (Page 390) Cirripedia nauplius

Class Branchiopoda (Page 381) Class Ostracoda (Page 410)

Facetotecta

Cyprid

nauplius

Class Malacostraca

Order Euphausiace (Page 505)

Order Decapoda (Page 420) nauplius

zoea

zoea

Megalopa

Nauplius

Calyptopis

Furcilia

Adult

Order Mysidacea (Page 471)

Order Stomatopoda (Page 465) zoea

FIGURE 4:  Continued

Order Amphipoda (Page 490)

146 ta xonom y contains information on the size and morphology of the family or genus. Key identification features are annotated on line drawings of the representative taxon. In addition there is information on the ecology and distribution of the taxa. The bathymetric and latitudinal distribution information is based on a number of sources ranging from specific textbooks to webbased sources (see section below for more details). The information provided on the morphology and length of the taxa described in this section is generally for preserved organisms (unless otherwise stated). Most plankton samples cannot be analysed immediately upon collection and therefore they need to be fixed immediately in order to prevent them from being consumed by predators present in the sample and to prevent degradation. Zooplankton are preserved in formalin, which bleaches the organisms in the long term. Protists are commonly preserved in an iodine solution, known as Lugol’s, that stains the samples dark brown depending on concentration of the fixative. Glutaraldehyde fixation better preserves the colour of specimens but can result in the loss of key features such as flagella. As most preservatives mask the true colour of plankton, colour is generally not used as an identification character in this book. Additionally, the reader should be aware that the size, i.e. total length, reported here for the zooplankton is for preserved specimens (unless otherwise stated). The zooplankton will shrink after fixation and over time with preservation.

1.4  Definition of the Geographical Habitat of the Species The vertical distribution of the species described in this book ranges from the surface to 1000 m depth (i.e. from the epipelagic

FIGURE 5: Longhurst’s biogeochemical provinces for the North

Atlantic Ocean and North Sea used to summarize the biogeography of the species described in this book. ARCT, Atlantic Arctic Province; CNRY, Canary Coastal Province; GFST, Gulf Stream Province; NADR, North Atlantic Drift Province; NECS, North East Atlantic Shelves Province; NASE, North Atlantic Subtropical Gyral Province; NASW, North Atlantic Subtropical Gyral Province; NWCS, North West Atlantic Shelves Province; SARC, Atlantic Subarctic Province.

to the mesopelagic zone) and latitudinally from the subarctic (i.e. approx. 70° N) to the subtropical regions (i.e. approx. 20° N) of the North Atlantic and the North Sea. For simplicity, and also due to the current lack of knowledge on the accurate distribution of many species, the biogeography of the planktonic organisms described in this section is based on broad oceanographic regions as defined by Longhurst (Longhurst, 1998) (see Fig. 5). Longhurst’s approach divides the oceans into biogeochemical provinces based on the role played by the physical forces that regulate the distribution and productivity of phytoplankton. Our assumption here is that such provinces also provide a framework to define ecologically meaningful habitats for plankton. Additionally, for the most common and abundant copepod species, we provide epipelagic distribution patterns based on the Continuous Plankton Recorder (CPR) survey data. Many of the species described are common to other ocean basins, but we do not give details as this is beyond the scope of this book.

1.5  Limitations of This Guide This guide focuses on eukaryotes so although bacteria, archaea and viruses vastly outnumber microbial eukaryotes and play a vital role in ocean ecology, it is beyond the scope of this book to include these groups. We refer readers to Schaechter (2009), Rosenberg et al. (2013) and Graham et al. (2009) and references therein as an introduction to these groups. The section dealing with the phytoplankton and protozooplankton is rather reduced because several excellent guides for these organisms already exist and we refer the readers to these more detailed books in the relevant chapters. Smaller heterotrophic organisms are less well characterized and are too numerous to cover here, but we have provided a summary of some common types in Figure 3 and the reader should consult Ohtsuka et al. (2015) for more detail on these organisms. Additionally we have omitted the Fungi many of which are hugely important to microbial food webs, and we refer readers to Richards et al. (2012) for more information on them and their ecological roles in marine systems. Overall, this section is focused on multicellular zooplanktonic organisms particularly crustacean zooplankton as these often dominate mesozooplankton communities and are generally the best preserved zooplankton in plankton samples. Several minor planktonic phyla such as the Cephalochordata could not be covered in this book. In any marine plankton sample, a great many unidentified objects may be found that are not planktonic organisms. These include seeds and pollen of terrestrial plants, transparent moults of crustaceans (particularly adult barnacles), fish scales, fragments of organisms (e.g. crustacean limbs, parts of adult bryozoan colonies, corals and sponges), butterfly scales and microplastic particles, including parts of fishing nets and fishing line. A large section could be written about these, but unfortunately it is beyond the scope of this book. We provide, however, some photographic illustrations as examples (see photos in colour plate section).

introduction to ta xonom y  147 R efer ences Adl, S., Simpson, A. G. B., and Lane, C. E. (2012). The Revised Classification of Protists. Journal of Eukaryotic Microbiology 59, 429–93. Appeltans, W., Ahyong Shane, T., Anderson, G., et al. (2012). The Magnitude of Global Marine Species Diversity. Current Biology 22, 2189–202. Boltovskoy, D. (1999). South Atlantic Zooplankton, Vol 1 and 2. Backhuys PublishersBy comparison the Metazoa (Animalia) has undergone little revision and remains relatively little changed, Leiden, Germany. Cavalier-Smith, T. (1998). A revised six-kingdom system of life. Biological Reviews of the Cambridge Philosophical Society 73, 203–66. Cavalier-Smith, T. (2010). Kingdoms Protozoa and Chromista and the eozoan root of the eukaryotic tree. Biol Letters 6, 342–5. Graham, L. E., Graham, J. M., and Wilcox, L. W. (2009). Algae. Pearson Benjamin Cummings, San Francisco, USA. Guiry, M. D. and Guiry, G. M. (2009). AlgaeBase. World-wide electronic publication. National University of Ireland, Galway. Keeling, P. J. and Palmer, J. M. (2008). Horizontal gene transfer in eukaryotic evolution. Nature Reviews Genetics 9, 605–18. Kraberg, A., Baumann, M., and Duerselen, C. -D. (2010). Coastal Phytoplankton Photo Guide for Northern European Seas. Verlag Dr Friedrich Pfeil Munich, Germany.

Longhurst, A.R. (1998). Ecological Geography of the Sea Academic press, San Diego, CA, USA. Ohtsuka, S., Suzaki, T., Horiguchi, T., Suzuki, N., and Not, F. (2015). Marine Protists: Diversity and Dynamics. Springer Japan, Tokyo, Japan. Raven, J. A. (2013). Cells inside cells: symbiosis and continuing phagotrophy. Current Biology 23, R530–R531. Richards, T. A., Jones, M. D. M., Leonard, G., and Bass, D. (2012). Marine fungi: their ecology and molecular diversity. Annual Review of Marine Science 4, 495–522. Rosenberg, E., DeLong, E. F., Lory, S., Stackebrandt, E., Thompson, F. (2013). The Prokaryotes: Prokaryotic Biology  and Symbiotic Associations. Berlin Heidelberg: Springer. Schaechter, M. (2009). Encyclopedia of Microbiology, 3e. Academic Press, Oxford. Shannon, R. R. Encyclopædia Britannica Online, s. v. “microscope”, accessed July 15, 2016, https://www.britannica.com/ technology/microscope. Throndsen, J., Hasle, G. R., Tangen, K. (2007). Phytoplankton of Norwegian Coastal Waters. Almatar Forlag As, Oslo, Norway. Tomas, C.R. (ed) (1997). Identifying marine phytoplankton. Academic Press, San Diego, CA, USA.

pa rt 1

Phytoplankton

PHYTOPLANKTON: DIATOMS A lex a ndr a K r a berg a nd Row ena Ster n 1 Introduction Diatoms belong to the Stramenopiles superphylum of protists and are one of the most abundant and diverse (morphologically and genetically) phytoplankton groups. They are responsible for at least 25% of global carbon dioxide fixation (Falkowski et al., 1998) and 20% of net primary production (Mann, 1999). One of the key characteristics of diatom cells is that they are enclosed in a silica ‘shell’. As a result they are well represented in the fossil record, appearing as early as the Jurassic period, with marine forms first recorded in the Cretaceous period. It is estimated that 20 000–100 000 diatom species exist worldwide in marine planktonic, benthic, brackish, freshwater and subaerial environments, depending on how a species is defined (Guiry, 2012; Mann and Vanormelingen, 2013), and genetic studies are revealing much undiscovered diversity (de Vargas et  al., 2015). Traditional morphological classification of diatoms has been used by taxonomists for over a hundred years. However, it has been noted that this does not necessarily reflect phylogenetic relationships (Medlin et  al., 1996; Kaczmarska et  al., 2000; Williams 2007).

FIGURE 6:  Arachniodiscus diatom (credit O. Larink, Plankton*net).

2  Life Cycle The vegetative cells of diatoms are diploid (in contrast to most other phytoplankton). Reproduction is for the most part asexual, by mitotic divisions (see Fig. 7), with the valves moving away from each other, and each of the two valves of the theca producing a new smaller valve to fit into that of the parent. This means that in 50% of the offspring, the diameter of individual cells will become successively smaller with each division, while the size of the offspring inheriting the parent’s hypovalve will remain unchanged. Commonly the length:width ratio also increases in this process. Before cell size reduction reaches a critical limit, they undergo sexual reproduction with the production of sperm and egg which fuse to form a specialized zygote called an auxospore. The auxospore sheds its frustrule, allowing the auxospore to expand. Inside the auxospore a large cell, termed initial cell, is then produced (for more details see Round et al., 1990). Different species also produce a range of resting stages that look very dissimilar from the vegetative stages (McQuoid and Hobson, 1995, 1996).

Vegetative cell division

Fertile cell size

Meiosis Vegetative cell

Diploid Haploid

Initial cell Fusion Auxospore

FIGURE 7:  Diatom life cycle: the dotted line separates diploid sexual

phase from the haploid vegetative stages.

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

152 ta xonom y

3  General Morphology

3.2  Centric diatoms

The diatom’s silica shell is termed the theca or frustule. The basic construction of the theca is that of a ‘Petri dish’ with a smaller valve (hypovalve) fitting into the larger valve (epivalve) (Fig. 8). An incredible diversity has evolved based on this basic design, but traditionally diatoms have been classed into the pennate and centric diatoms (Fig. 8). Cells can look different according to their orientation so they are often described in valve (end-on) or girdle (side-on) view. Diatom cell height is described as the perivalvar axis (P), the width as the apical axis (A), and its depth as the transapical axis (T) (see Fig. 8). While the pennate diatoms are essentially bilaterally symmetrical, the centric diatoms have a roughly radial symmetry, which does not mean, however, that they are always round in cross section. They can also be oval (with two poles = bipolar) or triangular (tripolar) for instance. In round centrics that are columnar in shape, the apical and transapical axes are identical (e.g. Coscinodiscus spp.). Diatoms can be unicellular (held together by processes) or colonial (attached by polysaccharide bonds) (Round et  al., 1990). The frustules of both centric and pennate diatoms have a large (and sometimes confusing) number of characteristics to separate species and genera. While some of these characters are discernible with light microscopy, scanning or transmission electron microscopy will be necessary to confirm the identity of difficult taxa. If live material is available, the morphology of the chloroplast can also be used to delineate genera (Cox, 1996).

These are radially symmetrical in valve view (Fig. 8) and are more abundant in marine pelagic systems. The arrangement of pores and the number, type and arrangement of processes on the valve face are particularly important for species identification (see Fig. 9). Processes on the valve face include: the strutted process (Fig. 9a), which is restricted to the order Thalassiosirales, the labiate process (Fig. 9b), and the occluded process, which is only found in Thalassiosira (see Fig. 9). The processes can bear long external tubes that are easily visible in light microscopy and can also aid in identification. In addition there are a variety of structures such as solid spines (e.g. Rhizosolenia, Fig. 9), horns (Odontella, Fig. 9), marginal ridges (e.g. Ditylum, Fig. 9) and pili (hair-like structures that rise from the valve pole and curve towards the central valve face in some Cymatosiraceae).

4  Ecology and Distribution Diatoms are found in almost all marine and freshwater habitats that receive enough light energy for growth. To grow they require light and nutrients (see Chapter 3), in particular silica (for their cell walls), nitrogen (for protein and DNA formation) and iron (to help fix nitrogen). As such, they have a strong seasonal growth cycle, often forming blooms particularly in spring when the water column mixes and delivers nutrients to the photic zone while the increased temperature creates a thermocline that minimizes cell sinking. In summer, diatom populations collapse due to nutrient depletion, grazing pressure from zooplankton, or parasitism. Different species have their own seasonality and nutrient preferences. Diatoms are more abundant at mid and high latitudes compared to lower latitudes, which have little seasonal changes. Many taxa are considered cosmopolitan, but some groups also exhibit very restricted modes of life, for example attached to the underside of ice sheets. Others are epiphytic, ­living on the surface of m ­ acroalgae. Species composition is also

3.1  Pennate diatoms Pennate diatoms are bilaterally symmetrical in valve view (see Fig. 8). The morphological features that have diagnostic value are the number and position of chloroplasts, the presence and shape of the raphe system or sternum (the latter in araphid diatoms), and ornamentation, such as striation ­patterns, pores, girdle bands and the presence and position of labiate processes.

A

B

Valve view

Valve view

Key to annotations

P Girdle view Radial centric diatom P

Bipolar centric diatom

T T

A

1 2 Pennate diatom

3

A Multipolar centric diatom

Capital letters with arrows indicate the direction of major axes in a diatom frustule: P: Perivalvar axis (height of the cell), T: Transapical axis (depth of the cell), A: Apical axis. Apical and transapical axes are identical in round centric, but not in bipolar/multipolar centric diatoms. 1: Striae, 2: Central nodule, 3: Raphe.

FIGURE 8:  Generalized diatom cell in girdle view showing the epi- and hypovalves’ girdle bands and the position of the mantle (A). A generalized valve face (view) of centric, bi- and multipolar centric diatoms and a pennate diatom (B) indicating the geometry of the valve structures: radial around a central point in the centric diatom and bilateral on either side of a central line. Valve views are generally more informative. Radial centrics belong to Coscinodiscophyceae while bi- and multipolar and radial Thalassiosirales belong to Mediophyceae. Pennate diatoms belong to Bacillariophyceae. Pictures have been adapted from Kraberg et al. (2010).

ph y topla nkton: di atoms  153 silicified elements. Three main classes are described here, whose main morphological difference is the valve outline. Classification is based on Medlin and Kaczmarska (2004) and AlgaeBase. After taxonomic revisions based on both morphological and phylogenetic evidence (Medlin and Kaczmarska, 2004), the centric diatoms are now split into two classes of Coscinodiscophyceae (radial centrics) and Mediophyceae (polar centrics and radial Thalassiosirales). Pennate diatoms are found within Bacillariophyceae (almost always bipolar). For more information we refer you to Kraberg et al. (2010) and Tomas (1997). For a recent combined view of diatom taxonomy and morphology see Medlin (2016). A taxonomic list of common North Atlantic diatom taxa is shown below: Box 1  Diatomea

FIGURE 9:  Types of processes in diatoms. Generalized valve face of a Thalassiosira cell showing strutted (a) and labiate (b) processes. The labiate process is also found in other genera and families of centric and pennate diatoms.Valve views of Chaetoceros and Bacteriastrum showing setae processes (c). On other diatoms, spines (g) and marginal ridges (h) can be found. Picture adapted from Kraberg et al. (2010).

fundamentally different between marine and freshwater habitats. Common marine genera include Coscinodiscus, Thalassiosira, Rhizosolenia, Proboscia. As primary producers they play an important role in aquatic food webs serving as food for larger zooplankton, such as copepods, and also the microzooplankton. Diatoms are also of increasing commercial interest for the production of biofuels as they can accumulate considerable amounts of lipids and other important hydrocarbons (Hildebrand et al., 2012). A number of diatom species are known to be toxic. These belong mainly to the genus Pseudo-nitzschia and they have been demonstrated to produce domoic acid which can cause amnesic shellfish poisoning (ASP) in humans (Lundholm et al., 1994; Turrell et al., 2008). Other harmful diatoms can cause physical damage: various Coscinodiscus species produce large amounts of mucilage clogging fish gills, while the spines of Chaetoceros may clog or cause bleeding to fish gills (Hallegraeff, 2004). The geographical distribution of species described in this section is based on sources cited in this chapter and online taxonomic databases such as AlgaeBase (http://algaebase.org) and the World Register of Marine Species (WoRMS).

5 Systematics Diatoms are classified as Stramenopiles, as some centric diatom sperm contain unequal flagella (also called cilia). A recent revision of protists by Adl et al. (2012) defines diatoms as unicellular organisms lacking in flagella (except for diatom male gamete stages) and comprised of two valves containing tightly bound

Supergroup SAR (Stramenopiles, Alveolates, Rhizaria) Superphylum Stramenopiles   Phylum Diatomea   Class Coscinodiscophyceae    Order Coscinodiscales     Family Coscinodiscaceae      Genus Coscinodiscus    Order Rhizosoleniales     Family Rhizosoleniaceae      Genus Rhizosolenia      Genus Pseudosolenia      Genus Guinardia      Genus Probosia   Class Mediophyceae    Order Triceratiales       Family Triceratiaceae      Genus Odontella,       Genus Triceratium    Order Chaetocerotales       Family Chaetocerotaceae      Genus Chaetoceros      Genus Bacteriastrum    Order Thalassiosirales       Family Thalassiosiraceae      Genus Thalassiosira      Genus Planktoniella      Genus Minidiscus      Genus Porosira   Class Bacillariophyceae    Order Bacillariales     Family Bacillariaceae      Genus Pseudo-nitzschia      Genus Bacillaria      Genus Fragilariopsis      Genus Cylindrotheca    Order Fragilariales     Family Fragilariaceae      Genus Asterionellopsis Only genera shown in bold type are covered in this chapter, but many more exist.

154 ta xonom y Family Chaetocerotaceae Genus Chaetoceros This is a large genus with roughly 200 described species. The many forms and varieties within this genus are characterized by long setae on the valve margins (Fig. 10). Cells are usually united into chains with a gap (aperture or foramen) formed between adjacent cells. Aperture shape depends on the geometry of the adjacent valve faces but is often oval to nearly round in girdle view. There are one, two or many chloroplasts per cell. Each valve (with few exceptions) carries two setae, one per valve end (pole). In some, but not all, a labiate process is present on the valve face. The point

A

B

of insertion of the setae (from exactly or slightly inside the valve margin to a central insertion point), their orientation with respect to the long chain axis (Brunel, 1972), the geometry of the valve face (concave, convex, flat) and the shape of the resulting aperture between cells are important diagnostic features. Chaetoceros chains have two setae, one at each end of the valve as in Figure 10. In contrast, Bacteriastrum, a similar species, has several setae regularly arranged around the margin, and these setae bifurcate some distance away from the valve margin.

Chaetoceros densus

T

(Cleve) Cleve, 1901 A

A

S

Size: 10–55 µm apical axis (A) diameter, 8–46 µm transapical axis (T) diameter, 12–45 µm height Distribution: NECS, NADR, NASE, NASW, NWCS, GFST, ARCT, SARC Harmful: yes, at high densities, clogging fish gills

T 30 μm

FIGURE 10:  A, Girdle view of generalized Chaetoceros. B, Valve view of C. densus. The angle of the setae (S) is an important diagnostic

character.

Family Coscinodiscaceae Genus Coscinodiscus Coscinodiscus cells have a cylindrical valve shape and with the exception of Coscinodiscus wailesii are more broad than high in girdle view, i.e. when looking at them sideways. The valve face has a marginal ring of labiate processes (rimoportulae). The larger processes are termed macrorimoportulae. The valve face has a pronounced radial areolation, i.e. pore-like surface marks, on the

valve surface. Two processes are enlarged and the distance (angle) between them is a diagnostic feature. Sometimes the areolae in the central valve face are enlarged and have a different shape. Coscinodiscus concinnus can either be drum shaped or discoidal (seen in Fig. 11A). In C. wailesii and C. concinnus there is a central, hyaline area that is unornamented.

Coscinodiscus concinnus A

3 1 2

C

3

Ehrenberg, 1844 Size: 110–500 µm diameter, 60-500µm height Distribution: colder water species. NECS, NADR, NWCS, GFST, NASW, NASE, SARC, ARCT. Blooms in winter and spring Harmful: yes, causing anoxia at high densities

1 2 4 B

D

Coscinodiscus wailesii 1 2 4

Gran and Angst, 1931 Size: 250–500µm diameter, 120–500 µm height Distribution: invasive from Pacific. Now cosmopolitan in the North Atlantic. NECS, NADR, NWCS, GFST, NASW, NASE, SARC, ARCT. Blooms in winter and autumn Harmful: yes, causing anoxia at high densities and clogs fish nets

FIGURE 11:  Girdle views of Coscinodiscus concinnus (discoid form) (A) and Coscinodiscus wailesii (B); the former has a more rounded girdle outline. C and D show valve views with radial aereolation in this genus (C), while D shows general valve features for Coscinodiscus. 1, Rimportula; 2, macrorimoportula; 3, aereola; 4, central hyaline area.

ph y topla nkton: di atoms  155 Family Triceratiaceae Genus Odontella This is a genus with a small but relatively easily recognizable number of species. Twenty species are currently accepted. Cell size varies considerably between species. Cells are bipolar in valve view, while they are roughly rectangular in broad girdle view with the four valve poles drawn out into distinct horns, with an ocellus at their distal ends. Two labiate processes are present per valve face, positioned between the process and the central valve face. Their

external tubes can be longer than the marginal horns. The valve face is almost flat in Odontella sinensis but can have different curvature patterns in other species. Cells contain numerous small, plate-like chloroplasts. The nucleus is located in the central part of the cell. Important diagnostic characteristics are the curvature of the valve face, the length and orientation of the labiate processes and the distance between the horn and labiate process.

A

Odontella sinensis Horns Labiate process P

P

N

(Greville) Grunow, 1884 Size: 80–260 µm apical axis (A) diameter, 35–120 µm transapical axis (P) diameter, 80–400 µm height Distribution: invasive from Pacific. NECS, NADR, NASE, NASW, NWCS, GFST, ARCT, SARC. Blooms spring and autumn Harmful: no

A FIGURE 12:  Odontella sinensis broad girdle view from Kraberg et al. (2010). N represents the nucleus.

Family Thalassiosiraceae Genus Thalassiosira A

Central strutted process with central thread Organic threads Marginal strutted process (MSP) Single labiate process

B

Thalassiosira gravida Cleve, 1896 (=Thalassiosira rotula (Meunier) Fryxell & Hasle, 1977) Size: 8–60 µm diameter, 5–20 µm height Distribution: cosmopolitan. NECS, NADR, NASE, NASW, NWCS, GFST. Blooms in spring and summer Harmful: no

C Marginal strutted process Central cluster of processes connecting central thread Single labiate process

FIGURE 13:  Girdle view of generalized Thalassiosira cell (A) that may have one or more central strutted processes. Panels B and C show Thalassiosira gravida in valve and girdle view respectively. The central strutted processes (B) producing a bundle of central threads connecting adjacent cells in a chain (C). Picture adapted from Tomas (1997) and Kraberg et al. (2010). (continued)

156 ta xonom y Family Thalassiosiraceae Genus Thalassiosira (Continued) Thalassiosira comprises more than 100 species. Thalassiosira species usually have small plate-like chloroplasts. Their frustules are round in valve view and cylindrical with straight or convex valve face in girdle view. Thalassiosira species have three types of processes. Two of these – the labiate process (also found in Coscinodiscus species) and the strutted process penetrate the theca and have an opening to the interior of the valve. The differences are most evident when viewing the inside of the valve, where strutted processes carry so-called satellite pores, while the labiate process ends in an interior lip-like structure. The strutted processes are arranged in one or more marginal rings on the valve face, with an additional one or more central strutted processes. The central strutted

processes unite cells into chains of varying lengths. One labiate process, or in some cases two labiate processes, is usually located in the marginal ring of strutted processes. The exact location and number of the different processes is used for species identification. A third type of process, the occluded process, which does not have an opening to the interior of the valve, is found exclusively in the Thalassiosiraceae and is very conspicuous in Thalassiosira punctigera. Thalassiosira is a common genus and often an important component of spring diatom blooms (e.g.  Thalassiosira nordenskioeldii ). Thalassiosira gravida has been shown to be a taxonomic synonym of Thalassiosira rotula (Sar et al., 2011).

Family Rhizosoleniaceae Genus Rhizosolenia Contiguous area Clasper

Otarium

Girdle segment

Rhizosolenia imbricata Brightwell, 1858

Process

Size: 15–80 µm diameter, 10–40 µm height Distribution: NECS, NADR, NASE, NASW, NWCS, GFST, ARCT, SARC. Blooms in spring and summer Harmful: no

FIGURE 14:  Generalized Rhizosolenia cell, showing the valve with diagnostic features labelled.

A

B

C

D

E

F

G

H

Rhizosolenia species are cylindrical and much higher than wide (i.e. a very long pervalvar axis). Cells contain many plate-like chloroplasts often arranged into several rows. Valve ends are unipolar, asymmetric cones, ending in a spine, often with small wings (otaria) at their base (exception: Rhizosolenia setigera), as shown in Figure 14. On one side of the cone, a depression (contiguous area) is located to accommodate the valve end of the adjacent cell in a chain. Internally, the base of the spine can have a labiate structure. Cells can form chains that can be long, for example in Rhizosolenia imbricata, but others, for example R. setigera, are more commonly found as individual cells or in pairs (see Fig. 15 for a comparison of different species). The girdle bands in Rhizosolenia are very distinct and discernible with the light microscope. The morphology of the spine, and the presence and shape of otaria are all diagnostic features. They are not always easy to see in light microscopy.

FIGURE 15:  Valve views of common North Atlantic species of Rhizosoleniaceae: A, Rhizosolenia pungens; B, Rhizosolenia setigera;

C,  Rhizosolenia imbricata; D, Rhizosolenia styliformis; E, Rhizosolenia hebitata f. semispina cone detail; F, Rhizosolenia bergonii with cone detail. For comparison, similar genera: G, Pseudosolenia calcar-avis; H, Proboscia alata. Picture adapted from Tomas (1997).

ph y topla nkton: di atoms  157

Pennate diatoms Family Bacillariaceae Genus Pseudo-nitzschia of striae and fibulae bands across the valve (see Fig. 16) are important characters for species identification but are rarely discernible in the light microscope (Skov et al., 1999). Up to four groups of species, called ‘morpho-species’ can be identified by light microscopy, based on cell and valve dimensions; however, species-level identification requires genetic or electron microscopy. For more details on identification we recommend Hallegraeff (2004) and Tomas (1997).

A small but important genus of pennate diatoms consisting of 44 species and containing toxic species to varying degrees. Species in the genus form stepped chains by overlapping valve ends of adjacent cells. Cells are small and elongate with pointed or rounded ends. Valve surface is structured by striae with rows of poroids, with interstriae separating the striae. The raphe system (so-called canal raphe) is strongly eccentric running along one valve margin. Cells contain two plate-like chloroplasts. The number

A T Interstriae Fibulae FIGURE 16:  Pseudo-nitzschia pungens valve view showing internal characteristics used to identify species. Striae and fibulae can sometimes be visible by light microscopy but poroids are only visible by electron microscopy. Very similar to Pseudo-nitzschia multiseries and Pseudonitzschia seriata; this species can only be distinguished by its heavier silicified valves by higher magnification using oil immersion. Otherwise it should be identified within the ‘Pseudo-nitzschia seriata complex’ along with the aforementioned species.

Central interspace

Pseudo-nitzschia pungens (Grunow ex Cleve) Hasle, 1965

Poroids

Size: 75–140 µm apical axis (A), 3.4.5 µm transapical axis (T), 5–8 µm height Distribution: NECS, NADR, NASE, NASW, NWCS, GFST. Blooms spring, summer Harmful: yes

Interstriae striae

Family Fragilariaceae Genus Asterionellopsis This genus belongs to Fragilariaceae, which contains 36 genera and 324 species; many of the genera consist of only a few species. Asterionellopsis only contains two species: Asterionellopsis glacialis (Fig. 17) and Asterionellopsis socialis. Valves are heteropolar with a  broadened foot-pole and a needle-like head pole. Cells are

commonly found united into star-shaped or zig-zag colonies. Adjacent cells are attached to each other by their valve faces, i.e. cells in a colony are usually seen in girdle view. In this view, the foot-pole is triangular in outline, as opposed to an oval outline in valve view. Fixation can separate the chain into single cells.

Asterionellopsis glacialis (F. Castracane) F.E. Round, 1990 Size: 30–150 µm apical axis, 7–18 µm basal transapical axis width, 5–15 µm basal transapical axis height Distribution: NECS, NWCS, GFST, ARCT. Blooms spring and summer Harmful: no

FIGURE 17:  Asterionellopsis glacialis, girdle view.

158 ta xonom y

References Adl, S., Simpson, A. G. B., Lane, C. E., et al. (2012). The Revised Classification of Protists. Journal Eukaryotic Microbiolgy 59(5), 429–93. Brunel, J. (1972). Orientation of setae in the genus Chaetoceros, in regard to the apical axis. Journal of the Marine Biological Association of India 14, 315–27. Cox, E.J. (1996). Identification of freshwaterdiatoms from live material. Chapman and Hall. de Vargas, C., Audic, S., Henry, N., et  al. (2015). Eukaryotic plankton diversity in the sunlit ocean. Science 348(6237), doi:10.1126/science.1261605. Falkowski, P. G., Barber, R. T., and Smetacek, V. (1998). biogeochemical controls and feedbacks on ocean primary production. Science 281, 200–6. Guiry, M. D. (2012). How many species of algae are there? Journal of Phycology 48(5), 1057–63. Hallegraeff, G.M. (2004). Manual on harmful marine microalgae, pp. 25–49. UNESCO, Paris. Hildebrand, M., Davis, K., A., Smith, S. R., Traller, J.C., and Abbriano, R. (2012). The place of diatoms in the biofuels industry. Biofuels 3(2), 221–40. Kaczmarska, I., Ehrman, J. M., and Bates, S. S. (2000). A review of auxospore structure, ontogeny and diatom phylogeny. Proceedings of the 16th International Diatom Symposium, pp. 153–69. Kraberg, A., Baumann, M., and Duerselen, C. -D. (2010). Coastal phytoplankton photo guide for Northern European seas. Verlag Dr Friedrich Pfeil Munich, Germany. Lundholm, N., Skov, J., Pocklington, R., and Moestrup, Ø. (1994). Domoic acid, the toxic aminoacid responsible for amnesic shellfish poisoning, now in Pseudonitzschia seriata (Bacillariophyceae) in Europe. Phycologia 33(6), 475–8. Mann, D. G. (1999). The species concept in diatoms. Phycologia 38(6), 437–95. Mann, D. G., and Vanormelingen, P. (2013). An inordinate fondness? The number, distributions, and origins of diatom species. Journal of Eukaryotic Microbiology 60, 414–20.

McQuoid, M. R., and Hobson, L. A. (1995). Importance of resting stages in diatom seasonal succession. Journal of Phycology 31, 44–50. McQuoid, M. R., Hobson, L. A. (1996). Diatom resting stages. Journal of Phycology 32, 889–902. Medlin L. K. (2016). Evolution of the diatoms: major steps in their evolution and a review of the supporting molecular and morphological evidence. Phycologia 55(1), 79–103. Medlin, L. K., and Kaczmarska, I. (2004). Evolution of the diatoms: V. Morphological and cytological support for the major clades and a taxonomic revision. Phycologia 43(3), 245–70. Medlin, L. K., Kooistra, H. C. F., Gersonde, R., and Wellbrock, U. (1996). Evolution of the diatoms (Bacillariophyta). II. Nuclear-encoded small subunit rRNA sequence comparisons confirm a paraphyletic origin for centric diatoms. Molecular Biology and Evolution 13(1), 67–75. Round, F. E., Crawford, R. M., and Mann, D. G. (1990). Diatoms: Biology and morphology of the genera. Cambridge University Press, Cambridge, UK. Sar, E. A, Sunesen, I., Lavigne, A. S., and Soledad L. (2011). Thalassiosira rotula, a heterotypic synonym of Thalassiosira gravida: morphological evidence. Diatom Research 26(1), 109–19. Skov, J., Lundholm, N., Moestrup, Ø., and Larsen, J. (1999). ICES identification leaflets for plankton, leaflet No. 185, 4. The diatom genus Pseudo-nitzschia, pp. 23. Tomas, C. R. (ed.) (1997). Identifying marine phytoplankton. Academic Press, San Diego, CA, USA. Turrell, E., Bresnan, E., Collins, C., et  al. (2008). Detection of Pseudo-nitzschia (Bacillariophyceae) species and amnesic shellfish toxins in Scottish coastal waters using oligonucleotide probes and the Jellet Rapid Test (TM). Harmful Algae 7(4), 443–58. Williams, D. M. (2007). Classification and diatom systematics: the past, the present and the future, In: Brodie, J., Lewis, J. (eds.), Unravelling the algae: the past, present, and future of algal systematics. CRC Press, Taylor and Francis group, Boca Raton, pp. 57–91.

PHYTOPLANKTON: DINOFLAGELLATES A lex a ndr a K r a berg a nd Row ena Ster n 1 Introduction Dinoflagellates occur in marine, brackish and freshwater habitats, although they are most diverse in the marine environment (Taylor et al., 2008a). About 90% of dinoflagellate species are planktonic and are responsible for a large proportion of primary productivity, but benthic species have also been described. Dinoflagellates are protists in the size range 5 µm to approximately 2000 µm diameter. About 2000 species have been described, and another 2500 fossil species have been recorded. However, genetic studies have shown many more species exist, especially parasitic dinoflagellates (Guillou et al., 2008; Kim et al., 2008; de Vargas et al., 2015). Due to their evolutionary history and ability to acquire genes horizontally from unrelated organisms, dinoflagellates are a functionally diverse and ecologically important group (Keeling, 2004). Photosynthetic dinoflagellate morphospecies such as Prorocentrum minimum and Protoperidinium pallidum tend to be cosmopolitan in temperate regions of the northern and southern hemisphere, while tropical regions tend to have endemic species such as Ornithocercus spp. or Pyrodinium bahamense, although true endemicism is rare (Taylor et al., 2008b). Several photosynthetic dinoflagellates are harmful, forming toxic blooms, with detrimental consequences to human and animal health and to the economy. Parasitic dinoflagellates often infect other dinoflagellate species. A variety of symbiotic dinoflagellates, such as Symbiodinium sp., form relationships with corals, sponges and foraminifera.

2  General Description Dinoflagellates are unicellular but can form chains and are usually split on the basis of morphology into two major groups (although as in diatoms these are not phylogenetically consistent) (Daugbjerg et al., 2000): the thecate and athecate dinoflagellates. The thecate dinoflagellates are covered in a series of cellulose plates, which form the theca. The number and arrangement of the thecal plates (also known as tabulation) as well as structures on the plates, such as spines, are a further diagnostic character for the identification of dinoflagellates. Athecate dinoflagellates lack these plates. Dinoflagellates are characterized by the presence of two flagella (with a different morphology). In the majority of dinoflagellate groups, the flagella are inserted in two grooves: the

FIGURE 18:  The dinoflagellate Protoperidinium subinerme (Morozova, T., Source: http://planktonnet.awi.de).

transverse groove (cingulum) and the longitudinal groove or sulcus (Fig. 9). The side of the cell on which the sulcus is located is defined as the ventral side of the cell. The one ­exception to this rule is the flagellar arrangement in the Prorocentraceae. In this group the flagella are inserted in a small area (the so-called periflagellar area) at the cell apex (Taylor, 1987). A further character typical of dinoflagellates is their nucleus, the dinokaryon. The dinokaryon is usually very large and the chromosomes are permanently condensed, i.e. visible as discrete strands. The nucleus is large and visible by light microscopy. This section reviews only a few common North Atlantic genera. For more information, we refer you to Steidinger and Tangen (1997), Kraberg et al. (2010) and Hoppenrath et al. (2009).

2.1  Thecate dinoflagellates The plate tabulation in dinoflagellates is a diagnostic feature. Although several systems exist, the most common one divides the plates into six series: two on the epitheca (apical and ­precingular series) and two on the hypotheca (postcingular and antapical). All of these series are bordering either the cingulum or the apex/antapex of the cell (Fig. 19). Plates not doing so are called intercalary plates. The last two series are the plates in the

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

160 ta xonom y A

B Partially inherited parental theca

Schizont gamete Schizont fusing gametes

Sexual

Asexual

Parental theca

Pellicle cyst (temporary)

Pellicle cyst (temporary)

. . ... .... ..

Planozygote

. . . . ... .. . .

Excystment

. . . ... .. .. . Hypnozygote

Resting cyst FIGURE 19:  Generalized dinoflagellate life cycle – the main picture (B) denotes the sexual cycle; most planktonic growth occurs asexually (A). Note cyst stages may not be planktonic, many sinking to sediments. The schizonts are haploid gamete stages without a theca cell wall in order to fuse.

sulcus and cingulum. Some examples of thecate dinoflagellate genera follow, as well as an indication of tabulation patterns.

2.2  Athecate dinoflagellates Also called unarmoured or naked dinoflagellates. These often have plates that are very thin and delicate and cannot be seen by light microscopy. Identification is based on living specimens, as preservatives tend to distort or destroy them. Cell size, shape (e.g. flattened, lobed), position of cingulum, position of sulcus, number and position of chloroplasts, and cell movement are used to distinguish between species.

This can then start dividing again or produce a resting cyst – the hypnozygote. While the planozygote has a morphology that is very similar to the haploid vegetative stage, the hypnozygote is morphologically dissimilar and many resting stages of known dinoflagellate species have been assigned different names as they were not recognized as a life-cycle stage but a separate taxon.

4  How to Identify Dinoflagellates Apex

Epitheca

3  Life Cycle Dinoflagellates have haplontic life cycles with cell division mostly occurring asexually by binary fission with each of the daughter cells inheriting a portion of the parental theca or valve. In other groups, for example the genus Protoperidinium, two daughter cells are produced within the parent theca, and each daughter cell produces an entirely new theca (Fig. 19A). Sexual reproduction has also been observed in many species (see Fig. 19B). Here two cells fuse to form a planozygote.

Epicone

Apex

Cingulum

Antapex

Hypotheca

Thecate dinoflagellate

Hypocone

Sulcus

Antapex Athecate dinoflagellate

FIGURE 20:  General overview of dinoflagellate terminology. Figure from Kraberg et al. (2010). The schematic is showing the ventral view side of the cell (on which the sulcus is visible).

ph y topla nkton: dinoflagellates  161 *

1

2a

2b

3

4

5

6

FIGURE 21:  Plate tabulation patterns in a generalized thecate dinoflagellate: 1. Apical series, 2a. Precingular series (dorsal view side of cell),

2b. precingular series (ventral view side of cell), 3. postcingular series, 4. antapical series, 5. cingular series, 6. sulcal series ; * = intercalary plate. Figure from Kraberg et al. (2010).

5  Ecology and Distribution Dinflagellates can occupy stratified waters, and several species, particularly in the genus Tripos, have also been observed to  show vertical migration. Most larger dinoflagellates prefer coastal, nutrient-rich waters, but smaller dinoflagellates are found in open waters (Cullen et al., 2002). Many dinoflagellate parasites have been found in open waters (Guillou et al., 2008). The toxic Dinophysis species grows offshore and is advected by wind to coastal locations. An increasing number of the species previously considered autotrophic are now discovered to be capable of ingesting prey (mixotrophy). These mixotrophic and also heterotrophic species exhibit a considerable diversity of feeding mechanisms, including direct engulfment, pallium feeding and feeding by means of a feeding tube which penetrates the prey cell. The feeding rates on diatoms are considerable, even exceeding those of mesozooplankton such as copepods ( Jeong et al., 2004). Dinophysis species are able to acquire chloroplasts from other species temporarily in order to photosynthesize and have an important relationship with ciliates and other flagellates such as cryptophytes. Dinoflagellate parasites such as the marine syndiniales group (class Syndinea) (Park et  al., 2004; Kim et al., 2008) tend to be very small and infect other dinoflagellates (Gunderson et  al., 2002), copepods (Skovgaard et  al., 2005) and commercial crab (Stentiford and Shields, 2005) and play an important role in carbon recycling and shaping the diversity of their hosts. They can represent up to 50% of the picoand nanoplankton (< 10 µm) community (de Vargas et  al., 2015). They therefore play a very important but often still underestimated role in marine food webs.

6  Harmful Species A considerable number of dinoflagellate taxa produce potent toxins that can cause disease in humans via consumption of contaminated shellfish or fish (ciguatera poisons). Common toxins are okadaic acid produced by species of the genus

Dinophysis, which causes diarrhetic shellfish poisoning (DSP), and saxitoxin (e.g. in species of the genus Alexandrium). The latter toxin is the causative agent of paralytic shellfish poisoning (PSP). Further examples are brevetoxin produced by the dinoflagellate Karenia brevis, and the ichthyotoxins produced by Gambierdiscus species. However, toxicity is not the only way in which dinoflagellates can cause harmful effects on ecosystems and local economies. Mass occurrences of dinoflagellate species can also regularly cause problems. The large dinoflagellate Noctiluca forms surface blooms causing significant discoloration of the water. Mass blooms of this species can also result in de-oxygenation in the water. Mass blooms are also formed by Akashiwo sanguinea (brown tides), Lepidodinium chlorophorum (green tides) and also Karenia species (red tides). For more information on harmful dinoflagellate species we refer the reader to Hallegraeff (2004).

7 Systematics Dinophyceae belong to Alveolata superphylum and are broadly related to the Stramenopiles, Alveolates, Rhizaria (SAR) group (see Fig. 1 of the Introduction to taxonomy). Dinoflagellate taxonomy has undergone some recent revisions. Basal dinoflagellates such as Oxyrrhis sp. and dinoflagellate parasite taxa such as Perkinsus sp. and the class Syndinea are classified under Protalveolata. The taxa described here belong to Dinoflagellata. Prorocentrum species are considered to be basal taxa. Thecate dinoflagellates tend to have more robust classification, but athecate taxa have more than one common ancestor and are likely to be taxonomically revised. Here we use higher level classification according to Adl et  al. (2012) and AlgaeBase (http://algaebase.org) (Guiry and Guiry, 2009) for family and lower level classifications. Commonly occurring dinoflagellate taxa are shown here to family level. Genera in bold type are described in this section.

162 ta xonom y BOX 1  Classification of Dinoflagellata

  Family Gonyaulacaceae   Genus Gonyaulax      Alexandrium   Family Gymnodiniaceae   Genus Akashiwo      Gyrodinium   Family Kareniaceae   Genus Karenia   Family Polykrikaceae   Genus Polykrikos   Family Prorocentraceae   Genus Prorocentrum   Family Amphidomataceae   Genus Azadinium   Family Ooidiniaceae   Genus Pfiesteria

Supergroup SAR (Stramenopiles, Alveolates, Rhizaria) Superphylum Alveolata Phylum Dinoflagellata Class Dinophyceae   Family Peridiniaceae   Genus Scrippsiella   Family Heterocapsaceae   Genus Heterocapsa   Family Protoperidiniaceae   Genus Protoperidinium   Family Ceratiaceae   Genus Tripos   Family Dinophysiaceae   Genus Dinophysis      Phalacroma

Thecate dinoflagellates Family Protoperidiniaceae Genus Protoperidinium This is a large genus of thecate dinoflagellates comprising more than 300 valid species. Protoperidinium species are heterotrophic, and many are known to feed by way of a pallium. The size range within the genus is approximately 15 µm to 250 µm. Cell outline and symmetry are variables. In different species the cells can be dorso-ventrally compressed or compressed in an apical-antapical direction. The cingulum is usually placed in a median position

with only slight displacement either clockwise or anticlockwise but, for example, in some species the cingulum is strongly inclined with the portion on the dorsal side of the cell strongly displaced towards the cell apex. The sulcus is slightly excavated. Usually, apical and antapical horns (antapical spines) are present. The thecal plates are often reticulate or otherwise ornamented (Dodge, 1983).

Protoperidinium claudicans (Paulsen) Balech, 1974 Size: 50–105 µm length, 48–75 µm diameter Distribution: NECS, NADR, NASE, NASW, GFST, NWCS Harmful: no Trophism: heterotrophic

Ventral FIGURE 22:  Protoperidinium claudicans ventral and dorsal views aspects. Figure adapted from Tomas (1997).

ph y topla nkton: dinoflagellates  163 Family Ceratiaceae Genus Tripos This is a species-rich genus with more than 100 described species formerly called Ceratium and mostly now called Tripos. All known species contain chloroplasts, but an increasing number of species are discovered to be mixotrophic, i.e. they can actively consume prey, for instance by direct engulfment. Cells are large, with very pronounced apical and antapical horns. The curvature and alignment of these horns with the main cell axis (and relative to each other) are a diagnostic feature that is used in the identification of different species. Cells are slightly dorso-ventrally compressed. The new name Neoceratium was recently proposed to replace the name Ceratium. However, this name is now deemed invalid and the genus has been renamed again: the currently accepted name for the genus is Tripos.

FIGURE 23:  Tripos furca, ventral view aspect. Figure from Kraberg

et al. (2010).

Apex

Tripos furca (Ehrenberg) F. Gómez, 2013 Size: 150–230 µm length, including horns, 30–35 µm diameter Distribution: NECS, NADR, NASE, NASW, GFST, NWCS, EACB, ARCT, SARC Harmful: no Trophism: mixotrophic

Antapex

Family Dinophysiaceae Genus Dinophysis This is a genus of thecate dinoflagellates showing extreme lateral compression, so that the sulcus is rarely visible as cells are always seen in lateral view. The cingulum is located near the apical margin, i.e. the epitheca is much smaller than the hypotheca. Both cingulum and sulcus are bordered by pronounced lists giving the cells their characteristic appearance. In some species, antapical horns or small protuberances (e.g. in Dinophysis acuminata) are also present. Several species are mixotrophic ( Jacobsen and Anderson, 1994). Many species of Dinophysis are known producers of okadaic acid, the cause of DSP (Larsen and Moestrup, 1992).

Cingular list

Dinophysis acuta Epitheca Right sulcal list Left sulcal list with ribs (R1-R3) Hypotheca

Ehrenberg, 1839 Size: 54–94 µm length, 43–60 µm diameter Distribution: NECS, NADR, NASE, NASW, GFST, NWCS, EACB, ARCT, SARC Harmful: yes Trophism: mixotrophic

FIGURE 24:  Dinophysis acuta with features. The shape of the hypotheca and the angle of the ribs are diagnostic features. Figure adapted from Kraberg et al. (2010).

164 ta xonom y Family Gonyaulacaceae Genus Gonyaulax of the cingulum on the ventral side is also obvious in most species with the right cingular margin overhanging the left cingular margin. As a result, the sulcus is also not straight but twisted posteriorly. Species in the genus Gonyaulax are photosynthetic.

This is another large genus of thecate dinoflagellates with more than 100 described species. Apical horns are usually well developed. The antapex exhibits pronounced spines in many species. The cingulum shows strong displacement, and overlap of the two ends

Gonyaulax spinifera (Claparède & Lachmann) Diesing, 1866 Size: 30–45 µm length, 30–40 µm diameter Distribution: NECS, NWCS, NASW, NASE, NADR Harmful: no Trophism: mixotrophic

FIGURE 25:  Gonyaulax spinifera ventral view. This species has distinct pore ornamentation on its surface theca. A key distinguishing feature

of this family is the displaced cingulum. Figure adapted from Throndsen et al. (2007).

Family Gonyaulacaceae Genus Alexandrium important diagnostic characters as is the morphology of the sulcal plates. Many species have been confirmed to produce saxitoxin, the causative agent of PSP. Many described species form complexes of almost indistinguishable species; for example, the Alexandrium tamarense complex comprises species such as A. tamarense itself, Alexandrium fundyense and Alexandrium catenella. This species complex has now been revised formally ( John et al., 2014).

This is a small genus with about 30 species. Cell size ranges from 20 µm to 80 µm. Cells are usually rounded. The cingulum is in a median position and only weakly displaced. Thecal plates lack any obvious ornamentation and are difficult to resolve in light microscopy. No spines or apical/antapical horns are present. The first apical plate is distinct, elongate with a pronounced pore. The exact morphology of the plate and location of the pore are

A

B 6’

6th precingular

C

Alexandrium tamarense (Lebour) Balech, 1995

1’ sa

1st apical

Size: 17–44 µm length, 21–51µm diameter Distribution: NECS, NWCS, NASE. Coastal Harmful: yes Trophism: autotrophic

Anterior sulcal

FIGURE 26:  Alexandrium tamarense. Each panel (A–C) shows the diagnostic plates (grey shading) and the plate name (named under each

panel) used for species identification. Figure from Kraberg et al. (2010).

ph y topla nkton: dinoflagellates  165

Athecate Dinoflagellates Family Gymnodiniaceae Genera Akashiwo and Gyrodinium Two genera, Gymnodinium and Gyrodinium, were originally dis­ tinguished on the basis of the degree of displacement of the cingulum. In the heterotrophic genus Gyrodinium, cell surfaces are usually striated. The cingulum is often displaced by several times

its width , while in Gymnodinium it was not., But this is no longer considered a diagnostic character. The family has now been re-examined (Daugbjerg et al. 2000) and many more genera have been described.

Akashiwo sanguinea (Hirasaka) Hansen & Moestrup, 2000 Size: 40–80 µm length Distribution: NECS, NWCS, NASE. Coastal, estuarine Harmful: yes Trophism: autotrophic

FIGURE 27:  Akashiwo sanguinea lateral and ventral views aspects. Figure from Kraberg et al. (2010).

Lateral

Ventral

Gyrodinium instriatum (Lebour) Balech, 1995 Size: 17–44 µm length, 21–51 µm diameter Distribution: NECS, NWCS, NASE Harmful: yes Trophism: autotrophic Lateral FIGURE 28:  Gyrodinium instriatum lateral and ventral views aspects. Figure from Kraberg et al. (2010).

Ventral

Note: currently accepted name is Levanderina fissa (Levander) Ø. Moestrup, P. Hakanen, G. Hansen, N. Daugbjerg & M. Ellegaard, 2014

References Adl, S., Simpson, A. G. B., Lane, C. E., et al. (2012). The Revised Classification of Protists. Journal of Eukaryotic Microbiology 59(5), 429–93. Cullen, J. J., Franks, P. J. S., Karl, D. M., and Longhurst, A. (2002). Physical influences on marine ecosystem dynamics, In: Robinson, A. R., McCarthy, J. J., Rothschild, B. J. (eds), The Sea, pp. 297–336. John Wiley and Sons, New York. Daugbjerg, N., Hansen, G., Larsen, J., and Moestrup, Ø. (2000). Phylogeny of the major genera of dinoflagellates based on  ultrastructure and partial LSU rDNA sequence data, including the erection of three new genera of unarmoured dinoflagellates. Phycologia 39(4), 302–17. de Vargas, C., Audic, S., Henry, N., et  al. (2015). Eukaryotic plankton diversity in the sunlit ocean. Science 348(6237), doi:10.1126/science.1261605.

Dodge, J. D. (1983). Ornamentation of thecal plates in Protoperidinium (Dinophyceae) as seen by scanning electron microscopy. Journal of Plankton Research 5(2), 119–27. Guillou, L., Viprey, M., Chambouvet, A., et al. (2008). Widespread occurrence and genetic diversity of marine parasitoids belonging to Syndiniales (Alveolata). Environmental Microbiology 10(12), 3349–65. Guiry, M. D. and Guiry, G. M. (2009). AlgaeBase. World-wide electronic publication. National University of Ireland, Galway. Gunderson, J., John, S. A., Boman, WC, and Coats, D. W. (2002). Multiple strains of the parasitic dinoflagellate Amoebophrya exist in Chesapeake Bay. Journal of Eukaryotic Microbiology 49(6), 469–74. Hallegraeff, G. M. (2004). Manual on harmful marine microalgae. UNESCO, Paris.

166 ta xonom y Hoppenrath, M., Elbrachter, M., and Drebes, G. (2009). Marine Phytoplankton: selected microphytoplankton species from the North Sea around Helgoland and Sylt. E. Schweizerbart’sche Verlagsbuchhandlung, Stuttgart, Germany. Jacobsen, D.M. and Anderson, R.A. (1994). The discovery of  mixotrophy in photosynthetic species of Dinophysis (Dinophyceae): light and electron microscopical observations of food vacuoles in Dinophysis acuminata, D. norvegica and two heterotrophic dinophysoid dinoflagellates. Phycologia 33, 97–110. Jeong, H.J., Yoo, Y.D., Kim, S.T., and Kang, N.S. (2004). Feeding by the heterotrophic dinoflagellate Protoperidinium bipes on the diatom Skeletonema costatum. Aquatic Microbial Ecology 36, 171–9. John, U., Wayne Litaker, R., Montresor, M., et al. (2014). Formal revision of the Alexandrium tamarense species complex (Dinophyceae) taxonomy: The introduction of five species with emphasis on molecular-based (rDNA) classification, Protist 165, 779–804. Keeling, P.J. (2004). Diversity and evolutionary history of plastids and their hosts. American Journal of Botany 91, 1481–93. Kim, S., Park, M. G., Kim, K. Y., et al. (2008). Genetic diversity of parasitic dinoflagellates in the genus amoebophrya and its relationship to parasite biology and biogeography. Journal of Eukaryotic Microbiology 55(1), 1–8. Kraberg, A., Baumann, M., and Duerselen, C. -D. (2010). Coastal Phytoplankton Photo guide for Northern European Seas. Verlag Dr Friedrich Pfeil Munich, Germany. Larsen, J. and Moestrup, Ø. (1992). Potentially toxic phytoplankton. 2. Genus Dinophysis, ICES Identification

leaflet for plankton International Council for the exploration of the sea, Copenhagen, p. 12. Park, M. G., Yih, W., and Coats, D. W. (2004). Parasites and phytoplankton, with special emphasis on dinoflagellate infections. J Eukaryot Microbiol 51(2), 145–55. Skovgaard, A., Massana, R., Balagué, V., and Saiz, E. (2005). Phylogenetic Position of the Copepod-Infesting Parasite Syndinium turbo (Dinoflagellata, Syndinea). Protist 156(4), 413–23. Steidinger, K. A. and Tangen, K. (1997). Dinoflagellates, In: Tomas, C.R. (Ed.), Identifying marine phytoplankton. Academic Press, San Diego, CA, USA pp. 387–584. Stentiford, G. D. and Shields, J. D. (2005). A review of the parasitic dinoflagellates Hematodinium species and Hematodiniumlike infections in marine crustaceans. Diseases in Aquatic Organisms 66(1), 47–70. Taylor, F. J. R. (1987). The biology of dinoflagellates. Blackwell Scientific Publications. Taylor, F. J. R., Hoppenrath, M., Saldarriaga, J. F. (2008a). Dinoflagellate diversity and distribution. Biodiversity and conservation 17(2), 407–18. Taylor, F. J. R., Hoppenrath, M., Saldarriaga, J. F. (2008b). Dinoflagellate diversity and distribution. Biodivers. Conserv. 17, 407–18. Throndsen, J., Hasle, G. R., and Tangen, K. (2007). Dinoflagellates-Dinophyta. In: Throndsen, J., Hasle, G. R., Tangen, K. (Ed.), Phytoplankton of Norwegian coastal waters, pp. 41–110. Oslo, Norway, Almater Forlag As.

PHYTOPLANKTON: FLAGELLATES Row ena Ster n, Heather Esson, a nd Cecili a Ba lestr er i 1 Introduction Marine flagellates is an all-inclusive term to describe a plethora of different protist species scattered throughout different eukaryotic lineages that move using their flagella. Many flagellates also have other morphological life stages, but this chapter primarily addresses those taxa for which the flagellated form is predominant. This heterogeneous group is a numerous and important part of the marine ecosystem that can be heterotrophic (feed on other prey), phototrophic (photosynthetic) or mixotrophic (cells that can photosynthesize and engulf prey). Many phototrophic taxa also have non-photosynthetic relatives that have heterotrophic or parasitic lifestyles (see Figs 2 and 3 of Introduction), some of which are covered here. However, this section deals mostly with phototrophic flagellates. Dinoflagellates (Dinophyceae) are also considered flagellates but are covered in Chapter 2. There is comparatively less information on phototrophic flagellates compared to larger phytoplankton, with the exception of calcifying haptophytes, because their small size conceals distinguishing features with light microscopy. Furthermore, most studies on marine phytoplankton work on preserved material, which destroys many flagellates that lack a cell wall which will affect estimates of their abundance. As such, estimates of diversity and/or abundance for this group are likely to be underestimated, especially as several taxa have only been discovered from unique DNA signatures and have little or no morphological records. In this chapter, we show examples of common phototrophic genera found in the North Atlantic, but this is not exhaustive. We refer readers to Tomas (1997) and Throndsen (1997) for detailed species. It is beyond the scope of this book to deal with many of the heterotrophic flagellates, as outlined in Figure 4 of the Introduction. We refer readers to Ohtsuka and Suzaki (2015) for more detailed information.

2  Life Cycle There is a variety of life cycles of flagellates, which are too numerous to recount here. Most flagellates undergo vegetative reproduction, but some cells have sexual stages. In many cases, the life cycle is unknown. The vegetative stages are generally the

F. Jouenne FIGURE 29:  Coccolithophore and Meringosphaera flagellates. Source: Left panel L. Chakravarti and C. Balestreri (2013) (personal communication). Right panel F. Jouenne (2005).

most common. Chlorophytes have a life cycle that includes flagellate fusion (e.g. Chlamydomonas) or alternating unicellular flagellate and multicellular palmelloid stages (e.g. Halosphaera). Cryptophytes display alternate forms, which were previously confused as separate species, and some have a colonial sessile palmelloid stage and cysts to resist harsh conditions. Sexual reproduction has been observed in Proteomonas and Chroomonas cryptophytes. Haptophytes have amoeboid forms with haploid and diploid stages that reproduce asexually by binary fission (Young et al., 2003). The predominant phase of Emiliania huxleyi is diploid, non-motile, lacking flagella and haptonemas and coccolith bearing (Young et al., 2003), but evidence exists of sexual reproduction in coccolithophores (Billard and Inouye, 2004). In Chrysophyceae, Dictyocha also undergoes sexual reproduction with multinucleate stages. Pelagococcus has been observed to undergo budding (Lewin et al., 1977). For more detailed information on life cycles, we refer the reader to Graham et al. (2009).

3  General Description Most Flagellate cells (excluding Dinophyceae) come in a variety of sizes up to 20 µm often termed picoflagellates (~ 0.8–2 µm) and nanoflagellates (~ 2–20 µm). Most cells are defined by cell shape, cell size, subsurface features, cell covering, flagella ­number, position and shape, chloroplasts (plastids) and their pigment, characteristic swimming motion and any feeding

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

168 ta xonom y structures. A sizeable proportion—principally the Bolidophyceae, Picophagophyceae, Mamiellophyceae and Eustigmatophyceae— are between ~ 1 µm and 2.5 µm which are too small to speciate by light microscopy, although live imaging allows for some diagnostic movement identification. The s­ ection describes features as seen by light microscopy. Cells can be ovoid or elongated. Some may be colonial, such as Chrysophyceae (e.g. Dinobryon sp.), or palmelloid (colonial encased in a mucilaginous casing), such as Prasinophyceae (e.g. Halosphaera sp.) and Haptophyceae (Phaeocystis sp.). Diagnostic cell surface features include calcified/silicified plates (Haptophyta), spines (Dictyochophyceae, Chrysophyceae) or a lorica (basket-like casing) (e.g. Dictyophyceae). Diagnostic intracellular characters are a gullet-like structure and ejectosomes (Cryptophyceae), a flagella-like feeding structure (such as a haptonema in haptophytes), chloroplast arrangement/features (Raphidophyceae, Cryptophyceae) and/or pigment signature (Cryptophyceae). Other subsurface features that are evident upon staining are the  proteinaceous pellicular covering of Euglenophyta or the periplasts of Cryptophyceae. In many cases, flagella number and arrangement are distinguishing features. Pelagophyceae and Dictyochophyceae have one flagellum; Cryptophyceae and Stramenopiles taxa have two unequal flagella but the tiny hairs on stramenopile flagella can only be distinguished by electron microscopy. By contrast, Prasinophyceae can have two, four, eight and occasionally 16 flagella (Tomas, 1997). Some features are symmetrical and have a characteristic motility; for example, some Euglenophyta have a flexible peristaltic movement, while cryptophytes twist in a spiral motion. Phototrophic cells can exhibit autofluorescence from red to yellow, depending on the suite of phytopigments they contain. This allows them to be distinguished using epifluorescence microscopy (Kemp et  al., 1993). Surface or intracellular features can be visualized by staining. Acridine orange can reveal scales with fluorescence microscopy, oil droplets are visible using Nile red solution

under fluorescence light microscopy, cellulose is visible with Calcofluor-white stain, and the mucocysts in Raphidophyceae and Euglenophyceae can be better observed using a neutral red vital stain under the coverslip. For detailed information on autofluorescence or stain-based detection using epifluorescence microscopy, we refer you to Chapter 26 of Kemp et al. (1993).

4  Ecology and Distribution While long-term marine studies have provided much information on protist spatio-temporal distribution, it is worth noting that most of this information comes from coastal, preserved samples that present a biased picture of the overall flagellate community. For those well-studied flagellates, Haptophyceae are mostly mixotrophic, for example Prymnesium patelliferum. They occur at all latitudes and are ubiquitous in open and coastal regions, although are not numerically abundant. Some species have preference for open or coastal regions. Many Haptophyceae (called coccolithales) have calcified plates on their cell wall and form large blooms that sink to form calcareous deposits that have substantially contributed to the chalky White Cliffs of Dover (Saruwatari et al., 2008). However, these cliff formations represent a small section of calcareous mineral deposits that extend into large parts of north-western Europe (Håkansson et al., 1975). Coccolithales are responsible for significant carbon dioxide (CO2) sequestration from the atmosphere and are sensitive to changes in ocean pH (Witty, 2011). Other haptophytes exist in different forms depending on their environment; for example, colonial Phaeocystis antarctica predominate in colder waters, while Phaeocystis globosa blooms in temperate or tropical waters (Lancelot et  al., 1998). Some ­haptophytes produce relatively high levels of a foul-smelling chemical called Dimethylsulphoniopropionate (DMSP) that gets converted to Dimethyl sulphide (DMS) by enzymic action.

F F

F H

Fu E

Fu F

E

Py P

P

Pl

P

Py Euglenophyta

Chlorophyceae

P

Haptophyta

Cryptophyta

FIGURE 30:  General flagellate forms as seen by light microscopy, with key to internal structure.

H Py Pl P Fu E F

= Haptonema = Pyrenoid = Pellicle = Plastid (chloroplast) = Furrow/gullet- cell invagination = Eyespot or stigma = Flagella

ph y topla nkton: flagellates  169 This is released to the atmosphere in aerosol form and eventually converts to sulphuric acid, which contributes to the formation of cloud droplets (Lee and de Mora, 1999; Alcolombri et al., 2015). Several flagellates have recently been found to be symbionts of microzooplankton. Haptophytes (Phaeocystis) ­ and Prasinophyceae (Tetraselmis) are symbionts living in Radiolaria, while Pelagophyceae (Pelagococcus) are symbionts of Foraminifera (van den Hoek et al., 1997; Gast and Caron, 2001; Decelle et  al., 2012). Marine Cryptophyceae are mostly phototrophic and tend to inhabit coastal and brackish waters year round in temperate waters, but are not abundant. Their photosynthetic pigment suite allows them to take advantage of low light, and they often appear at subsurface photic zone levels and have a diurnal vertical migratory pattern. Many flagellates are predated on by zooplankton; Phaeocystis is food for krill in Arctic regions (Haberman et  al., 2003), and Cryptophyceae plastids are indirectly consumed by dinoflagellates (Hackett et al., 2003). The green algae taxa have a varied distribution. Prasinophyceae are phototrophic and generally less than 2.5 µm with a preference for oligotrophic conditions, i.e. ocean water with few nutrients, where they are the main producers, as they have reduced nitrogen requirements in comparison to larger flagellate species. Some are photosynthetically adapted to different light levels and exist at different subsurface layers in the photic zone (Foulon et  al., 2008; Demir-Hilton et  al., 2011). Many ­prasinophyceans are endosymbionts of other marine organisms such as Radiolaria or Noctiluca (Gast and Caron, 2001). Chlorophyceae are almost exclusively freshwater but a few examples, such as Chlamydomonas, occur in brackish, coastal environments (Graham et al., 2009). Chrysophyceae and Synurophyceae are freshwater species that have extended their range to brackish water and nearcoastal waters, and Ochromonas (Chrysophyceae) is common in Norwegian coastal waters (Throndsen et al., 2007). Dinobryon commonly occurs in freshwater, while Dinobryon balticum is common in cold waters. Euglenophyceae and Raphidophyceae are often found in eutrophic, sheltered inland waters and rock  pools in temperate waters and can bloom in summer. Dictyochophyceae (often called silicoflagellates) are ubiquitous with species-specific preferences for temperature and nutrients. Trebouxiophyceae, Eustigmatophyceae, Xanthophyceae and Euglenophyceae have few marine representatives (Graham et al., 2009) but Nannochloropsis (Eustigmatophyceae) has a wide oceanic distribution. For further information on flagellate distribution, we refer the readers to Graham et  al. (2009) and Kooistra et al. (2007).

5  Toxic Species Toxic haptophytes such as Chrysochromulina polylepis, Chrysochromulina leadbeateri and Prymnesium parvum have been reported to destroy fish. Large Phaeocystis blooms are also harmful to fish as Phaeocystis mucus can clog fish gills. All

Raphidophyceae taxa are also toxic, causing damage to aquaculture through the production of mucus that clogs gills, and through free fatty acids, superoxide and hydroxyl radicals that destroy cell membranes and neurotoxic brevotoxins. Dictyocha (Dictyochophyceae) is associated with farmed fish deaths, and Aureococcus (Pelagophyceae) frequently forms toxic blooms in the Eastern seaboard of the USA. For more information on toxic flagellate species, we refer the  readers to the Manual on Harmful Marine Microalgae (Hallegraeff, 2004).

6 Systematics Flagellates are a diverse group of taxa from multiple lineages (see Fig. 1 in Introduction to taxonomy), mainly from Stramenopiles, cryptophytes (Cryptophyceae) and haptophytes (Haptophyceae). Our taxonomy is based on that used by Adl et al. (2012), while we used AlgaeBase (http://algaebase.org) (Guiry and Guiry, 2009) for lower classification levels that have remained relatively stable. There has been extensive taxonomic revision at higher classification levels. The superkingdom Chromalveolata containing Chromista and Alveolata is retained but is controversial. Haptophyta (and classes within) and Cryptophyta were formerly in Chromista along with diatoms, but recent molecular evidence has placed doubt on this grouping, and  they are currently incertae sedis (Adl et  al., 2012). Dictyochophyceae were formerly part of Chrysophyceae, and the phylum Ochrophyta has been removed but is still commonly used elsewhere. All descriptions of taxa are given by their light microscopic characteristics.

BOX 1  Classification of Flagellates Stramenopiles (supergroup SAR)   Class Bolidophyceae   Class Chrysophyceae   Class Dictyophyceae   Class Pelagophyceae   Class Pinguiophyceae   Class Raphidophyceae   Class Eustigmatales Chloroplastida (supergroup Archaeplastida)   Class Prasinophyceae Haptophyceae (incertae sedis)   Class Prymnesiophyceae   Class Pavlovaphyceae   Cryptophyta (incertae sedis) Discoba (supergroup Excavata)   Class Euglenophyceae

170 ta xonom y Class Prymnesiophyceae (incertae sedis) Yellow-brown Cells spherical (e.g. Chrysochromulina hirta, E. huxleyi), oval (e.g. Prymnesium parvum), elongated (Isochrysis spp.) or saddle shaped. Calcified/silicified liths of various shapes (1) that are a diagnostic feature in coccolithophores such as E. huxleyi (likely to be synonymous with Gephyrocapsa oceanica), which has several layers of liths. Spine scales (2) are present in Chrysochromulina and one to two plastids per cell. The haptonema (3) in coccolithophores is emergent in Coccolithus, Calyptrosphaera sphaeroidea, Syracosphaera pulchra, Algirosphaera robusta, Helicosphaera carteri (Billard and Inouye, 2004). E. huxleyi has a vestigial haptonema in its haploid phase; the diploid phase is  predominant. The haptonema is short, non-coiling as in

Prymnesium spp. (Fig 31B) or long and sometimes coiling, such as Chrysochromulina spp. (Fig.  31D). Two equal flagella (4) of variable length, not present in diploid coccolithophores, although present in its haploid phase (Paasche, 2002) (Fig.  31A). Tiny flagellated cells can form larger colonies encased in mucous, for example Phaeocystis spp. (Fig. 31C). Note: approx. 400 species, single cells 2–20 µm. Chrysochromulina hirta is similar to Chrysochromulina ericina using light microscopy. Coccolithophores can be seen with Lugols preservation under neutral or alkaline conditions to preserve the liths. Flagella, haptonema often lost on preservation. Cells should be observed live

A Emiliania huxleyi

B 1

(Lohnmann) Hay & Mohler, 1967

Prymnesium parvum

C Scherffel. 1899 Size: flagellate cells 3–8 µm, colonial >2 mm Distribution: NECS, NADR, NWCS, GFST, NASW, NASE, SARC, ARCT. Coastal, spring/summer bloom

4

Carter, 1937 Size: 6–12 µm, haptomema is 0.25–0.5 times cell length Distribution: NECS, NWCS. Brackish, coastal

Size: 5–10 µm Distribution: NECS, NADR, NWCS, GFST, NASW, NASE, SARC. Cosmopolitan, summer bloom

Phaeocystis globosa

3

D 5

3 4

Chrysochromulina hirta Manton, 1978 Size: 6–7 µm, flagella 20 µm, haptomema >20 µm Distribution: NECS, SARC, ARCT, NWCS. Cosmopolitan, coastal

FIGURE 31:  A, Emiliania huxleyi; B, Prymnesium parvum; C, Phaeocystis globosa; D, Chrysochromulina hirta.

2

ph y topla nkton: flagellates  171 Class Pavlovophyceae (incertae sedis) Yellow-brown Pavlova gyrans

Very small cells, ovoid, and two unequal flagella (1), one much longer than the other (twice as long in Pavlova gyrans) and, in live samples, the longer one ‘pulls’ the cell along. The haptonema is short. Cells have an eyespot (2) and one plastid per cell often as two lobes so can appear as two plastids. Flagella and haptonema inserted subapically in Pavlova genus. Diacronema species are compressed, asymmetrical and have a laterally inserted flagella.

Butcher, 1952 Size: 4–10 µm Distribution: NECS, NWCS.

1 2

Brackish

Note: approx. 12 species, 4–10 µm length. Cells not visible with preservation. Haptonema difficult/impossible to observe by light microscopy

FIGURE 32:  Pavlova gyrans.

Class Cryptophyceae (incertae sedis) Commonly red or olive green (live samples) Marine species are oval to ovoid with two flagella (1). The cells have a gullet (or furrow) (2) lined with ejectosomes (3) that can fire toxins. The cells have one to two acquired plastids per cell of variable shape and contain pyrenoids (4) and sometimes eyespots. Faint striation in periplast can be seen in Rhodomonas (5). Goniomonas is a non-photosynthetic cryptophyte, containing a leucoplast (plastid without pigment). They have a wide range of pigments from brick red, blue-green to olive-green dye to phycoerythrin and phycocyanin pigments. Cryptophytes trace a large, distinctive spiral in their swimming motion that can be used to identify them in living specimens. They can form nonmotile palmelloid stages. Note: approx. 75 species, 5–30 µm length. Lugols preservative only allows flagella and cell outline to be seen. Due to inconsistent taxonomic features and life-form variants, it is only possible to identify Cryptophyceae to genera using light microscopy

Rhodomonas salina

1

(Wislouch) Hill and Weatherby, 1989 Size: length 10–14 µm, width 5–7 µm Distribution: NECS, NWCS, coastal or 2 brackish 3 4 5

FIGURE 33:  Rhodomonas salina.

172 ta xonom y Class Euglenaceae (Discicristata) Green Eutreptiella marina da Cunha, 1914 Size: length 40–50 µm Distribution: NECS, NWCS, coastal 5

6

7 2 3 4

Pear or spindle shaped. Some are rigid in their movement; others appear flexible with a peristaltic movement, for example Eutreptiella gymnastica. A striped pellicle (1) covers the cell. Plastids (2) are arranged in two clusters or are scattered stellate, discoid or ribbon shaped and may contain pyrenoids (3). Carbohydrate bodies (4), made of paramylon, are present in some species. A lightsensing eyespot or stigma (7) in some species near base of flagella. Two to four flagella (5); one commonly emerges from furrow (6) (also called canal or reservoir). Eutreptiella flagella emerge from a furrow, and Eutreptiella marina possess heterodynamic flagella (they beat with different patterns with respect to each other). Note: approx. 1160 species, 40–100 µm length. Euglenaceae and Eutreptiaceae (13 species) are only marine representatives. Very few marine species, all coastal. Best viewed alive. Preservation changes cell shape. Lugols may obscure features but retains flagella, pyrenoids and mucous bodies, whereas flagella are lost with formaldehyde

1

FIGURE 34:  Eutreptiella marina.

Class Raphidophyceae (Stramenopiles) Yellow/brown Chattonella antiqua (Hada) Ono, 1980 1

Size: length 40–50 µm (largest of this genus) Distribution: NECS, NWCS, coastal

2

Round to ovoid or potato shaped, asymmetric, and dorso-ventrally flattened. Heterosigma akashiwo has a lumpy surface. Two unequal flagella (1), the forward-pointing flagellum (flimmer) has a rapid movement, while the second often points backwards. Flagella groove present subapically in Heterosigma, a flagellar furrow can be observed in Fibrocapsa and Chattonella spp. Mucocysts are present in Heterosigma, Fibrocapsa (prominent in posterior cell) and Chattonella. Multiple, densely packed plastids in Chattonella, 3–27 peripheral plastids in Heterosigma. Gullet (2) is present in all except Heterosigma spp. Note: approx. 11 marine species, 8–100 µm length. Many marine members are toxic. Identification is only possible with live cells. Cell shapes can vary according to environmental conditions. Cells are sensitive to microscopy and flagella may be lost or groove no longer apparent

3

4

FIGURE 35:  Chattonella antiqua.

ph y topla nkton: flagellates  173 Class Chrysophyceae (Stramenopiles) Yellow/brown Round or pyriform in shape either naked (Ochromonadales) or with organic/silica scales. Two unequal flagella at apex, the shorter one not always visible by light microscopy. One to two plastids per cell. Cells can be free-swimming or sessile. Cells contain chrysolaminarian as a storage product, with a metallic hue.

Ochromonas cosmopoliticus Carter 1937 Size: length 8–10 µm Distribution: NECS, NWCS, SARC, ARC. Coastal and oceanic

Note: approx. 487 species, 3.5–35 µm length. Chrysolaminarian is visible as cherry-red coloration when stained with brilliant cresyl blue

FIGURE 36:  Ochromonas cosmopoliticus.

Class Dinobryaceae (Stramenopiles) Yellow/brown Round oval cells and many species are attached by cytoplasmic strand (5) to a cellulose lorica (2) urn-shaped envelope. Cells have two unequal flagella (1). Photosynthetic species have plastids (3) with an eyespot (4). Cells have storage bodies made of chrysolaminarian storage that has a shiny, metallic appearance. Many are epiphytic on diatoms or microparticles.

1

Dinobryon belgica Meunier, 1910

2 3

Note: approx 175 species. Cell size: 3.5–25 µm, lorica: 24–60 µm. Lugols preservative distorts cell shape, so identification is only possible through lorica and flagella structures

Size: lorica length 24 µm, lorica width 7 µm Distribution: ARCT, SARC, NWCS. Likely coastal. Summer bloomer

4 5

FIGURE 37:  Dinobryon belgica.

Class Dictyochophyceae (Stramenopiles) Yellow/brown Round/pyriform and radially symmetrical. Cells are either naked or covered with external scales (1), spine scales (2) as in Apedinella (B) in various forms or various star shaped silicon skeletons (5), for example Dictyocha. Cell moves by a pulling motion from the single, flimmer flagellum (4) emerging from an anterior pit. Cells can produce thin pseudopodial extensions, also called rhizopodia (6), arranged radially that can be confused with Heliozoa, for example

in Meringosphaera (Fig. 29). Photosynthetic species have three, six or many plastids (3) typically arranged in a ring. Mucocysts prominent in some species, not shown here. Note: approx. 52 species. Size ~ 4–80 µm, excluding spines/ actinopodia. Dictyocha may be toxic. Pseudopodia and naked cell shape lost with Lugols preservation (continued )

174 ta xonom y Class Dictyochophyceae (Stramenopiles) Yellow/brown (Continued) Pseudopedinella pyriforme

A

Apedinella radians (Lohmann) Campbell, 1973

Carter, 1937

Size: cell length 6–10 µm, Distribution: NECS, ARCT, SARC, NWCS, NASE. Coastal/brackish

4

Size: length 4–5.5 µm, Distribution: NECS, NADR, ARCT, SARC, NWCS, GFST, NASW

4 1 3

3

NASE. Coastal/brackish

Dictyocha speculum

B

2

6

C 4

Ehrenberg, 1839 Size: cell length 19–34 µm plus spines. Distribution: NECS, NADR, NASW, GFST, NASE, ARCT, SARC, NWCS, NASE. Coastal/oceanic

5

3

FIGURE 38:  A, Pseudopedinella pyriforme; B, Apedinella radians; C, Dictyocha speculum.

Class Bolidophyceae (Stramenopiles) Yellow/brown Bolidomonas mediterranea Guillou & Chrétiennot-Dinet, 1999 Size: cell diameter 1–1.7 µm Distribution: NECS, NADR, NASW, GFST, NASE, NWCS, NASE. Likely oceanic

Round/pyriform with one plastid, lacking a cell wall. Two unequal flagella facing different orientations. Moves rapidly. Note: only two species, 1–2 µm in size. Difficult to detect by light microscopy alone

FIGURE 39:  Bolidomonas mediterranea.

ph y topla nkton: flagellates  175 Class Pinguiophyceae (Stramenopiles) Yellow/brown Spherical or pyriform cell shape, no cell wall and one ovoid plastid (2) with pyrenoid (3). Vacuoles (1) are prominent. No stigma. The two unequal flagella (4), oriented in opposing direction. Motile stages are light dependent. Some stages lack flagella. Cytoplasmic processes can be seen in some species. Note: only six species, 1.5–5 µm in size. Difficult to identify by light microscopy

Pinguiococcus pyrenoidosus Anderson, Potter, Bailey, 2002 4

1

Size: cell diameter 3–8 µm Distribution: NECS, NADR, NASW, GFST, NASE, NWCS, NASE. Few light microscopy reports, likely oceanic

2 3

FIGURE 40:  Pinguiococcus pyrenoidosus.

Class Pelagophyceae (Stramenopiles) Yellow/brown Cells are round, ovoid, sarcinoid, filamentous and can be naked or have a cell wall or thecae covering (made of cellulose). Sarcinochrysis possess two unequal flagella which orient in opposite directions; other members lack flagella. Usually one to two plastids but always one in Pelagococcus and two in Sarcinochrysis. Palmelloid stages exist in Sarcinochrysis. No flagellate stages in Pelagococcus but they can form colonies in which cells are elongated.

A

Sarcinochrysis marina Geitler, 1930 Size: cell length 5–7 µm Distribution: NECS, NADR, NASW, GFST, NASE, NWCS, NASE. Likely oceanic

Note: 12 species ranging from 2.5 to 7 µm. Difficult to identify by light microscopy. Species can only be identified by DNA and/or electron microscopy

B

Pelagococcus subviridis Norris in J. Lewin et al., 1977 Size: cell length 2.5–5.5 µm Distribution: SARC. Likely cosompolitan and oceanic

FIGURE 41:  A, Sarcinochrysis marina, flagellate (left) and palmelloid stage (right); B, Pelagococcus subviridis.

176 ta xonom y Class Eustigmatophyceae (Stramenopiles) Yellow/brown Nannochloropsis granulata Karlson & Potter, 1996 Size: cell length 2–4 µm Distribution: NECS but likely to be widely distributed in coastal and oceanic regions

1

Coccoid or rounded cylindrical with thin cell wall containing one plastid lacking a pyrenoid, and many have refractile bodies (1). N. oculata has a red refractile body. Genus level identification only possible. No flagella in this genus. Note: only six marine species, ranging from 2 to 4 µm

FIGURE 42:  Nannochloropsis granulata.

Chlorophyceae (Chlorophyta) Green/olive A

1

Chlamydomonas coccoides Butcher, 1959

2 3

Size: cell length 4.5–5 µm Distribution: NECS, NASE, NWCS, Coastal/brackish, in rock pools

4 B

Dunaliella salina (Dunal) Teodoresco, 1905

5

Cells can be round to ovoid or lobed shape either naked or with a cellulose cell wall. Smooth flagella (1) that number between 1, 2, 4 and 8. One plastid (2) bell shaped, lobed or reticulated with pyrenoid (4) surrounded by starch (5) and an eyespot (3). Cells swim with a pushing breaststroke motion. Note: approx. 2960 species, size: 5–48 µm. Most Chlorophyceae are freshwater species. Cell wall composition may be determined by applying a hypertonic solution to the cells causing the cell contents to contract; staining with Calcofluor-white or staining with zinc chloride and sulphuric acid to detect cellulose cell wall. Lugols obscures much of the cell contents, but pyrenoids may be visible

Size: cell length 16–24 µm Distribution: NECS, NASE, NWCS Coastal/brackish, in rock pools

FIGURE 43:  A, Chlamydomonas coccoides and B, Dunaliella salina.

ph y topla nkton: flagellates  177 Pedinophyceae (Chlorophyta) Green Resultomonas moestrupii (formerly Pedinomonas mikron)

Oval, flattened cells with one long flagellum (2 to 5 times cell size) moving with a jumping motion, flagellum emerging from cell depression. One bean-shaped plastid. This genus has one species and has been found as an endosymbiont in dinoflagellates. Lugols obscures cell contents. Plastid shape best distinguished using epifluorescence microscopy.

Marin, 2012 Size: cell length 1.5–2.5 µm Distribution: SARC, NECS, NASE, NWCS. Coastal/brackish

Note: approx. 21 species

FIGURE 44:  Pedinomonas micron.

Class Prasinophyceae (Chloroplastida) Green/Olive Coccoid, oval, bean-shaped cells. Some species have different life stages; coccoids are phycoma stages (Fig.  45D,E) of flagellated forms (not shown here). Phycomas are much larger with single or multiple ala (1). Flagella (1) are thick, sometimes attached to hair and number between 0, 1, 4, and 8. Scale-theca cell covering appears as cell wall that can be used to speciate cells but only through electron microscopy. One bell or lobed-shaped plastid Pyramimonas cirolanae

A

Pennick, 1982 Size: cell length 3.6–4 µm

with/without pyrenoid (3) that can be covered in a thick starch band. Red stigma (2). Flagellated (Fig.  45A,B) forms may have characteristic swimming motion in a straight line. Note: approx 108 species. Size: Flagellates 1.5–7 µm, Phycoma 33–200 µm. Cell con­tents obscured with Lugols preservation but pyrenoids may be visible

1

Micromonas pusilla

2

(Butcher) Manton & Parke, 1960

B

Size: cell length 1–3 µm

Distribution: NECS, SARC.

3

Likely to be widely distributed

Distribution: NECS, NADR, NASE, NASW, GFST, NWCS,

in coastal regions

SARC, ARCT. Coastal , less abundant in oceanic regions

Mamiella gilva

C

Halosphaera viridis Schmitz, 1878

(Parke & Rayns) Moestrup, 1984

Size: phycoma cell

Size: cell length 4–6.5 µm

400–800 µm

Distribution: NECS, NADR,

Distribution: NWCS NADR.

NASE, NASW, GFST, NWCS.

Likely to be widely distributed in

Mainly oceanic

coastal and oceanic regions

Pterosperma porosum

D

E

Parke, 1978 Size: cell length 45–55 µm (including ala) Distribution: NECS. Likely to be widely distributed in coastal and oceanic regions

1 FIGURE 45:  A, Pyramimonas cirolanae;

B, Micromonas pusilla; C, Mamiella gilva; D, Halosphaera viridis phycoma stage; E, Pterosperma porosum phycoma stage.

178 ta xonom y

References Adl, S., Simpson, A. G. B., Lane, C.E., et al. (2012). The Revised Classification of Protists. Journal of Eukaryotic Microbiolgy 59(5), 429–93. Alcolombri, U., Ben-Dor, S., Feldmesser, E., et  al. (2015). Identification of the algal dimethyl sulfide–releasing enzyme: A missing link in the marine sulfur cycle. Science 348(6242), 1466–9. Billard, C. and Inouye, I. (2004). What is new in coccolithophore biology? In Thierstein, H. R. and Young, J.R. (eds) Coccolithophores. From Molecular Processes to Global Impact, pp. 1–29. Springer, New York, NY. Decelle, J., Probert, I., Bittner, L., et  al. (2012). An original mode of symbiosis in open ocean plankton. Procedings of the National Academy of Sciences 109(44), 18000–5. Demir-Hilton, E., Sudek, S., Cuvelier, M. L., et al. (2011). Global distribution patterns of distinct clades of the photosynthetic picoeukaryote Ostreococcus. The ISME Journal 5(7), 1095–107. Foulon, E., Not, F., Jalabert, F., et al. (2008). Ecological niche partitioning in the picoplanktonic green alga Micromonas pusilla: evidence from environmental surveys using phylogenetic probes. Environmental Microbiology 10(9), 2433–43. Gast, R. J. and Caron, D. A. (2001). Photosymbiotic associations in planktonic foraminifera and radiolaria. Hydobiologia 462, 1–7. Graham, L. E., Graham, J. M., et  al. (2009). Algae. Pearson Benjamin Cummings: San Francisco, CA. Guiry, M. D. and Guiry, G. M. (2009). AlgaeBase. World-wide electronic publication. National University of Ireland, Galway. Haberman, K. L., Quetin, L. B., and Ross, R. M. (2003). Diet of the Antarctic krill (Euphausia superba Dana): I. Comparisons of grazing on Phaeocystis antarctica (Karsten) and Thalassiosira antarctica (Comber). Journal of Experimental Marine Biology and Ecology 283(1–2), 79–95. Hackett, J. D., Maranda, L., Yoon, H.S., and Bhattacharya, D. (2003). Phylogenetic evidence for the cryptophyte origin of  the plastid of Dinophysis (Dinophysiales, Dinophyceae). Journal of Phycology 39(2), 440–8. Hallegraeff, G. M. (2004). Manual on harmful marine microalgae. UNESCO: Paris. Håkansson, E., Bromley, R., et al. (1975). Maastrichtian chalk of North-West Europe: a Pelagic Shelf Sediment. In Hsu, K. J. and Jenkyns, H.C. (eds) Pelagic sediments on land and under the sea. Wiley-Blackwell: Hoboken, NJ. Kemp, P. F., Sherr, B. F., Sherr, E. B., Cole, J. J. (1993). Handbok of methods in aquatic microbial ecology. Lewis Publishers: Boca Raton, FL.

Kooistra, W., Gersonde, R., et  al. (2007). The origins and evolution of the diatoms: Their adaptation to a planktonic existence. In Falkowski, P. G. and Knoll, A. (eds) Evolution of primary producers in the sea, pp. 210–39. Elsevier: Burlington, MA. Lancelot, C., M. D. Keller, et  al. (1998). Autecology of the marine haptophyte Phaeocystis sp. In Anderson, D. M., Cembella, A.D., and Hallagraeff, G.M. (eds). Physiological ecology of harmful algal blooms, 41, 209–24. SpringerVerlag: Berlin. Lee, P. A. and de Mora, S. J. (1999). INtracellular Dimethylsulfoxide (DMSO) in unicellular marine algae: speculations on its origin and possible biological role. Journal of Phycology 35, 8–18. Lewin, J., Norris, R. E., Jeffrey, S. W., and Pearson, B. E., (1977). An aberrant chrysophycean algal Pelagococcus subviridis Gen. Nov. et Sp. Nov. from the North Pacific ocean. Journal of Phycology 13, 259–66. Ohtsuka, S. and Suzaki, T. (2015). Marine protists: Diversity and dynamics. Springer Japan: Tokyo. Paasche, E. (2002). A review of the coccolithophorid Emiliania huxleyi (Prymnesiophyceae), with particular reference to growth, coccolith formation, and calcification-photosynthesis interactions. Phycologia 40, 503–29. Saruwatari, K., Ozaki, N., Nagasawa, T., and Kogure, T. (2008). Comparison of crystallographic orientations between living (Emiliania huxleyi and Gephyrocapsa oceanica) and fossil (Watznaueria barnesiae) coccoliths using electron microscopes. American Mineralogist 93, 1670–7. Throndsen, J. (1997). The planktonic marine flagellates. In  Tomas, C. R. (ed.) Identifying marine phytoplankton, pp. 633–44. Academic Press: San Diego, CA. Throndsen, J., Hasle, G.R., Tangen, K. (2007). Chrysophyceaegolden algae. In: Throndsen, J., Hasle, G.R., Tangen, K. (eds) Phytoplankton of Norwegian coastal waters, pp. 201–10. Almatar Forlag As: Oslo. Tomas, C. R. (ed.) (1997). Identifying marine phytoplankton. San Diego, CA, USA, Academic Press: San Diego, CA. van den Hoek, C., Mann, D.G., Jahns, H.M. (1997). Algae. An Introduction to Phycology. In van den Hoek, C., Mann, D.G., and Jahns, H.M. (eds) Algae. An introduction to phycology, pp. 219–34. Press Syndicate of the University of Cambridge: Cambridge. Witty, M. (2011). The White Cliffs of Dover are an example of natural carbon sequestration. Ecologia 1, 23–30. Young, J., Geisen, M., Cros, L., et al. (2003). A guide to extant coccolithophore taxonomy. Journal of Nannoplankton Research Special Issue 1, 1–125.

ph y topla nkton: flagellates  179

Appendix 1: Terminology used in the description of phytoplankton flagellates Term

Description

Ejectosomes

An explosive organelle composed of a coiled structure within a vesicle that can be rapidly discharged to release toxins.

Eyespot

A dark orange-red, photoreceptive organelle capable of light detection and composed of bright orange-red pigments. Often called a stigma

Filopodium/Filopodia

Thin, spine-like extensions of the cell

Flimmer flagella

The longer of the two unequal sized flagella in Stramenopiles with one or two rows of hair-like appendages (not visible by light microscopy), and sometimes whip flimmer flagella that are ‘winged’ or broader for locomotion, lacking hairs.

Gullet

An invagination into a cell, also called furrow or

Haptonema

A fine tube-like feeding structure in Haptophytes

Heterodynamic flagella Flagella on the same cell that beat with different patterns Heterotrophic

Obtain nutrition solely through feeding on other organisms

Lith

Calcerous or silicious structures covering Coccolithophore (Haptophyceae) cells

Longitudinal flagellum

One of two flagella pointing towards the Antapex in Dinophyceae

Lorica

Basket-like structure encasing cell

Mastigoneme

The longer flagellum in stramenopile cells, with tripartite hairs (see Fig. 2)

Mixotrophic

Obtains nutrition both by photosynthesis and feeding on other organisms.

Naked

Without a cell wall

Pellicle

The striped, proteinaceous cell wall cover of Euglenophyta

Phototrophic

Obtains nutrition solely through photosynthesis. Also called autotrophic or photolithotrophic

Plastid

Chloroplast

Pseudopodium/ Pseudopodia

Thick, ‘foot-like’cytoplasmic extensions of cells

Smooth flagella

Smaller of the two unequal sized flagella in Stramenopiles (see Fig. 2)

Spicule

Spine-like structures covering

Test

Calcerous shell excreted by and covering the surface of foraminifera cells.

Transverse flagellum

One of two flagella coiled round the circumference of Dinophyceae

Valve

Silicious structure of Diatom cells. Diatoms have two valves fused together.

Winged Flagella

Thicker width flagella , ribbon shaped

pa rt 2

Zooplankton

Protozooplankton: Ciliates A lex A NDR A K r a berg, Row ena Ster n, a nd Mich a ela Strüder-K y pk e 1 Introduction Ciliates are unicellular eukaryotic organisms ranging in size from 10 µm in small spheroid forms to 4500 µm in highly elongate and contractile ones. Over 8000 morphospecies of ciliates including about 200 fossil forms and close to 3000 symbiotic species have been described (Lynn, 2008), although it can be assumed that this represents a gross underestimate (Foissner et al.,  2008). Ciliates can be found in almost every environment; the majority of species, however, are aquatic and occur in marine, brackish and freshwater habitats. They play a major role in nutrient cycling in the food web and although most are heterotrophic, some are also capable of photosynthesis through acquisition of chloroplasts from their prey, which they temporarily keep for photosynthesis. Phylogenetic analysis of ciliates has altered some taxa groupings and their systematics continue to be revised.

2  Life Cycle The life cycles of ciliates are complex with many variants on the general scheme outlined in Figure 47. Sexual reproduction usually happens under unfavourable environmental conditions; the common way for ciliates to proliferate is (asexual) division. Unlike other protists, ciliates possess two types of nuclei (nuclear dimorphism). The micronucleus is diploid and is the generative nucleus, which divides either by mitosis or meiosis (sexual stage) with haploid daughters fusing to form new micro- and macronuclei (Fig.  47). The macronucleus is polyploid (contains several copies of each chromosome) and controls the physiological and biochemical functions of the cell (vegetative nucleus) and can divide by mitosis (binary division), although after several hundred generations deteriorates and needs to be regenerated from the micronucleus in the sexual stage. The life cycle of a ‘typical ciliate’ involves 1) a vegetative (asexual) cycle with growth and division of the cell, 2) a sexual cycle with conjugation and exchange of genetic material, and 3) a cryptobiotic cycle with a resting cyst. However, this basic model has been adapted in many different ways. During conjugation, two cells fuse partially and temporarily, and exchange a so-called migratory micronucleus, which subsequently fuses with the stationary micronucleus to form a synkaryon and allows for genetic recombination.

Figure 46:  Planktonic ciliate Laboea strobila (credit F. Neidl, source: Plankton*net (http://planktonnet.awi.de)).

3  General Morphology Ciliates are unicellular and vary in size, cell shape, and ciliation. Most ciliates have a cytostome (cell mouth), located apically, subapically, or posteriorly. In bacterivorous ciliate species, oral ciliature (arrangement of ciliates) and the cytostome form an effective filter feeding apparatus. Some ciliate groups can build a lorica (a cage-like structure), which is either attached to the substrate or free (i.e. the tintinnids). Some can be contractile, while others attach to a substrate through a stalk or a lorica. These features can mostly be distinguished in live or Lugol’s fixed samples, although exact species identification in most cases still requires protargol staining. The ciliation is one of the key features. The cilia are morphologically similar to the flagella but are much shorter and usually occur in much greater numbers. Individual cilia are arranged in rows (called kineties) or ‘clusters’ (membranelles, cirri). The

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

184 ta xonom y

Figure 47:  Typical ciliate life cycle.

cortex, which is the main interface between the ciliate and its environment, can be divided into a somatic region and an oral region. The somatic region functions in locomotion, protection, defensive response and attachment to the substrate. The oral region functions in the acquisition and ingestion of food particles (Lynn,  2008). Somatic ciliation can be holotrichous (uniformly ciliated, cilia arranged in rows = kineties, e.g. Tetrahymena) or greatly reduced to just a few kineties (e.g. Strobilidium), or it can be entirely missing in certain stages of some taxa (e.g. Acineta). The oral ciliation can be missing (e.g. Suctoria), very ‘simple’ (e.g. Didinium), or extremely conspicuous (e.g. Strobilidium). Tintinnid ciliates have a lorica and are cylindrical. The cell is attached to the lorica with a posterior, stalk-like peduncle. Tintinnids are conspicuous in the marine plankton, and over 1200 species have been described based on lorica morphology. Other features of ciliates are contractile vacuoles which can be star shaped; however, these are usually absent in marine species. There is a large range of variation in number and shape of macronuclei among different ciliates. The shape, size and number of the nuclei (macro- and micronucleus) can be species or at least genus specific. Many ciliates have photosynthetic symbionts, for example Mesodinium rubrum, and some display colouration due to pigments in the cortex, for example Blepharisma.

4  Ecology and Distribution Ciliates have representatives in all major habitats from freshwater to fully saline waters and soils, although tintinnids are predominantly marine (Foissner et al., 2008; Foissner, 2008). Their geographic distribution is truly global, ranging from Arctic to Antarctic. Most marine species are free-swimming planktonic, and others are attached to particles in plankton such as marine snow. Ciliates are an important component of the microzooplankton. Most ciliates are heterotrophic and feed on prey including bacteria, various kinds of phytoplankton and other ciliates. Some ciliates are mixotrophic, such as M. rubrum, Laboea strobila and some species in the genera of Strombidium and Tontonia, temporarily retaining the functional chloroplasts of their photosynthetic prey (Stoecker et al., 1988) and thus able to photosynthesize and capture prey. Several taxa are parasitic and some live symbiotically/commensalistically in or on animals, for example on the surface of crustaceans. Some suctorians also colonize the surface of the thecae of diatoms. Both M. rubrum and Tiarina fusus are harmful species and can form extensive blooms (red tides) (Dale and Dahl,  1987). The geographical distribution of species described in this section is based on a number of sources including the World Register of Marine Species (WoRMS).

protozoopla nkton: cili ates  185

5  How to Identify Ciliophora Ciliates are a monophyletic group and are united by three characteristic features: (a) presence of cilia; (b) nuclear dimorphism (two types of nuclei); and (c) conjugation (sexual reproduction). The kinds and arrangements of ciliary structures are important in the identification of the major groups of ciliates. In some species the ciliature is easily discernible in light microscopy and allows species identification (examples are L. strobila and M. rubrum). Often, however, staining (e.g. with Protargol; Skibbe, 1994) is necessary to visualize kineties and internal structures. A major distinguishing characteristic of ciliates is the presence of two types of nuclei (nuclear dimorphism). The micronucleus is typically smaller and spherical, while the macronucleus is much larger and occurs in many different shapes (spherical, ellipsoidal, reniform, C- or J-shaped, moniliform, multiple small ones). There is a large range of variation in number and shape of macronuclei among different ciliates. Of the major ciliate groups that are present in marine ­plankton, spirotrichs are probably the most easily recognizable group (Fig. 48). Their oral ciliation consists of a large band of membranelles (compound cilia that are called polykinetids or  trichea) either in an open or closed circle (Oligotrichia, Choreotrichia), or as an inverted J-shaped band (Euplotia, Hypotrichia, Licnophoria) around their mouth (oral cavity). The ciliature resembles bristles rather than fine hairs. The somatic ciliation (oral kineties proper) is reduced and several taxa possess cirri (Euplotes, Stylonychia). Other marine planktonic groups with holotrichous ciliation are the prostome ciliates (Urotricha, Tiarina), and many scuticociliates (Uronema, Pleuronema). Their cell size varies and different groups can be distinguished by the location and overall appearance of the oral region (apical to subapical with circumoral cilia in prostomes, and anterior or posterior cell region with undulating ­membrane

Anterior adoral membranelles Ventral adoral membranelles Girdle kinety Subpellicular platelet layer (seen with fixation) Macronuclear nodules

Distended cell surface

Figure 48:  Example of a spirotrich ciliate – Laboea strobila.

and oral membranelles on the left and right side, respectively, of the oral region in scuticociliates).

6 Systematics Ciliates belong to the Alveolata superphylum along with dinoflagellates, and they are broadly related to the SAR group (see Fig. 1, Introduction). This section presents a reduced classification of the most common marine planktonic ciliate taxa. For more information on the current classification of ciliates (Ciliophora) see Adl et al. (2012) and Lynn (2008). Detailed systematics of oligotrich and choreotrich taxa can be found in Agatha and Strüder-Kypke (2013, 2014) and in Dolan et al. (2013) that focuses on tintinnid ciliates.

Box 1  Classification of ciliates Supergroup SAR (Stramenopiles, Alveolates, Rhizaria) Superphylum Alveolata Phylum Ciliophora Class Spirotrichea   Subclass Oligotrichia   Order Strombidiida    Family Tontoniidae     Genus Laboea     Species Laboea strobila   Subclass Choreotrichia   Order Choreotrichida   Suborder Lohmanniellina    Family Lohmanniellidae     Genus Lohmaniella     Species Lohmanniella oviformis   Order Tintinnida    Family Codonellidae     Genus Tintinnopsis     Species Tintinnopsis cylindrica Class Litostomatea    Incertae sedis    Family Mesodiniidae     Genus Mesodinium     Species Mesodinium rubrum Class Prostomatea    Order Prorodontida    Family Colepidae     Genus Tiarina     Species Tiarina fusus Only taxa in bold are covered in this chapter.

186 ta xonom y Class Spirotrichea Subclass Oligotrichia Order Strombidiida Family Tontoniidae The most abundant group of marine planktonic ciliates belongs to the class Spirotrichea and, more specifically, oligotrich and choreotrich genera. Spirotrichea are characterized by numerous, prominent oral membranelles (1 and 2). Oligotrich ciliates are always aloricate, have a globular to obconical cell shape, a prominent adoral zone of membranelles, and are arranged in an open circle and in two morphologically distinct regions, the anterior and the ventral zone of membranelles (‘collar and lapel’). They possess only two somatic kineties (so-called girdle and ventral kinety) with possibly sensory function. Arrangement and direction of these kineties are important taxonomic characters. We currently recognize two families: Tontoniidae and Strombidiidae. The latter is a large and speciose group, which likely consists of several families. L. strobila is an obligate mixotrophic ciliate, 50–150 µm long. The cell outline is conical. The girdle kinety (3) running in several whorls around the body makes this a distinct species. The oral cavity (4) is narrow with distinct oral cilia extending into the oral groove. More than 40 macronuclear nodules (5) per cell can be present (Agatha et al., 2004).

1 4 2

3

5

Figure 49:  Laboea strobila.

Note: many species are mixotrophic and sequester chloroplasts of ingested prey algae

Class Spirotrichea Subclass Choreotrichia Order Choreotrichida Suborder Lohmanniellina Family Lohmanniellidae 1 2

3 4 Figure 50:  Lohmanniella oviformis.

Choreotrich ciliates very much resemble oligotrich ciliates in cell size and shape. The distinct adoral zone of membranelles forms a closed circle, with an internal and an external zone of membranelles (1). The somatic ciliature of aloricate choreotrichs is reduced, but not as much as in oligotrich species – it usually consists of three or more somatic kineties, arranged mostly longitudinally but in varying patterns, which are species specific. Cells of Lohmanniella oviformis are 10–25 µm long and heterotrophic. The body is subspherical with the oral cavity (2) located anteriorly but acentrically. One kidney-shaped, lateral macronucleus (3) is present. Five short somatic kineties (4) are present near the posterior end of the cell radiating from the posterior pole.

Class Spirotrichea Subclass Choreotrichia Order Tintinnida Family Codonellidae External Polykinitid zone Oral cavity Ventral kinety Macronucleus

posterior kinety

3 1

2

Choreotrich ciliates that are mostly encased in a lorica (a proteinaceous shell) are called tintinnnids and are cylindrical in shape. The cell is attached to the lorica by a stalk-like peduncle (3). Some tintinnids have hyaline loricae (e.g. Favella), while other tintinnids possess agglutinated loricae (1) on the surface of which small fragments of minerals are incorporated (e.g. Tintinnopsis). Lorica morphology, however, shows a high degree of phenotypic plasticity, depending on environmental conditions, or life stages. The most reliable lorica features are general outline, details of opening rim, opening diameter, and wall texture

Figure 51:  Tintinnopsis cylindrica.

(Laval-Peuto and Brownlee,  1986). The genus Tintinnopsis comprises a collection of several, not necessarily closely, related species (Agatha and Strüder-Kypke, 2013) (see Fig. 52). Tintinnopsis cylindrica (Fig. 51) is 135–180 µm long with an aperture width of 45–50 µm, and it possesses an agglutinated lorica (1), posteriorly tapered, merging into a straight cylindroidal process (2) about 20 µm long and 10–15 µm wide. Cells elongate, obconical and highly contractile. Posterior end narrowed and always forming a short stalk-like peduncle (3), with which cell adheres to inside of lorica.

B

A

D

C

Figure 52:  Selected tintinnid lorica forms. A, Epiplocylis mucronata

(formerly Codonella mucronata); B, Dictyocysta lata, C, Cymatocylis convallaria (formerly Coxliella cymatiocoides); and D, Rhabdonella chavesi. Redrawn from Marshall (1969).

Class Litostomatea (Incertae Sedis) Family Mesodiniidae The family Mesodiniidae comprises several freshwater species and M. rubrum. The genus Mesodinium is easily recognizable due to its particular cilia arrangement in two ciliary belts. M. rubrum is a mixotrophic species, 10–30 µm long. Cell outline is roughly oval, a subcentral constriction (1) seemingly bearing a dense ring of cilia that actually consists of two belts of polykinetids (2, 3). Two macronuclei are located in a central position (4). The cytoplasm has a pinkish colour resulting from the presence of chloroplasts that Mesodinium acquires from feeding on cryptophytes. Importantly the ingested cryptophyte plastids and nuclei remain functional for several days (Gustafson et al., 2000; Johnson et al., 2007). After Lugol fixation the cells appear dark brown. Live cells are highly motile, performing jumping movements with periods of rest between jumps. Jumps can exceed the body length tenfold. Notes: Taxonomic placement is highly uncertain. Some morphological and molecular features unite Mesodinium with the litostome clade, and specifically with the free-living Haptoria (Strüder-Kypke et al., 2006; Moestrup et al., 2012). Other characteristics, however, separate it clearly from any other ciliate class. We therefore list it here as uncertain taxon within the Class Litostomatea.

2 3 1 4

Oral tenticles

Figure 53:  Mesodinium rubrum.

Class Prostomatea Order Prorodontida  Family Colepidae Prostome ciliates are mostly small in size, with ovoid to cylindroid cell shape. The somatic ciliation is holotrichous but often reduced in the posterior region. The cytostome is located apically to subapically, with circumoral ciliation. Microtubular ribbons support the cytopharynx. T. fusus has a fusiform body, i.e. tapering towards the posterior end. Five rows of calcareous plates (1) with lateral teeth (2) are present in the cortex and are visible in light microscopy. Approximately 15 rows of somatic kineties (3) are located between the plates. Cells contain one macronucleus (4) and one micronucleus (5). The oral cavity is located apically. The posterior end of the cell can bear spines (6).

Oral ciliature

Wing-like structure

5 4

1

2

3

Spiculum

Figure 54:  Tiarina fusus diagram based on Chen et al. (2012).

6

Caudal cilium

188 ta xonom y

References Adl, S., Simpson, A. G. B., Lane, C.E., et al. (2012). The Revised Classification of Protists. Journal of Eukaryotic Microbiolgy 59(5), 429–93. Agatha, S. and Strüder-Kypke, M. C. (2013). Systematics and evolution of tintinnid ciliates. In Dolan, J. R., Montagnes, D. J. S., Agatha, S., Coats, D. W. & Stoecker, D. K. (eds) The biology and ecology of tintinnid ciliates—Models for marine plankton, pp. 42–84. John Wiley & Sons, Ltd: Oxford, Chichester. Agatha, S. and Strüder-Kypke, M. C. (2014). What morphology and molecules tell us about the evolution of Oligotrichea (Alveolata, Ciliophora). Acta Protozoology 53, 77–90. Agatha, S., Strüder-Kypke, M. C., and Beran, A. (2004). Morphologic and genetic variability in the marine planktonic ciliate Laboea strobila Lohmann, 1908 (Ciliophora, Oligotrichia), with notes on its ontogenesis. Journal of Eukaryotic Microbiology 51(3), 267–81. Chen, X., Gao, S., Liu, W., et al. (2012). Taxonomic descriptions of three marine colepid ciliates, Nolandia sinica spec. nov., Apocoleps caoi spec. nov. and Tiarina fusa (Claparède & Lachmann, 1858) Bergh, 1881 (Ciliophora, Prorodontida). International Journal of Systemic Evolutionary Microbiology 62(3), 735–44. Dale T. and Dahl, E. (1987). Rodt vann ved Tvedestrand masseforekomst av flimmerdyret Tiarina fusus. Fauna 40, 98–103. Dolan, J. R., Montagnes, D. J. S., Agatha, S., Coats, D. W., and Stoecker, D. K. (eds) (2013). The biology and ecology of tintinnid ciliates—Models for marine plankton. John Wiley & Sons, Ltd: Oxford, Chichester. Foissner, W. (2008). Protist diversity and distribution: some basic considerations. Biodiversity and Conservation 17, 235–42.

Foissner, W., Chao, A., and Katz, L. A. (2008). Diversity and geographic distribution of ciliates (Protista: Ciliophora). Biodiversity and Conservation 17, 345–63. Gustafson, D. E., Stoecker, D. K., Johnson, M. D., Van Heukelem, W. F., and Sneider, K. (2000). Cryptophyte algae  are robbed of their organelles by the marine ciliate Mesodinium rubrum. Nature 405, 1049–52. Johnson, M. D., Oldach, D., Delwiche, C. F., and Stoecker, D. K. (2007). Retention of transcriptionally active cryptophyte nuclei by the ciliate Myrionecta rubra. Nature 445, 426–8. Laval-Peuto, M. and Brownlee, D. C. (1986). Identification and  systematics of the Tintinnina (Ciliophora): evaluation and suggestions for improvement. Annales de l’Institut Océanographique 62, 69–84. Lynn, D. H. (2008). The ciliated protozoa. Characterization, classification, and guide to the literature. Springer: Dordrecht. Marshall, S. M. (1969). Zooplankton: Protozoa. Fiches Ident. Zoopl. C. I. p. l. e. d. l. mer: 117–23. Moestrup, Ø., Garcia-Cuetos, L., Hansen, P. J., and Fenchel, T. (2012). Studies on the genus Mesodinium I: ultrastructure and description of Mesodinium chamaeleon n. sp., a benthic marine species with green or red chloroplasts.’ Journal of Eukaryotic Microbiology 59, 20–39. Skibbe, O. (1994). An improved quantitative protargol stain for ciliates and other planktonic protists.’ Archiv für Hydrobiologie 130(3), 339–47. Stoecker, D. K., Silver, M. W., Michaels, A. E., and Davis, L. H. (1988). Obligate mixotrophy in Laboea strobila, a ciliate which retains chloroplasts. Marine Biology 99, 415–23. Strüder-Kypke, M. C., Wright, A. -D. G., Foissner, W, Chatzinotas, A., Lynn, D. H. (2006). Molecular phylogeny of litostome ciliates, (Ciliophora, Litostomatea) with emphasis on the free-living haptorian genera. Protist 157, 261–78.

Protozooplankton: R adiolaria Row ena Ster n, Cla ir e Tay lor, Fa br ice Not, a nd Joh a n Decelle 1 Introduction Radiolaria are amoeboid unicellular eukaryotes with mineral skeletons, often forming elaborate symmetrical lattice structures with spines. They form an abundant part of marine holoplankton from surface to deep waters. This is an ancient group that has existed for at least 600 million years based on fossil records (Kling and Boltovskoy, 2002), often forming microfossils useful to geologists and micropaleontologists for palaeoclimatic dating. Often classed as microzooplankton, Radiolaria are larger, buoyant cells of 50–5000 µm size that  are solitary or colonial. They feed on different prey, and many radiolarian species can also live symbiotically with microalgae such as haptophytes and dinoflagellates. Radiolaria have undergone extensive taxonomic revision that is still in progress,  and this term now encompasses five major orders of organisms – Acantharia, Taxopodia, Collodaria, Nassellaria and  Spumellaria (the latter three are also together called Polycystinea) (Suzuki and Aita, 2011). A key feature of Radiolaria is their compartmentalized internal cell structure. They possess axopodia (a type of pseudopodia) – stiff cytoplasmic, retractable processes radiating out from the cell used for capturing prey. Acantharia (see Plate 18 for examples) have spicules (spike-like structures) made uniquely of strontium sulphate (celestite) that is very soluble and rapidly dissolves upon the organism’s death by fixatives, making taxonomic studies and geological dating of them rare (Decelle et al. 2012a). Other Radiolaria are either naked or have amorphous silica (opal) cell walls (see Plate 19 for examples). Even if their species abundance and diversity in the environment have been recently proved to be significant, based on both molecular and in situ imaging technologies, they are likely to be underestimated. Radiolaria are best collected in fine mesh nets or in water samples, due to their delicate nature. So far, no culture of Radiolaria has been possible so very little is known about their life cycle.

2  Life Cycle Due to the difficulty in maintaining Radiolaria (no one has managed to culture them), little is known about their reproduction. A variety of different life cycles have been reported, although none have been completely proven or observed ex situ

Figure 55:  Radiolaria. Photo courtesy of Savalli (2015).

(Dolven et al., 2009). It is thought that there are two modes of reproduction: vegetative binary and multiple fission (Ishitani et  al., 2016). In the former, daughter cells divide within the ­skeleton in polycystinean radiolarians – one inherits the parent skeleton and the other forms a new test. Adult acantharians can divide sexually, releasing thousands of 2–3 µm biflagellated swarmer cells that are probably gametes after encystment hypothesized to occur in deep water or directly from the adults in shallow water, depending on the taxa, shown in Figure 56 (Anderson, 1983; Decelle et al., 2012a, 2013). Radiolaria are often infected with intracellular parasites that release similar-­ looking flagellated cells post infection, and these can erroneously be attributed as swarmer cells (Anderson, 1983). A variety of cysts have also been observed for Acantharia, and some studies such as Decelle et al. (2013) have tentatively inferred their life cycles and evolution from morphogenetic and molecular studies of environmental marine samples (Fig. 56).

3  Ecology and Distribution Radiolaria (including Acantharia) are active predators and can feed on bacteria, phytoplankton and some zooplankton (including younger copepod stages) and in turn are fed on by larger organisms such as salps (Ishitani et al., 2016). Radiolaria are ­cosmopolitan, although some have niches that can be used as indicator species. They are numerically dominant in oligotrophic

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

190 ta xonom y release of swarmers Sinking cyst

Deep water

Encystment clades A, B, C

Algal symbiont acquisition (clades E,F)

unicellular algae (dinoflagellates, haptophytes) benefiting from photosynthetic primary production that photosynthesis brings (Probert et al., 2014; Decelle and Not, 2015). Their diurnal cycle is likely to be influenced by their algal ­symbionts. Because of their mixotrophy, symbiosis with a photosynthetic partner and mineralized skeletons, Radiolaria play multiple roles in the ecology of marine ecosystems. For instance, Acantharia contribute largely to the carbon and strontium biogeochemical cycles. (Decelle et al., 2013) They also contribute to other trace metal recycling such as barium. Other Radiolaria, composed of silicon, are better preserved and date back to the Cambrian, and their skeletons contribute to sedimentary cycles (Lazarus, 2005).

Adult cell Shallow water

4  General Morphology release of swarmers

Figure 56:  Hypothesized life cycle of Acantharia (Decelle et al.

2013) showing vegetative (1) and cyst (2) life-cycle stages that are thought to occur in the surface photic zone and the deep ocean. Juvenile forms aquire algal symbionts later in their development. Alternatively cells encyst, sink to greater depths and release swarmers. There may be sexual stages that fuse and form juvenile Acantharia that rise to the ocean surface.

waters in equatorial and subtropical waters and are rarer in temperate and polar waters and in near-shore environments (Suzuki and Not, 2015a). Radiolaria have been observed in large blooms from the surface downwards (on calm days) where they can contribute a significant fraction of plankton biomass in intertropical regions (Biard et al., 2016). They can also descend to several hundreds to thousands of metres below the surface in disturbed weather conditions. Many of them have endosymbiotic

By light microscopic identification, Radiolaria cells are large and spherical, ovoid or lenticular, with or without spicules (see  Fig.  57). Many are encased in elaborate mineral shell forms. Acantharia generally contain spicules radiating out from a merged point at the cell centre. The internal cell body of radiolarians consists of an internal central capsule (often pigmented brown, black or red) containing the endoplasm that is separated from the outer ectoplasm by a fibrillar capsular cell wall. Surrounding the ectoplasm is the extracapsular cortex. The central capsule contains organelles and performs all essential functional roles of respiration and reproduction. It also contains a structure called the axoblast from which axopodia originate and extend outside the cell, resembling a sun burst. The shape of the skeleton and central capsule are the main diagnostic determinants for taxonomic identification. Both ecto- and endoplasm may contain algal endosymbionts. The extracapsular cortex is joined to  spicules by contractile myoneme organelles in Acantharia. The periplasmic cortex has a frothy appearance that can hold captured prey.

Cephalus Filopods Zooxanthellae Ectoplasm (Calymma) Nuclei Endoplasm Radial spicule

Endoplasm with zooxanthellae Thorax Stricture

Ectoplasmic cortex

Abdomen

Axopod

Apical spine

Myoneme

Post-abdomenal segment

Rib Pore Aperture

Figure 57:  Schematic organization of Radiolaria showing typical acantharian (left) and on the right, polycystine morphology belonging to the

Nassellaria order.

protozoopla nkton: r a diola r i a  191

5  How to Identify Radiolaria Descriptions in this section are given for class and orders only as their detailed taxonomy are beyond the scope of this book. We  refer readers to the Radiolaria chapters referred to at the  end of this section (Suzuki and Not, 2015a,b), Radiolaria (Ishitani et al., 2016) and Acantharia (Decelle and Not, 2015) for more information. Order Acantharia: Generally spherical with 10 or 20 spicules. Comprised of four suborders of about 50 genera and 150 species (Decelle et al., 2012b; Decelle and Not, 2015). Phylogenetic analysis has placed them into nine main clades (groups) (Decelle et  al., 2012b; Decelle and Not, 2015) that p­ ossess a delicate strontium sulphate skeleton. Suborders Holacanthida, Chaunacanthida and Arthracanthida have multiple, small nuclei in the endoplasm. Diagnostic characters are presence/absence of a latticed skeleton and its morphology, presence/absence of apophyses (small lateral celestite extensions on spicules), internal structure of cell, presence of symbionts, and most importantly the form and central junction of spicules (Decelle et al., 2012b). Spicules in  Chaunacanthida are in a dodecahedral arrangement that can contract but Holacanthida, a basal group, only have ten spicules simply crossing the cell without any junctions. This skeleton arrangement very likely explains why encystment mainly occurs in these acantharia. Arthracanthida almost always have microalgal endosymbionts and have solid spicules that cannot retract sometimes with a thick capsule visible by light microscopy. It is not certain that the order Symphiacanthida exists in current systematics (Decelle and Not, 2015). Colour Plate 18A–G shows several examples of acantharians. Order Taxopodia have a changing heart shape and bear characteristic oar-like spicules that are used for motility. Only one species, Sticholonche zanclea, has been described to date, but environmental DNA analysis suggests that many diverse species exist. Panel 18H shows Sticholonche zanclea. Class Polycystinea are solitary or colonial cells, 20–300 µm in length, with a robust silica skeleton. Order Spumellaria are typically spherical with uniformly distributed skeletal pores but concentric internal cell structure. There may be a single spicule or several arranged in multiple or concentric arrangements. The endoplasm and ectoplasm are separated by a central capsule wall. Ectoplasm surrounded by extracapsular cortex of foamy, web-like appearance. Common genera are Centrocolla, Spongosphaera and Rhizosphaera. Order Nassellaria (from the word ‘nass’ in Italian, meaning fish basket) are smaller (100–500 µm) and tend to be ovate in shape with a silicon skeleton of various shapes that is multisegmented with one opening containing a pore of variable shape. Algal symbionts contain pores that are distributed evenly over the cell in spherical forms, or are located at poles of ovate forms. Skeleton absent or composed of siliceous material that forms a variety of shapes from spherical to beautiful elaborate latticed structures with spicules of single, multiple or concentric arrangements in Spumellaria, or tripod, ring or helmet-like structures in Nassellaria. Skeleton shape

and central capsule are the main diagnostic features. Common genera include Cladoscenium, Pterocorys and Eucyrtidium. Plate 19C shows an example of a nassellarian. Order Collodaria are round, have absent skeletons or scattered silicious spines (spicules) or irregular silica shells. They are  the only colonial radiolarians—embedded in a gelatinous sheath connected by cytoplasmic strands—a few mm up to 3 m in length! Plate 19B shows an example of a Collodaria. For a more comprehensive review of Collodaria we refer the readers to Biard et al. (2015).

6 Systematics Radiolaria were originally classified by the skeleton and the type of pseudopodia they formed and were classified under amoeboid protists (Sarcodina). The systematics of each ­r adiolarian group is under revision with the help of molecular tools. Analysis using a combination of morphological (ultrastructural) characters and phylogenetic analysis using ­conserved ribosomal genes like the 18S ribosomal DNA marker have now led to their reclassification (Burki et al., 2007). They belong to the superphylum Rhizaria that is broadly related to the Stramenopiles, Alveolates and Rhizaria (SAR) supergroup (see Introduction to taxonomy, Fig. 1). Radiolaria formerly included Phaeodaria that is now classified under Cercozoa (Polet et al., 2004), while Taxopodida, previously classified under Heliozoa, has now been found to  be a radiolarian. We present the most recent class-level classification (Adl et al., 2012) based on phylogenetic analysis  combined with taxonomy. It differs from traditional morphological classification, although there are still classification anomalies, and new studies are likely to lead to further ­taxonomic revisions.

Box 1  Classification of Radiolaria Supergroup SAR (Stramenopiles, Alveolates, Rhizaria) Superphylum Rhizaria   Phylum Radiolaria   Order Acantharia     Family Acanthometridae      Genus Acanthometron   Order Taxopodia   Class Polycystinea   Order Symphiacanthida   Order Spumellaria   Order Nassellaria   Order Collodaria     Family Thalassicollidae      Genus Thalassicola

192 ta xonom y Genus Thalassicola Solitary, large spherical cells with a granular appearance, lacking a skeleton or spicules, with a single nucleus at the centre of the intracapsulum containing oil droplets. Many foamy looking vesicles in the ectoplasm.

Ectoplasm

Central capsule Filopodia Figure 58:  Thalassicola nucleata.

Genus Acanthometron Spherical with 20 long spicules of the same thickness and length, or with two to four spicules that are longer and thicker than the others. Spicules form into a central junction. The outer shell is absent. Colourless central capsule containing algal symbionts.

Figure 59:  Acanthometron pellucida.

References Adl, S., Simpson, A. G. B., Lane, C.E., et al. (2012). The Revised Classification of Protists. Journal of Eukaryotic Microbiology 59(5), 429–93. Anderson, O. R. (1983). Radiolaria. Springer-Verlag: New York. Biard, T., Pillet, L., Decelle, J., et al. (2015). Towards an integrative morpho-molecular classification of the Collodaria (Polycystinea, Radiolaria). Protist 163, 374–88. Biard T, Stemmann L, Picheral M, Mayot N, Vandromme P, et  al. (2016). In situ imaging reveals the biomass of giant protists in the global ocean. Nature 532, 504–7. Burki, F., Shalchian-Tabrizi, K., Minge, M., et al. (2007). Phylogenomics reshuffles the eukaryotic supergroups.’ PLoS ONE 2(8), e790. Decelle, J. and Not, F. (2015). Acantharia. John Wiley and Sons: Chichester, UK.

Decelle, J., Probert, I., Bittner, L., et al. (2012a). An original mode of symbiosis in open ocean plankton. Proceedings of the National Academy of Sciences 109(44), 18000–5. Decelle, J., Suzuki, N., Mahé, F., et al. (2012b). Molecular phylogeny and evolutionary history of the Acantharia (Radiolaria). Protist 163, 435–50. Decelle, J., Martin, P., Paborstava, K., et al. (2013). Diversity, ecology and biogeochemistry of cyst-forming Acantharia (Radiolaria) in the oceans. PLoS one 8(1), e53598. Dolven, J. K., K. R. Bjørklund, et al. (2009). Polycystinea: Polycystine radiolarians. http://tolweb.org/Polycystina/ 121189 Accessed 22 February 2016. Ishitani, Y., Febvre-Chevalier, C., Febvre, J. (2016). Radiolaria. Encyclopedia of Life Science. John Wiley and Sons: Chichester, UK.

protozoopla nkton: r a diola r i a  193 Jahn, T., Bovee, E. C. et al. (1949). How to know the Protozoa. WCB/McGraw-Hill: Dubuque, Iowa. Japanese Association for Marine Biology (2015). Sticholonche zanclea. Jambio: Regionally Integrated Marine Database (RINKAI). Available from https://www.shimoda.tsukuba. ac.jp/~marinelife-db/?marinelifedata=sticholonche-zanclea (accessed 10 March 2016). Kling, S. A. and Boltovskoy, D. (2002). What are Radiolarians? Available from Radiolaria.org. (accessed 22 February 2016). Lazarus, D. (2005). A brief review of radiolarian research. Paläeontologische Zeitschrift 79(1), 183–200. Lenz, J. (2000). Introduction. In Harris, R. Wiebe, P., Lenz,  J.,  Skjoldad, H.R., Huntley, M. (eds) ICES Zooplankton  Methodology Manual, pp. 1–30. Academic Press: London. Polet, S., Berney, C., Fahrni, J., Pawlowski, J. (2004). Smallsubunit ribosomal RNA gene sequences of Phaeodaria challenge the monophyly of Haeckel's Radiolaria. Protist 155(1), 53–63.

Probert, I., Siano, R., Poirier, C. et al. (2014). Brandtodinium gen. nov. and B. nutricula comb. Nov. (Dinophyceae), a dinoflagellate commonly found in symbiosis with polycystine radiolarians. Journal of Phycology 50(2), 388–99. Savalli, U. M. (2015). Radiolarians, phylum Radiolaria. Arizona State University: Tempe, AZ. Suzuki, N. and Aita, Y. (2011). Radiolaria: achievements and unresolved issues: taxonomy and cytology. Plankton & Benthos Research 6(2), 69–91. Suzuki, N. and Not, F. (2015a). Biology and ecology of Radiolaria. In Ohtsuka, S. Suzaki, T., Horiguchi, T., Suzuki, N., Not, F. (eds) Marine protists: Diversity and dynamics, pp. 179–222. Springer Japan: Tokyo. Suzuki, N. and Not, F. (2015b). Radiolaria. In Ohtsuka, Suzaki, S. Horiguchi, T., Suzuki, N., Not, F. (eds) Marine protists: diversity and dynamics. Springer Japan: Tokyo. Webber, M. (2009). Acanthochiasma fusiforme BioSearch: Marine Biodiversity Database of India. Available from http://www. biosearch.in/ (accessed 10 March 2016).

Protozooplankton: For aminifer a Row ena Ster n, Cla ir e Tay lor, a nd Sa eed Sa dr i 1 Introduction Foraminifera are composed of a well-hidden, amoeboid body commonly enclosed within a test (shell) that ranges from 0.9  mm to several mm in size. They are one of the most common shelled marine organisms and date back to the Cambrian era where they are responsible for the colour of the sediment on some shorelines. About 7500 living species are estimated (Appeltans et al.,  2012), but genetic analysis suggest greater diversity. The only visible part of the body are the filopodia – a thread-like cytoplasmic processes forming a branching, joined (anastomizing) network (reticulopodia) that extends out from the test and is used for locomotion and feeding. Filopodia are the main point of contact between the organism and its environment, and it is involved in most of the life processes such as respiration, attachment or movement, feeding, test construction and reproduction (Murray,  2006). This section only briefly touches on the diversity of this group, and we refer readers to Neil et al. (2005) and a review by Darling and Wade (2008).

2  Life Cycle Foraminifera alternate between asexual agamonts and sexually reproducing gamonts (see Fig. 61). The young agamont zygote develops into a spherical shaped first chamber and as a juvenile, the cytoplasm grows and additional chambers emerge into a spiral. Only smaller phytoplankton are consumed at this stage. The chambers develop and the aperture moves into the correct position. Pores develop all over the test and adult characteristics begin to materialize. Large prey such as zooplankton can now be consumed. The adult diploid agamont has established test patterns and any ­secondary apertures now develop. Additionally, spines are shed and the test wall thickens with calcite – this is termed gametogenic thickening. All of the cytoplasm is transformed during gamete production, leaving just an empty test. Haploid flagellated young are released from the test, which can grow into a haploid gamont that can reproduce asexually or return to the diploid through the release of gametes following meiosis.

Figure 60:  Photo of the Foraminiferan, Amphistegina lessonii, showing chambers (top panel) and filopodia (bottom panel). Photo courtesy of G. Langer (Marine Biological Association; 2016).

3  Ecology and Distribution Foraminiferans are widely distributed throughout the world’s oceans, although they do not inhabit enclosed water bodies and are rarely found in shelf areas. They account for 20% of carbonate formed globally, and play an important role in carbon cycling (Neil et al., 2005), although only 1% of foraminifera are

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

protozoopla nkton: for a minifer a  195 Diploid agamont (microsphaeric)

Young agamont (diploid zygote)

gamont formation Sexual haploid young

gametes

young gamont Asexual haploid gamont (megalosphaeric) Figure 61:  Generalized life-cycle of Foraminifera. Adapted from Laybourn-Parry (1984).

planktonic (Campbell, 2013). The distribution and diversity of planktonic species varies according to climate, current systems and seasonal upwelling, water properties and food availability, and there are species-specific preferences. Most are benthic but some are planktonic – mostly from the order Globigerinida, for example Globigerina bulloides, Turborotalita quinqueloba (syn. Globigerina quinqueloba), Globorotalia inflata (syn. Globigerina inflata) and Neogloboquadrina pachyderma (syn. Globigerina pachyderma). Some benthic species, although rare, will also have stages of their life cycle in the plankton. Recent studies also suggest that vertical migration may also occur. Foraminifera

c­ ontribute significantly to Carbon drawdown and nearly 70% of sea-floor sediment is composed of foraminifera. Planktonic foraminiferans extend their filopodia into networks to feed on other protists such as diatoms, while the benthic organisms consume detritus material and can burrow with their filopodia (Campbell, 2013). Studies have also shown that they can be parasitic on each other. Foraminifera are preyed upon by shrimps, molluscs, worms, gastropods, fish and echinoderms. Some Foraminifera species host symbionts  such as pigmented dark red dinoflagellates, which may increase their overall size and alter the colour of the body to brown-red. Foraminifera have a variety of test forms which hold the body (see Fig. 62). Reticulopodia (a type of filopodia), used for feeding, can be observed coming out of the test in live samples. Within the test there is a single opening or aperture. The aperture allows the flow of cytoplasm between chambers. In the majority of genera there are several chambers (locula) that change as the organism grows (Fig. 62A). Chambers are ­connected by large pores (foramina) from the oldest chamber (proloculus) to the newest, which is the aperture. The test is actually internal and covered in a thin layer of cytoplasm, and the form and number of chambers of the test are the principal features used for classification. Athalamids have no shell or a single-­ chamber shell (commonly called unilocular). Polythalamous forms make shells composed of chambers added in a coil like a snail shell (also called trochospiral) as in Figure 62A, but can be other forms (Figs 62B, C) or globular (Fig. 64). The test surface is commonly composed of calcareous material of clear or opaline appearance, but planktonic forms mostly have a rough, opaque texture as they are composed of agglutinated material such as calcite, aragonite and silica bound into a fibrous matrix (e.g. Leptohalysis and Globigerina bulloides).

A

B

C

Aperture

21 20

3 12

19 11 18 . . .. . . . . . . .. . . .. Perforations

10

5 6 4 P 3 2 7 9

17

13

8

14

15 16

2 Chamber Foraminaseparating chambers

P

1

2

3

1

P

Figure 62:  Representations of Foraminifera. A–C, Morphological types. A, Spiral (e.g. Elphidium sp.); B, leaf-like (e.g Frondicularia sp.). Note reticulopodia in A. C, Linear. Diagram adapted from Jahn et al. (1949).

196 ta xonom y

4  How to Identify Foraminifera

helically as it grows). Occasionally the tests are tri-, bi- or uniserial. Globular or c­ rescent-shaped chambers are observed in juveniles.

4.1  Class Monothalamea These are early lineage forms that either lack a test or have a single-­ chambered test (round/ovate) with an organic or agglutinated wall that has a smooth, transparent orange to tan appearance. This class possesses a distinct collar in aperture and reticulopodia extend 0.5–2 mm around the test. Two chambered forms also group with s­ ingle-chambered forms. This class requires revision.

5 Systematics Foraminifera belong to the superphylum Rhizaria, which is broadly related to the Stramenopiles, Alveolates and Rhizaria supergroup. Foraminifera have now been classified according to

4.2  Class Tubothalamea

Box 1  Classification of Foraminifera

Bi- or multi-chambered test with tubular chambers at least in juvenile stages. Calcareous and agglutinated test.

Supergroup Stramenopiles, Alveolates, Rhizaria Superphylum Rhizaria Phylum Foraminifera Class Monothalamea Class Tubothalamea   Order Milliolida   Order Spirillinida Class Globothalamea   Order Globigerinida   Family Globigerinidae    Genus Globigerina   Order Textulariida   Family Reophacidae    Genus Leptohalysis

Order Milliolida: Agglutinated or calcareous test with high magnesium content. Test of porcelain appearance (shiny, smooth, white) due to agglutinized crystal reflectance. The wall commonly lacks perforations. This group contains tubular or elongate chamber walls, coiled in a disc-shaped spiral (planispiral), with internal structures to accommodate endosymbionts. Order Spirillinida: These possess a low-magnesium calcite test. The proloculus test has a rolled tubular chamber. Members of this order have no or few chamber divisions per whorl.

4.3  Class Globothalamea Members possess multi-chambered agglutinated or calcareous test, generally t­rochospirally enrolled (spiral that is extended

Taxa in bold type are described in this section as common planktonic Atlantic taxa.

Order Textulariida Leptohalysis scotti (syn. Reophax scotti) (Chaster, 1892) Dimensions: 800–1000 µm in length Distribution: NECS, SARC, ARCT, NWCS, NADR, NASE, GFST. Coastal, nearshore species P 1 2

These have agglutinated tests with foreign particles attached to the organic lining or low-magnesium calcite. The type of test material (organic with particles or magnesium calcite) is a taxonomic feature. Two families are not taxonomically resolved. This order is also identified by the presence and type of perforations. The family Carterinidae have calcareous spicules. Members can be spiral in form but the genus Leptohalysis has a linear arrangement of thin, angular test chambers increasing in size from the globular proloculus (Holzmann, 2010; Camacho et al., 2015). The cell wall is thin and flexible. Note: nine species found in the North Atlantic

3 4 5

6

Figure 63:  Leptohalysis scotti showing poloculus (P) and sequential tests growths from it, numbered 1–6.

protozoopla nkton: for a minifer a  197 phylogenetics, and in this chapter we use the classification system described by Adl et al. (2012) to describe higher taxon groups and the World Register of Marine Species (WoRMS) database for lower taxonomic classifications. Unlike earlier taxonomic c­ lassifications,

phylogenetics classify Foraminifera into three classes that relate to the number of chambers in their tests and forms. There is some complexity in how the Textulariida and Globigerinida orders relate to the common ancestor of foraminiferans.

Order Globigerinida Members possess a test of low-magnesium calcite, bilamellar, some covered in fine, elongated spines (spine-bearing forms are termed spinose) and with large (macro) or small (micro) perforations. Globigerina chambers are round, of different sizes and are clustered together or spiral-shaped, but without the aperture, and they range from 100 µm to 1000 µm in diameter ( Johnson and Allen, 2012). Three morphotypes exist that correspond with phylogenetic clades: Heterohelicoidea (micro-perforate non-spinose), Globorotaloidea (macro-perforate non-spinose) and Globigerinoidea (spinose) (Darling and Wade, 2008). The tests of G. bulloides are covered in agglutinated minerals. Note: 47 morphospecies found in the North Atlantic (Darling and Wade, 2008)

Figure 64:  Globigerina bulloides.

References Adl, S., Simpson, A. G. B., Lane, C.E., et al. (2012). The Revised Classification of Protists. Journal of Eukaryotic Microbiolgy 59(5), 429–93. Appeltans, W., Ahyong, S. T., Anderson, G., et al. (2012). The Magnitude of Global Marine Species Diversity. Current Biology 22(23), 2189–202. Camacho, S.G., de Jesus Moura, D.M., Conner, S., Scott, D.B., and Boski, T. (2015). Taxonomy, ecology and biogeographical trends of dominant benthic foraminifera species from an Atlantic-Mediterranean estuary (the Guadiana, southeast Portugal). Palaeontologica Electronica 18(1.17A), 1–27. Campbell, D. (2013). Foraminifera, Brief Summary, 2013 ed. Encyclopedia of Life. Darling, K.F. and Wade, C.M. (2008). The genetic diversity of planktic foraminifera and the global distribution of ribosomal RNA genotypes. Marine Micropaleontology 67(3–4), 216–38. Holzmann, M. (2010). ForamBarcoding: Molecular Database of Foraminifera. University of Geneva, Geneva, Swizerland. Available from http://forambarcoding.unige.ch/ (accessed 11 November 2016).

Jahn, T., Bovee, E.C., and Jahn, F. (1949). How to know the Protozoa, 2 ed. WCB/McGraw-Hill: Dubuque, Iowa. Johnson, W.S. and Allen, D.M. (2012). Protozooplankton, 2nd ed. John Hopkins University Press, Baltimore, USA. Laybourn-Parry, J. (1984). A functional biology of free living protozoa. Springer, USA. Murray, J. (2006). Ecology and applications of benthic foraminifera. Cambridge University Press, Cambridge, New York, Melbourne. Neil, H., Cooke, P. and Northcote, L. (2005). The life and death of planktonic foraminifera. Water & Atmosphere 13(1), 18–19. Pearson, P. (2013). Planktonic Foraminifera. University of Cardiff. Available from https://paleonerdish.wordpress. com/2013/09/16/to-see-the-world-in-a-grain-of-sandplanktonic-foraminifera-and-evolution/ (accessed 31 March 2016). Savalli, U.M. (2015). Radiolarians, phylum Radiolaria. Arizona State University, Tempe, USA.

CNIDARIA: SCYPHOZOA AND NON-COLONIAL HYDROZOA Pr iscilla Lica ndro, Astr id Fischer, a nd Dhuga l J. Lindsay 1 Introduction The Phylum Cnidaria (in old classification grouped with Porifera and Ctenophora as Coelenterata) assembles highly diverse primitive invertebrates that carry stinging cells called cnida. The presence of cnida, which are organized in specialized structures called cnidocysts (or nematocysts), makes the organisms of this group venomous to varying degrees. Cnidarians are aquatic, mostly marine invertebrates, that are characterized by a great variety of forms and sizes; they may live as free-living plankton or settle as sessile generally benthic forms, being either solitary or colonial. They are ubiquitous, occurring at all latitudes and depths. Cnidarian populations are characterized by a regular seasonal cycle, but they also undergo interannual fluctuations, with much larger populations in some years than in others. Due to the plasticity of their physiological response, cnidarians may quickly achieve extremely high abundances (up to ten– hundred individuals per cubic metre of water) with favourable environmental conditions (Mills, 2001; Purcell et al., 2007; Boero et al., 2008). During such events, called outbreaks, swarms of Cnidaria can be spread over large regions of the sea (Pitt and Lucas, 2014). This chapter is focused on pelagic non-colonial cnidarians, while siphonophores, which are pelagic colonial hydrozoans, have been described in a separate chapter.

2  Life cycle Cnidaria Medusozoa are typically dimorphic, i.e. they may assume completely different morphologies at different stages of their life cycle (Bouillon et al. 2006) (Fig. 66). In meroplanktonic species (Fig. 66A) the ciliated free-swimming early larva (planula) ­metamorphoses into a generally benthic polyp (or hydroid) that in turn produces by budding several to many free-living ­immature medusae (called ephyra in Scyphozoa or actinula in some Hydrozoa). Polyps can be ‘thecate’, i.e. surrounded by a ­chitinous perisarc that may extend around hydranths (­hydrotheca), reproductive organs (gonotheca), and dactylozooids (dactylotheca), which serve the colony for both defense and food capture. Normally the polyp and the medusa are, respectively, the asexual and sexual stages.

FIGURE 65: Hydrozoa Anthoathecata Bougainvillia muscus. Photo

courtesy of C. Carré.

In holoplanktonic species (Fig. 66B) the planula directly develops in an ephyra or actinula-like stage. Sexual reproduction is usually external, with male and female medusae releasing sperm and eggs into the water where the fertilization occurs. It is only in some Cubomedusae that the fertilization is mainly internal, with the male using the manubrium to directly transfer its spermatophore (i.e. packed sperm) to a tentacle of the female that, in turn, transfers it into the gastric cavity (Lewis and Long,  2005). Some Hydromedusae may reproduce asexually, either by direct budding of young medusae or by longitudinal fission. In most Scyphozoa and Cubozoa the medusa phase is shorter than the polyp phase. Cnidaria are able to adjust their life cycle entering into dormancy to survive when resources are limiting; with favourable environmental conditions the resting stage or cyst will hatch, releasing a larva that will restart the reproductive cycle, leading to a new peak in the population.

3 Ecology Cnidarians are essentially carnivorous and, depending on their size, can eat a variety of prey (Purcell, 1991; Purcell and Arai, 2001). Hydrozoans eat mainly copepods but also ostracods, molluscs, chaetognaths, euphausiids, and early fish larvae and eggs, with the exception of the Narcomedusae, which prey on

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  199 A

medusa

fertilised egg

young medusa polyp

planula

stolon

B fertilised egg

planula developing ephyra

ephyra

medusa FIGURE 66:  Life cycle of Cnidaria. Examples of life cycles in mero­

planktonic Anthomedusae (A) and holoplanktonic Scyphomedusae (B). After Naumov (1960).

other gelatinous organisms. Big Scyphozoa can also prey on smaller cnidarians and small fish. There a few examples of Cnidaria (e.g. Cassiopea and Mastigias) hosting symbiotic zooxanthelle algae, which can take up dissolved organic matter. Although different species may peak at different times of the year, generally Cnidaria tend to be more abundant between spring and autumn. High densities of medusae with calm seas and no wind tend to remain aggregated in swarms. Meroplanktonic Cnidaria are mainly distributed in coastal waters. On the contrary, holoplanktonic medusae can be distributed inshore and offshore. In medusae the rhythmic contraction of the umbrella empties the water from beneath the subumbrella and allows the animal to move by jet-propulsion. Cnidarians are able to adjust their buoyancy through osmoregulation to some degree. Due to the difficulty in their cultivation, basic information on their physiology and environmental preferences are not yet available for most cnidarian species.

4  General morphology The main morphological traits of the three pelagic groups of Medusozoa and of the main orders of Hydrozoa are summa-

rized in Tables 1 and 2. Pelagic cnidarians are characterized by a relatively simple structure (Fig. 67) consisting of an ectodermal epidermis and an endodermal gastrodermis separated by gelatinous, mainly acellular, mesoglea. Free-swimming medusae are mainly characterized by a tetramerous radial symmetry, typically bell- or lens-shaped, with a swimming bell (umbrella) of variable thickness and height The outer aboral surface of the umbrella (exumbrella) of ectodermal origin is usually convex while the inner oral surface (subumbrella) is concave and delimits the internal gastrovascular cavity (or coelenteron) that serves for circulation, as well as digestion and distribution of food. Most Hydromedusae have a velum, i.e. a thin membrane of ectoderma and mesoglea that, within the bell margin, partially closes the space under the subumbrella. The velum is absent in the Scyphomedusae, whereas in the Cubomedusae it is replaced by a velarium, which has a similar function but different origin. The subumbrella is extended in an appendix called a manubrium containing the gastric cavity, which in some species is mounted upon a gelatinous gastric peduncle; the mouth, which often carries lobes called lips, is either suspended at the end of the manubrium or in the centre of the subumbrella. Manubria greatly vary in shape and size. In some species of Hydrozoa (e.g. Leuckartiara octona) the manubrium is perradially attached to the subumbrella through a layer of tissue called mesentery. In most Scyphomedusae the manubrium is tubular and ends in four elongated structures, i.e. oral arms, which in some Rhizostomeae are covered by strapped structures named epaulettes (or scapulettes) (Fig. 68N). In Hydromedusae the coelenteron ramifies radially along the bell to the margin into radial canals, which in most Hydromedusae join a ring canal lying within the rim of the bell. In some species radial canals are branched in other centripetal canals (Fig. 68K) that rise upwards from the circular canal without reaching the gastrovascular cavity. In Scyphomedusae and Cubomedusae, the coelenteron is divided by four longitudinal oral–aboral mesenteries and the pattern of radial canals is often complex. As in the Hydromedusae, the four mouth lips are perradial and there are four interradii and eight adradii. In some Hydrozoa (e.g. Narcomedusae) and in the Scyphomedusae the umbrella margin is interrupted by notches forming separate marginal lobes called lappets (Fig.  67E). In Hydromedusae and  Scyphomedusae, tentacles armed with nematocysts, sometimes grouped in knobs or specialized capsules (i.e. ­cnidophores) (Figs 68G and I) are typically located at the ­margin of the umbrella (marginal tentacles) and sometimes around the mouth (oral or gastric tentacles). In some Hydromedusae (e.g. Narcomedusae) they are inserted on the exumbrella. Tentacles and similar organs called cirri (Fig. 68A,B,F) that are  placed between marginal tentacles, greatly vary in shapes and numbers. The margin of the bell may be provided with specialized sense organs (Figs 67A,B,F and 68A–E). Those include: (i) cordyli, minute club-shaped structures situated between the tentacles; (ii) gravity receptors containing one or  more statolyths called statocysts; and (iii) rhopalia, i.e.  club-shaped sensory centres, each containing a statocyst,

200 ta xonom y A HYDROZOA - Trachymedusae

B SCYPHOZOA - Semaeostomeae

Aboral Pole

GO RC

Ex_U M

Sub_U

RC MO

GO ISP CC Sub_U OC V

T_BU marginal T

CC

RO

Oral arms

Oral Pole

D SCYPHOZOA - Coronatae

C HYDROZOA- Narcomedusae

RS

CG

MAP

GO CM LAP

Nausithoe globifera STA

marginal T

T

E HYDROZOA - Narcomedusae

Solmundaegina nematophora

CC LAP

F CUBOZOA - Carybdeida MO

MAP

M Solmissus incisa GO

PE RO Carybdea alata

marginal T

Key to annotations CC CG CM GO ISP LAP M MAP MO

= Circular Canals = Coronal Groove = Coronal Muscle = Gonad = Interradial Space = Lappet = Manubrium = Manubrial Pouches = Mouth

OC PE RC RO RS STA T T_BU U V

= Ocellus/ocelli = Pedalia = Radial canals = Rhopalium = Radial Septa = Statocyst = Tentacle/s = Tentacle Bulb = Umbrella = Velum

FIGURE 67:  General morphology of Cnidarian medusae. A, Hydrozoa Trachymedusae, lateral view. B, Scyphozoa Semaeostomeae, lateral

view. C, Hydrozoa Narcomedusae Solmundaegina nematophora, lateral view. D, Scyphozoa Coronatae Nausithoe globifera, ventral view. E, Hydrozoa Narcomedusae Solmissus incisa, ventral view. F, Cubozoa Carybdeida Carybdea alata, lateral view. A, Modified from Trégouboff and Rose (1957); B, modified from Naumov (1960); C, Lindsay et al. (in press); D and E, Russell (1970); E, edited from Fewkes, 1886; F, © Corbera in Pagès et al. (1992).

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  201 TABLE 1:   Main morphological traits and life cycles in Medusozoa. Pelagia noctiluca after Mayer (1910); Carybdea marsupialis after © Corbera, in Mianzan & Cornelius (1999); Liriope tetraphylla after Russell (1953).

Known Species

Scyphozoa

Cubozoa

‘true jelly’

‘sea wasps and box jelly’

Pelagia noctiluca

Carybdea marsupialis

200

3700

Fairly small (up to 25 cm tall); cubic With velarium: colourless Lower edge of umbrella extended in a ‘pedalium’ that supports tentacles

Mostly small (up to 10 cm in diameter) though siphonophore colonies can reach 40m length With velum: transparent

Divided by four longitudinal oral-aboral septa

Undivided

Endodermal origin

Ectodermal origin, if present

Strobilation if not holoplanktonic, i.e. transverse fission of the polyp (scyphistoma)

Polyp directly metamorphoses into one medusa

Asexual budding, sometimes by longitudinal fission

External

Internal

External

Usually small and simple

Usually colonial, with individual specialized polyps. Male and female colonies are usually separated

saucer-shaped or hemispherical (except Coronatae) Without velum: distinct pigmentation and thick mesoglea

Sense organs

Origin

Reproduction

Liriope tetraphylla

45

MEDUSA (sexual pelagic stage) Umbrella Big (up to 2 m in diameter);

Gastrovascular cavity

Hydrozoa

Divided by four mesenteries Endodermal origin. Eight aradial (Coronatae) and four interradial (Semaeostomeae and Rhizostomeae)

POLYP (asexual benthic stage) Structure

Usually small and simple, mainly solitary. Sometimes absent (e.g. in Pelagia, Atolla, and Peryphylla)

epidermal neurons, a pair of sensory pits and often a photoreceptor ­(ocellus). The number and shape of statocysts is often used to separate different groups of Hydromedusae: ectodermal statocysts, open or closed, are small marginal pockets or open pits that develop in the velum and can be found in the Antho/Leptomedusae (Fig. 68A,B,D); endodermal statocysts are sensory clubs, small tentacle-like structures hanging freely out of the umbrella margin or enclosed in the umbrella (Fig. 68C,D), that characterize Limnomedusae, Trachymedusae, and Narcomedusae. The shape of the manubria, types of ­tentacles (Fig. 68M), position of the gonads, as well as the presence of distinctive canals (Fig. 68K–L) and of other structural characters, such as cirri, otoporpae, warts, and gelatinous papillae (Fig. 68A,B and F–J) are also used for taxonomic ­identification.

5 Systematics The classification of the Cnidaria has seen many changes over the years, with recent updates following the advent of molecular phylogenetic analyses. Here we adopt the classifications proposed by Schuchert (2012), Daly et al. (2007), and Mianzan and Cornelius (1999) that revised previous classifications from Bouillon et al (2006) and Kramp (1959a and 1961). As the focus of this book is the North Atlantic plankton, the taxonomic list and keys are limited to the families of cnidarian medusae. Key morphological features of the main medusa species and of the planktonic hydroid colony of the Porpitidae Velella velella are also included. Considering the recent changes in their systematics (Lindsay et al., in press), the taxonomic key of hydrozoan Narcomedusae has been detailed up to the genus level.

202 ta xonom y TABLE 2:   Main morphological traits in non-colonial Hydrozoa. Sarsia tubulosa after Naumov (1960); Lovenella clausa after Russell (1963); Rhopalonema velatum after Mayer (1910); Solmaris corona after Pagès et al. (1992).

Anthomedusae Sarsia tubulosa

Umbrella

Bell-shaped, higher/ as high than wide

Circular canals Yes. Can have centripetal

Leptomedusae

Trachymedusae

Narcomedusae

Lovenella clausa

Rhopalonema velatum

Solmaris corona

Lens-shaped, wider/ as wide than high

Hemispherical, quite high

Flat with lobes

canals (e.g. Timoides)

Yes

Yes. Can have centripetal canals

Absent or looped into the marginal lappets

Yes

Yes

Yes

No

On/around manubrium, might be attached to subumbrella by mesenteries

On radial canals, sometimes extending into manubrium

On radial canals

On manubrium and/or on manubrial pouches

Marginal tentacles

Hollow/solid, mainly tentacular bulbs

Usually hollow, with tentacular bulbs

Sense organs

No statocysts/cordyli, might have ocelli

Might have ectodermal statocysts, cordyli, and/or ocelli

Radial canals Position of gonads

Solid or solid and hollow, If present, solid. Primary tentacles with no real tentacular bulbs on exumbrella No tentacular bulbs, only solid roots Endodermal statocysts, generally free but also enclosed. No ocelli/cordyli

Mostly free ecto-/endodermal statocyts. Can have otoporpae. No ocelli or cordyli

Life cycle

Meroplanktonic

Meroplanktonic

Holoplanktonic

Holoplanktonic

Polyp

Athecata polyp

Thecata polyp

No polyp stage

No polyp stage but can have parasitic stolon phase

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  203

A

B

STRUCTURAL CHARACTERS AND SENSE ORGANS OPEN D C T_BU

T_BU

lateral CI

marginal WA

CC

open STA CO

E

ECTODERMAL

closed STA

CLOSED

OC

F

T_BU

CO-like structure

open STA

marginal CI

T

G

CLOSED

ENDODERMAL

OPEN

H PA

K

GO

L

AD-

R

PER-R

J

OT

IN

TE

I

R-

R

N_KN

CEC

PER-R

AD-

R

CNI

CC

T

GAP PERO

M

M

CC

Key to annotations N = Nematocysts AD-R = Adradial N_KN = Nematocyst knobs BU = Bulb OC = Ocellus/ocelli CC = Circular canal (peripheral canal in Narcomedusae) OT = Otoporpae CEC = Centripetal canal PA = Gelatinous papilla/ae CI = Cirri PERO = Peronia CNI = Cnidophores PER-R = Perradial CO = Cordylus/cordyli RO = Ropalium GAP = Gastric peduncle SCA = Scapulette/s GO = Gonad STA = Statocyst/s INTER-R = Interradial T = Tentacle/s M = Manubrium WA = Warts

N

FIGURE 68:  Structural characteristics of cnidarian medusa. Part of the marginal bell of A, Eutima coerulea; B, Eucheilota maculata; C, Laodicea

spp.; E, Parateclaia norvegica; and F, Cosmetira pilosella. D, Types of statocysts; G, Turritopsis nutricula, mouth and detail of mouth margin with nematocyst knobs; H, Halicreas minimum, detail of umbrella and of gelatinous papilla; I, Zanclea costata, lateral view and detail of cnidophores; J, Pegantha rubiginosa, marginal lappets; K, Liriope tetraphylla, some details; L, diagram to define the radii of a medusa; M, Obelia spp., portion of umbrella margin, showing bases of solid marginal tentacles and their endodermal roots; N, Rhizostoma pulmo, scapulette. All lateral views. A, K and M, Mayer (1910); B, D, F, G, I, and L, Russell (1953); C, Kramp (1919); E, Hosia and Pagès (2007); H (detail of umbrella), Vanhöffen (1902); H (detail of gelatinous papilla), Bigelow (1909); J, Kramp (1959a); N, Russell (1970).

TABLE 3:   Outline classification of marine cnidarian medusa. Families in bold are those treated in more detail.

Phylum Cnidaria

Orders

Suborders and families

Carybdeida

Alatinidae; Carukiidae; Carybdeidae; Tamoyidae; Tripedaliidae

Chirodropida

Chirodropidae; Chiropsalmidae; Chiropsellidae

Trachymedusae

Geryoniidae; Halicreatidae; Petasidae; Ptychogastriidae; Rhopalonematidae

Narcomedusae

Aeginidae; Cuninidae; Solmarisidae; Solmundaeginidae; Pseudaeginidae

Leptothecata

Aequoreidae; Barcinidae; Blackfordiidae; Campanulariidae; Cirrholoveniidae; Dipleurosomatidae; Eirenidae; Hebellidae; Laodiceidae; Lovenellidae; Malagazziidae; Melicertidae; Mitrocomidae; Orchisotomatidae; Octocannoidae; Phialellidae; Phialuciidae; Sugiuridae; Teclaiidae; Tiarannidae; Tiaropsidae

Siphonophorae*

* group described in separate chapter

Anthoathecata

[Suborder: Aplanulata] Tubulariidae; Corymorphidae; Margelopsidae; Aplanulata incerta sedis

Class: Cubozoa

Class: Hydrozoa Subclass: Trachylinae

Subclass: Hydroidolinae

[Suborder: Capitata sensu stricto] Moerisiidae; Cladonematidae; Corynidae; Porpitidae; Cladocorynidae; Zancleidae [Suborder: Filifera] Oceaniidae; Bougainvilliidae; Rathkeidae; Pandeidae; Bythotiaridae; Hydractiniidae; Proboscidactylidae; Ptilocodiidae; Eucodoniidae; Protiaridae; Trichydridae Class: Scyphozoa

Subclass: Discomedusae

Coronatae

Atollidae ; Atorellidae; Linuchidae; Nausithoidae; Paraphyllinidae; Periphyllidae

Semaeostomae

Cyaneidae; Drymonematidae; Pelagiidae; Ulmaridae

Rhizostomeae

[Suborder: Kolpophorae] Cassiopeidae; Cepheidae; Mastigiidae; Thysanostomatidae; Versurigidae [Suborder: Daktyliophorae] Lychnorhizidae; Catostylidae; Lobonematidae; Rhizostomatidae ; Stomolophidae

TRACHYMEDUSAE

Yes

T with adhesive disk (1)

numerous T arranged in groups, both along Yes and above U rim. M broad with 8 radial lobes, without GAP. With CEC

PTYCHOGASTRIIDAE

1 Ptychogastria asteroides

No

Yes with well developed CEC (2)

generally with long GAP (3)

GERYONIIDAE

Yes GAP Liriope tetraphylla 4

Yes

4 RC

Yes

T with terminal club-shaped N_KN (3), pendant STA (4)

Petasus atavus

PETASIDAE

3

5

No

8 (rarely more) RC

Yes

M and RC wide (5), T with stiff distal tips

Yes

HALICREATIDAE

Halicreas minimum 6

Yes

M and RC both narrow (6)

Yes

RHOPALONEMATIDAE Rhopalonema velatum

FIGURE 69:  Trachymedusae: Ptychogastria asteroides, whole figure and detail of umbrella margin (1), after Gili et al. (1999); Liriope tetraphylla, whole figure and detail of umbrella

(2), after Trégouboff and Rose (1957); Petasus atavus and Rhopalonema velatum after Mayer (1910); Halicreas minimum after Vanhöffen (1902); H. minimum, ventral view (5), after Kramp (1947); Pantachogon haeckeli, ventral view (6), after Russell (1953). All lateral views, unless noted otherwise.  

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  205

2

No

1

No

U bell-shape

Vermiform shape, 4 swimming flaps (1)

TETRAPLATIIDAE

Yes

Yes

206 ta xonom y

NARCOMEDUSAE GROUP I

Genus Tetraplatia

T. volitans SOLMARISIDAE No

with MAP (2)

No MAP. GO on manubrial wall/ diverticula. Numerous T inserted into ex_U, at level of M attachment to sub_U (3)

Yes

Yes

Without peripheral canal (here CC) and without OT

3 S. flavescens

Genus Solmaris

No

Yes

Solmaris flavescens

Yes

With CC and OT

P. rubiginosa

Genus Pegantha

CUNINIDAE MAP (2) PER-R and undivided. Primary T PER-R between marginal LAP, with origin on centre of MAP (4) 2

Yes

With secondary T (5)

Yes

With CC, without OT

Genus Sigiweddellia

S. bathypelagica

No

5

Cunina fowleri Without CC, without OT

Yes

Genus Solmissus S. incisa

4

NARCOMEDUSAE GROUP II

With/without CC, with OT

Genus Cunina

Yes C. globosa

FIGURE 70:  Narcomedusae group I: Tetraplatia volitans after Russell (1970); Solmaris flavescens after Mayer (1910); S. flavescens, ventral view, after Russell (1953); Pegantha rubiginosa after Pagès et al. (1992); Sigiweddellia bathypelagica after Bouillon et al. (2001); Solmissus incisa, ventral view, modified after Kramp (1959a); Cunina globosa after Bigelow (1909); Cunina fowleri, ventral view, after Browne (1906); All lateral views, unless noted otherwise.  

NARCOMEDUSAE GROUP II AEGINIDAE

MAP INTER-R, divided in two parts (1). Primary T are PER-R between marginal LAP, with origin above MAP (2)

Yes

Yes

with secondary T (3)

2

Yes

with 8 primary T

Genus Aeginura

Yes 1

with 4 primary T

A. grimaldii

Yes

No Genus Bathykorus

B. bouilloni 3 Aeginura grimaldii Yes

with 32 MAP

Genus Aeginodiscus*

Yes

< 32 MAP

without CC

with orallypointing T roots

Yes

Yes

with 2 T

Genus Solmundella

Yes S. bitentaculata

No with CC

8 T and OT

Yes

Genus Otoporpa

Yes

T roots apically-pointing

with 4 T and 8 MAP Yes

Genus Tetraotoporpa*

Yes

T roots orally-pointing

Yes

Genus Aeginopsis A. laurentii

O. polystriata 4 T and OT

Yes

with 4 T and 16 MAP

Yes

Genus Pseudaegina* (new Genus, type-species P. rhodina)

Genus Aegina A. citrea

with N clusters on external side of U. exumbrellar grooves above T

without N clusters on external side of U. no exumbrellar grooves above T

Genus Solmundaegina* (new Genus)

Yes

Yes

Genus Aeginona* (new Genus, type-species A. brunnea)

(*Figure in Lindsay et al., 2017)

FIGURE 71:  Narcomedusae group II: Aeginura grimaldii, ventral (1) and lateral views, after Maas (1905); Bathykorus bouilloni after Wang et al. (2014); Solmundella bitentaculata and Aeginopsis laurentii after Mayer (1910); Otoporpa polystriata after Xu and Zhang (1978); Aegina citrea after Lindsay et al. (2017). All lateral views, unless noted otherwise.  

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  207

No

LEPTOMEDUSAE GROUP I - Order: LEPTOTHECATA up to 6 M (1), no CEC (2)

1

Yes

2

SUGIURIDAE

Yes

with STA (3) or CO (4)

Sugiura chengshanense No

GAP broad; ≥ 8 RC; many filiform, solid tentaculiform arms without marginal BU unconnected to CC

Yes

with GAP (5)

Yes

No

Yes

ORCHISTOMATIDAE

Orchistoma pileus Yes

base of M attached over its whole surface (6)

RC simple or bifurcated

6

Yes

MELICERTIDAE

7 Melicertum octocostatum

No

5

base of M narrow, RC branched (7) or, if simple, irrregularly arranged

Yes

Cuvieria carisochroma

9 CO (4) or CO-like structures (8) No

Yes

M with 4 perradial lobes (9)

Yes

DIPLEUROSOMATIDAE

Yes

GO on M extending on perradial lobes

TIARANNIDAE

No Chromatonema rubrum with CO (4), without STA (3)

Yes

Yes

4 or 8 simple RC

LAODICEIDAE Staurostoma mertensii

No

4

Yes

≥ 4 branched RC

No

HEBELLIDAE Hebella muscencis

8 CO-like structures (8), GO elongated forming linear sacs on RC, separated from M with or without open STA (3)

Yes TECLAIIDAE

LEPTOMEDUSAE GROUP II

3

Parateclaia euromarge

FIGURE 72:  Leptomedusae group I- Leptothecata: Sugiura chengshanense after Sugiura (1973); detail of umbrella (ventral view) of Gastroblasta raffaelei (2), after Lang (1886); Orchistoma

pileus, whole figure and detail of manubrium (5), and Cuvieria carisochroma after Mayer (1910); Melicertum octocostatum after Kramp (1933); Chromatonema rubrum and detail of umbrella margin of Laodicea spp. showing tentacle bulbs, ocelli and cordili (4), after Kramp (1919); details of gonads and manubrium of Margalefia intermedia (9) after Pagès et al., 1991; Staurostoma mertensii after Hartlaub (1897); Hebella muscencis after Boero et al. (1997); Parateclaia euromarge, whole figure and detail of umbrella margin (3 and 8), after Bouillon and Boero (2000). All lateral views, unless noted otherwise.

208 ta xonom y

No

1M

LEPTOMEDUSAE GROUP II- Order: LEPTOTHECATA Yes

open STA (1)

Yes

with OC (2)

TIAROPSIDAE 2

No No

Octogonade mediterranea

1 Yes

without OC

MITROCOMIDAE Mitrocomella brownei

mainly closed STA (3) Yes

Yes with (adaxial) OC (4)

Yes 4 RC

BARCINIDAE

4 No

with GAP (5)

Barcino foixensis

Yes

≥ 8 STA (3)

Yes

4–6 RC

EIRENIDAE

Yes

5 Eutima gracilis

No

M narrow, usually 4-8 RC

No

Yes

M broad, > 16 RC

AEQUOREIDAE Aequorea tenuis

Yes

T_BU with lateral CI (6) 6

Yes

4 RC

LOVENELLIDAE

Eucheilota paradoxica

No with marginal CI (7)

7

Yes

Yes

4 RC

Yes CIRRHOLOVENIIDAE

No Cirrholovenia tetranema

LEPTOMEDUSAE GROUP III FIGURE 73:  Leptomedusae group II-Leptothecata: Octogonade mediterranea, Aequorea tenuis and detail of umbrella margin of Eutima coerulea (1), after Mayer (1910); details of umbrella margin of Tiaropsis multicirrata (2), E. viridula (3) and of Eucheilota maculata (6), after Russell (1953); Mitrocomella brownei and Eutima gracilis after Kramp (1959a); Barcino foixensis, whole figure and detail of umbrella margin (4), after Gili et al. (1999); Eucheilota paradoxica after Carré and Carré (1990); detail of manubrium of Eirene viridula (5), Cirrholovenia tetranema, whole figure and detail of umbrella margin (7), after Kramp (1959b). All lateral views.  

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  209

3

LEPTOMEDUSAE GROUP III- Order: LEPTOTHECATA

with marginal tentaculae (2)

Yes

Yes

Yes

Many STA, 4 RC

OCTOCANNOIDAE 2

No No

1

Octocannoides ocellata Yes

8 STA, 4 RC

PHIALELLIDAE

Phialella quadrata

GO completely surrounding RC (3)

Yes 4

endodermal core of T (4) extending into bell mesoglea

Yes

Yes

4 RC

BLACKFORDIIDAE

No

3

Blackfordia virginica No (except in Obelia)

4 RC

Yes

CAMPANULARIIDAE

Clytia hemisphaerica Yes

T_BU with excretory papillae (5)

5

4–8 (sometimes 12) RC, no permanent rudimentary T

Yes MALAGAZZIIDAE

No Malagazzia condensum

4-8 RC, numerous BU that will not develop T

Yes

PHIALUCIIDAE Phialucium mbenga

FIGURE 74:  Leptomedusae group III-Leptothecata: Octocannoides ocellata after Bouillon et al. (1991); Phialella quadrata after Kramp (1959a); detail of gonads of P. quadrata (1) and tentacle bulb with excretory papillae of Eutima gegenbauri (5), after Russell (1953); Blackfordia virginica and Clytia hemisphaerica after Mayer (1910); detail of gonads (ventral view) of C. hemisphaerica (3) after Pagès et al. (1992); detail of umbrella margin of B. virginica (4) after Moore (1987); Malagazzia condensum after Bouillon (1984a); Phialucium mbenga after Bouillon (1984b). All lateral views, unless noted otherwise.  

210 ta xonom y

GO divided in two lateral parts (1) separated by a median groove

ANTHOMEDUSAE GROUP I - Suborders: CAPITATA sensu stricto (*), FILIFERA Yes

Marginal T branched

Yes

T with some adhesive ends (1). Usually > 4 RC. Mainly benthic medusa.

* CLADONEMATIDAE

1

No

Eleutheria dichotoma

No T branched through CNI (2). Outer U covered with N_KN.

Yes

* ZANCLEIDAE

2 Zanclea sessilis

3 Yes

Yes

MO large, flaring, rim with continuous row of N_KN (4) No

OCEANIIDAE

6

Turritopsis nutricula

M tubular, MO small, circular or quadratic (5) 4

5

Yes

No

Yes

T_BU with 1 T (6), MO with clusters of N_KN

HYDRACTINIIDAE

No PER-R T_BU with ≥ 2 T. M with oral T/arms (3,7,8) ANTHOMEDUSAE GROUP II Yes

7

Oral T inserted above MO rim (7)

Podocoryna carnea Yes

BOUGAINVILLIDAE

Bougainvillia britannica Oral T very near MO rim (8), 8 groups of marginal T. Medusa buds on M (9)

9 8

RATHKEIDAE

Yes Rathkea octopuncata

FIGURE 75:  Anthomedusae group I-Suborder Capitata sensu stricto: Eleutheria dichotoma after Hincks (1868); Zanclea sessilis (whole figure), detail of mouth of Turritopsis nutricula (4) and of Bougainvillia britannica (7), after Russell (1953); particular of tentacles of Z. sessilis (2), mouth of Bougainvillia pyramidata (3), mouth (ventral view) of Podocoryna carnea (5 and 6) after Schuchert (2012); T. nutricula and B. britannica (whole figures) after Kramp (1968); P. carnea after Edwards (1972); Rathkea octopuncata, whole figure and detail of manubrium and mouth (8 and 9), after Naumov (1960). All lateral views, unless noted otherwise.  

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  211

M with oral T/arms (3,7,8) or N_KN (4)

ANTHOMEDUSAE GROUP II - Suborders: APLANULATA (**),

CAPITATA sensu stricto (*), FILIFERA

Yes

BYTHOTIARIDAE

No

Marginal T with BU

Bythotiara murrayi 1

Yes

2

** MARGELOPSIDAE

Yes

PER-T_BU with ≥ 2 T (1)

Clusters/tracks of N (3) on outer U between T. 4-6 branched RC (4).

Yes

T_BU with 1 T (2)

Yes

Margelopsis haeckelii Yes * PROBOSCIDACTYLIDAE

3

5 4

No

Proboscidactyla stellata 6

MOERISIIDAE

Yes

M extending in PER-lobes bearing the GO (5) No M flaring and cruciform (6) or very large and with rim folded (7)

Yes

8 * TRICHYDRIDAE

No

Yes ANTHOMEDUSAE GROUP III 9

Odessia maeotica

T_BU present. Many fine branched CEC (8) connecting RC and T_BU

T_BU non clearly separated by T. 2 or > 4 marginal T. Usually with interradial BU or T.

No

7

Yes

MO with 4 simple/crenulated/complexly folded L (7). RC often ribbon-like, with jagged margin (9).

7

4 marginal T, no INTER-T_BU, MO cruciform, rim unfolded.

Trichydra pudica * PANDEIDAE

Yes

8

Neoturris pileata

Yes

MO with 4 simple L, 4 T arising from large, hollow BU (10)

Yes

* PROTIARIDAE 10 Paratiara digitalis

FIGURE 76:  Anthomedusae group II-Suborders Aplanulata, Capitata sensu stricto, Filifera: Margelopsis haeckeli after Werner (1954); Bythotiara murrayi and Proboscidactyla stellata after

Pagès et al (1992); particular of tentacles of Climacocodon ikarii (1) after Kubota (1993); detail of umbrella margin of P. stellata (3) after Russell (1953); detail of tentacle of Phialellidae (2) after Russell (1963); Odessia maeotica after Morri (1981); detail of umbrella of Moerisia lyonsi (5) after Boulenger (1908); Trichydra pudica after Edwards (1973); Neoturris pileata, whole figure and detail of umbrella (7 and 9), after Hartlaub (1914); Amphinema rugosum, ventral view (6) and detail of gonads and manubrium of N. pileata (7), after Schuchert (2012); Paratiara digitalis after Kramp (1959a). All lateral views, unless noted otherwise.  

212 ta xonom y

M without oral T, oral arms or N_KN

ANTHOMEDUSAE GROUP III - Suborders: APLANULATA (**), CAPITATA sensu stricto (*), FILIFERA

Yes

MO small, with circular rim (1)

with OC (2)

Yes

Marginal T_BU with dark, conspicuous, abaxial OC (2)

No

Yes * CORYNIDAE

2

2 Sarsia tubulosa

1 No

3 4 marginal T, all with distinct terminal swelling (3)

Plotocnide borealis

No

Yes

1-4 marginal T, without distinct terminal swelling but may end in N_KN (4)

EUCODONIIDAE Eucodonium brownei

5

3 Yes Yes

Exumbrella with N tracks (5)

1, 2 or 4 marginal T equally long

Yes

** TUBULARIIDAE

4 Ectopleura dumortierii

Hybocodon prolifer

No

No M sausage-shaped, sometimes with sac-like processes

Yes

1 to 4 marginal T of different length

Yes

Corymorpha nutans

** CORYMORPHIDAE

FIGURE 77:  Anthomedusae group III-Suborders Aplanulata, Capitata sensu stricto, Filifera: Sarsia tubulosa after Edwards (1978); Plotocnide borealis, Ectopleura dumortierii, Hybocodon

prolifer and Corymorpha nutans after Kramp (1959a); detail of manubrium and mouth of Sarsia spp. (1) and Eucodonium brownei after Russell (1953); detail of tentacle bulb of Codonium proliferum (2) modified after Schuchert (2012). All lateral views.  

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  213

With GAP & medusae budding on M

Yes

**APLANULATA incerta sedis

U with a CG (1)

No

15 cm

Yes

> 8 marginal T and RO > 16 LAP

Yes

ATOLLIDAE

Atolla wyvillei

Yes

U usually less tall than wide

8 marginal T and RO 16 LAP and MAP

1

Yes

NAUSITHOIDAE

Atolla chuni

Nausithoe punctata 6 marginal T and RO 12 LAP

ATORELLIDAE

No Atorella subglobosa

RO PER-R, 4–12 marginal T

U more/as tall than/as wide

Yes

Large. CG (1) near mid U

Yes

Yes

4 RO

PARAPHYLLINIDAE

Paraphyllina ransoni

RO INTER-R, 4-28 marginal T

Yes 1

PERIPHYLLIDAE

No Periphylla periphylla

Small. CG (1) near top U

Yes

1 LINUCHIDAE

SCYPHOMEDUSAE GROUP II Linuche unguiculata FIGURE 78:  Scyphomedusae group I- Order Coronatae: Atolla chuni (lateral view), Atolla wyvellei (lateral view), Nausithoe punctata (lateral and ventral views), Atorella subglobosa (lateral view), Paraphyllina ransoni (detail of ventral view), Periphylla periphylla (lateral view) and Linuche unguiculata (lateral and ventral views) after Mayer (1910); Atolla wyvellei (detail of ventral view) after Russell (1970); P. ransoni (lateral view), modified after Russell (1970).  

214 ta xonom y

SCYPHOMEDUSAE GROUP I - Order: CORONATAE

SCYPHOMEDUSAE GROUP II - Orders: SEMAEOSTOMEAE, RHIZOSTOMAE (Scapulatae) Medusa with 4 oral arms (1) Yes Yes

with MAP (2)

T arising from edge of U. MAP separate and unbranched. Long, pointed oral arms (3)

2

Yes

PELAGIIDAE

No

Pelagia noctiluca

3 Yes

T arising under the U Pelagia noctiluca 1

CYANEIDAE

No

4

No Cyanea capillata Yes

Yes

with marginal T (5)

5

Aurelia aurita ULMARIDAE - II

No

Aurelia aurita

Stygiomedusa gigantea Medusa with 8 pairs of SCA (6)

Yes

Oral arms fused proximally (7), without a primary MO opening.

Yes RHIZOSTOMATIDAE 7

No

Rhizostoma pulmo

No

6 Medusa without SCA

Oral arms fused along whole lenght (8). With a primary MO opening (9).

8 9

Yes SCYPHOMEDUSAE GROUP III

STOMOLOPHIDAE

Stomolophus fritillarius

FIGURE 79:  Scyphomedusae group II-Orders Semaeostomeae, Rhizostomae (Scapulatae): Pelagia noctiluca, Rhizostoma pulmo, whole figure and detail of umbrella (6), detail of umbrella margin of Cyanea capillata (ventral view) and Stomolophus fritillarius, whole figure and details of manubrium (8 and 9) and of umbrella margin (ventral view), after Mayer (1910); P. noctiluca, ventral view (1 and 2) and detail of umbrella margin (ventral view), after Russell (1970); C. capillata (whole figure) modified from picture of © Jason Gregory, www. britishmarinelifepictures.co.uk; Aurelia aurita and detail of umbrella margin of R. pulmo (ventral view), after Stiasny (1923); A. aurita ventral view (4), after Russell (1968); Stygiomedusa gigantea (whole figure and detail of umbrella margin, ventral view), composite by Elias after Mianzan and Cornelius (1999). All lateral views, unless noted otherwise.  

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  215

with RC (4)

ULMARIDAE - I

SCYPHOMEDUSAE GROUP III - Order: RHIZOSTOMAE (Inscapulatae)

Network of RC (1) arising from CC (2)

Yes

1

Yes

CEC (3) between16 RC, usually blindly ending and NOT anastomosing. Broad, much folded oral arms

LYCHNORHIZIDAE

Lychnorhiza arubae

Lychnorhiza lucerna No

No

4 Yes

Network of anastomosing circular CEC (4), communicating with CC (2). Pyramidal oral arms

2

CATOSTYLIDAE

Catostylus tagi

5 1 Network of RC (1) arising from MO area (5)

Yes

Numerous RC between RO (6). Ex-U smooth, oral arms with long filaments (7)

Yes

No CC

CEPHEIDAE

Yes 7 6 Cotylorhiza tuberculata

No

6

5 8 With CC (8). Ex-U finely granular and dome-shaped

Yes

1 MASTIGIIDAE

7

Phyllorhiza punctata

FIGURE 80:  Scyphomedusae group III-Order Rhizostomae (Inscapulatae): Lychnorhiza lucerna whole figure and Cotylorhiza tuberculata, whole figure and detail of umbrella margin (1, 5 and 6, ventral view), after Mayer (1910); detail of umbrella margin of Lychnorhiza arubae (1, 2 and 3, ventral view), after Stiasny (1921); Catostylus tagi whole figure, after Grenacher and Noll (1876); Phyllorhiza punctata, whole figure and detail of umbrella margin (1, 5 and 8, ventral view) and detail of umbrella margin (2 and 4, ventral view) of C. tagi, after Stiasny (1923). All lateral views, unless noted otherwise.  

216 ta xonom y

2 3

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  217 Trachymedusae  Family Geryoniidae 4-6 (sometimes more) RC; With CEC. M with GAP, GO flat and leaf-shaped on RC. Marginal T solid and hollow. Ecto-endodermal STA enclosed in mesoglea. Distribution and ecology: L. tetraphylla is a cosmopolitan neritic species that performs daily vertical migrations. The sting of G. proboscidalis is considered highly irritating for humans.

Liriope tetraphylla (Chamisso & Eysenhardt, 1821)

A

Size: 10–30 mm W RC: 4 GO: 4, midway on RC CEC: 1–3 per ISP GAP: twice the M length (1) T: 4 long and 4 short STA: 8 Note: juveniles have GAP reduced/absent and more T Distrib: epipelagic in CNRY, GFST, NASE, NASW, NECS, NWCS

B

C

1

D

Geryonia proboscidalis (Forsskål, 1775) Size: 35–80 mm W RC: 6 GO: 6 CEC: up to 7 per ISP GAP: twice the size of M T: 6 long and 6 short STA: 12 Distrib: epipelagic in CNRY, NASE, NASW

Trachymedusae  Family Halicreatidae Usually ≥ 8 broad RC; without CEC. M broad and circular, without GAP. MO without distinct lips. Numerous marginal T, different in size but all structurally alike, i.e. flexible proximally and stiff distally. T that can be arranged in a continuous row (e.g. in Halicreas and Haliscera) or in groups (e.g. in Botrynema). Might have PA on exumbrella (genus Halicreas). Free ecto-endodermal STA.

Botrynema brucei Browne, 1908

E

2

Haliscera conica F

Size: up to 25 mm W U: wide, ends in a distinct knob RC: 8 GO: midway on RC M: short and wide T: arranged in 16 groups of 11–12 (2) + 8 solitary perradial STA: 3 per ISP +1–2 on either side of perradial T Distrib: meso-bathypelagic in ARCT, NADR, NASE, NASW, NECS

Vanhöffen, 1902 Size: up to 18 mm W U: wide, with conical projection RC: 8 GO: midway on RC M: short and wide T: 8–9 per ISP STA: 2 in each ISP Distrib: meso-bathypelagic in NASE, CNRY, NASW

Halicreas minimum Fewkes, 1882

G

3

H

Size: 30–40 mm W U: wide, with conical projection RC: 8 PA: 8 (3) GO: flat, along greater part of RC M: short and wide T: Very numerous, up to 640 STA: 3 in each ISP +1–2 on either side of the perradial T Distrib: meso-bathypelagic in ARCT, GFST, NADR, NASE, NASW, NECS

FIGURE 81:  Liriope tetraphylla: A, adult, lateral view; B, C, juvenile, lateral and latero-ventral views. Geryonia proboscidalis: D, adult, lateral

view. Botrynema brucei: E, adult, lateral view; Haliscera conica: F, adult, lateral view; Halicreas minimum: G, H, adult, lateral and ventral views. A and D Mayer (1910); B, C and E Russell (1953); F and G, Vanhöffen, (1902); H, Kramp (1947).

218 ta xonom y Trachymedusae  Family Rhopalonematidae 8 (rarely more) RC; no CEC. M narrow, with distinct lips, might have GAP. GO globular, linear or pendent on RC or as a ring around M, extending on RC. Marginal T evenly distributed, sometimes of two kinds, generally of uniform structure throughout. STA are free, or rarely enclosed in the U.

Aglaura hemistoma

Aglantha digitale

A

GAP (O. F. Müller, 1776)

GO

Size: 10–40 mm H RC: 8 GO: 8 sausage-shaped pendent from sub-U portion of RC, at the base of GAP GAP: slender, as long as U T: ≥ 80 solid, all alike M STA: 8 free, club-shaped Note: U twice higher than wide, with small conical projection Distrib: epipelagic in ARCT, GFST, NADR, NASE, NASW, NECS, NWCS, SARC

C

1

D

H

M

V

(Kramp, 1913)

E

Vanhöffen, 1902 GO

Pantachogon haeckeli Maas, 1893

Size: 10 mm H, 22 mm W RC: 8 GO: 8 sausage-shaped on RC, at the base of M GAP: absent T: up to 350 STA: number unknown Note: Ex_U of Crossota spp. with numerous meridional furrows (2). Red-brown U, M, GO and T in alive specimens. Distrib: bathypelagic in ARCT, NADR, SARC

F

Size: 20 mm H/W RC: 8 GO: linear, on 2/3 of RC GAP: absent T: 64 solid, all alike STA: 64 free, club-shaped

GO

Crossota norvegica

GO

Vanhöffen, 1902 Size: 18 mm H, 20 mm W RC: 10–14 T: 200-250 GO, GAP, STA, Note: see Crossota rufobrunnea Distrib: bathypelagic in ARCT, SARC

2

Note: sub-U with circular muscular fields at the apex (3) Distrib: epi-bathypelagic in ARCT, NASE, NADR, NASW

I

Rhopalonema velatum GO

4

Gegenbaur, 1857

Size: 6 mm H, 8–10 mm W RC: 8 GO: 8 oval, mid- of RC GAP: absent T: 8 PER-R long, club-shaped, T V up to 24 AD-R short, cirrus-like STA: 8 enclosed, beside T Note: U with apical knob (4) and very broad V Distrib: epipelagic in NADR, NASE, NASW, NECS, NWCS, SARC

Note: U with flat apex. Broad V Distrib: epipelagic in NASE, NWCS

Crossota rufobrunnea

Colobonema sericeum

3

Size: 4–6 mm H, 3-4 mm W RC: 8 GO: 8 sausage-shaped on GAP GAP: shorter than U and slender T: 48–85, all alike STA: 8 free, club-shaped

GO

Size: 35 mm H, 45 mm W RC: 8 GO GO: 8 linear, extending on whole RC GAP: absent T: 32 all alike, developing in succession STA: free, club-shaped Note: sub-U with muscular fields typically star-shaped (1). Broad V Distrib: bathypelagic in ARCT, NADR, NASE

G

Péron & Lesueur, 1810

GAP

B

J GO

Persa incolorata

GAP

McCrady, 1857

M

Size: 4 mm H, 3 mm W RC: 8 GO: 2 oval/sausage-shaped pendent from sub-U mid of opposite RC GAP: small T: 48 long, ending with N_KN STA: 8 free, club-shaped

Distrib: epi-bathypelagic in NASE, NWCS

FIGURE 82:  Aglantha digitale: A, adult, lateral view; Aglaura hemistoma: B, adult, lateral view. Colobonema sericeum: C, D, adult, lateral and ventral views. Crossota rufobrunnea, E, and C. norvegica, F: adults, lateral views. Pantachogon haeckeli: G, H, adult, lateral and ventral views. Rhopalonema velatum: I, adult, lateral view. Persa incolorata: J, adult, lateral view. A, B, I and J Mayer (1910); C, G, Vanhöffen (1902); D, H Russell (1953); E, Kramp (1913); F, Kramp (1959a).

Narcomedusae  Narcomedusae Families Aeginidae, Pseudaeginidae, and Solmundaeginidae Following the recent revision of Lindsay et al. (2017) some genera previously attributed to the family Aeginidae have now been assigned to the new family Solmundaeginidae, which comprises the species of Solmundaegina, Solmundella, Aeginopsis and Solmundus, and others to the family Pseudaeginidae. Aeginidae might have CC, while CC are present in Pseudaeginidae and absent in Solmundaeginidae. All families have MAP INTER-R bearing the GO, undivided or divided into two to four parts; PER-R primary T on ex-U, inserted above MAP and between marginal LAP. Aeginidae might have secondary T, while Pseudaeginidae and Solmundaeginidae have only secondary T_BU. Only Aeginidae might have OT.

A

B

Aeginura grimaldii MAP

1

2

Solmundella bitentaculata

MAP

Maas, 1904

(Quoy & Gaimard, 1833)

Size: up to 45 mm W GO: 8 MAP: 16 T: 8 large primary T/PER (1). 3–5 small secondary T in each ISP (2) STA: 1–2 between adjacent secondary T Note: without CC or OT Distrib: meso-bathypelagic in ARCT, NADR, NASE, NASW, NWCS, SARC

3

Size: up to 12 mm W GO: 8 MAP: 8 T: 2 opposite and long (3), 4 PERO STA: 8–16 (up to 32) Note: without secondary T, CC and OT. V well developed. With vestigial T_BU. T roots small and curved orally Distrib: epi-bathypelagic in NASE, NECS, NWCS, SARC

Narcomedusae  Family Cuninidae Might have CC. MAP PER-R, undivided and bearing the GO. PER-R primary T on ex-U, inserted on to the central margin of each MAP. Might have secondary T. Might have OT. Distribution and ecology: S. incisa preferably prey on gelatinous plankton, particularly on ctenophores, cnidarians and salps.

Cunina globosa Eschscholtz, 1829

C

4

OT

4

D

Solmissus incisa (Fewkes, 1886) Size: up to 100 mm W GO: up to 10–14 MAP: 20–40, with oval outline (4) LAP: rectangular, as long as large T: 20–40 stiff and tapering STA: 2–5 on each LAP Note: flat disk-like U. Without OT. Probably a species complex Distrib: meso-bathypelagic in ARCT, GFST, NADR, NWCS

Size: up to 18 mm W GO: up to 10–14 MAP: 10–14, wide and quadratic (4) LAP: short and broad T: 10–14 relatively short STA: 3 on each LAP Note: with CC. OT short/oval on LAP margin Distrib: meso-bathypelagic in NWCS

Narcomedusae  Family Solmarisidae Might have CC. Without MAP. GO on M wall/diverticula. Numerous T on U at level of periphery of M. Might have OT.

E

Solmaris corona (Keferstein & Ehlers, 1861)

F

Size: 12–15 mm W GO: forming a broad ring on M wall LAP: ≥30–36 rectangular, up to twice as long as broad T: ≥30–36 STA: usually 2 (1–4) on large cushion Note: without CC and OT Distrib: meso-bathypelagic in NECS, NADR, NASW

Pegantha rubiginosa (Kölliker, 1853)

G

Size: up to 16 mm W GO: without radial diverticulae LAP: 12–16 rectangular, with rounded corners. 2 mid OT longer than LAP, 2 lateral short T: ≥30–36 STA: 4–6 on each LAP Note: with CC and OT Distrib: meso-bathypelagic in NASW

FIGURE 83:  Aeginura grimaldii: A, adult, lateral view. Solmundella bitentaculata: B, adult, lateral view. Cunina globosa: C, adult, lateral view.

Solmissus incisa: D, adult, ventral view. Solmaris corona: E, adult, lateral view. Pegantha rubiginosa: F, adult, lateral view; G, detail of umbrella margin showing otoporpae and CC. A, Maas, (1905); B and F, Mayer (1910); C, Bigelow (1909); D, edited from Fewkes (1886); E, Pagès et al., (1992); G, Kramp (1959).

220 ta xonom y Leptomedusae  Family Aequoreidae Many simple or branched RC. M very wide, circular, usually without GAP. GO on RC, separated from M and divided longitudinally. Marginal T hollow, fine, about 1/2 to 3 times the number of RC. No CI. With excretory pores, might have PA and OC. STA closed. Note: Aequorea spp. have simple and undivided RC and no PA. Distribution and ecology: Aequorea spp. eat mainly soft-bodied prey. 1

A

Aequorea forskalea

2

Péron and Lesueur, 1810

B

C

Size: 175 mm W RC: usually 60–80, sometimes fewer or up to 160 GO: along whole length of RC T: generally less than RC, can be 1/2 or twice the number of RC (1). STA: 5–10 between successive T Distrib: neritic in NECS, NWCS, SARC

Aequorea vitrina Gosse, 1853 Size: 100–170 mm W RC: 60–100 GO: along whole length of RC T: >3 times than RC (2) STA: 1–2 between successive T

Distrib: neritic in NECS, NWCS

Leptomedusae  Family Blackfordiidae 4 simple RC. M narrow and short, without GAP. MO with 4 long curved lips (3). GO completely surround RC. 80–250 hollow marginal T, with endodermal core extending from margin into bell mesogela. Numerous closed STA. Distribution and ecology: B. virginica preys not only on zooplankton but also on phytoplankton, ciliates and detritus.

D

Blackfordia virginica Mayer, 1910

4

E

3

Size: 14 mm W RC: 4 GO: linear, >1/2 length of RC from M T: 80, long, with diverticula into U margin (4) STA: 1 (rarely 2) between successive T (5) Note: introduced species from the Black Sea Distrib: neritic in NASE, NWCS

5

Leptomedusae  Family Campanulariidae Usually 4 RC. Might have V. M short, without GAP. GO separated from M, completely surrounding RC. Marginal T usually hollow (except in Obelia). Without excretory pores, PA, CI (except in Paralovenia) and OC. 16–200 closed STA. Note: the classification of this family is unsatisfactory. Divisions are not always well defined.

Clytia hemisphaerica

F

(Linnaeus, 1767)

6

7

9

G

Size: up to 20 mm W RC: 4 (sometimes up to 12) GO: oval/linear, 1/2–3/4 of length of RC, opposite M (6) T: 32, with prominent globular T_BU (7) STA: 1–3 between successive T Distrib: neritic in CNRY, NASE, NECS, NWCS, SARC

Obelia spp. Péron & Lesueur, 1810

Size: 2.5–6 mm W RC: 4 GO: sac-like, on middle-end of 8 RC (8) T: numerous short, stiff solid, with diverticula into U margin (9) STA: 8, under T_BU Note: very flat, no V Distrib: neritic in CNRY, NASE, NECS, NWCS, SARC

FIGURE 84:  Aequorea forskalea: A–B, adult, lateral and ventral views. Aequorea vitrina: C, adult, ventral view. Blackfordia virginica: D, adult lateral view; E, detail of umbrella margin showing endodermal diverticula of tentacle bulbs and statocyst. Clytia hemisphaerica: F, adult lateral view. Obelia spp.: G, adult dorsal view. A, D, E and F, Mayer (1910); B, C, Russell (1953); G, Kramp (1933).

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  221 Leptomedusae  Family Eirenidae 4–6 simple RC. M small, usually on well-developed GAP. GO separated from M, in each species on well-defined part(s) of RC. Hollow marginal T (except in Eugymnanthea). Might have marginal WA or CI, excretory pores or PA. Without OC. 8 or more closed STA.

A

B

Eutima gracilis

Eutonina indicans

3

(Romanes, 1876)

(Forbes & Goodsir, 1853) Size: up to 13 mm W GAP: much longer than U (1) GO: 4, restricted to GAP T: 2–4 PER-R (at times more) (2) STA: 8 Note: 40–80 WA. With CI on WA/T Distrib: neritic in CNRY, NASE, NASW, NECS

2 1

Size: 25–35 mm W GAP: no longer than U GO: linear and long, on sub-U part of RC (3) T: > 100 (4) STA: 8 Note: without CI/ WA

4

Distrib: neritic in NECS, SARC

Leptomedusae  Family Laodiceidae 4 or 8 simple RC. M might have lobes or pouches, without GAP. GO on RC, on RC and M-lobes, or into M-pouches. Hollow marginal T. With marginal CO. Might have CI and OC. Without STA.

C

Laodicea undulata

E

D

Size: up to 90 mm W, 30 mm H RC: 4 with 20–30 lateral diverticula attached to sub_U, at each side of RC GO: 4, inside the diverticula (8) T: 300–500, with club shaped CO Note: M with funnel-shaped PER-R lobes. No CI/OC Distrib: meso-bathypelagic in GFST, NWCS, SARC

Size: up to 37 mm W RC: 4 straight 5 GO: 4 long, sinuous along RC (5) close to M T: up to 400–600 (6). Some/all T_Bu with OC Note: MO with crenulated lips (7). With CI and CO Distrib: neritic CNRY, NASE, NECS, NWCS, SARC

7

6

Ptychogena lactea Agassiz, 1865

(Forbes & Goodsir, 1853) 5

8

Leptomedusae  Family Lovenellidae 4 simple RC. M small, without GAP. GO on RC, separated from M. Hollow marginal T with lateral CI*. Without marginal CI, OC and excretory pores. With closed STA. (*Except in Paralovenia).

Lovenella clausa

F

G

(Lovén, 1836) 9

10

Size: 5–9 mm W RC: 4 GO: oval, longitudinally divided, on RC very close to CC (9). T: 16–24, with large conical T_BU (10) CI: 1–3 on either side of each T STA: 16–32 Distrib: neritic in NECS

12

Eucheilota paradoxica 11 Mayer, 1900 Size: 4 mm W RC: 4 GO: swollen, on the middle of RC T: 4 large PER-R, with 2 lateral CI STA: 8 Note: medusa buds on RC (11). M flask-shaped. ≥4 rudimentary T_BU with CI (12) Distrib: neritic in NASE, NWCS

FIGURE 85:  Eutima gracilis: A, adult, lateral view. Eutonina indicans: B, adult, lateral view. Laodicea undulata: C, adult, lateral view. Ptychogena

lactea: D, adult, ventral view; E, adult, lateral view. Lovenella clausa: F, adult, lateral view. Eucheilota paradoxica: G, adult, lateral view. A, B, and D, Kramp (1959a); C, Hyman (1940); E, Naumov (1960); F, Russell (1963); G, Carré and Carré (1990).

222 ta xonom y Leptomedusae  Family Melicertidae 8 simple/bifurcated RC. Base of M attached over its whole surface, without GAP. GO on RC. Hollow marginal T. Without STA, CI and CO. Might have OC.

A

Melicertum octocostatum

1

Size:10–14 mm W/H GO: 8 sinuous separated from M, covering almost whole length of RC and thicker near U (1) T: up to 88 large T, alternating with small ones OC: None Note: 3–7 fine lines of N in each ISP on sub-U

(M. Sars, 1835)

Distrib: neritic in ARCT, GFST, NASE, NECS, NWCS

Leptomedusae  Family Mitrocomidae ≥ 4 simple/S-shaped RC. Base of M attached to Sub-U along edges of RC. GO linear/oval on RC only. Hollow marginal T. Open STA. Might have CI. Without OC.

B

Cosmetira pilosella

C

Size: 20–48 mm W RC: 4 straight GO: 4 linear with median division, along 1/2–3/4 of RC T: 64–100 with large T_BU (2) STA: 8 Note: 6–10 CI between adjacent T (3) Distrib: neritic in NECS, SARC

2

3

Mitrocomella polydiademata

D

Forbes, 1848

(Romanes, 1876)

2

Size: 12–30 mm W RC: 4 straight GO: 4 straight, along 2/3–4/5 of RC

T: usually 36–48 with large T_BU (2) STA: 16 Note: 5–9 spiral CI with terminal CNI, arranged between adjacent T Distrib: neritic in NWCS, NECS, SARC

Leptomedusae  Family Phialellidae 4 simple RC. M small, without GAP. GO divided in the middle by a median groove (4), placed on RC, separated from M. Hollow marginal T smooth/moniliform. Without marginal/lateral CI, excretory pores or OC. 8 closed STA, usually each on a bulbous-like swelling.

E

F

Phialella quadrata

4

(Forbes, 1848)

4

5

6

Size: up to 13 mm W RC: 4 GO: elongated, on the outer third of RC T: 16–32, with globular T_BU (5) CI: 1–3 on either side of each T STA: 16–32 (6) Note: V well developed Distrib: neritic in NASE, NECS

FIGURE 86:  Melicertum octocostatum: A, adult, lateral view. Cosmetira pilosella: B, adult, lateral view; C, detail of umbrella and gonads, ventral

view. Mitrocomella polydiademata: D, adult, lateral view. Phialella quadrata: E, adult, ventro-lateral view. F, adult, detail of umbrella margin, radial canal and gonad. A, Kramp (1933); B, Cornelius (1995); C, Hartlaub (1909); D, Mayer (1910); E and F, Russell (1953).

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  223 Leptomedusae  Family Tiarannidae 4 simple RC. M wide, cross-shaped, with 4 PER-R lobes attached to Sub-U, without GAP. MO with 4 simple/crenulated lips. GO folded on M, extending on PER-R lobes. Numerous hollow marginal T. Hollow CO-like structures. Without OC.

B

A

Chromatonema rubrum Fewkes, 1882

Modeeria rotunda

C

(Quoy & Gaimard, 1827)

2

Size: up to 27 mm W/ 22 mm H GO: 10–16 sac-like, separated and hanging on each side of PER-R lobes (1) T: 20–24, with conical T_BU 1 Note: 2 CO among adjacent T (2) Distrib: meso-bathypelagic in ARCT, NADR, NASE, NWCS

3

Size: 20 mm W GO: transversally folded (3), on M and connected with PER-R lobes T: 16–28, with conical T_BU Note: 2–3 CO among adjacent T

Distrib: neritic-bathypelagic in ARCT, NASE, NADR, NECS, NWCS, SARC

Leptomedusae  Family Tiaropsidae 4–8 (exceptionally 16) simple RC. Marginal T long and rudimentary of 1 or 2 types, with T_BU. Open STA associated with OC.

Tiaropsis multicirrata E

M. Sars, 1835

D

4

5 6

F

5

Size: about 20 mm W RC: 4 straight GO: 4 kind of sinuous, along 1/2–2/3 of RC T: about 300 of one kind only, with swollen T_BU (4) OC: 8 STA: 8 ecto-endodermal (5) Note: MO with long, much folded and crenulated lips (6). Broad and flat GAP. Without CI Distrib: neritic in ARCT, NWCS, NECS, SARC

Anthomedusae  Family Aplanulata incerta sedis The polyp of Plotocnide was recently identified as Boreohydra simplex (Pyataeva 2016). Even though the medusa of this genus has been previously associated with Tubulariidae or Corymorphidae, following Schuchert (2012) we do not assign Plotocnide to any particular family.

Plotocnide borealis Wagner, 1885

G 7

8

9 Size: 1.5–3.5 mm H RC: 4 narrow and straight GO: ringlike encircle M (7) leaving lips and top portion of M free M/GAP: M 1/2 length of sub-U, with broad apical chamber with vacuolated endodermal cells T: 4 solid PER-R, up to twice U length. Each T has a small round T_BU and ends with an oval/swollen N_KN (8) OC: round, on T_BU, near origin of T Note: U with very thick mesoglea at the apex (9). N scattered on ex-U. No OC. Distrib: ARCT, NECS, SARC

FIGURE 87:  Chromatonema rubrum: A, adult, lateral view; B, detail of umbrella margin with marginal tentacles and cordyli. Modeeria rotunda: C, adult, lateral view. Tiaropsis multicirrata: D, adult, lateral view; E, detail of umbrella margin with marginal tentacles and statocyst; F, ectoendodermal statocyst. Plotocnide borealis: G, adult, lateral view. A, Kramp (1919); B, Kramp (1933); C, Kramp (1920); D, Agassiz (1849); E and F, Russell (1953); G, Kramp (1959a).

224 ta xonom y Anthomedusae  Family Bougainvilliidae 4 simple RC. MO circular with simple or dichotomously branched oral T clearly inserted above MO rim, ending with N_KN. Might have GAP. GO on M, either ring-like or in INTER/AD/PER-R position. Solid marginal T, all alike, either solitary or in groups on 4, 8 or 16 T_BU. Might have OC. 4 T_BU in PER-R position bearing ≥ 2 identical solid marginal T

Bougainvillia muscus

Bougainvillia britannica (Forbes, 1841)

A

Size: 5–8 (up to 12) mm W GO: 8 AD-R (1) separated by PER-R cleft 1 T: 4 groups of 12–17 (up to 30) T on PER-R triangular T_BU (2). Oral T divided up to 4–6 times, with long basal trunks (3). OC: oval/lenticular, at the base of T Note: without GAP Distrib: NECS, SARC

3

2

(Allman, 1863)

B

Size: 1–4 mm W GO: 4 INTER-R, at times slightly on RC T: 4 groups of 2–8 (usually 3–4) T on PER-R T_BU (4). Oral T divided 2 times, with moderate basal trunks (5). OC: round, on T_BU, near origin of T Note: GAP absent or very small Distrib: NECS, NWCS, SARC

5

4

Bougainvillia niobe Mayer, 1894

C 7 6

Size: 5 mm W GO: 8 ADT: 4 groups of 8 T. Oral T divided 4 times, with long basal trunks (6) OC: at the base of each T Note: M wide, flask-shaped, 1/2 length of sub-U, often with medusa buds (7) Distrib: NASW, NWCS

Anthomedusae  Family Corymorphidae 4 simple RC. M sausage-shaped generally not longer than U. Simple circular MO. GO surrounding M along its whole length. 1–4 marginal T, only 1 of which fully developed. 3–4 T_BU. Without OC. 8

D

Corymorpha nutans

Euphysa aurata

M. Sars, 1835 Size: 3 mm W/up to 6 mm H GO: around M, leaving GAP/MO free T: 1 moniliform with 40–80 N_KN. 3 rudimentary T_BU Note: U with pointed apex and canal (8). With small GAP Distrib: NECS, SARC

E

10

(Allman, 1863) Size: 2.5–6 mm H GO: around whole M T: 1 quite short PER-R, with 6–25 N_KN (9). 3 rudimentary T_BU (10) Note: M 2/3 or as long as sub-U, with oil drops in the upper half of M Distrib: NECS, NWCS, SARC

9

FIGURE 88:  Bougainvillia britannica: A, adult, lateral view. Bougainvillia muscus: B, adult, lateral view. Bougainvillia niobe: C, adult, lateral view. Corymorpha nutans: D, adult, lateral view. Euphysa aurata: E, adult, lateral view. A, C, D and E, Kramp (1959a); B, Russell (1953).

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  225 Anthomedusae  Family Corynidae 4 simple RC. M tubular, without lips. GO around M. 4 hollow marginal T. 4 T_BU with 1 OC each.

A

Coryne eximia

B

Sarsia tubulosa (M. Sars, 1835)

Allman, 1859 1

Size: 3 (up to 10) mm W GO: encircle M for almost entire length (1)

2

T: 4, 2–3 times as long as U, with 30 N_KN Note: length of M about 2/3 of

Size: 6–10 mm H GO: undivided, around M almost along its entire length (1), leaving both ends free T: 4 very long with many N_KN covering at least 1/2 of each T Note: length of M >2 times that of U,

sub-U. Without medusa budding on T_BU Distrib: NASE, NECS, NWCS

which is dome-shaped with mesoglea thickened at apex (2) Distrib: NECS, NWCS, SARC

1

Anthomedusae  Family Hydractiniidae 4 simple RC. M tubular/sac-shaped no longer than U, with/without GAP. MO ending with 4 N_KN, at times inserted on branched oral arms. GO on M, INTER-R, sometimes extending along basal, PER-R protrusions of the M. 4 or ≥8 marginal T ungrouped. Might have OC but not on T_BU.

Podocoryna borealis (Mayer, 1900)

C

3

Size: up to 4 mm W, 3–4 mm H GO: oblong, INTER-R T: 16–30 of different length. PER-R and INTER-R T_BU are largest. Note: oral arms branched 1–3 times (3). M without GAP, long about 2/3 times the length of sub-U. Without OC Distrib: NECS, NWCS, SARC

Anthomedusae  Family Oceaniidae 4 simple RC. MO margin fringed with numerous N_KN. GO on INTER-R walls of M. Solitary, solid T, numerous in adults. OC on AD-R side of T_BU. Distribution and ecology: Turritopsis eggs develop to planulae in the sub-U. Molecular analyses have proved that Turritopsis nutricula, which was considered a cosmopolitan species, is only distributed in the Western Atlantic. Historical records of T. nutricula in the eastern Atlantic likely refer to T. polycirrha.

D

Turritopsis nutricula McCrady, 1857 Size: up to 4–11 mm H GO: 4 pairs on INTER-R walls of M

4

T: 80–120. Note: pseudo-GAP formed by vacuolated endodermal cells separated in 4 blocks (4). In the congeneric species T. polycirrha vacuolated cells are contiguous. Distrib: NASE, NECS, NWCS

FIGURE 89:  Coryne eximia: A, adult, lateral view. Sarsia tubulosa: B, adult, lateral view. Podocoryna borealis: C, adult, lateral view. Turritopsis

nutricula: D, adult, lateral view. A, Mayer (1910); B, modified from Kramp (1959a); C, Edwards (1972); D, Kramp (1959a).

Anthomedusae  Family Pandeidae 4 (rarely 8) simple RC, often broadened/ribbon-like or with jagged margin. M large, with/without GAP. MO with simple/crenulated/ complexly folded lips. Might have mesenteries, rarely CEC. GO AD-/INTER-R on M, have a smooth or more commonly complexly folded/ pitted surface. GO might be extended on RC. 2 or >4 hollow marginal T with conical T_BU. Might have rudimentary T, i.e. small solid marginal T called tentaculae and OC. Distribution and ecology: Polyps of different species (e.g. Pandea conica) settle on shells of pteropods, bivalves or in the coelom of polychaetes. Thus their medusae are widely distributed in coastal and oceanic waters.

A

B

Amphinema dinema

Amphinema rugosum

(Péron & Lesueur, 1810) 1

3

Size: 4 mm W, up to 6 mm H RC: 4 broad GO: 8 AD-R pads, with smooth surface (1) M/GAP: M length 3/4 of sub-U, without GAP and with crenulated lips T: 2 opposing PER-R with large hollow T_BU (2). Without tentaculae Note: U with large conical apical process. 6–12 marginal WA. Without OC

2

4

(Mayer, 1900) Size: 5–6 mm H RC: 4 broad GO: 8 AD-R pads, each one with 3–4 folds (3) M/GAP: as in A. dinema T: as in A. dinema but with 14–24 tentaculae (4) Note: U with spherical/conical apical process. Without OC Distrib: NECS, NWCS

Distrib: NECS, NWCS

Leuckartiara octona C

D

Size: usually 10 (5–20) mm H RC: 4 broad often with jagged outline, joined to M by long mesenteries GO: INTER-R on whole M, horse-shoeshaped with PER-R folds and a few pits M/GAP: M flask-shaped no longer than U, without GAP and with crenulated lips T: Usually 16 (8–23) hollow, with T_BU clasping U to form a spur and with 1 OC. 1–3 rudimentary T_BU between adjacent T Note: U with round/conical apical process. Distrib: NECS, NWCS 8

E

Neoturris pileata

(Fleming, 1823)

6

Pandea conica Size: 10–20 mm H RC: 4 broad with smooth/jagged outline, forming mesenteries GO: INTER-R on 2/3 of M, separated perradially by mesenteries, forming coarse network of ridges and pits (7) M/GAP: M long 1/2 of sub-U, mainly attached to it by mesenteries. Without GAP and with highly folded/crenulated lips T: 16–24 (up to 45) hollow, with conical T_BU laterally compressed that clasp U forming spurs and have AD-R OC

Size: up to 20–35 mm H 5 RC: 4 broad, with short lateral diverticula joined to M by mesenteries (5) GO: INTER-R on whole M, with horizontal AD-R folds and with >20 isolated INTER-R pits (6) M/GAP: M flask-shaped no longer than U, with strongly folded L. Without GAP T: 60–90 hollow with T_BU clasping U but not forming spurs Note: U bell/bullet-shaped with solid apical process. Without OC Distrib: CNRY, NASE, NECS, NWCS, SARC

10

(Quoy and Gaimard, 1827) 7

(Forsskål, 1775)

11

F

9

Pandea rubra Bigelow, 1913 Size: up to 75 mm H and W RC: 4 very broad with jagged outline GO: INTER-R as in P. conica but with close network of ridges and pits (9) M/GAP: M large with broad base. Other characteristics as in P. conica. T: As in P. conica but T_BU have no OC Note: U round (10), without CNI tracks. Sub-U deeply pigmented red (11) Distrib: meso-bathypelagic in NASW, NECS

Note: U with round/conical apex (8), with CNI tracks from each T_BU Distrib: CNRY, NADR, NASE, NASW, NECS, NWCS, SARC FIGURE 90:  Amphinema dinema: A, adult, lateral view. Amphinema rugosum: B, adult, lateral view. Leuckartiara octona: C, adult, lateral view.

Neoturris pileata: D, adult, lateral view. Pandea conica: E, adult, lateral view. Pandea rubra: F, adult, lateral view. A, B and C, Kramp (1959a); D, Hartlaub (1914); E, Pagès et al. (1992); F, Lindsay et al. (2008).

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  227 Anthomedusae  Family Porpitidae The hydroid floating colony is composed of a chitinous internal skeleton with concentric air chambers (pneumatophore) and covering tissue (mantle). Central large feeding polyp (gastrozooid) surrounded by gastro-gonozooids (reproductive polyps) and dactylozooids (defensive polyps). Medusa has 4/8 simple RC and CC. M short and MO circular. GO PER-/INTER-R. 2/4 opposite T start from T_BU and end with round N_KN. At maturity U has 4/8 N tracks issued from marginal BU. Distribution and ecology: Hydroids float at surface while medusae are rarely seen. The hydroids of Velella velella prey on euphausiid larvae, copepods, fish eggs and larvae.

A

1

Hydroid

Velella velella

Size: up to 20 mm W and 10 mm long. Oval to elliptical colony, with a median

(Linnaeus, 1758)

Medusa B

sail (1) and deeply blue when alive

Size: 2 mm W and 3 mm H 3

RC: 4 GO: PER-R and INTER-R T: 1 short and 1 long on each side

2

2 PER-R T_BU without T (2)

Distrib: CNRY, GFST, NASE, NADR, NASW, NECS, NWCS

Note: 4 N tracks on ex-U (3)

Anthomedusae  Family Rathkeidae 4/8 RC. M cylindrical not longer than U, with GAP. MO with 4 lips elongated to form simple or branched oral arms/T armed with CNI knobs. GO completely encircling M. Mainly 8 (rarely 4) T_BU with ≥ 1 T. Without OC.

C

Lizzia blondina

D

E

Forbes, 1848

F

Rathkea octopunctata (M. Sars, 1835) Size: up to 3–4 mm H, 4 mm W RC: 4 T: 4 branched oral T (6). 4 perradial T_BU with

Size: 1–2 mm H RC: 4 T: 4 unbranched oral T (4). 4 perradial T_BU with 1–3 T each (5) and 4 interradial T_BU with 1 T 4 5

Note: U with small apical process. M with medusa buds Distrib: CNRY, NASE, NECS, NWCS, SARC

6 7

3–5 T each, 4 interradial T_BU with 2–3 T (7) Note: U round/dome-shaped. M with medusa buds Distrib: ARCT, NASE, NASW, NECS, NWCS, SARC

Anthomedusae  Family Tubulariidae 4 simple RC. M circular. GO covering M entirely. 1–4 T. Without OC. Ex-U with N tracks.

Hybocodon prolifer Agassiz, 1860

G

8

Size: 2–4 mm H RC: RC by T longer than the other RC M/GAP: M fusiform, shorter than U cavity. It might have GAP 10 T: 1 moniliform with broad T_BU (8). 3 9 smaller T_BU without T Note: medusa buds develop from T_BU (9). U has oblique margin and 5 lines of N on external side (10) Distrib: ARCT, NECS, NWCS, SARC

FIGURE 91:  Velella velella: A, hydroid, lateral view: B, medusa, lateral view. Lizzia blondina: C, adult, lateral view; D, mouth. Rathkea octopunctata: E, adult, lateral view; F, mouth. Hybocodon prolifer: G, adult with medusa buds, lateral view. A, Pagès et al. (1992); B, Brinckmann (1964); C and G, Kramp (1959a); D and F, Russell (1953); E, Naumov (1960).

228 ta xonom y Scyphomedusae  Family Atollidae ≥16 marginal LAP and >8 marginal RO alternating with an equal number of marginal T. 8 AD-R GO. Atolla spp. have 8 AD-R GO, 8 RO, a flat disk and a deep CG. They have a ‘hypertrophied’ T (HYP), larger than the others, that has a different position in different species.

Atolla chuni

Atolla parva

Atolla vanhoeffeni

Atolla wyvillei

(Vanhöffen, 1902)

Russell, 1958

Russell, 1957

Haeckel, 1880

Size: diam. up to 70 mm

Size: diam. circa 30 mm

Size: diam. circa 30 mm

Size: diam. up to 150 mm

T: 24. HYP_T in line

(up to 63 mm)

(up to 50 mm)

T: 22. HYP_T not in line

with interradius (1) Note: 7–9 small papillae on

T: 20–26. HYP_T in line with interradius (1)

T: 20. HYP_T in line with interradius (1)

with interradius (5) Note: RS divergent and

upper surface of each LAP

Note: RS straight, up to

Note: base of M cross-

extending beyond inner

inner margin of CM (2)

shaped (3), with 8 dark spots at outer corners (4)

margin of CM (6)

A

C

G

E

4 1

B

D

2

I F

5

3

6

H

Distrib: meso- but also epipelagic, recorded across the whole North Atlantic, particularly A. parva and A. wyvillei

Scyphomedusae  Family Cyaneidae Stomach divided in MAP containing the GO, divided peripherally in numerous branching canals ending blindly in marginal LAP. Without CC. Marginal T arising from sub-U distant from U margin. 4 oral arms with much folded lips. Cyanea spp. have typically 8 AD-R groups of marginal T arranged on >1 row and 8 RO. For the identification of the ephyra/post-ephyra stages of C. capillata and C. lamarckii see Holst (2012).

J

Cyanea capillata K

7

L

Cyanea lamarckii

(Linnaeus, 1758)

Péron & Lesueur, 1810

M

8

Size: diam. 300–500 mm T: 70–150 in each AD-R group Note: CM and RS with small

Size: diam. 60–150 mm T: 40–60 in each group Note: CM and RS usually without small pit-like intrusions (8). Yellowish-

pit-like intrusions (7) Usually yellow-brown colour Distrib: mainly recorded in NECS and NWCS

blue colour Distrib: mainly recorded in NECS

Scyphomedusae  Family Pelagiidae MAP separate and unbranched. Marginal T arising from U margin. Without CC. Long oral arms with frilled lips. For the identification of the ephyra/post-ephyra stages of Chrysaora hysoscella see Holst (2012). Distribution and ecology: C. hysoscella is often found in association with young carangid fish.

N

9

Chrysaora hysoscella (Linnaeus, 1767) Size: diam. up to 300 mm T: 24 in groups of 3, alternating with 8 RO Note: Ex-U typically with 16 V-shaped radial brown markings (9). 4 long oral arms with frilled edges Distrib: nearshore distribution in NECS

FIGURE 92:  Atolla chuni: A, adult, lateral view. Atolla parva and A. chuni: B, diagram showing position of hypertrophied tentacle (HYP), ventral view. A. parva: C, adult, lateral view; D, closeup of umbrella margin. Atolla vanhoeffeni: E, adult, lateral view; F, diagram showing position of HYP, ventral view. Atolla wyvillei: G, adult, lateral view; H, diagram showing position of HYP, ventral view; I, closeup of umbrella margin. Cyanea capillata: J, adult, lateral view; K, closeup of umbrella margin. Cyanea lamarckii: L, adult, lateral view; M, closeup of umbrella margin. Chrysaora hysoscella: N, adult, lateral view. A and G, Mayer (1910); B, F, H, Repelin (1965); D, K, M and N, Russell (1970); I, Russell (1959); C and E, composite by Elias, in Mianzan and Cornelius (1999); J and L, modified from pictures of © Jason Gregory, www. britishmarinelifepictures.co.uk.

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  229 Scyphomedusae  Family Pelagiidae MAP separate and unbranched (1). Marginal T arising from U margin. Without CC. Long oral arms with frilled lips.

A

Distribution and ecology: P. noctiluca has no polyp stage. It preferably preys on chaetognaths and mollusc larvae but also on copepods and fish larvae. It is bioluminescent.

Pelagia noctiluca

B

1

Forsskål, 1775 Size: diam. circa 120 mm T: 8, alternating with 8 RO Note: Ex-U covered by red-brown stinging warts (2). 4 thick oral arms

2

Distrib: epi-mesopelagic in CNRY, GFST, NADR, NASE, NASW, NATR, NECS, NWCS, SARC

Scyphomedusae  Family Periphyllidae 4 INTER-R RO and ≥ 4 (up to 28) marginal T Distribution and ecology: In the last two or three decades P. periphylla has become very abundant in some Norwegian fjords, where it is seen as a nuisance by the local fisherman. This species performs extensive diel vertical migrations.

3

Periphylla periphylla

C

(Péron & Lesueur, 1809) Size: diam. up to 250 mm T: 12 stiff and tapering, 3 between each pair of RO (4 RO in total) Note: U hemispherical/conical, with deep CG near mid U (3) Distrib: meso-bathypelagic but also found in the upper layers in the whole North Atlantic

Scyphomedusae  Family Rhizostomatidae 8 pairs of SCA on upper MO-arms (4), which are fused only proximally and terminate distally with a single terminal club and 3-winged appendages each (5). Without primary MO-opening. M with complex canal system.

D

E

Distribution and ecology: R. pulmo is mainly coastal, with hotspots in semi-enclosed areas that receive freshwater and nutrient input. It is often found in association with amphipods, crabs and young carangid fish. It is a favourite food of the leatherback turtle.

4 5

Rhizostoma pulmo (Macri, 1778) Size: diam. circa 400 mm T: without marginal T Note: U bell-shaped, translucent white-blue as the 8 thick oral arms. Edges of the 8 marginal LAP are violet. 8 RO, 16 SCA (4) Distrib: epipelagic in NASE, NECS, SARC

Scyphomedusae  Family Ulmaridae Unbranched and/or branched RC and a CC. With or without subgenital pits. The genus Aurelia has, in addition to the characters of the subfamily Aureliinae (i.e. small marginal T, GO invaginated, external subgenital pits and anastomosised RC), an U with 8–16 broad and shallow marginal LAP. Molecular studies have revealed that A. aurita, which is characterized by high morphological Aurelia aurita 6 variability, is a cryptic species. For the identification of the (Linnaeus, 1758) ephyra/post-ephyra stages of A. aurita see Holst (2012). F

Distribution and ecology: In A. aurita the planula develops in brood pouches located on the oral arms of the female medusa.

Size: diam. circa 250 mm T: filiform, very numerous (100–1000) Note: U flat and 4 long oral arms with crenulated lips. 4 horseshoe-shaped MAP (6) and purple/violet GO. 8 marginal LAP Distrib: cosmopolitan, usually found inshore in NASE, NECS, NWCS, SARC

FIGURE 93:  Pelagia noctiluca: A, adult, lateral view; B, detail of umbrella margin. Periphylla periphylla: C, adult, lateral view. Rhizostoma pulmo: D, adult, lateral view; E, detail of umbrella margin. Aurelia aurita: F, adult, lateral view. A, C and D, Mayer (1910); B, Russell (1970); E and F, Stiasny (1923).

230 ta xonom y

References Agassiz, L. (1849). Contributions to the natural history of the Acalephae of North America, pt. I. On the naked-eyed medusae of the shores of Massachusets in their perfect states of development. Memoirs of the American Academy of Arts and Sciences, n. ser. 4 (9): 221–316. Bigelow, H. B. (1909). Reports on the scientific results of the expedition to the eastern tropical Pacific, in charge of Alexander Agassiz, by the U. S. Fish Commission steamer ‘Albatross’ from October, 1904, to March, 1905. XVI. Medusae. Bulletin of the Museum of Comparative Zoology at Harvard University 37, 1–243. Boero, F., Bouillon, J., & Kubota, S. (1997). The medusae of some species of Hebella Allman, 1888, and Anthohebella gen. nov. (Cnidaria, Hydrozoa, Lafoeidae) with a world synopsis of the species. Zoologische Verhandelingen, Leiden, 310, 1–53. Courtesy Naturalis Biodiversity Center, Leiden, the Netherlands. Boero, F., Bouillon, J., Gravili, C., Miglietta, M. P., Parsons, T., and Piraino, S. (2008). Gelatinous plankton: irregularities rule the world (sometimes). Marine Ecology Progress Series 356, 299–310. Bouillon, J. (1984a). Hydroméduses de la Mer de Bismarck (Papouasie, Nouvelle-Guinée), pt. IV. Leptomedusae (HydrozoaCnidaria). Indo-Malayan Zoology, 1, 25–11. Bouillon, J. (1984b). Révision de la famille des Phialuciidae (Kramp, 1955) (Leptomedusae, Hydrozoa, Cnidaria), avec un essai de classification des Thecatae-Leptomedusae. IndoMalayan Zoology, 1, 1–24. Bouillon, J., Boero, F., and Seghers, G. (1991). Notes additionelles sur les méduses de Papouasie Nouvelle-Guinée (Hydrozoa, Cnidaria) IV. Cahiers de Biologie Marine, 32, 387–411. Bouillon, J. and Boero, F. (2000). Phylogeny and classification of Hydroidomedusae. Thalassia Salentina, 24, 1–296. Bouillon, J., Pagès, F. and Gili, J.-M. (2001). New species of benthopelagic hydroidomedusae from the Weddell Sea. Polar Biology, 24, 839–845. Bouillon, J., Gravili, C., Pagès, F., Gili, J., and Boero, F. (2006). An Introduction to Hydrozoa, Paris, France, Publications Scientifiques du Muséum. Brinckmann, A. (1964). Observations on the structure and development of the medusa of Velella velella (Linne 1758). Videnskabelige Meddelelser fra Dansk naturhistorisk Forening bd 126, 327–36. Courtesy Danish Natural History Society, Natural History Museum of Denmark. Browne, E. T. (1906). IX. Biscayan Plankton. Part IX.–The Medusæ. Transactions of the Linnean Society of London. 2nd Series: Zoology 10, 163–87. Carré, D. and Carré, C. (1990). Complex reproductive cycle in Eucheilota paradoxica (Hydrozoa, Leptomedusae): Medusae, polyps and frustules produced from medusa stage. Marine Biology 104, 303–10. Cornelius, P. F. S. (1995). North-west European thecate hydroids and their medusae (Cnidaria, Leptolida, Leptothecatae). Synopses of the British Fauna pt 1: 1–347; pt. 2: 1–386. Field Studies Council. Daly, M., Brugler, M. R., Cartwright, P., et al. (2007). The phylum Cnidaria: A review of phylogenetic patterns and diversity 300 years after Linnaeus. Zootaxa 1668, 127–82.

Edwards, C. (1972). The hydroids and the medusae Podocoryne areolata, P. borealis and P. carnea. Journal of the Marine Biological Association of the United Kingdom 52, 97–144. Edwards, C. (1973). The hydroid Trichydra pudica and its medusa Pochellapolynema. Journal of the Marine Biological Association of the UK, 53, 87–92. Edwards, C. (1978). The hydroids and medusae Sarsia occulta sp. nov., Sarsia tubulosa and Sarsia loveni. Journal of the Marine Biological Association of the UK, 58, 291–311. Fewkes, J. W. (1886). Report on the Medusae collected by the U.S. Fish Commission Steamer ‘Albatross’ in the Region at the Gulf Stream in 1883–84. Reports of the Commission of Fish and Fisheries U.S.A. for 1884. Gili, J.-M., Bouillon, J., Pagès, F., Palanque, A. and Puig, P. (1999). Submarine canyons as habitats of prolific plankton populations: three new deep-sea hydroidomedusae in the western Mediterranean. Zoological Journal of the Linnean Society of London 125, 313–29. Grenacher, H. and Noll, F.C. (1876). Beiträge zur Anatomie und Systematik der Rhizostomeen. Abhandlungen der Senckenbergischen Naturforschenden Gesellschaft 10, 119–80. Hartlaub, C. (1897). Die Hydromedusen Helgolands. Wissenschaftliche Meeresuntersuchungen, 2, 449–536, plates 14–23. Hartlaub, C. (1909). Méduses, in Orléan s L. P. R. DUC D’ (ed.), Croisière océanographique accomplie à bord de la ‘Belgica’ dans la mer du Grönland 1905. Imprimerie scientifique C. Bulens, Bruxelles, 463–78. Hartlaub, C. (1914). XII. Craspedote Medusen, I. Teil. 3. Lief.: Familie IV Tiaridae. Nordisches Plankton 6, 237–363. State Library Berlin- Preussischer Kulturbezitz. Shelf Mark: Lh 5028–6. Hincks, T. (1868). The History of the British hydroid zoophytes, volume 1: I–lXVII, 338 p.; volume 2: 67 plates. London, UK, John van Voorst. Holst, S. (2012). Morphology and development of benthic and pelagic life stages of North Sea jellyfish (Scyphozoa, Cnidaria) with special emphasis on the identification of ephyra stages. Marine Biology 159, 2707–22. Hosia, A. and Pagès, F. (2007). Unexpected new species of deepwater Hydroidomedusae from Korsfjorden, Norway. Marine Biology 151, 177–84. Hyman, L. H. (1940). The Invertebrates, vol. I. Protozoa through Ctenophora, McGraw-Hill: London. Kramp, P. L. (1913). Medusae collected by the ‘Tjalfe’ Expedition. Videnskabelige Meddelelser fra dansk Naturhistorisk Forening 65, 257–86. Kramp, P. L. (1919). Medusae, pt. 1. Leptomedusae. Danish Ingolf Expedition 5, 1–111. Kramp, P. L. (1920). Anthomedusae and Leptomedusae. Report on the Scientific Results of the ‘Michael Sars’ North Atlantic Deep-Sea Expedition 1910, pp. 1–14. Kramp, P. L. (1932). A revision of the medusae belonging to family Mitrocomidae. Videnskabelige Meddelelser fra Dansk naturhistorisk Forening 92, 305–83. Courtesy Danish Natural History Society, Natural History Museum of Denmark.

cnida r i a: scy phozoa a nd non-coloni a l h y drozoa  231 Kramp, P. L. (1933). Craspedote Medusen, III. Leptomedusen. Nordisches Plankton 22, 541–602. State Library BerlinPreussischer Kulturbezitz. Shelf Mark: Lh 5028–6. Kramp, P. L. (1947). Medusae, pt. III. Trachylina and Scyphozoa, with zoogeographical remarks on all the medusae of the northern Atlantic. Danish Ingolf Expedition 5, 1–66. Courtesy Danish Natural History Society, Natural History Museum of Denmark. Kramp, P. L. (1959a). The Hydromedusae of the Atlantic Ocean and adjacent waters. Dana-Report 46, 1–279. Kramp, P.L. (1959b). Some new and little known Indo-Pacific medusae. Videnskabelige Meddelelser fra dansk Naturhistorisk Forening, 121, 223–59. Courtesy Danish Natural History Society, Natural History Museum of Denmark. Kramp, P. L. (1961). Synopsis of the medusae of the world. Journal of the Marine Biological Association of the UK 40, 1–469. Lewis, C. and Long, T. A. F. (2005). Courtship and reproduction in Carybdea sivickisi (Cnidaria: Cubozoa). Marine Biology, 147: 477–83. Lindsay, D. J., Pagès, F., Corbera, J. et al. (2008). The anthomedusan fauna of the Japan Trench: preliminary results from in situ surveys with manned and unmanned vehicles. Journal of the Marine Biological Association of the United Kingdom 88, 1519–39. Lindsay, D.J., Grossmann, M.M., Bentlage, B., Collins, A.G., Minemizu, R., Hopcroft, R.R., Miyake, H., Hidaka-Umetsu, M. and Nishikawa, J. (2017) The perils of online biogeographic databases: A case study with the “monospecific” genus Aegina (Cnidaria, Hydrozoa, Narcomedusae). Marine Biology Research 13:5, 494–512. DOI: 10.1080/17451000.2016.1268261. Maas, O. (1905). Die Craspedoten Medusen der Siboga-Expeditie. Siboga Expedition, Monograph 10, 1–84, plates 1–14. Mayer, A. G. (1910). Medusae of the world. Publications, Carnegie Institution of Washington. Mianzan, H. W. and Cornelius, P. F. S. (1999). Cubomedusae and Scyphomedusae. In: Boltovskoy, D. (ed.) South Atlantic Zooplankton. Leiden, the Netherlands: Backhuys Publishers. Mills, C.E. (2001). Jellyfish blooms: are populations increasing globally in response to changing ocean conditions? Hydrobiologia 451, 55–68. Moore, S.J. (1987). Redescription of the leptomedusan Blackfordia virginica. Journal of the Marine Biological Association of the UK 67(1–2), 287–91. Morri, C. (1981). Idrozoi Lagunari. Guide per il riconoscimento delle specie animali delle acque lagunari e costiere italiane AQ1/94 6: 7–105. Naumov, D. V. (1960). Hydroids and Hydromedusae of the USSR. Keys to the fauna of the USSR. Zoological Institute of the Academy of Science of the USSR 70, 1–660 [Translated from Russian to English by Israel Program for Scientific Translations, 1969]. Naumov, D. V. (1969). Hydroids and Hydromedusae of the U.S.S.R. Fauna S.S.S.R. 70, 1–660, plates 1–30, figures 1–463 (Jerusalem, Israel Program for Scientific Translations Cat. No. 5108). Pagès, F., Bouillon, J. & Gili, J.-M. (1991). Four new species of Hydromedusae (Cnidaria, Hydrozoa) from the coast of south-western Africa. Zoologica Scripta 20, 89–98. Pagès, F., Gili, J. -M., and Bouillon, J. (1992). Medusae (Hydrozoa, Scyphozoa, Cubozoa) of the Benguela Current (southeastern Atlantic). Scientia Marina 56, 1–64.

Pitt, K. A. and Lucas, C. H. (eds) (2014). Jellyfish blooms. Springer Science and Business Media Dordrecht. Purcell, J. E. (1991). A review of cnidarians and ctenophores feeding on competitors in the plankton. Hydrobiologia 216, 335–42. Purcell, J. E. and Arai, M. N. (2001). Interactions of pelagic cnidarians and ctenophores with fish: a review. Hydrobiologia 451, 27–44. Purcell, J. E., Uye, S.-I. and Lo, T. (2007). Anthropogenic causes of jellyfish blooms and their direct consequences for humans: a review. Marine Ecology Progress Series 350, 153–74. Pyataeva, S. V., Hopcroft, R. R., Lindsay, D. J. and Collins, A. G. (2016). DNA barcodes unite two problematic taxa: the meiobenthic Boreohydra simplex is a life-cycle stage of  Plotocnide borealis (Hydrozoa: Aplanulata). Zootaxa 4150, 85–92. Repelin, R. (1965). Quelques méduses de l’Ile Anno Bon (Golfe de Guinée). O.R.S.T.O.M. Océanographiques 3, 73–9. Russell, F. S. (1953). The Medusae of the British Isles. Anthomedusae, Leptomedusae, Limnomedusae, Trachymedusae and Narcomedusae Cambridge University Press: Cambridge, UK. Russell, F. S. (1959). Some observations on the scyphomedusa Atolla. Journal of the Marine Biological Association of the United Kingdom 38, 33–40. Russell, F. S. (1963). Hydromedusae. Families: Campanulariidae, Lovenellidae, Phialellidae. ICES Identification Leaflets for Plankton No 101. Russell, F. S. (1970). The Medusae of the British Isles. II. Pelagic Scyphozoa with a supplement to the first volume on Hydromedusae. Cambridge University Press: Cambridge. Schuchert, P. (ed.) (2012). North-West European Athecate Hydroids and their Medusae, The Linnean Society of London: London. Stiasny, G. (1921). Studien über Rhizostomeen mit besonderer Berücksichtigung der Fauna des Malayischen Archipels nebst eine Revision des Systems. Capita Zool. 1(2), 1–179, figs. 1–17, pls. 1–5, tabs. 1–3, 1 ‘Schema’. (Copepoda: 60). Stiasny, G. (1923). Das Gastrovascularsystem als Grundlage fuer ein neues System der Rhizostomeen. Zoologische Anzeiten 57, 241–7, 17 text-figs. Sugiura, Y. (1973). On the polyp and medusa of the Hydromedusa, Gastroblasta chengshanensis Ling. In: Tokioka, T.  & Nishimura, S. (eds.) Recent trends in research in coelenterate biology. The Proceedings of the Second International Symposium on Cnidaria 20, 209–220. Publications of the Seto Marine Biological Laboratory. Tregouboff, G. and Rose, M. (1957). Manuel de planctonologie mediterranéenne. Tome I & II., Centre national de la recherche scientifique: Paris. Vanhöffen, E. (1902). Die Acraspeden Medusen der deutschen Tiefsee-Expedition 1898–1899. Die Craspedoten Medusen der deutschen Tiefsee Expedition 1898–1899, I. Trachymedusen. Wissenschaftliche Ergebnisse der Deutschen Tiefsee-Expedition auf dem Dampfer ‘Valdivia’ 1898–1899 3, 1–52, 55–86. Wang, C., Huang, J., Xiang, P. et al. (2014). Hydromedusae from the Arctic in 2010 during the 4th Chinese National Arctic Research Expedition (CHINARE 4). Acta Oceanologica Sinica 33(6), 95–102. Xu, Z. and Zhang, J. (1978). On the Hydromedusae, Siphonophores and Scyphomedusae from the coast of the east Guangdong Province and South Fujian Province, China. Acta Scientiarum Naturalium Universitatis Amoiensis 17, 19–64.

CNIDARIA: COLONIAL HYDROZOA (SIPHONOPHOR AE) Pr iscilla Lica ndro, Claude Ca r r é, a nd Dhuga l J. Lindsay 1 Introduction The order Siphonophorae encompasses highly polymorphic, colonial, mainly marine Hydrozoa. With very few exceptions, siphonophores are pelagic organisms that can be found the whole year round, sometimes in a characteristic season, inshore and offshore at all latitudes and depths (Alvariño, 1971). As in all hydrozoans, they carry tentacles equipped with stinging cells (nematocysts), which are used by the colony to immobilize and kill their prey. About two-thirds of the 160 currently known species belong to the suborder Calycophorae (Dunn et al., 2005a), which tend to dominate in samples collected by nets. More fragile Physonectae are often seriously damaged or destroyed by the nets, but the increased use of imaging systems or other in situ survey/sampling techniques has shown that the abundance and diversity of this group has been largely underestimated.

2  Life Cycle According to Totton (1965), the fully grown colonial siphonophore is an enlarged larval nurse carrier, which buds off the sexually mature individuals, called gonophores. Most siphonophores are monoecious, i.e. they release gonophores of both sexes, even though not necessarily at the same time. Few exceptions include Physalia physalis, which being dioecious releases gonophores of only one sex. In calycophoran siphonophores (e.g. Muggiaea kochi), the sexual stage (eudoxid) is released by the colony before the sexual maturation of the gonophore (Fig. 95A). Female/male eudoxids feed and develop independently. At full maturity they release the gametes for the external fertilization that produces a free-living planula larva, which matures into a calyconula larva and subsequently develops into a new colony (Fig.  95A). Physonect siphonophores have a similar development (Fig. 95B), even though their late larva, called a siphonula, is morphologically different from the calyconula, being equipped with an ­apical, gas-filled float (pneumatophore).

FIGURE 94:  Siphonophorae: Calycophorae Chelophyes appendiculata. Photo courtesy of C. Carré

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

cnida r i a: coloni a l h y drozoa  233 mature eudoxid

A

planula young calyconula

Eudoxid maturation

Larval development

young eudoxid Polygastric stage

mature calyconula

mature colony young colony

structure, which represents the ‘fishing posture’. This behaviour has been named the ‘veronica’, after the classic toreador movement that it resembles. Calycophoran siphonophores eat primarily small copepods, but also other plankton including ostracods, molluscs, chaetognaths, and larvae of euphausiids and fish (Mapstone, 2009 and references therein). In turn, they are prey of bigger cnidarians, ctenophores, heteropods, and of several fish species (Mapstone, 2009). Assimilation efficiencies of oceanic Calycophorae are typically higher than other planktonic carnivores: 87–90% for carbon and 90–96% for nitrogen (Purcell, 1983). Because of their colonial morphology, siphonophores may rapidly become dominant under favourable conditions, as they are able to release hundreds of eudoxids, each one producing a new colony. Swarms of siphonophores can have a significant predatory impact on the abundance of other planktonic organisms including small fish (Mackie et al.,  1987; Purcell,  1997), causing massive mortalities of farmed fish (Greve,  1994; Båmstedt et al., 1998).

B gonophore

4  General Morphology

egg

planula single cormidium

siphonula mature colony

young colony FIGURE 95: Life cycle of Siphonophorae. A, The calycophoran siphonophore Muggiaea kochi. B, The physonect Agalma sp. Modified from Carré and Carré (1991); B, Courtesy of A. Fischer.

3 Ecology Siphonophores are amongst the most abundant carnivores in the marine system (Mackie et al., 1987 and references therein; Mapstone, 2009). They are generally passive, ambush predators that deploy a network of tentacles and then remain motionless, capturing prey that come into contact with their nematocyst batteries (Mackie et al. 1987). The polygastric stage of M. atlantica employs a specialized swimming behaviour that spreads the  siphosome and tentacles into a three-dimensional helical

Each siphonophore colony, otherwise called the polygastric stage, is composed of a collection of zooids (Fig. 96A): (i) medusozoid zooids, i.e. asexual swimming bells (nectophores) and sexual gonophores, which alone or in clusters constitute the reproductive unit (gonozooid); (ii) polypoid zooids, which are used to regulate the buoyancy of the colony (pneumatophore), for feeding and digestion (gastrozooids) or for manipulation (dactylozooids and palpons). All zooids are arranged along a contractile stem, which is a tube surrounding the main gastrovascular canal. The apex of the stem, or anterior/aboral pole, carries the nectophores and is called the nectosome; the distal posterior/oral pole, which carries the remaining zooids and ends with the oldest larval zooids, is called the siphosome (Fig. 96A). The Siphonophora are divided into three suborders based on the presence/absence of the pneumatophore and of nectophores. Thus, Cystonectae siphonophores only have a pneumatophore, Calycophorae lack a pneumatophore but develop nectophores, whereas Physonectae have both (Fig. 96). The morphology of the nectosome and in particular of the nectophores is an important taxonomic character. Nectophores can be attached to either the dorsal/upper or ventral/lower side of the nectosome, depending on the family. Each nectophore, corresponding to a highly modified medusa, is composed predominantly of mesoglea, which is often characterized by ridges or folds useful for taxonomic identification. The internal cavity or nectosac corresponds to the sub-umbrella and bears the radial canals (two lateral, one upper, and one lower) that have different shapes and lengths depending on the species and can be connected through a pedicular canal to a pallial or mantle canal (MC) (Fig. 97). The mantle canal always has an ascending branch (AMC) and sometimes also has a descending branch (DMC), which is a feature of taxonomic importance to distinguish different groups of Physonectae. The nectosac is open externally

234 ta xonom y A

C AP ridge thrust block

PN

Anterior pole Nectosome

NE

E

AP wings

BR

F

PN

G

PH

Anterior pole ANT_NE NES

NES

LAT ridge

BR

GON

D Siphosome

RC

GA

GON

GA

SOM

STE basal_facet

T

MO_PL

female GON

STE

attachment point of the STE POST_NE

DA male GON

B

GA TEN Posterior pole

STE

OS with ostial teeth

Key to annotations

ANT AP BR DA BR GA GON HYD PAL LAM LAT MO NE NES

= Anterior = Apical = Bract = Dactylozooid = Gastrozooid = Gonophore = Hydroecium = Lamella = Lateral = Mouth = Nectophore = Nectosac

OS PAL PH PL PN POST RC SOM STE T TEN

HYD T

= Ostium = Palpon = Phyllocyst = Plate = Pneumatophore TEN = Posterior = Radial canal = Somatocyst siphosomal STE = Stem basal_LAM Posterior pole = Tentacle = Tentilla

FIGURE 96:  General structure of siphonophores. A–D, Physonectae. A, Agalma elegans, whole animal, lateral view. B, Distal/posterior part of

the siphosome. C, Nectophore, upper view. D, Nanomia bijuga, detail of stem bearing gonophores and dactylozooids. E, Cystonectae, Rhizophysa filiformis, lateral view. F,G, Calycophorae, F, Muggiaea kochi, young eudoxid, lateral view. G, Sulculeolaria quadrivalvis, polygastric stage, lateral view. A–C, Totton (1965); D, photo C. Carré; E, Pagès and Gili (1992); F, Carré and Carré (1991); G, Carré (1979).

to the ostium, which is surrounded by a circular or ring canal. On the lower side of the nectophore lies a cavity called the hydroecium that, in the Calycophorae, serves to protect the siphosomal budding zone when the whole or a part of the stem is retracted into it, for defense or during locomotion. In calycophorans, from the hydroecium runs the somatocyst, sometimes containing oil droplets, that is an extension of the original larval gastrovascular system and is connected with the radial canals. The mouth plate or basal lamella is a lower/ventral process below the ostium. Calycophorans usually have one or two nectophores only (i.e. the anterior and posterior nectophores, Fig. 96G), whereas physonects have variable numbers of nectophores, depending on the s­ pecies (Fig. 96A). The siphosome is generally much longer than the nectosome, up to many metres in some physonect species. Several units called cormidia are arranged along the ventral/lower side of the siphosome, and can be retained or progressively released free from the distal part of the siphosome once they have reached sexual maturity. Each cormidium is composed, at a m ­ inimum, of: a single gastrozooid, carrying a tentacle; a male/female

gonophore; and a bract (absent in cystonects hippopodids, most Clausophyids and some other scattered species), characterized by a reduced gastrovascular canal (phyllocyst), that has floatation functionality and may contain metabolic reserves. Tentacles are long and typically bear a number of branches or tentillae, armoured with nematocysts. The structure of the tentillum is taxonomically important, as it differs in different physonect taxa (Fig.  97). A tentillum typically comprises a ­pedicel, often an involucrum, a cnidosac with a cnidoband and one or more terminal filaments. In physonects the tentillum might include swollen vesicle/s called ampulla/ae. In physonects and in the calycophoran genus Stephanophyes, the cormidium also includes a few dactylozooids or palpons. Each palpon typically bears an unbranched tentacle called a palpacle, while dactylozooids have no palpacles. In the calycophorans the detached cormidium is called an eudoxid (or monogastric stage). Cystonect and a minority of physonect colonies (e.g. Apolemiidae, Erennidae, Marrus, Pyrostephidae, Rhodaliidae, Stephanomiidae) bear gonophores of only one sex, whereas in

cnida r i a: coloni a l h y drozoa  235 Families/genera

MFZ

DMC

NO

YES

Larval type TEN, without INV, without TFI in Cordagalma (except C. ordinatum). TEN bilobate, with INV and spiralled CNIB in Cardianecta.

NO

YES

TEN without INV, with straight CNIB. TFI large and stiff with central axial canal, without N.

YES

NO

Forskalia edwardsi

TEN with long pedicel, without INV, with coiled CNIB. TFI flexible.

NO

YES

Frillagalma vityazi

TEN made by 2 consecutive AM after CNIS. No INV, CNIB, TFI.

NO

YES

TEN with complete INV, with complexely coiled red CNIB. 8 TFI around large AM.

NO

YES

TEN without INV, with straight/ loosely coiled CNIB. TFI flexible.

YES

NO

TEN without INV, with straight CNIB. TFI flexible.

YES

NO

2 types of TEN with complete INV: with coiled CNIB proximally and zigzagged CNI distally.

NO

YES

TEN without INV, with loosely coiled CNIB. TFI long and flexible.

YES

NO

AM

TEN

TFI CNIB

AGALMATIDAE sensu strictu

A

INV

B

PE Agalma elegans

Nanomia bijuga

ST CORDAGALMATIDAE

Cordagalma spp.

C

PE

CNIS

CNIC

CNIB ERENNIDAE

photophore Erenna richardi

PE TFI

D CNIB

PE

FORSKALIIDAE

E

ST

Frillagalma spp.

PE

F TFI

AM

CNIB

Lychnagalma spp.

TFI

AM

CNIS

TEN with INV, tricornuate (i.e. with 2 TFI and AM) in Agalma. Unicornuate (1 TFI) in the other genera. CNIB tightly coiled.

Lychnagalma utricularia

G TFI Marrus spp.

Marrus orthocanna

PE CNIB

H

TFI

I PYROSTEPHIDAE

Bargmannia elongata ST

INV RESOMIIDAE

CNIB

J

CNIB TFI

L

MFZ

N

thrust block

Stephanomia amphytridis

thrust block

AMC DMC

PC

lateral RC lower RC

Stephanomia amphytridis

TFI

TFI

CNIB

STEPHANOMIIDAE

M

Resomia convoluta INV

PE PE

K

Key to annotations AM = Ampulla AMC = Ascending mantle canal CNIB = Cnidoband CNIC = Cnidocils CNIS = Cnidosac DMC = Descending mantle canal INV = Involucrum MFZ = Muscle free zone

N PC PE RC ST TEN TFI

= Nematocyst = Pedicular canal = Pedicel = Radial canals = Stenotele = Tentilla = Terminal filament

Halistemma rubrum

FIGURE 97:  Characteristics of taxonomic importance to identify Physonectae families in Group III (see Fig. 100). A–L, schematic representation

of tentilla, lateral views; M, N, nectophores, lower views. A–C, H, Mapstone (2009); D–F, I–K, Mapstone (2014); G, Pugh and Harbison (1986); L, modified from Pugh and Baxter (2014); M, N Pugh and Baxter (2014).

236 ta xonom y calycophorans and most physonects the colonies are hermaphrodite as they bear gonophores of both sexes.

5 Systematics Here we generally adopt the classification from the World Register of Marine Species, which is the most up-to-date classification of the group. Over the years the systematics of this group has been repeatedly revised and some authors have raised the order Siphonophora to a subclass of the class Hydrozoa (e.g. Pugh,  1999; Bouillon et al.,  2006). Molecular phylogenetic ­analyses show that Cystonectae are separated from  all other ­siphonophores (Dunn et al.,  2005a; Dunn and Wagner, 2006). The physonect family Agalmatidae is a polyphyletic group that is slowly undergoing taxonomic revision (e.g. Pugh, 2016). Taking into account recent updates, we consider Agalmatidae sensu strictu (Mapstone,  2009; Pugh,  2016) separately from Marrus spp., Frillagalma spp. and Lychnagalma spp. The keys for identification are based on the main morphological features of nectophores and bracts, as those are the components of the colony that are usually caught using ­ ­plankton nets.

Box 1  Classification of Siphonophorae Phylum Cnidaria   Class Hydrozoa   Order Siphonophorae    Suborder Cystonectae     Family Physaliidae     Family Rhizophysidae     Family Bathyphysidae    Suborder Physonectae     Family Agalmatidae     Family Apolemiidae     Family Cordagalmatidae     Family Erennidae     Family Forskaliidae     Family Physophoridae     Family Pyrostephidae     Family Resomiidae*     Family Rhodaliidae     Family Stephanomiidae    Suborder Calycophorae     Family Abylidae     Family Clausophyidae     Family Diphyidae     Family Hippopodiidae     Family Prayidae     Family Sphaeronectidae * North Atlantic species not known in this family. Families in bold are covered in this chapter.

SIPHONOPHORA GROUP I - Suborder: CYSTONECTAE (*), PHYSONECTAE PN

yes

Various NE organized to form a nectosome. Usually numerous BR

no

Large PN (1). yes Reduced NE, without nectosome

PN horizontal purplish (1)

yes * PHYSALIIDAE

1 no

Physalia physalis

no PN oval rounded (2) yes yes

SIPHONOPHORA GROUP II

young GA with wings (3)

2 no

with winged young GA (3)

yes

with simple GA

GA old GA without wings

yes

APOLEMIIDAE Apolemia uvaria

SIPHONOPHORA GROUP III 5

Nectosome elongated. Siphosome shortened to form a sac on which cormidia are arranged spirally. Many large PAL arranged in a ring (5) to protect the other cormidial elements

PHYSOPHORIDAE

yes

Physophora hydrostatica

B

6

Nectosome and siphosome contracted, forming a solid corm (6)

yes

RHODALIIDAE

Stephalia corona, (a) expanded and (b) contracted FIGURE 98:  Siphonophora group I: suborder Cystonectae(*), Physonectae. Physalia physalis, Stephalia corona (a,b), after Totton (1965); Rhizophysa filiformis, after Pagès

and Gili (1992); Bathyphysa sibogae, after Biggs and Harbison (1976); Apolemia uvaria, Physophora hydrostatica, after Kirkpatrick and Pugh (1984).

cnida r i a: coloni a l h y drozoa  237

yes

no 6

* BATHYPHYSIDAE

4

no Nectosome and siphosome elongate, with a narrow STE

A

Rhizophysa filiformis

yes

Bathyphysa sibogae

T present between NE on nectosome. NE deeply hollowed axially (4), small/delicate BR

* RHIZOPHYSIDAE

yes

NE flattened in upper/lower direction.

yes

BR absent. Up to about 16 similar NE, bearing protuberances/spines (1), with large and shallow NES.

Hippopodius hippopus

no yes

NE and BR rounded, smooth, with thick mesoglea

NE

yes

2 NE very different in size, with reduced SOM. NES of smaller NE is reduced

Amphicaryon acaule

HYD yes

1 rounded fragile larval NE, with simple SOM. Small BR with a single canal NE and BR pointed, toothed/ of irregular shape

POST_NE with SOM

SPHAERONECTIDAE Sphaeronectes koellikeri

no PRAYIDAE Prayinae

yes

2 NE of approx. equal size, SOM simple/branched. BR have canals with 5-6 branches

yes

yes

PRAYIDAE Amphicaryoninae

no

no

2 NE (ANT, POST)

HIPPOPODIIDAE

1

Rosacea cymbiformis no

1 NE, usually large, with simple/toothed ridges. SOM usually branched (2). Large BR with branched canals. Clausophyes tropica, Dimophyes arctica, Muggiaea spp. also only ANT NE. yes

yes

2

ANT_NE with large, ventral/lower opening of HYD (3). Small BR with PH and 2 canals or BR absent (Clausophyes spp.)

yes

HYD

4

Chuniphyes multidentata 3

yes POST_NE Bassia bassensis

no ANT_NE of similar size/bigger than POST_NE and often with shallow HYD. Conical BR

CLAUSOPHYIDAE

ANT_NE

no ANT_NE angular, larger than POST_NE and with a deep HYD (4). Rigid, angular BR.

PRAYIDAE Nectopyramidinae

Nectadamas diomedeae

yes

ABYLIDAE

HYD Chelophyes appendiculata

DIPHYIDAE

FIGURE 99:  Siphonophora group II: suborder Calycophorae. Hippopodius hippopus, Amphicaryon acaule, Sphaeronectes koellikeri, Chuniphyes multidentata, Chelophyes appendiculata, after Kirkpatrick and Pugh (1984); Rosacea cymbiformis, after Tregouboff and Rose (1957); Nectopyramis diomedeae, after Totton and Fraser (1955a); Bassia bassensis, after Totton and Fraser (1955b).

238 ta xonom y

SIPHONOPHORA GROUP II - Suborder: CALYCOPHORAE

SIPHONOPHORA GROUP III - Suborder: PHYSONECTAE 1 yes

NE arranged in spiral. Each NE usually asymmetrical (1), flattened in upper/lower direction

FORSKALIIDAE 3

no

Forskalia edwardsi NE bilaterally symmetrical, arranged in 2 series on nectosome

NE with lower LAT wings (2). Much enlarged triangular thrust block (3)

yes

yes

PYROSTEPHIDAE

4

no NE with AP wings (4)

2

yes

NE with MFZ

no

yes

TEN with straight CNIB, without INV. TFI straight

Bargmannia elongata

yes

ERENNIDAE Erenna richardi

no TEN with loosely coiled/straight CNIB, without INV. TFI coiled

yes

yes

LAT-RC with major loops (5)

5

no

STEPHANOMIIDAE Stephanomia amphytridis

LAT-RC straight (6)

yes

Marrus spp.

6 Marrus orthocanna

yes

NE heart-shaped

yes

Frillagalma spp.

no

Frillagalma vityazi yes

yes

2 types of TEN on the same T

RESOMIIDAE

no

Resomia ornicephala

TEN tightly coiled, uni/tricornuate

AGALMATIDAE sensu strictu Agalma elegans

no TEN complexely coiled, 8 TFI

yes

yes

Lychnagalma spp. Lychnagalma utricularia

FIGURE 100:  Siphonophora group III: suborder Physonectae. Forskalia edwardsi, Bargmannia elongata, Agalma elegans after Kirkpatrick and Pugh, (1984); Erenna richardi, after Pugh (2001); Stephanomia amphytridis, after Pugh and Baxter (2014); Cordagalma ordinatum, after Pugh (2016); Frillagalma vityazi, after Pugh (1998); Resomia ornicephala, after Pugh and Haddock, (2010); Lychnagalma utricularia, after Pugh and Harbison (1986); Marrus orthocanna, after Dunn et al. (2005b); Agalma elegans, after Kirkpatrick and Pugh (1984).

cnida r i a: coloni a l h y drozoa  239

larval-type TEN, without CNIB

coiled TEN

CORDAGALMATIDAE

Cordagalma ordinatum

no

Chelophyes appendiculata medium size SOM medium size HYD

quandrangular section

Muggiaea kochi

long SOM deep HYD

Hydroecium well developed

Muggiaea atlantica

NE spirally twisted pentagonal section

Eudoxoides spiralis

sub-spherical SOM on long pedicel Lensia subtilis

oval section

slightly 5-ridged

can be pigment spots Lensia campanella SOM club-shaped globular SOM with no pedicel

round apex

round section

vertical MO_PL

Lensia meteori

tapered SOM on short peduncle Lensia conoidea

pentagonal section

Hydroecium virtually absent or shallow

globular SOM Lensia fowleri quite thin SOM pointed apex

polygonal section

FIGURE 101:  Key to main Calycophorae species.

Lensia multicristata

240 ta xonom y

KEY TO MAIN CALYCOPHORAE SPECIES

Order Siphonophora Suborder Calycophorae  Family Abylidae Rigid, angular NE. POST_NE without SOM, usually much larger than the ANT_NE, with serrated ridges and teeth. SOM in ANT_NE usually curved ventrally.

Abyla spp. A

C

B

NES

HYD

SOM

2

Abyla bicarinata

E

D

PH

1

ANT_NE: with 10/11 facets. AP facet divided by a transverse ridge, with ridges often serrated NES: long, up to the apex of NE HYD: long, up to the apex of NE SOM: large, oval, lies on lower side POST_NE: with long AP apophysis. It has only 4 ridges, lower ones highly serrated basally. Toothed comb on left wing of HYD (1). 5 serrated OS teeth BR: prismatic, with 6 facets, the upper one rectangular. PH very large Note: ANT_NE of A. bicarinata has wing-like processes (2) and edges of facets rounded. In A. trigona most of the ridges are heavily serrated

F

A. trigona Quoy & Gaimard, 1827

Quoy & Gaimard, 1827 Size: ANT_NE ~6 mm H, BR~ 3 mm

Size: ANT_NE ~6.5 mm H, POST_NE ~30 mm Distribution: Epipelagic species, both found in NASE and CNRY. A. trigona also in NASW, NASDR

NES

G

H

AP peduncle

I

K

Abylopsis spp.

L

ANT_NE: 7 facets, without AP facet. Pentagonal upper/lower facets NES: cylindrical shape HYD: partly between SOM and NES. SOM: with AP peduncle POST_NE: short, curved AP apophysis. 5 ridges. Toothed comb on both wings of HYD BR: with 7 facets. PH with 2 swollen LAT branches, 1 narrow descending branch and 1 AP peduncle Note: in the ANT_NE of A. tetragona LAT_RC form an ascending loop (3), absent in A. eschscholtzii

J PH HYD

SOM

3

Abylopsis eschscholtzii

A. tetragona

(Huxley, 1859)

(Otto, 1823)

Size: ANT_NE ~5 mm H, Size: ANT_NE up to 5 mm H, POST_NE 6.5 mm H, BR 3 mm H POST_NE 20 mm H, BR 4 mm H Distribution: Both species found in epipelagic waters in NATR and NWCS. A. tetragona also common in CNRY, NASE, NASW

M

SOM

Bassia spp.

N

PH 4

Bassia bassensis (Quoy & Gaimard, 1833)

O Size: ANT_NE up to 4 mm H, POST_NE 15 mm H, BR 5 mm H Distribution: Epipelagic in CNRY, NASE, NADR, NASW, NATR, NECS, NWCS

Ceratocymba spp.

ANT_NE: polyhedric. NES: cylindrical shape HYD: does not reach SOM and NES SOM: without AP peduncle POST_NE: 4 ridges, with basal teeth BR: with 7 facets, rhomboidal on upper face and pentagonal on lower. PH is a long tube swollen at the apex Note: in the ANT_NE of B.bassensis the HYD does not reach the zone between SOM and NES (4)

Q

P

R 5

Ceratocymba dentata (Bigelow, 1918)

SOM

C. sagittata (Quoy & Gaimard, 1827)

ANT_NE: 7 facets, AP facet undivided POST_NE: long and narrow, without wing-like expansions. Short upper ridge ends on an upper tooth BR: characteristic shape, roughly triangular with a concave AP facet, prominent LAT horns and a median dorsal ridge. PH with 2 thin ventro-LAT branches Note: ANT_NE of C. sagittata has pointed apex and NES much longer than SOM (5). ANT_NE of C. dentata has LAT_margins deeply bowed and serrated Size: C. dentata ANT_NE ~12.5 mm H, BR ~13 mm Size: C. sagittata ANT_NE ~19 mm H, BR~ 21 mm Distribution: both species found in the epipelagic in CNRY, NASE, NASW, NATR. C. sagittata also in NADR

FIGURE 102:  Abyla bicarinata: A, B, anterior nectophore, lower and lateral views; C, eudoxid, lateral view. A. trigona: D, E, anterior nectophore,

lower and lateral views; F, posterior nectophore, lateral view. Abylopsis eschscholtzii: G, anterior nectophore, lower view; H, posterior nectophore, lateral view; I, bract, lateral view. A. tetragona: J, anterior nectophore, lower view; K, posterior nectophore, lateral view; L, bract, lateral view. Bassia bassensis: M, polygastric stage, lower view; N, bract, lateral view. Ceratocymba dentata: O, anterior nectophore, lower view; P, bract, lateral view. C. sagittata: Q, anterior nectophore, lower view; R, bract, lateral view. A–E, G, I–J, L, O–R, Pugh, (1999); F, Totton (1965); H, modified from Gili (1986); K, Kirkpatrick and Pugh (1984); M-N, Pagès and Gili (1992).

Order Siphonophora Suborder Calycophorae  Family Clausophyidae Chuniphyes spp.

ANT_ and POST_NE have a SOM and are streamlined. HYD more prominent than in the diphyids. BR, when present, usually has PH with 2 fine basal branches extending down into the neck shield.

ANT_NE: with 4 ridges at the pointed apex and 8 ridges at the base, ending in prominent teeth. POST_NE: 3 ridges at the apex and 6 at

Distribution and ecology: Siphonophores from this family mainly live in deep waters.

base, ending in distinct teeth. HYD extends the whole length, with 2 large asymmetrical flaps in its upper half (1) BR: flattened, not identifiable at species level. PH with 2 LAT branches Note: Ch. multidentata is characterized by the crossshaped SOM (2), while in Ch. moserae it is linear

A

B

C

1

3

Kephyes spp.

D

ANT/POST_NE: smooth, ridgeless, laterally compressed. NES >2/3 of NE length has slightly looped RC. SOM reaches ANT_NE apex (3). PC with a descending branch. In ANT_NE the extensive HYD reaches the OS (4). BR: conical, with rounded apex and long neck2

shield. PH reaches the apex

Clausophyes spp.

4

Chuniphyes multidentata

Kephyes ovata

Lens & van Riemsdijk, 1908

(Keferstein & Ehlers, 1860)

Size: ANT_NE up to 35 mm H, POST_NE 40 mm H, BR 2 mm Distribution: epi-mesopelagic in ARCT, CNRY, NADR, NASE, NASW, NECS,

Size: ANT/POST_NE up to 14 mm H, BR 8 mm Distribution: meso-bathypelagic in CNRY, NASE, NECS, SARC

NWCS, SARC

ANT/POST_NE: without ridges, laterally compressed. Nes with lateral RC with major loop before joining OS ring canal. PC with no descending branch. BR: absent, no eudoxids developed. Note: C. moserae and C. galeata are common in the North Atlantic and C. laetmata was recently reported (Hosia et al 2008).

Order Siphonophora Suborder Calycophorae  Family Diphyidae ANT and POST_NE have similar size (or POST is smaller), both streamlined but dissimilar. ANT_NE has usually large NES and often

E

G

F

5 6

HYD

Chelophyes appendiculata (Eschscholtz, 1829)

SOM with oil droplets. Diphyidae is a polyphyletic group assembling species with many different combinations of characters.

Chelophyes spp.

neck shield

ANT_NE: rigid, with 5 ridges, the upper one only shortly above ostium. Only 3 ridges reach the apex. MO_PL divided HYD: talon-shaped SOM: ~ 1/2 of NE length, spindle-shaped, on a peduncle. POST_NE: apically pointed, with 4 serrated ends. MO_PL divided in 2 strong, asymmetric teeth (5) BR: conical with small/rounded neck shield HYD: deep, ~ 1/2 of whole L. Cylindrical PH, reaching apex of BR NOTE: C. appendiculata is characterized by ANT_NE with a straight SOM (6) and POST_NE with characteristic MO_PL Size: ANT_NE up to 20 mm H, POST_NE 8 mm H, BR 4 mm H Distribution: Epi-mesopelagic in CNRY, NASE, NASW, NATR, NECS, NWCS

H

I

7

Dimophyes arctica (Chun, 1897)

J

neck shield

PH

Dimophyes spp. Presently monotypic genus for D. arctica

ANT_NE: smooth, without ridges. MO_PL undivided (7) HYD: large, extending above OS SOM: carrot-shaped, ~ 2/3 of NE length POST_NE: reduced. NES opens upper-basally BR: conical, with extensive neck shield. PH with AP- and LAT horns Size: ANT_NE up to 15 mm H, BR 10 mm H Distribution: epi-mesopelagic in ARCT, CNRY, GFST, NADR, NASE, NASW, NECS, NWCS, SARCS

FIGURE 103:  Chuniphyes multidentata: A, anterior and B, posterior nectophores, lateral views. Kephyes ovata: C, anterior and D, posterior

nectophores, lateral views. Chelophyes appendiculata: E, anterior and F, posterior nectophores, lateral views; G, eudoxid, lateral view. Dimophyes arctica: H, anterior and I, posterior nectophores, lateral views; J, eudoxid, lateral view. A–B, E–J, Kirkpatrick and Pugh (1984); C–D, Pugh (2006).

Order Siphonophora Suborder Calycophorae  Family Diphyidae Diphyes spp. A

C

B

1

1

Diphyes bojani

D

D. dispar

D. chamissonis

(Eschscholtz, 1825) Size: ANT_NE ~10 mm H, POST_NE 7 mm H, BR 4 mm H

1

E

Huxley, 1859

Chamisso & Eysenhardt, 1821

Size: ANT_NE~12 mm H, BR 5 mm H

Size: ANT_NE~36 mm H, POST_NE 27 mm H, BR 8 mm H

ANT_NE: with 5 ridges and 3 prominent ostial teeth. HYD: deep. SOM: spindle-shaped POST_NE: with long AP apophysis and 3 ostial teeth (unknown for D. chamissonis) BR: helmet-shaped Note: ANT_NE of different species characterized by depth of HYD (1/3 NE length in D. bojani, 1/2 NE length in the other species), SOM length and shape of NES apex (1) (rounded in D. chamissonis, with a diverticulum in D. dispar, pyramidal in D. bojani)

Eudoxoides spp. F

H

G

I

2

2

E. spiralis

Eudoxoides mitra (Huxley, 1859)

(Bigelow, 1911)

Size: ANT_NE ~8 mm H, BR 4 mm H

Size: ANT_NE up to 12 mm H, BR 4 mm H

K

J

ANT_NE: rigid, with 5 ridges, usually serrated, not always reaching the apex (4 AP-ridges in E. spiralis). Might be spirally twisted. MO_PL divided (2). No ostial teeth HYD: 1/2 NES length, less deep than in Chelophyes POST_NE: with curved furrow between apex and apical apophysis. Absent in E. spiralis BR: hood-shaped, with PH long reaching the apex. Long neck-shield. GON twisted with serrated edges Note: ANT_NE and GO of E. spiralis typically spirally twisted. SOM carrot-shaped in E. spiralis, pear-shaped in E. mitra Distribution: both species are epipelagic in CNRY, NASE, NASW, NATR, NWCS. E. spiralis also found in NADR and NECS

M

L

SOM

3

M. bargmannae

M. kochi

Cunningham, 1892

Totton, 1954

(Will, 1844)

Size: ANT_NE ~5 mm H, BR 2 mm H

Size: ANT_NE ~9 mm H

Size: ANT_NE ~4 mm H, BR 2 mm H

Muggiaea atlantica

Muggiaea spp. ANT_NE: pyramidal, with 5 ridges. Oblique, divided MO_PL. HYD: deep, not open ventrally SOM: very close to NES wall POST_NE: not developed BR: small/conical, with asymmetrical base. Shallow HYD, PH club-shaped Note: species identified by different shape and length of SOM. ANT_NE of M. bargmannae with LAT folds in place of ridges (3) Distribution: epipelagic. M. atlantica found in CNRY, NASE, NECS, NWCS, SARC; M. bargmannae in ARCT, NARC, SARC; M. kochi in NASE, NASW, NECS

FIGURE 104:  Diphyes bojani: A, anterior nectophore, lateral view; B, eudoxid, lateral view. D. chamissonis: C, anterior nectophore, lateral view. D.dispar: D, anterior nectophore, lateral view; E, eudoxid, lateral view. Eudoxoides mitra: F, anterior nectophore, lateral view; G, eudoxid, lateral view. E. spiralis H, anterior nectophore, lateral view; I, eudoxid, lateral view. Muggiaea atlantica: J, anterior nectophore, lateral view; K, bract. M. bargmannae: L anterior nectophore, lateral view. M.kochi: M, anterior nectophore, lateral view. A–M, Pugh (1999).

Order Siphonophora Suborder Calycophorae  Family Diphyidae A

B

Lensia spp.

D

C

E

SOM HYD

L. campanella

L. conoidea

Totton, 1941

(Moser, 1917)

(Keferstein & Ehlers, 1860)

Size: ANT_NE up to 15 mm H Distribution: mesopelagic in ARCT, CNRY, NADR, NASE, NASW, NATR, NECS, SARC

Size: ANT_NE ~6 mm H, BR 1.5 mm H Distribution: epipelagic in CNRY, NADR, NASE, NASW, NATR, NECS, NWCS

Size: ANT_NE up to 20 mm H, BR 3 mm H Distribution: epi-mesopelagic in ARCT, CNRY, NADR, NASE, NASW, NATR, NECS, SARC

Lensia achilles

F

G

H

I

ANT_NE: pyramidal, mainly with 5 to many ridges. Small divided MO_PL. No ostial teeth HYD: shallow, rarely extending above OS POST_NE: AP truncated, rounded MO_PL. Often undescribed, e.g. for L. hardy, L. hunter, L. meteori BR: helmet-shaped Note: species shown here with 5 ridges, except L. meteori and L. subtilis that have no ridges. Species characterized by number/ shape of ridges, SOM shape/length and depth of HYD

J

L. fowleri

L. hotspur

L. hunter

L. meteori

(Bigelow, 1911)

Totton, 1941

Totton, 1941

(Leloup, 1934)

Size: ANT_NE ~20 mm H Distribution: epi-mesopelagic in CNRY, NADR, NASE, NASW, NATR, NECS, NWCS

Size: ANT_NE Size: ANT_NE ~9 mm H ~6 mm H Distribution: epipelagic Distribution: in NATR epipelagic in CNRY, NASE, NASW, NATR, NECS

M

O

P

SOM Ostial tooth

Size: ANT_NE up to 26 mm H

L. subtilis (Chun, 1886)

Size: ANT_NE ~4.5 mm H Distribution: mesopelagic in CNRY, NADR, NASE, NASW, NECS

1

(Sars, 1846)

L

Size: ANT_NE ~6 mm H Distribution: epipelagic in CNRY, NADR, NASE, NASW, NATR, NECS, NWCS

Sulculeolaria spp.

N

Sulculeolaria biloba

K

S. monoica (Chun, 1888) Size: ANT_NE up to 10 mm H

S. quadrivalvis de Blainville, 1830 Size: ANT_NE up to 18 mm H, BR 4 mm H

ANT_NE: rounded apex, without ridges. Divided MO_PL HYD: virtually absent POST_NE: extensively looped LAT-RC (1) (distinctive character of the genus) BR: small, leaf-like Note: replacement ANT/POST_NE often have different characters. Species characterized by the presence/absence of ostial teeth and by the shape of SOM Distribution: all species epipelagic in CNRY, NASE, NASW, NATR. S. biloba also found in ARCT, NADR, NECS, SARC. S. quadrivalvis also in NWCS

FIGURE 105:  Lensia achilles: A, anterior nectophore, lateral view. L. campanella: B, anterior nectophore, lateral view; C, eudoxid, lateral view. L. conoidea: D, anterior nectophore, lateral view; E, eudoxid, lateral view. L. fowleri: F, anterior nectophore, lateral view; G, eudoxid, lateral view. L. hotspur: H, anterior nectophore, lateral view. L. hunter : I, anterior nectophore, lateral view. L. meteori: J, anterior nectophore, lateral view. L. subtilis: K, anterior nectophore, lateral view; L, bract, lateral view. Sulculeolaria biloba: M, anterior nectophore, lateral view. S. monoica: N, anterior nectophore, lateral view; S. quadrivalvis: O, anterior and P, posterior nectophore, lateral views. A–P, Pugh (1999).

cnida r i a: coloni a l h y drozoa  245 Order Siphonophora Suborder Calycophorae  Family Hippopodiidae can be distinguished by the shape of SOM, curving smoothly over mid-dorsal surface of HYD (1).

Up to 16 NE flattened in upper-lower axis, arranged in series of two. NE bear protuberances or spines. BR absent, GO directly arise from the siphosome. Larval NE similar to that of Prayidae-Prayinae,

Hippopodius spp. NE: horseshoe-shaped, with 4 rounded protuberances (2) forming an arc above ostium; Larval NES with only 2 RC

A

Distribution: epi-mesopelagic NE: with protuberances (3), in CNRY, NADR, NASE, spines or ridges (4). Larval NASW, NATR, NECS, NES with 4 RC NWCS

C

B

2

SOM

Vogtia spp.

3

Distribution: both mesopelagic in ARCT, CNRY, NADR, NASE, NASW, NATR, NECS. V. glabra also in NWCS

D

4

1

RC

Vogtia glabra

V. serrata

(Forsskål, 1776)

Bigelow, 1918

(Moser, 1925)

Size: NE diam. up to 20 mm. larval NE ~5 mm H

Size: NE diam. up to 30 mm

Size: NE diam. up to 40 mm

Hippopodius hippopus

Order Siphonophora Suborder Calycophorae  Family Prayidae NE quite large and usually rounded. Larval NE sometimes retained in the polygastric stage (e.g. in the subfamily Amphicaryoninae) or replaced by 1–4 definitive NE that have a SOM often complexly

branched. In subfamily Prayinae specimens have usually 2 (up to 4) smooth NE, while only 1 large NE characterizes the subfamily Nectopyramidinae. BR rounded and unridged.

F

E 5

6

5

6 7

7

A. peltifera

Amphicaryon acaule Chun, 1888

(Haeckel, 1888)

Size: larger NE diam. ~10 mm

Size: larger NE diam. ~4 mm

DPC

DPC HYD RC

RC

HYD

NE: 2 different. Larger NE, possibly the retained larval NE (5), in A. acaule partly encloses the reduced vestigial definitive NE (6), which has a NES without an ostium BR: with 2 LAT HYD-canals Note: species can be identified by the size of the NE and shape of the vestigial RC (7) Distribution: both epipelagic in NADR, NASE, NASW, NATR. A. acaule also NECS, NWCS

Rosacea spp.

H

G

Amphicaryon spp.

RC

Rosacea cymbiformis

R. plicata

(Delle Chiaje, 1830)

Bigelow, 1911

Size: NE ~ 11 mm H

Size: NE up to 30 mm H

8

NE: 2 medium, rounded NE with simple SOM without side branches. Sinuous LAT RC on NES BR: kidney-shaped, with characteristic arrangement of canals Note: species can be identified by the extension of the HYD, that in R. plicata does not reach the base of NE (8) Distribution: R. cymbiformis epipelagic, R. plicata mesopelagic. Both in NADR, NASE, NECS. R. cymbiformis also in CNRY, C. plicata also in NATR

FIGURE 106:  Hippopodius hippopus: A, definitive nectophore, lower view; B, larval nectophore, lateral view. Vogtia glabra: C, definitive

nectophore, upper view. V. serrata: D, definitive nectophore, upper view. Amphicaryon acaule: E, polygastric stage, lateral view. A. peltifera: F, polygastric stage, lateral view. Rosacea cymbiformis: G, definitive nectophore, lateral view. R. plicata: G, definitive nectophore, lateral view. A–D, Kirkpatrick and Pugh (1984); E–H Pugh (1999).

246 ta xonom y Order Siphonophora Suborder Calycophorae  Family Sphaeronectidae Small, fragile rounded/conical larval NE, the only one in the polygastric stage. BR small, rounded. See recent revision of the genus in Pugh (2009) and Grossmann et al. (2012).

A

B 1

C

HYD

SOM

1

SOM

HYD

Sphaeronectes koellikeri Huxley, 1859 Size: ~6 mm (NE diameter), ~2 mm (BR)

S. irregularis (Claus, 1873) Size: ~3 mm W

Sphaeronectes spp. See family characteristics. Note: species can be identified by the shape of SOM, position of the intersection and form of RC (1), NES height and HYD extension Distribution: S. koellikeri epipelagic in GFST, NASE, NECS, NWCS; S. irregularis epi-pelagic in SARC

Order Siphonophora Suborder Cystonectae  Family Physaliidae This family is presently monotypic for Physalia physalis, the Portuguese Man O’War. Distribution and ecology: Physalia floats on the surface of the sea. Its float responds actively to wind, adopting a characteristic ‘sailing posture’ together with erection of the crest.

D

Physalia physalis 2 (Linnaeus, 1758)

T with N

GA

Huge, asymmetric, horizontal purplish-blue PN (2), with an erectile diagonal ‘sail’ at the top. Cormidia attached to one side of the float. T can be more than 10 m long Size: float up to 30 cm in length Distribution: epipelagic in CNRY, NADR, NATR, NECS, NWCS

FIGURE 107:  Sphaeronectes koellikeri: A, definitive nectophore, lateral view; B, bract, lateral view. S. irregularis: C, definitive nectophore,

lateral view. Physalia physalis: D, whole animal. A, B, Kirkpatrick and Pugh (1984); C, Carré (1968), D, Totton (1965).

cnida r i a: coloni a l h y drozoa  247 Order Siphonophora Suborder Physonectae  Family Agalmatidae sensu strictu It includes the genera Agalma, Halistemma, Nanomia, Athorybia and Melophysa, the last two being short-stemmed forms with, in the genus Athorybia, the total suppression of the nectosome.

B

A

Agalma spp.

D

C

Dorsal nectosome (i.e. NE budded off on dorsal side of the stem). NE have DMC and no MFZ. Adult TEN are involucrate tricornuate in Agalma, or unicornuate in the other genera.

E

F

G

Agalma clausi

A. elegans

A. okenii

Bedot, 1888

(Sars, 1846)

Eschscholtz, 1825

Size: NE ~16 mm H

Size: NE ~5-7 mm H. Colony up to 1 m L

H

Size: NE ~4.5 mm H

thrust block

I upper LAT ridges vertical LAT ridge Size: NE ~ 7 mm H

lower LAT ridge

Halistemma rubrum

OS

(Vogt, 1852)

J

NE: V-shaped. Triangular NES, Tshaped (A. elegans) or Y-shaped (A. clausi and A. okenii). LAT-RC distinctly looped. TEN: tricornuate. BR: foliaceous Note: A. clausi and A. okenii without LAT_ridges Distribution: epi-mesopelagic. A. elegans and A. okenii in CNRY, NATR, NWCS. A. elegans also in GFST, NADR, NASE, NASW, NECS. A. clausi in NWCS

Halistemma spp.

NE: large thrust block. Characteristic arrangement of ridges, i.e. ≥1 pair of vertical LAT ridges, 1 pair of upper and lower LAT and 2 pairs of LAT ridges. Typical sinuous arrangement of RC TEN: unicornuate, with very reduced INV BR: foliaceous, of 2 different types Note: H. rubrum characterized by 1 pair of incomplete vertical LAT_ridges, not joining upper and lower LAT_ ridges. MO_PL absent. Size: Colony up to a few meters length Distribution: epi-mesopelagic in ARCT, CNRY, NADR, NASE, NASW, NATR, NECS, NWCS

Nanomia spp. K

1

L

2

N

M

LAT ridge

Nanomia bijuga

N. cara

(Delle Chiaje, 1844)

Agassiz, 1865

Size: NE ~3 mm H. Colony up to 10–30 cm L

Size: NE up to 10 mm H

NE: Upper LAT_ridges incomplete (1). LAT_ridges complete. LAT_RC form loops. TEN: unicornuate, with incomplete INV. BR: thin and leaf-like. Note: N. bijuga has squared NES hollow in the middle, NE with AP wings folded (2). N. cara has Y-shaped NES and NE with only 1 LAT_ridge on each side. Distribution: epi-mesopelagic. Both species found in NECS, NWCS; N. bijuga also in NASE and NASW, while N. cara in ARCT, NADR, NECS, SARC

FIGURE 108:  Agalma clausi: A, definitive nectophore, upper view; B, bract, upper view. A. elegans: C, definitive nectophore, upper view; D

and E, two bracts of different shape, upper views. A. okenii: F, definitive nectophore, upper view; G, bract, upper view. Halistemma rubrum: H, definitive nectophore, lateral view; I, definitive nectophore, upper view; J, bract, upper view. Nanomia bijuga: K, definitive nectophore, upper view; L, young nectophore, lower view; M, bract, upper view. N. cara: N, definitive nectophore, upper view. A, B, Bedot (1888); C, L, N, Totton (1965); D, E, J, K, M, Pugh (1999); F, G, Bigelow (1911); H, I, Pugh and Baxter (2014).

248 ta xonom y Order Siphonophora Suborder Physonectae  Family Apolemiidae One T or groups of T present between NE on nectosome. NE deeply hollowed axially, forming a pair of large AP wings. NES large with S-shaped LAT_RC.

A

B

Apolemia uvaria (Lesueur, 1815) Groups of 5–6 T on nectosome, between each pair of NE. LAT_RC with short branches on the upper loop. BR covered by opaque spots.

Size: NE 15–20 mm H. BR ~6 mm. Colony up to 20–30 m L Distribution: epipelagic in NATR, NECS, SARC

Order Siphonophora Suborder Physonectae  Family Forskaliidae Cylindrical/cone-shape nectosome, with NE arranged spirally. NE flattened in upper-lower axis, often asymmetrical. NES restricted to

C

basal half, with straight RC. BR of variable shapes of four types: stem, bolster and two kinds of knee shaped.

Forskalia spp.

E

D

See family characters Note: left AP wing is large in F. contorta and small in F. edwardsi, without apical incision in both species. In F. contorta NES has marked LAT wings (1), while F. edwardsi has small yellow spots on OS where RC meet ring canal.

1

Forskalia edwardsii

F. contorta

Kölliker, 1853

(Milne Edwards, 1841)

Size: NE up to 7 mm H

Size: NE up to 10 mm H

Size: colony up to 5–10 m long Distribution: F. edwardsi epipelagic in NASE, NATR, NECS. F. contorta epi-mesopelagic in CNRY, NASE, NASW, NATR

Order Siphonophora Suborder Physonectae  Family Physophoridae NE apparently ridgeless, with an extensive NES, which has characteristic looped LAT_RC. Upper and lower canals are sinuous. Siphosome shortened to form a sac on which cormidia are arranged spirally. Each cormidium has one large PAL. Two species have been described: Physophora hydrostatica (without BR) and P. gilmeri (with BR)

F

Physophora spp. See family characters Size: NE up to 20 mm H. Colony up to 10 cm long Distribution: mesopelagic in ARCT, CNRY, NADR, NASE, NASW, NECS, NWCS, SARC

Physophora hydrostatica Forsskål, 1775 FIGURE 108 (CONTINUED):  Apolemia uvaria: A, definitive nectophore, upper view; B, bract, lateral view. Forskalia edwardsi: C, definitive

nectophore, upper view; D, bracts, upper views. F. contorta: E, definitive nectophore, upper view. Physophora hydrostatica: F, definitive nectophore, upper view. A, E, Pugh (1999); B, Totton (1965); C, D, F, Kirkpatrick and Pugh (1984).

cnida r i a: coloni a l h y drozoa  249

References Alvariño, A. (1971). Siphonophores of the Pacific with a review of the world distribution. Bulletin of the Scripps Institute of Oceanography Technical series 16, 14–32. Båmstedt, U., Fosså, J.H., Martinussen, M.B., Fosshagen, A. (1998). Mass occurrence of the physonect siphonophore Apolemia uvaria (Lesueur) in Norwegian waters. Sarsia 83, 79–85. Bedot, M. (1888). Sur l’Agalma clausi n. sp. Recueil Zoologique Suisse Geneve 5, 73–92. Bigelow, H. B. (1911). Reports of the scientific results of the expedition to the eastern tropical Pacific, in charge of Alexander Agassiz, by the U.S. Fish Commission Steamer ‘Albatross’, from October, 1904, to March, 1905, Lieut.Commander L.M. Garret, U.S.N., commanding. XXIII. The Siphonophorae. Memoirs of the Museum of Comparative Zoölogy, Harvard College 38, 173–401, 32 pls. Biggs, D. C. and Harbison, G. R. (1976). The siphonophore Bathyphysa sibogae Lens & van Riemsdijk 1908 in the Sargasso Sea, with notes on its natural history. Bulletin of Marine Science 26, 14–18. Bouillon, J., Gravili, C., Pagès, F., Gili, J., and Boero, F. (2006). An Introduction to Hydrozoa, Publications Scientifiques du Muséum: Paris. Carré, C. (1968). Contribution à l’étude du genre Sphaeronectes Huxley, 1859. Vie Milieu 19, 85–94. Carré, C. (1979). Sur le genre Sulculeolaria Blainville, 1834 (Siphonophora, Calycophorae, Diphyidae). Annales de l’Institut océanographique de Paris 55, 27–48. Carré, C. and Carré, D. (1991). A complete life cycle of the calycophoran siphonophore Muggiaea kochi (Will) in the laboratory, under different temperature conditions: ecological implications. Philosophical Transactions of the Royal Society London B 334, 27–32. Dunn, C. W., Pugh, P. R., and Haddock, S. H. D. (2005a). Molecular phylogenetics of the siphonophora (Cnidaria), with implications for the evolution of functional specialization. Systematic Biology 54, 916–35. Dunn, C. W., Pugh, P. R., and Haddock, S. H. D. (2005b). Marrus claudanielis, a new species of deep-sea physonect siphonophore (Siphonophora, Physonectae). Bulletin of Marine Science 76, 699–714. Dunn, C. W. and Wagner, G. P. (2006). The evolution of colonylevel development in the Siphonophora (Cnidaria: Hydrozoa). Development Genes and Evolution 216, 743–54. Gili, J. -M. (1986). Estudio sistemático y faunístico de los Cnidarios de la costa catalana. Ph.D. Thesis, University of Barcelona: Barcelona. Greve, W. (1994). The 1989 German bight invasion of Muggiaea atlantica. ICES Journal of Marine Science 51, 355–8. Grossmann, M.M., Lindsay, D.J., and Fuentes, V. (2012). Sphaeronectes pughi sp. nov., a new species of sphaeronectid

calycophoran siphonophore from the subantarctic zone. Polar Science 6(2), 196–199. Kirkpatrick, P.A. and Pugh, P.R. (1984). Siphonophores and Velellids. The Linnean Society of London and The Estuarine and Brackish-Water Sciences Association Publishers, Bath, Great Britain. Mackie, G. O., Pugh, P. R., and Purcell, J. E. (1987). Siphonophore Biology. Advances in Marine Biology 24, 97–262. Mapstone, G. M. (2009). Siphonophora (Cnidaria: Hydrozoa) of Canadian Pacific waters. Ottawa, Ontario, Canada: NRC Research Press. 302 p. Mapstone, G. M. (2014). Global Diversity and Review of Siphonophorae (Cnidaria: Hydrozoa). PLoS ONE 9:e87737. doi:10.1371/journal.pone.0087737 Pagès, F. and Gili, J. -M. (1992). Siphonophores (Cnidaria, Hydrozoa) of the Benguala Current (southeastern Atlantic). In: Pagès, F., Gili, J.-M., and Bouillon, J. (eds), Planktonic Cnidarians of the Benguela Current. Scientia Marina, 56(Suppl. 1), 65–112. Pugh, P. R. (1998). A re-description of Frillagalma vityasi Daniel, 1966 (Siphonophorae Agalmatidae). Scientia Marina, 62, 233–45. Pugh, P. R. (1999). In: Boltovskoy (ed), Siphonophora of the Indian Ocean with systematic and biological notes on related species from other oceans. Discovery Report 27, 1–161. Pugh, P. R. (2001). A review of the genus Erenna Bedot, 1994 (Siphonophora, Physonectae). Bulletin of the Natural History Museum, Zoology Series (London) 67, 169–82. Pugh, P. R. (2006). Reclassification of the clausophyid siphonophore Clausophyes ovata into the genus Kephyes gen. nov. Journal of the Marine Biological Association of the United Kingdom 86, 997–1004. Pugh, P. R. (2009). A review of the family Sphaeronectidae (Class Hydrozoa, Order Siphonophora), with the description of three new species. Zootaxa 2147, 1–48. Pugh, P. R. (2016). A synopsis of the Family Cordagalmatidae fam. nov. (Cnidaria, Siphonophora, Physonectae). Zootaxa, 4095,1–64. Pugh, P. R. and Baxter, E. J. (2014). A review of the physonect siphonophore genera Halistemma (Family Agalmatidae) and Stephanomia (Family Stephanomiidae). Zootaxa 3897, 1–111. Pugh, P. R. and Harbison, G. R. (1986). New observations on a rare physonect Siphonophore, Lychnagalma utricularia (Claus, 1879). Journal of the Marine Biological Association of the United Kingdom 66, 695–710. Pugh, P. R. and Haddock, S. H. D. (2010). Three new species of remosiid siphonophore (Siphonophora: Physonectae). Journal of the Marine Biological Association of the United Kingdom 90, 1119–43.

250 ta xonom y Purcell, J. E. (1983). Digestion rates and assimilation efficiencies of siphonophores fed zooplankton prey. Marine Biology 73, 257–61. Purcell, J. E. (1997). Pelagic cnidarians and ctenophores as predators: selective predation, feeding rates, and effects on prey populations. Annales de l’Institut océanographique 73, 125–37. Totton, A. K. (1965). A Synopsis of the Siphonophora. British Museum (Natural History), London, UK.

Totton, A. K. and Fraser, J. H. (1955a). Siphonophora. Sub-order Calycophorae. Family: Prayidae. Conseil Internat. pour l’Exploration de la Mer, Sheet No. 58. Totton, A. K. and Fraser, J. H. (1955b). Siphonophora. Sub-order Calycophorae. Family: Abylidae. Conseil Internat. pour l’Exploration de la Mer, Sheet No. 60. Tregouboff, G. and Rose, M. (1957). Manuel de planctonologie mediterranéenne. Tome I & II., Centre national de la recherche scientifique: Paris.

CTENOPHOR A Pr iscilla Lica ndro a nd Dhuga l J. Lindsay 1 Introduction The ctenophores, or comb jellies, are gelatinous metazoans belonging to a small and entirely marine phylum of about 150 species (Mills and Haddock, 2007), and perhaps 50–100 more that are still undescribed. They are mostly planktonic, with the exception of the benthic order Platyctenida, where only the larvae are planktonic. Ctenophora were initially associated with the Porifera and Cnidaria in the group Coelenterata. Subsequently they were recognized as three distinct phyla of primitive invertebrates. Molecular studies are still inconclusive as to the nearest relatives of ctenophores, though the most recent studies suggest that they evolved from an organism that was also the common ancestor of the Porifera and/or Placozoa (Bridge et al., 1995; Collins, 1998; Podar et al., 2001; Pisani et al., 2015). Due to their fragile nature, ctenophores are easily destroyed when collected with conventional plankton nets. The recent development of in situ observation, in particular of remotely operated vehicles equipped with underwater cameras, has shown that ctenophores are much more abundant and diverse than previously thought.

2  Life Cycle Most ctenophores are simultaneous hermaphrodites, capable of self-fertilization. Only the genus Ocyropsis is known to definitely be dioecious (Harbison and Miller, 1986). Exceptionally, the Platyctenida are protandrous and they can perform asexual reproduction (Harbison, 1985). In ctenophores, gonads are usually located in the wall of the meridional canals, and eggs and sperm are released through separate gonoducts (Harbison, 1985) or the mouth. The cydippid larva, (i.e. the ctenophore primary larva) is with or without tentacles in the Tentaculata and Nuda, respectively. In Cydippida and Beroida the cydippid larva is morphologically very similar to the adult (Fig. 110). In Cestida, Lobata and Platyctenida instead, the larva undergoes a sort of metamorphosis during its development.

FIGURE 109:  Ctenophora Lobata Bolinopsis vitrea. Photo courtesy of

C. Carré.

3 Ecology Ctenophora have traditionally been thought to be carnivorous, feeding on copepods, amphipods, euphausiids, appendicularians, fish eggs, and larvae of various sizes, although some species have also been observed to feed on marine snow in situ (D.J.L., personal observation) and it is probable that they can also uptake at least a subset of the available dissolved organic matter in seawater directly. Some ctenophores (e.g. Beroida) feed also on gelatinous zooplankton including medusae, salps, and other ctenophores (Kremer, 1979; Mianzan and Sabatini,

Marine Plankton. Edited by Claudia Castellani and Martin Edwards, Oxford University Press (2017). © Oxford University Press. DOI: 10.1093/acprof:oso/9780199233267.001.0001

252 ta xonom y eggs

adult

early cydippid larva

late cydippid larva FIGURE 110:  Ctenophore life cycle – Pleurobrachia pileus. Modified from Liley (1958); Greve (1975).

1985; Purcell, 1985 and references therein; Monteleone and Duguay, 1988). Lobate ctenophores capture prey on large mucous-laden, muscular surfaces known as lobes, with the assistance of auricles and/or fine tentacles deployed within the lobes. Others (e.g. Beroids, Haeckeliids, Lampeiids) instead are able to feed on very large prey engulfing them with their very flexible and large mouth. Recent works show that ctenophore species can adapt to a wide range of environmental conditions, surviving and reproducing successfully for weeks without food (Gambill et al., 2015; Granhag and Hosia, 2015; Jasper et  al., 2015). Broad environmental tolerances have allowed ctenophores such as Mnemiopsis leidyi, native to the western Atlantic, to invade northern Europe, the Black and the Baltic Seas, as well as the Mediterranean, once introduced via ballast waters (Shiganova, 1998; Purcell et  al., 2001; Javidpour et al., 2006; de Oliveira, 2007).

4  General Morphology The body of ctenophores (Fig. 111) is composed of gelatinous mesoglea, sandwiched between an external (ectoderm) and an internal (endoderm) epithelium, and covered by eight rows (comb rows) of fused macrocilia (comb plates or ctenes). Mesodermderived tissue (e.g. muscles) is also present. In ctenophores, literally comb bearers (from the Greek cten, comb, and phero, to bear), the ctenes move with a metachronal beating coordinated by an apical sense organ (statocyst), allowing the locomotion of the animal. All ctenophores bear comb plates at some stage in

their life, even though they may lose them at maturity, as in adult Platyctenida that become creeping forms as an adaption to a benthic life. Some ctenophore species can also move by flapping their oral lobes. The gastrovascular system in ctenophores is composed of an axial and a peripheral part. The axial part consists of the mouth, that in the order Lobata is provided with two oral lobes and four slender gelatinous appendages (auricles), a large pharynx (stomodaeum or gastric cavity), the gut (infundibulum), the infundibular and excretory canals and anal pores. The peripheral part, which is connected to the infundibulum, comprises a complex net of canals: eight meridional canals, each one situated under a comb row, perradial, interradial, and adradial canals, and the tentacular and paragastric canals. Depending on the species, any of these canals can be missing, except for the adradial and meridional canals. The connections, morphology and number of the peripheral canals are important taxonomic features. Most ctenophores (with the exception of the order Beroidea) have paired tentacles that may be simple or have side branches (i.e. tentilla). The base of the tentacle (tentacle bulb) is often deep inside the animal, so that the tentacles can be retracted within tentacle sheaths (absent in the order Lobata). As opposed to Cnidaria, ctenophore tentacles are not equipped with stinging cells but usually carry colloblasts, i.e. specialized very sticky adhesive cells, which are used to capture the prey (Fig. 111G). Ctenophores can be rounded (e.g. Cydippida) (Fig. 111A) or ­ribbon-shaped (e.g. Cestida) (Fig. 111E). They are considered bi-radially symmetrical.

5 Systematics Here we adopt the classifications of Ctenophora proposed by Harbison (1996), updated by Mianzan (1999) and more recently by Mills (Wrobel and Mills, 1998; Mills and Haddock, 2007). The main diagnostic characters are those suggested by Mayer (1912), Liley (1958), Greve (1975), Harbison (1985, 1996), O’Sullivan (1986), and Harbison and Madin (1982). Recent work (Simion et al., 2015), using molecular genetic techniques to study relationships between ctenophore species, has shown that the present phylogenetic framework which we use for the classification of known ctenophore species needs to be radically revised. Here we adopt the taxonomic list recently revised by Mills (1998-present). Because of the confused state of ctenophore taxonomy it is highly recommended that all future published reports should include photographs of the various morphotypes under discussion, to allow correct interpretation of the data once their ­taxonomy has been adequately clarified.

ctenophor a 253 A Cydippida type - Hormiphora spp.

B Cydippida type (Bathyctenidae/Pleurobrachiidae)

MO substomodeal MER-C

Oral Pole

STO

subtentacular MER-C

PAR-C

CT T_BU

STO

T_BU

T

T_SH AD-C

INT-C

T-C

C Lobata type - Mnemiopsis leidyi

CT

MER-C

OLO

T-C TEN AD-C T

MO

AUR

INF-C

INF AN-C

CT PAR-C

STA

MO

STO STA

MER-C diverticulum of MER-C

E Cestida type - Cestum veneris INF

MO

F

G

T

CT PAR-C

Aboral Pole STA

papillae

D Beroida type - Beroe spp.

subtentacular MER-C

INF

Key to annotations AD-C AN-C AUR CT INF INF-C

INT-C = Interradial canal = Adradial canal MER-C = Meridional canal = Anal canal MO = Mouth = Auricle OLO = Oral Lobe = Ctenes (comb plates) PAR-C = Paragastric canal = Infundibulum (gut) STA = Statocyst = Infundibular canal

STO T T_BU T-C T_SH TEN

= Stomodaeum (pharynx) = Tentacle = Tentacle bulb = Tentacular canal = Tentacle sheath = Tentilla

FIGURE 111:  General morphology of Ctenophora. A–B, Cydippida. A, Hormiphora spp., lateral view. B, schematic view of the internal canal

structure from the aboral pole. C, Lobata, Mnemiopsis leidyi, lateral view. D, Beroida, Beroe spp., lateral view. E, Cestida, Cestum veneris, lateral view. F, ctenes; G, colloblast. A, Mills and Haddock (2007); B, Lindsay and Miyake (2007); C, de Oliveira et al. (2007); D, F–G, Mianzan (1999); E, Harbison and Madin (1982).

254 ta xonom y Box 1  Classification of Ctenophora Phylum Ctenophora Class Tentaculata Order Cydippida Family Haeckeliidae Family Ctenellidae Family Bathyctenidae Family Aulacoctenidae Family Lampeidae Family Pleurobrachiidae Family Pukiidae Family Euplokamididae Family Cryptocodidae Family Mertensiidae Family Dryodoridae Order Platyctenida Family Ctenoplanidae Family Tjalfiellidae Family Lyroctenidae Family Savangiidae Family Coeloplanidae Order Cambojiida Family Cambojiidae Order Ganeshida Family Ganeshidae Order Cryptolobiferida Family Cryptolobatidae Order Thalassocalycida Family Thalassocalycidae Order Lobata Family Bathocyroidae Family Bolinopsidae Family Leucotheidae Family Ocyropsidae Family Eurhamphaeidae Family Lampoctenidae Family Lobatolampeidae Order Cestida Family Cestidae Class Nuda Order Beroida Family Beroidae Families in bold are included in this chapter.

1

CTENOPHORA Presence of T or lobes

No

Yes

Body like a sac, MO opens wide (1). STO occupies nearly entire interior

BEROIDA

Yes Beroe ovata Yes

Body like an airplane wing

Yes

Body long and flat, with fine T (2) along the entire leading edge that sweep back over the body surface

CESTIDA Cestum veneris 2

No

Body like a shower cap

Yes

3

Body medusiform with 8 short CT (3) on the aboral surface and two small branched T hanging into the open cavity

Yes

THALASSOCALYCIDA

Thalassocalyce inconstans

No

OLO

Body with two rounded OLO at the oral pole No

Yes

Usually with small, inconspicuous T between the OLO

Yes LOBATA

Without OLO, with a pair of T arising on opposite sides of the body Yes

Mnemiopsis leidyi

4 Cydippiid larva of

T_BU midway/closer to external surface of the body than to the STO (4), positioned aborally of the oral end of comb rows. T with TEN not exiting aborally

Yes

LOBATA, CESTIDA or

STO T_BU

Cydippid larva of Bolinopsis vitrea

Not all those characteristics Hormiphora plumosa

CYDIPPIIDA

FIGURE 112:  Key to Ctenophora. Beroe ovata, after Mianzan (1999); Cestum veneris, after Harbison and Madin (1982); Thalassocalyce inconstans, drawn from picture of Michigan Science Art (arthttp://michiganscienceart.com/); Mnemiopsis leidyi, after Mianzan (1999); Bolinopsis vitrea, drawn from picture of Claude Carré; Hormiphora plumosa, after Chun (1880).

ctenophor a 255

THALASSOCALYCIDA

T without TEN, exiting close to the oral pole/mid of the body. MO able to open very wide

Yes

T_BU small and round, far from STO (1). Very short T_SH (2) prevents full retreat of T

DRYODORIDAE

No Dryodora glandiformis Type B internal canal structure

Yes

T_BU very long (3), >1/4 of the body length. T with TEN, exiting close to aboral pole/mid of the body

1 2

3

No

Yes

HAECKELIIDAE

No Type C internal canal structure

Hormiphora spp.

Yes

AULACOCTENIDAE

4

T_SH exits mid of the body (4). T_BU short.

Yes

STO voluminous. MO very mobile and extensible. Body up to several centemeters long, circular in cross-section. CT rows shorter than MER-C

Yes

LAMPEIDAE

TEN

No

No

Lampea pancerina

STO

T_SH exits towards the aboral pole (6). T_BU short/ long

Yes

STO small, extending 1/4 of total body length. T_BU small and angled obliquely (5). CT rows short, with only a few large CT, ending below the midline

MERTENSIIDAE (Genus Charistephane)

5 Charistephane fugiens

Yes 7

Yes

Type A internal canal structure

Yes

T attached middle of T_BU (7).

Hormiphora plumosa

No T attached at the aboral end of T_BU (8). No

PLEUROBRACHIIDAE

6

Yes

TEN lightly coiled. Often with aboral keels of CT rows on ridges. Type C internal canal structure

Yes

MERTENSIIDAE (Genera Mertensia, Callianira)

8 Mertensia ovum

6 T_BU

T_BU parallel to STO. T with TEN strongly coiled, no conspicuous keels.

Yes

Type C internal canal structure

9

Yes TEN

EUPLOKAMIDIDAE

Euplokamis dunlapae

FIGURE 113:  Key to Ctenophora. Cydippida. Dryodora glandiformis, after Seravin (1995); Hormiphora spp., after Mills and Haddock (2007); internal canal structures of Haeckeliidae, Aulacoctenidae, Pleurobrachidae, Mertensiidae, Euplokamididae, after Lindsay and Miyake (2007); Lampea pancerina and Mertensia ovum, after Mianzan (1999); Charistephane fugiens and Hormiphora plumosa, after Chun (1880); Euplokamis dunlapae, after Mills (1987).

256 ta xonom y

CTENOPHORA - CYDIPPIDA (Main families)

ctenophor a 257 Order Cydippida  Family Dryodoridae T_BU small and round, mid-body and far from STO, very close to the outer body wall. T_SH very short prevent full retreat of T, which are fine and unbranched, without colloblasts. Oral pole opening on a large INF-C. Short STO, not extending >1/4 to oral end. Voluminous vestible distal to MO. Distribution and ecology: D. glandiformis is a boreal species, likely a species complex. It feeds on appendicularians. Eggs emerge in strings of 20 to 30

Dryodora glandiformis (Mertens, 1833)

A

T_BU

Size: up to 10 mm CT rows: circa 1/2 body L MER-C: extended up to oral end where they form hooked branches T: short/smooth without TEN Colour: body transparent, T_SH purple. Sometimes T-C, MER-C and oral pole are reddish-brown Note: oral end opens very wide Distribution: SARC

Order Cydippida  Family Euplokamididae T_SH exit at about 1/4 of body L, aboral of the base of STO. T_BU parallel to STO. T with widely spaced TEN, held tightly coiled when relaxed, but stretch out when in contact with prey. Distribution and ecology: E. dunlapae eats copepods of various sizes, wrapping the prey in its tentilla. It might release bioluminescent globules when disturbed

Euplokamis dunlapae Mills, 1987 Size: up to 20 mm CT rows: all circa 2/3–3/4 body L MER-C: extend slightly behind and above the CT rows T: each with typically 10–20 TEN Colour: body transparent. Might have red pigment at the end of CT rows, base of T and TEN Note: body elongate/ovoid. Might have gelatinous extensions below the STA Distribution: mesopelagic in NWCS, SARC and NECS

B

TEN

FIGURE 114:  Dryodora glandiformis: A, adult, lateral view. Euplokamis dunlapae: B, adult, lateral view. A, Seravin (1995); B, Mills (1987).

258 ta xonom y Order Cydippida  Family Haeckeliidae T_BU very long, usually more than half the body length. Opening of T_SH between MO and the end of CT rows. T without TEN. MO widely expandable. Distribution and ecology: Haeckelia eats narcomedusae, storing their nematocysts in its tentacles

Haeckelia beehleri

Haeckelia rubra

(Mayer, 1912)

A

Size: ≤ 10 mm CT rows: 8 nearly equal in length, from aboral end to < 1/2 body L MER-C: 8, equal in length extending nearly the entire body T: Long T_SH exiting near the MO Colour: sligthly opaque. Might have red pigment along canals Distribution: epipelagic in NWCS

C CT rows

Haeckelia bimaculata B

(Kölliker, 1853) Size: ≤ 7 mm CT rows: 8 unequal in length, from aboral end to 3/5 and 1/2 body L MER-C: 4, all 3/4 body L T: Long T_SH exiting near the MO Colour: Transparent, often green. 2 pairs of orange/red spots on T and T_SH. Note: very large STO. Pointed oral end Distribution: epipelagic in western Atlantic

Carré and Carré, 1989 Size: ≤ 3 mm CT rows: 8 nearly equal in length, from aboral end to middle of body L MER-C: 4, all 3/4 of body L T: Long T_SH exiting at 1/4 body L, below the MER-C Colour: transparent. Large orange spots on canals, at the base of STO and T Small red spots along CT rows Note: rounded oral end Distribution: epipelagic in western Atlantic

FIGURE 114 (CONTINUED):  Haeckelia beehleri: A, adult, lateral view. H. bimaculata: B, adult, lateral view. H. rubra: C, adult, lateral view.

A, modified after Mayer (1912); B, modified after Carré and Carré (1989); C, Chun (1880).

Order Cydippida  Family Lampeidae Long, cylindrical body. T_BU short, mid-body and parallel to STO. T_SH exit mid-body. T with numerous, long TEN. Highly eversible STO.

Lampea pancerina (Chun, 1879)

A

Size: up to 70 mm CT rows: of equal length, extend from the aboral end to the opening of T_SH MER-C: same length as CT rows T: with relatively few short TEN Colour: milky white body. CT rows might have pink pigment. Note: very large STO, occupying 4/5–5/6 of body L. T_SH exit horizontal Distribution: epi-mesopelagic in NASW, NATR, NWCS

Distribution and ecology: L. pancerina eats salps, engulfing them with its large mouth. Young stages can flatten on salp body, looking like a parasite

Order Cydippida  Family Mertensiidae Body oval or strongly compressed along pharyngeal axis. T_BU from small to large, parallel or oblique to STO. T_SH exit mid-body or towards the aboral pole. T with TEN. Distribution and ecology: M. ovum can feed on various prey, from small copepods to amphipods and krill. Thanks to its robust tentacles this species is an efficient predator on large prey

Charistephane fugiens

Mertensia ovum

Chun, 1879

B

C

Size: up to 20 mm CT rows: short (each made by ≤7 CT), ending oral to the midline MER-C: extend 3/4 of body L to oral pole T: with short, widely spaced TEN Colour: very transparent Note: T_BU small and oblique. Small STO, occupying 1/4 of body L. T_SH short, exit at the aboral end. Body laterally compressed Distribution: mesopelagic in NWCS

(Fabricius, 1780) Size: up to 55 mm CT rows: 4 substomodeal are longer (4/5 of body L) than 4 subtentacular MER-C: same length as CT rows T: long with short, simple TEN Colour: T, CT rows pink Note: T_BU banana-shaped, at the aboral pole, with TEN attached at the aboral end of T_BU Distribution: epipelagic in ARCT, NWCS, SARC

Order Cydippida  Family Pleurobrachiidae Body spherical, ovoid or cylindrical, slightly compressed along pharyngeal axis. T_SH exit towards aboral pole. T_BU short/long. T attached in the middle of T_BU. In Hormiphora the T_BU lie close to the STO while in Pleurobrachia they are clearly separated. Distribution and ecology: H. plumosa has large batteries of colloblasts on its tentacles. Pleurobrachia species are not bioluminescent. P. bachei is an ambush predator that mainly feeds on small copepods

Hormiphora plumosa (M. Sars, 1859)

D

Hormiphora cucumis (Mertens, 1833) Size: up to 100 mm CT rows: extend nearly the entire body MER-C: as long as CT rows T: long, with TEN all alike Colour: very transparent Note: T_BU long > 1/4 of body L, narrow, close to STO. T_SH angle abruptly outward at 1/3 of body L from aboral end Distribution: epipelagic in NWCS

E

Size: up to 15–20 mm CT rows: all long 2/3 of body L MER-C: as long as CT rows T: long, with 2 types of TEN (filamentous and roughly hand-shaped) and TB with cock's-comb shaped expansions Colour: very transparent. TEN and STO sligthly yellow/red Note: pear-shaped body. AD-C open onto MER-C at level with STO. Oral end of T_BU extends further oralwards than oral end of MER-C Distribution: epipelagic in NASW, NWCS

FIGURE 114 (CONTINUED):  Lampea pancerina: A, adult, lateral view. Charistephane fugiens: B, adult, lateral view. Mertensia ovum: C, adult,

lateral view. Hormiphora cucumis: D, adult, lateral view. H. plumosa: E, adult, lateral view. A and C, Mianzan (1999); D, Wrobel and Mills (1998); B, Chun (1880); E, Mayer (1912).

260 ta xonom y Order Cydippida  Family Pleurobrachiidae

A

Pleurobrachia bachei

Pleurobrachia brunnea

A. Agassiz, 1860

Mayer, 1912

Size: up to 15 mm CT rows: extend nearly the entire L MER-C: as long as CT rows T: with numerous TEN. T_BU short, close to STO. T_SH angle down, open in the lower 1/4 Colour: transparent, yellow/red on T, might have red spots on STO Note: STO 2/3 of body L MER-C: extend well past ends of CT rows T: each with circa 32 TEN, often coil. T ending with a large knob. Colour: STO and TEN yellow/brown, purple pigment spots on T & TEN in proximal half of each Note: MER-C extend more oralwards than T_BU Distribution: epipelagic in NWCS

B

Pleurobrachia rhodopis

Pleurobrachia pileus C

(O. F. Müller, 1776)

Chun, 1879

D

Size: up to 25 mm CT rows: extend >3/4 the body L MER-C: as long as CT rows T: black spots at the base of TEN, which are numerous and when expanded can be 15–20 times the body L Colour: STO and T are milky or yellow/dull orange. Note: AD-C open onto MER-C above INF and below T_SH. STO >1/2 of body L. T_BU far from STO Distribution: epipelagic in NECS, NWCS, SARC

Size: up to 10 mm CT rows: less than 1/2 body L MER-C: as long as CT rows T: with one type of TEN. Colour: transparent, base of T might be pink Note: AD-C open onto MER-C at level of INF, which is 2000 m), or anchialine caves. Monstrilloida: parasites of marine polychaetes and molluscs. Mormonilloida: found in oceanic waters deeper than 400 m; only four known species. Poecilostomatoida: mostly parasitic but several free-living families are common in the plankton. Siphonostomatoida: mostly fish parasites. The most speciose order of copepods to be found in the plank­ ton is the Calanoida. Over 800 calanoid species are found in the North Atlantic alone, with its members forming the bulk of the

taxa described in this guide. A dichoto­mous key to the copepods of the North Atlantic would be lengthy, frustrating, and of limited practical use: indeed reliance on one feature in a ­ ­decision-making process lends itself to error making and inefficiency, and can be summed up by the phrase, keys are ‘compiled by those who do not need them for those who cannot use them’ (Lobanov, 2003). Nevertheless, when confronted with a large group such as the Copepoda, and within it the Calanoida, a  means of navigation is needed to nar­row one’s search in reaching a correct identification. For this reason, a summary table directing the user to the level of superfamily within the Calanoida is included (Table 1). For a detailed review of the Copepoda, including a key to families, please see Boxshall and Halsey (2004) or Bradford-Grieve (1999a). For an interactive multistate key to the Calanoida and Oncaeidae please see http://www.crustacea.net/crustace/calanoida/index.htm and http://www.rb-schnack.de/index.php/contact.html, respectively. Descriptions of species in this guide apply to the mor­phology of adults only, as most species can only reliably be identified in  the adult stage. For some taxa we provide maps of their epipelagic distribution based on Continuous Plankton Recorder survey data. Species abundances were spatially interpolated by applying the inverse squared distance procedure with a search radius of 200 km with a spatial grid resolution of 1° latitude x 1° longitude (–75°W to 30°E and 35°N to 80°N). Interpolation was carried out for each month and each year over the period 1958–2014. For those species where the abundance is low across the basin occurrence is symbolised by gray crosses. For brevity only the most common and abundant taxa are described.

5.1  Key characters for identification of copepods 1. Total body length (measured from head to end of caudal rami, not including caudal setae). 2. Body tagmosis/general shape: gymnoplean (calanoid) or podoplean (cyclopoid/harpacticoid, etc.) (see Fig. 123). 3. Number of prosome and urosome segments; relative length of somites. 4. A1 length and form of male geniculate antennule, if present. 5. Mx2 and Mxp strength and ornamentation (setose or scythe-like spines). 6. Presence/absence of metasome projections. 7. Shape and size of genital somite and seminal receptacles. 8. Size and shape of P5, if present. 9. Length of caudal rami. 10. Pigmentation: colouration in body or limbs. 11. Geographical location: many species have distinct environmental tolerances e.g. warm or cold water; oceanic, shelf or coastal regions.

272 ta xonom y Box 1  Classification of marine planktonic copepods Kingdom Animalia Phylum Arthropoda Subphylum Crustacea Superclass Multicrustacea Subclass Copepoda Superorder Gymnoplea   Order Calanoida (Table 1)   Superfamily Arietelloidea    Family Arietellidae    Family Augaptilidae    Family Discoidae    Family Heterorhabdidae    Family Hyperbionychidae     Family Lucicutiidae    Family Metridinidae    Family Nullosetigeridae   Superfamily Calanoidea    Family Calanidae    Family Megacalanidae    Family Paracalanidae   Superfamily Clausocalanoidea    Family Aetideidae    Family Clausocalanidae     Family Diaixidae     Family Euchaetidae    Family Mesaiokeratidae    Family Phaennidae    Family Pseudocyclopiidae    Family Scolecitrichidae    Family Stephidae    Family Tharybidae   Superfamily Diaptomoidea    Family Acartiidae    Family Candaciidae     Family Centropagidae

   Family Fosshageniidae    Family Parapontellidae    Family Pontellidae    Family Pseudodiaptomidae    Family Sulcanidae     Family Temoridae     Family Tortanidae Superfamily Eucalanoidea    Family Eucalanidae    Family Rhincalanidae Superorder Podoplea   Order Cyclopoida    Family Oithonidae    Family Thaumatopsyllidae   Order Harpacticoida    Family Aegisthidae    Family Clytemnestridae    Family Ectinosomatidae    Family Euterpinidae    Family Miraciidae    Family Peltidiidae    Family Thalestridae    Family Tisbidae   Order Mormonilloida    Family Mormonillidae   Order Monstrilloida    Family Monstrillidae   Order Poecilostomatoida    Family Corycaeidae    Family Lubbockiidae     Family Oncaeidae    Family Paralubbockiidae    Family Sapphirinidae    Family Urocopiidae NB: Only taxa in bold are covered in this chapter

Orders of Copepoda found in North Atlantic plankton samples Most commonly found Least commonly found

Calanoida Gymnoplean, i.e. Prosome-urosome articulation located between 5th pedigerous seg and GS.

Cyclopoida Podoplean, i.e. Prosome-urosome articulation located between 4th and 5th pedigerous segs.

Poecilostomatoida Podoplean, i.e. Prosome-urosome articulation located between 4th and 5th pedigerous segs.

Harpacticoida Podoplean, i.e. Prosome-urosome articulation located between 4th and 5th pedigerous segs.

Siphonostomatoida Podoplean, i.e. Prosome-urosome articulation located between 4th and 5th pedigerous segs.

Monstrilloida Podoplean, i.e. Prosome-urosome articulation located between 4th and 5th pedigerous segs.

A1 longer than head with typically a maximum of 25 segs.

A1 in marine planktonic forms, usually longer than head. ♀ A1 up to 26 segs, ♂ A1 up to 17 segs and geniculate on both sides.

A1 shorter than head; ♂ non-geniculate, ♀ at most 7 segs.

Distinction between prosome and urosome typically unclear.

A1 shorter than head, up to 21 segs in ♀.

A1 maximum of 5 segs and anteriorly projected.

Claw-like Mxp, sexually dimorphic.

Urosome as long as body.

Body shape highly variable.

A1 shorter than head; at most 14 segs in ♂ and 9 in ♀.

Urosome shorter than prosome.

Body shape extremely variable; from highly modified to resembling a cyclopoid.

A2, Md, Mx1, Mx2 and Mxp absent in adults. Adults are free-swimming, juveniles are parasitic. Only one family in the order, Monstrillidae.

Figure 123:  Generalized diagram of common copepod orders and summary of simple identification features used to separate copepod orders. Arrow denotes location

of GS.

crustacea: copepoda  273

May have a wide range of body forms: Typical planktonic form as seen in figure above.

All members of this order are parasitic and possess an oral cone.

Superfamily

Arietelloidea

Calanoidea

Clausocalanoidea

Diaptomoidea

Eucalanoidea

♂ A1

Geniculate on left

Non-geniculate (geniculate on right in some Megacalanidae)

Non-geniculate

Geniculate on right

Non-geniculate

P1 endopod

Usually 3 segments, 2 segments in Lucicutiidae

Usually 3 segments, 2 segments in Paracalanidae

1 segment

2–3 segments

2 segments

♂ P5

Typically claw-like, or with right exopod recurved

Resembling other swimming limbs, or uniramous, slender, with left leg longer than the right in Paracalanidae

Highly modified and strongly assymetrical, long and slender; may have one leg longer than the other

Highly modified and strongly assymetrical, claw-like or recurved

Highly modified and strongly asymmetrical, usually long and slender, right leg may be absent in Eucalanidae

♀ P5

Biramous, somewhat resembling other swimming limbs; uniramous in Metridinidae, Discoidae, Hyperbionychidae, and Nullosetigeridae

Biramous, somewhat Uniramous or absent resembling other swimming limbs, uniramous in Paracalanidae

Biramous; uniramous in Acartiidae, Candaciidae, Fosshageniidae, Temoridae and Tortanidae

Uniramous in Rhincalanidae; absent in Eucalanidae

Genera

Heterorhabdus (p. 275), Heterostylites (p. 278), Lucicutia (p. 279), Metridia (p. 280), and Pleuromamma (p. 283)

Calanoides (p. 288), Calanus (p. 289), Calocalanus (p. 296), Mecynocera (p. 298), Mesocalanus (p. 292), Nannocalanus (p. 293), Neocalanus (p. 294), Paracalanus (p. 299), and Undinula (p. 295)

Acartia (p. 324), Anomalocera (p. 336), Calanopia (p. 337), Candacia (p. 328), Centropages (p. 331), Eurytemora (p. 346), Isias (p. 334), Labidocera (p. 338), Paracartia (p. 327), Parapontella (p. 335), Pontella (p. 340), Pontellina (p. 342), Pontellopsis (p. 343), Pseudodiaptomus (p. 345), Temora (p. 346), and Tortanus (p. 349)

Eucalanus (p. 350), Pareucalanus (p. 351), Rhincalanus (p. 355), and Subeucalanus (p. 352)

Aetideopsis (p. 301), Aetideus, (p. 302), Amallothrix (p. 320), Clausocalanus (p. 308), Ctenocalanus (p. 310), Diaixis (p. 313), Euchaeta (p. 314), Euchirella (p. 303), Gaetanus (p. 306), Microcalanus (p. 311), Paraeuchaeta (p. 317), Pseudoamallothrix (p. 320), Pseudocalanus (p. 312), Scaphocalanus (p. 321), Scolecithricella (p. 322), Scolecithrix (p. 323), and Undeuchaeta (p. 307)

274 ta xonom y

Table 1:   Summary table of characteristics used to separate the order Calanoida into the common planktonic superfamilies with a list of genera described in this chapter.

crustacea: copepoda  275 Order Calanoida Superfamily Arietelloidea  Family Heterorhabdidae Urosome: ♀ 4 segs; ♂ 5 segs. Posterior dorsal edge of segs with small spines (8). ♀ GS with genital operculum rounded in lateral view (9) (c.f. flat in Heterostylites). Caudal rami with oblique outer edge (10); L-ramus fused with AS (11) and longer than R-ramus; 2nd inner seta thickest and longer than body (12)

Genus: Heterorhabdus Giesbrecht, 1898 Total length: ♀ 1.1–4.6 mm; ♂ 1.6–4.2 mm Cephalosome: head rounded with a small papilla on forehead (1). A1 first seg long and stiff (3 fused segs) (2). A1 reaching from middle of urosome to end of caudal rami; proximal segs slightly serrated and hairy in appearance. ♂ L-A1 geniculate. Rostrum with 2 filaments. Mx2 with more than 2 long scythelike spines (c.f. two in Heterostylites); 5th lobe with 2 large spines (3) (c.f. one spine in Heterostylites and Paraheterorhabdus); Enp with short setae (c.f. long in Heterostylites) (4)

Notes: nine species reported for the study area. A comprehensive study of Heterorhabdidae can be found in Park (2000) where he splits the genus into four groups: ‘spinifrons’, ‘papilliger’, ‘fistulosus’, and ‘abyssalis’. Prior to this work there have been numerous misidentifications within the Heterorhabdidae, calling into question the distribution of certain species

Metasome: body oval, 4 segs; 4th and 5th segs fused. End of metasome rounded

Similar taxa: Paraheterorhabdus spp., Mesorhabdus spp., and Lucicutia spp.

Legs: noticeably broad and transparent (5). ♀ P5 symmetrical, biramous, 3 segs; resembling other swimming limbs, 2nd Exp seg with a well-developed inner-spine (6). ♂ P5 biramous, asymmetrical; basis with hairy inner lobe, well developed on right; Exp and Enp with 3 segs; right Exp 2nd seg with inner projections (7), 3rd seg elongate and claw-like

A

1

spp.,

Distribution and ecology: epi-abyssopelagic. Carnivorous. CNRY, NASE, NWCS, GFST, NASW, NADR, NECS, SARC, ARCT

E

1

G R

F

L

C ♀ 

B

2

3

 ♂

7

5

D

6

H

4

12

Heterostylites

9

I

8

12

11

12

10

Figure 124:  Heterorhabdus papilliger. A, ♀ dorsal view; C, ♂ dorsal view; F, ♀ P5; G, ♂ P5; H, ♀ urosome lateral view; I, ♀ urosome dorsal

view. Heterorhabdus spinifrons. B, ♀ lateral view; D, Mx2; E, P3. A, C, F, Rose (1933); B, D, E, G-I, Park (2000).

Total length

Heterorhabdus papilliger

Heterorhabdus spinifer

Heterorhabdus spinifrons

Heterorhabdus oikoumenikis

Heterorhabdus norvegicus

(Claus, 1863) Papilliger group

Park, 1970 Papilliger group

(Claus,1863) Spinifrons group

Park, 2000 Abyssalis group

(Boeck, 1872) Abyssalis group

♀ 1.1–1.3 mm, ♂ 1.6–1.9 mm

♀ 2.1–4.0 mm, ♂ 2.0–3.8 mm

♀ 1.6–2.7 mm, ♂ 1.6–2.7 mm

Spiniform process

LV

VV

♀ 2.6–3.5 mm, ♂ 2.4–3.3 mm

VV

Sigmoid on left

VV

Operculum width=length

♀ lateral and GS

Straight

LV

Operculum width>length

Concentric ridges

VV

LV

Hump

(Park, 2000)

(Park, 2000) 5th lobe spines lacking serrations and spinules

5th lobe unserrated spine as long as serrated spine

Hump

Long A1

Long A1 (modified from Sars 1924, Park 2000) 5th lobe unserrated spine as long as serrated spine

Keyhole shaped

Operculum asymmetrical LV

LV Hump

♀ 2.7–4.6 mm, ♂ 2.6–4.2 mm

Straight (Park, 2000) 5th lobe unserrated spine shorter than serrated spine

(Park, 2000) 5th lobe unserrated spine much shorter than serrated spine

(Park, 2000)

(Park, 2000)

swollen

LV

♀ Mx2 (Park, 2000)

(Park, 2000)

(Park 2000, as H. guineansis) one seta on 1st endopod seg

seta on 1st endopod seg absent

one seta on 1st endopod seg

one seta on 1st endopod seg

one seta on 1st endopod seg

♀ P5 (Park 2000, as H. guineansis) Short L lobe R

(Park, 2000) flat

Elongate lobe

(Park, 2000)

R

L

Lobed

Finger-like lobe

(Park, 2000) R

Very elongate lobe

L

(Park, 2000) R

L

Very elongate lobe R

Conical process < spine

♀ P5 Conical process > spine

Very long & thin

L Short spine

Small lobe (Park, 2000)

Long spine (Park, 2000)

Long lobe

(Park, 2000)

(Park, 1970)

(Park, 2000)

Similar spp.

H. spinifer, H. spinifrons

H. papilliger, H. spinifrons

H. papilliger, H. spinifer H. oikoumenikis

H. norvegicus, H. spinifrons

H. oikoumenikis

Distribution

Epi-bathypelagic. CNRY, NASE, NASW, NWCS, GFST, NADR, NECS, SARC.

Epi-bathypelagic. NASW, NASE.

Epi-bathypelagic. CNRY, NASE, NWCS, NASW, NADR, GFST, SARC,

Epi-bathypelagic. NADR, NASE, NASW, GFST, NWCS.

Epi-bathypelagic. ARCT, SARC, NWCS, NASW, NECS, NASE, CNRY, NADR, GFST.

276 ta xonom y

Table 2:   Heterorhabdus species.

DISTRIBUTION: Heterorhabdus papilliger

DISTRIBUTION: Heterorhabdus oikoumenikis

DISTRIBUTION: Heterorhabdus spinifer

DISTRIBUTION: Heterorhabdus norvegicus

278 ta xonom y Order Calanoida Superfamily Arietelloidea  Family Heterorhabdidae Genus: Heterostylites Sars, 1920

R-Exp 2nd seg with enlarged projection (8); 3rd seg elongate and claw-like

Total length: ♀ 2.3–5.8 mm; ♂ 2.6–5.5 mm Cephalosome: head rounded with a small papilla on forehead (1). A1 first seg long and stiff (3 fused segs). A1 reaching beyond end of caudal rami (2); proximal segs slightly serrated and hairy in appearance. ♂ L-A1 geniculate. Rostrum with 2 filaments. Mx2 with 2 scythelike spines, one on 4th and 5th lobe (c.f. more than 2 in Heterorhabdus); 5th lobe elongated (3); Enp with long setae (4) (cf. short in Heterorhabdus) Metasome: body oval, 4 segs; 4th and 5th segs fused. End of metasome rounded Legs: somewhat broad and transparent. ♀ P5 symmetrical, biramous, with 3 segs; resembling other swimming limbs, Exp seg 2 with a long thin inner-spine (5), and a serrated distal margin (6) (unlike in Heterorhabdus). ♂ P5 biramous, asymmetrical; basis with hairy inner lobe, elongate on right (7); Exp and Enp with 3 segs.

Heterostylites longicornis A

B

1

Urosome: ♂ 5 segs. ♀ 4 segs. ♀ GS with genital operculum elongated and flat in lateral view (9) (c.f. short and rounded in Heterorhabdus). Caudal rami with oblique outer edge (10); L-ramus fused with AS (11) and longer than R-ramus; 2nd inner seta thickest and as long as body (12) Notes: two species of Heterostylites reported for the study area. A  comprehensive study of Heterorhabdidae can be found in Park  (2000). Prior to this work there have been numerous misidentifications within the Heterorhabdidae, calling into question the distribution of certain species Similar taxa: Heterorhabdus spp., Paraheterorhabdus spp. and Lucicutia spp.

C

H

(F. Dahl, 1894) ♀ 4.0–5.8 mm ♂ 3.8–5.5 mm Meso-bathypelagic CNRY. NASE, NWCS, NASW, GFST, NADR, SARC, NECS

E

F

L

R

7

6

G 5

♀

2

I

K

J

4

Heterostylites major

D

♂

♀ 2.3–4.3 mm ♂ 2.6–4.3 mm Epi-bathypelagic CNRY. NASE, NWCS, NASW, GFST, NADR, SARC, NECS

spp.,

Distribution and ecology: epi-bathypelagic. CNRY, NASE, NASW, GFST, NWCS, NADR, SARC, NECS

♀

(Giesbrecht, 1889)

Mesorhabdus

R

11

L

L

9

M 3

12

10

8

Figure 125:  Heterostylites longicornis. A, ♀ head dorsal view; B, ♀ lateral view; C, ♂ dorsal view; D, ♀ P5; E, ♂ P5; F, ♀ GS lateral view; G, ♀ GS ventral view. Heterostylites major. H, ♀ lateral view; I, Mx2; J, ♂ urosome dorsal view; K, ♂ P5; L, ♀ GS lateral view; M, ♀ GS ventral view. A, B, D–M, Park (2000), C, Rose (1933).

crustacea: copepoda  279 Order Calanoida Superfamily Arietelloidea  Family Lucicutiidae Urosome: ♀ 4 segs. ♂ 5 segs. ♀ GS with a pronounced ventral knob  (7). Caudal rami long and slender (8), sometimes with an elongated terminal seta on each ramus (9)

Genus: Lucicutia Giesbrecht & Schmeil, 1898 Total length: ♀ 1.1–5.4 mm; ♂ 1.1–5.2 mm Cephalosome: A1 extending to or beyond caudal rami; very ‘hairy’ in appearance at base, with numerous obvious aesthetascs (1) particularly in ♂; ♂ L-A1 geniculate. Rostrum with paired filaments. Front of head sometimes with papilla (2) or lateral projections (3) in some deeper living species

Notes: 12 species found in the study area; however, many other species are found at greater depths. A comprehensive key to the genus can be found in Boxshall and Halsey (2004). Reported differences in size ranges and morphology of Lucicutia flavicornis may indicate the presence of an undescribed sibling species (Bradford-Grieve 1999b). According to Herring (1988) all Lucicutia spp. are bioluminescent even in copepodid stages

Metasome: body oval and slender, 4 segs, 4th–5th segs fused. End of metasome rounded Legs: ♀ P5 biramous, resembling other swimming limbs; Exp 3 segs, 2nd seg with styliform inner seta (4); Enp with 2–3 segs. ♂ P5 asymmetrical, biramous: L-leg basis often toothed on inner margin (5); L-leg Enp and Exp with 3 segs: R-leg Exp with 2 segs, last Exp seg recurved (6); R-leg Enp 2–3 segs

Lucicutia flavicornis (Claus, 1863) ♀ 1.3–2.5 mm ♂ 1.1–1.9 mm Epi-bathypelagic CNRY, NASE, GFST, NASW, NADR, NWCS, SARC, NECS

A

Distribution and ecology: mostly meso-abyssopelagic, with a few epipelagic species. Circumglobal, oceanic: CNRY, NASE, GFST, NASW, NADR, NWCS, ARCT, SARC, NECS

1

1

B

Similar taxa: Temora spp., Heterostylites spp. and Heterorhabdus spp.

2

C

♀

H

G

♂

♀

♂

Lucicutia gemina Farran, 1926

D

♀ 1.4–1.9 mm ♂ 1.3–1.7mm Epi-mesopelagic CNRY, NASE, GFST, NASW, NADR, NWCS, NECS

8 7 9

3

Lucicutia clausi (Giesbrecht, 1889)

L

R

5

E

4

F 6

♂

♀

♀ 1.6–2.2 mm ♂ 1.6–1.9 mm Epi-mesopelagic CNRY, NASE, GFST, NASW,

I

J

NADR, NWCS

K

Figure 126:  Lucicutia flavicornis. A, ♀ dorsal view; B, ♀ A1; C, ♂ dorsal view; D, ♀ urosome lateral view; E, ♀ P5; F, ♂ P5. Lucicutia gemina.

G, ♀ dorsal view; H, ♂ dorsal view. Lucicutia clausi. I, ♀ dorsal view; J, ♂ dorsal view; K, distribution map of Lucicutia spp. A, B, D, G, H, Bradford-Grieve (1999b); C, Rose (1933); E, F, Grice (1962); I, J, Park (1970).

280 ta xonom y Order Calanoida Superfamily Arietelloidea  Family Metridinidae Genus: Metridia Boeck, 1864 Total length: ♀ 1.5–4.5 mm; ♂ 1.3–3.3 mm Cephalosome: A1 reaching from end of metasome to AS; proximal segs strongly toothed (1). ♂ A1 geniculate on the left side, rarely on the right. Rostrum with paired filaments Metasome: body elongate, 4 segs, 4th and 5th segs fused. End of metasome rounded or with a small point (2) Legs: P2 first Enp seg is hollowed on inner margin and has 2 or more basally-directed hooks (characteristic of family) (3). ♀ P5 with 2-4 segs, and 2-3 long plumose setae on terminal seg (4). ♂ P5 asymmetrical and uniramous, 4-5 segs: R- or L-leg with long inner projection on 3rd seg (5) (absent in M. venusta); the other leg with inner spinous process (6)

A

1

C

Urosome: long and narrow. ♀ 3 segs, ♂ 5 segs. ♀ GS characteristically long and slender (7). AS widening distally, overhanging caudal rami (like in Pleuromamma) (8) Notes: five species reported for the study area. M. lucens and M. longa are bioluminescent. Several other species, up to 10 mm in length, are found in the N. Atlantic but are bathypelagic Similar taxa: Pleuromamma spp. Distribution and ecology: omnivorous. Undergo strong diel vertical migration. Epi-abyssopelagic. CNRY, NASE, NASW, NWCS, GFST, NADR, ARCT, SARC, NECS

♂

♂

♀

F

E

D

3

B ♀

2 2

8

H G

7 6

5

4

Figure 127:  Metridia lucens. A, ♀ A1; B, ♀ dorsal view; C, ♀ lateral view; D, ♂ dorsal view; E, ♂ lateral view; F, P2; G, ♀ P5; H, ♂ P5.

A, B, D, F, G, Bradford-Grieve, 1999b; C, Sars, 1903; E, H, Vidal, 1971.

Table 3:   Metridia species.

Total length

Metridia brevicauda

Metridia curticauda

Metridia longa

Metridia lucens

Metridia venusta

Giesbrecht, 1889

Giesbrecht, 1889

(Lubbock, 1854)

Boeck, 1854

Giesbrecht, 1889

♀ 1.5–2.3 mm, ♂ 1.3–1.7 mm

♀ 2.3–3.8 mm, ♂ 1.8–3.3 mm

♂

♀

♀

♀ 1.6–4.5 mm, ♂ 1.6–4.0 mm ♂

♀

♂

♀ 1.9–4.0 mm, ♂ 1.5–3.0 mm ♂

♀

♀ 2.7–3.2 mm, ♂ 2.4–2.8 mm ♂ urosome DV

♀

♀ and ♂ Blunt tip

Small knob

♀ urosome short cf. other Metridia spp.

♀ A1 reaching to AS (Bradford-Grieve 1999b; Vervoort, 1951)

(Bradford-Grieve, 1999b)

A1 reaching to end of metasome

♀ A1 reaching to middle of GS

2 segs

3 segs

3 segs

♀ P5

L-caudal ramus longer than right (Sars, 1924; Bradford-Grieve 1999b)

(Sars, 1901; Bradford-Grieve, 1999b)

(Sars, 1903; Vidal, 1971) 4 segs

4 segs

A1 reaching to AS

acute point

♂

A1 reaching to AS

long and smooth setae

L-leg longer than R-leg

2 setae (Vervoort, 1951)

(Scott, 1909) 5 segs. Terminal segs broad (1), with short spine on R-leg (2); L-leg with long inner spine (3)

♂ P5

(Sars, 1903)

5 segs. Terminal segs broad (1), with short spine on R-leg (2); L-leg with long inner spine (3) R

L

R

L

1

3 (Wilson, 1932 )

2

3

(Vervoort, 1951)

(Bradford-Grieve, 1999b)

(Sars, 1924)

5 segs. Terminal seg oblong (1). L-Leg with spiniform process (2), R-leg with long inner spine (3)

L R

4 segs. R-leg terminal seg long with 4 folds (1); with short inner hairy process (2). L-leg short, R L last 2 segs hairy (3). 3

L

R 2

2

2

seta

3

3

1 (Sars, 1903)

1

(Vidal, 1971)

Similar spp.

M. curticauda

M. brevicauda, M. lucens, M. longa

M. lucens, M. curticauda

M. longa, M. curticauda

Distribution

Meso-abyssopelagic. CNRY, NASE, NASW, NWCS, GFST, NADR, NECS, SARC, ARCT

Meso-abyssopelagic. CNRY, NASE, NASW, NECS,

Epi-bathypelagic. CNRY, NASE, NASW, NWCS, NADR, ARCT, SARC, NECS

Epi-mesopelagic. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS, SARC, ARCT

1

(Sars, 1924)

Meso-bathypelagic. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS, SARC

crustacea: copepoda  281

1

2

5 segs. Terminal segs oblong with distal part narrow (1), short spine on Lleg (2), R-leg with long inner spine (3)

DISTRIBUTION: Metridia longa

DISTRIBUTION: Metridia lucens

crustacea: copepoda  283 Order Calanoida Superfamily Arietelloidea  Family Metridinidae Genus: Pleuromamma Giesbrecht, 1898 Total length: ♀ 1.5–5.9 mm; ♂ 1.4–6.4 mm Cephalosome: ♀ A1 reaching from end of metasome to AS, proximal segs strongly toothed (1); segs 1 and 2 may bear strong spines (2). ♂ A1 geniculate on the left side, rarely on the right. Head with short acute apical process (3). Rostrum very large with two hairy filaments (4). May have red-pink pigmentation Metasome: 4 segs, 4th and 5th seg fused. 1st metasome seg with one characteristic lateral black pigmented spot (5) in ♀ and ♂ Legs: P2 Enp seg 1 (on one or both legs) hollowed on inner margin, with basally-directed hooks (characteristic of family) (6). ♀ P5 of two forms: 4 segs, distal seg with 3 long setae (7); or 2–3 segs, distal seg with 3 spines (8). ♂ P5 asymmetrical and uniramous with

A

3

2

1

♀

3

5  segs: R-leg last seg broad, strongly curved and prehensile (9), preceding seg with curved spine (10); L-leg with inner projection on 3rd seg, shape of projection is species-specific (11) Urosome: ♀ 3 segs, ♂ 5 segs. May be strongly asymmetrical and hairy in ♂ (12). ♀ GS elongate and ventrally swollen (13); shape and pigmentation vary between species (14). AS widening distally, overhanging caudal rami (characteristic of family) (15) Notes: seven species reported for study area. Bioluminescent, black pigmented spot is thought to be a luminescent organ Similar taxa: Metridia spp. Distribution and ecology: epi-bathypelagic; oceanic, typically abundant in warm waters, undergo strong vertical migration. CNRY, NASE, NASW, NWCS, GFST, NADR, SARC, ARCT, NECS

D

B

F

E

4

♂

C 6 7

5

8

G

H

I L

R

15

10

12

11

14 13

9

Figure 128:  Pleuromamma abdominalis. A, ♀ dorsal view; B, ♂ dorsal view; C, rostrum lateral view; D, P2; E, ♀ P5; G, ♀ urosome ventral

view; H, ♀ GS lateral view; I, ♂ P5. Pleuromamma borealis. F, ♀ P5. A, B, E, I, Bradford-Grieve (1999b); C, Giesbrecht (1893 [‘1892’]); D, F-H Steuer (1932).

Total length

Pleuromamma borealis

Pleuromamma gracilis

Pleuromamma piseki

Pleuromamma abdominalis Pleuromamma robusta

Pleuromamma xiphias

F. Dahl, 1893

Claus, 1863

Farran, 1929

(Lubbock, 1856)

(Giesbrecht, 1889)

♀ 1.7–2.5 mm, ♂ 1.4–2.1 mm

♀ 1.5–2.6 mm, ♂ 1.5–2.3 mm

A1 without hooks

A1 without hooks

♂

♀

♀

(F. Dahl, 1893)

♀ 1.7–2.4 mm, ♂ 1.6–1.9 mm ♀ 2.4–4.5 mm, ♂ 2.4–4.3 mm ♀ A1 with 2 strong recurved hooks

A1 without hooks

♀

♂

♂

♂

♀

♀ and ♂ dorsal

♀ 2.3–4.7 mm, ♂ 2.1–4.0 mm ♀ 3.3–5.9 mm, ♂ 3.9–6.4 mm A1 without hooks

♂

♀

♀ A1 with strong recurved hook elongate projection

♀

♀

♂

hairy hairy urosome asymmetrical (Chen and Zhang, 1965; Bradford-Grieve, 1999b)

Prosome pigmentation

(Bradford-Grieve, 1999b)

None key-hole shape, yellow

♀ urosome ventral

L

R

(Rose, 1933) knobbed process (1) L

1

♂ P5

1

(Chen and Zhang, 1965)

(Chen and Zhang, 1965)

Similar spp.

P. gracilis

Distribution

Epi-mesopelagic. Epi-mesopelagic. CNRY, NASE, NASW, NWCS, GFST, CNRY, NASE, NASW, NWCS, GFST, NADR, SARC, NECS NADR, SARC, NECS

P. piseki

L

R

1

4 segs

3 long spines

(Bradford-Grieve, 1999b) R

(Steuer, 1932)

4 segs

3 long spines

3 short stout spines

(Chen and Zhang, 1965) knobbed process (1)

(Sars, 1924) Red horseshoe shape in cephalosome

(Steuer, 1932)

4 segs

2 segs

3 short stout spines

(Bradford-Grieve, 1999b) blunt process (1)

(Steuer, 1932)

(Rose, 1933)

2 segs

3 slender spines

(Bradford-Grieve, 1999b) Red horseshoe shape in cephalosome

round shape, large dark spot

(Chen and Zhang, 1965)

3 segs

(Bradford-Grieve, 1999b) Diffuse red-pink in metasome

None

round shape, small dark spot

(Rose, 1933)

♀ P5

(Bradford-Grieve, 1999b)

None

urosome asymmetrical

spiniform process (1)

3 long spines

(Sars, 1903) R

L

(Steuer, 1932) L

R 1

toothed process (1)

1

1

tongue-like process (1) (Bradford-Grieve, 1999b)

(Bradford-Grieve, 1999b)

(Sars, 1924)

(Bradford-Grieve, 1999b)

P. gracilis

P. xiphias, P. quadrungulata

P. abdominalis

P. abdominalis

Epi-mesopelagic. CNRY, NASE, NASW, NWCS, GFST, NADR, NECS

Epi-bathypelagic. NASE, NASW, NWCS, GFST, NADR, SARC, NECS

Meso-bathypelagic. CNRY, NASE, NASW, NWCS, GFST, NADR, ARCT, SARC, NECS

Epi-bathypelagic. CNRY, NASE, NASW, NWCS, GFST, NADR, SARC, NECS

284 ta xonom y

Table 4:   Pleuromamma species.

DISTRIBUTION: Pleuromamma borealis

DISTRIBUTION: Pleuromamma piseki

DISTRIBUTION: Pleuromamma gracilis

DISTRIBUTION: Pleuromamma abdominalis

DISTRIBUTION: Pleuromamma robusta

DISTRIBUTION: Pleuromamma xiphias

crustacea: copepoda  287 Order Calanoida Superfamily Calanoidea  Family Calanidae Genus: Calanoides Brady, 1883

asymmetrical, R-Exp without inner setae, R-Enp with 4 setae, L-Enp vestigial and without setae (6)

Calanoides carinatus (Krøyer, 1848)

Urosome: ♀ 4 segs. ♂ 5 segs. ♀ spermathecal sac long and thin in lateral view (unlike Calanus) (7)

Total length: ♀ 1.7–3.2 mm, ♂ 2.0–2.8 mm Cephalosome: ♀ A1 not geniculate with segs 1–2 and 3–5 fused (unlike in Calanus) (1). Anterior head sharply keeled in ♀ (2), rounded in ♂ (3). Rostrum in the form of paired filaments. Mxp with fewer setae than other calanids

Notes: one species found in the study area. Pointed head present from stage CIV (Conway, 2006). For a summary of features used to distinguish members of the Calanidae please see Table 6 Similar taxa: Calanus spp., Neocalanus gracilis, Mesocalanus tenuicornis

Metasome: 5 segs. End of metasome somewhat broadly tapering to a point in ♀ (4)

Distribution and ecology: epi-bathypelagic associated with cold upwelling water. CNRY, NASE, NADR, NECS. Developmental stages are restricted to deeper waters according to Vervoort (1963)

Legs: P5 unmodified similar to P1–P4. P5 coxa lacking teeth along inner border (5) (unlike Calanus). ♀ P5 Enp with 6 setae. ♂ P5

A

B

2

♀

C

D

3

1

♀

♂

♂

4

H

R

5

R

L

5

G 7

E

F

6

Figure 129:  Calanoides carinatus. A, ♀ lateral view; B, ♀ dorsal view; C, ♂ lateral view; D, ♂ dorsal view; E, ♀ P5; F, ♂ P5; G, ♀ GS lateral view; H, distribution map of C. carinatus. A, G, Brodsky (1972) C, redrawn from Giesbrecht (1893) [‘1892’]; B, Zheng et al. (1982); D, Corral Estrada (1970); E, F, Dakin and Colefax, (1940).

288 ta xonom y Order Calanoida Superfamily Calanoidea  Family Calanidae distal corner of the Enp (both legs in ♀, R-leg in ♂) can also be used in species id (6)

Genus: Calanus Leach, 1816 Total length: ♀ 1.9–10.0 mm, ♂ 2.4–7.0 mm Cephalosome: ♂ A1 segs 1–2 fused and bulbous (1); not geniculate. Rostrum in the form of paired filaments (2)

Urosome: ♀ 4 segs. ♂ 5 segs. ♀ seminal receptacles separate (7) (unlike Nannocalanus)

Metasome: 5 segs, end of metasome rounded (n.b. small sharp point in Calanus hyperboreus)

Notes: four species found in the study area. Species easily misidentified due to their morphological similarity and overlap in size. This is particularly true for Calanus finmarchicus and Calanus glacialis (see review by Frost (1974)). For a summary of features used to distinguish members of the Calanidae please see Table 6

Legs: P5 relatively unmodified in both sexes, similar to P2-P4, except for toothed inner border on coxa (3). Size of teeth and shape of inner border of coxa used to id ♀ species. ♀ P5 with 7–8 setae on Enp. P5 slightly asymmetrical in ♂, L-Exp longer than R-Exp (4) (same length in C. hyperboreus). Position of the distal margin of the 3rd Enp seg relative to the second Exp seg of the L-P5 is used to id ♂ species (5). ♂ P5 R-leg: without inner setae on Exp (unlike Nannocalanus); with 8 setae on R-Enp. The length of the

A

2

1

B

C

♀

♀

D

Similar taxa: Nannocalanus minor, Neocalanus gracilis, Calanoides carinatus, Mesocalanus tenuicornis Distribution and ecology: epi-bathypelagic. Common and abundant copepod genus. Known to diapause (except Calanus helgolandicus). ARCT, SARC, NECS, NADR, NWCS, GFST, NASW, NASE, CNRY

E

♂

F

♀ R

♂

L

♂ L

R

3

3

L-Enp reaching to ⅓ length of 2nd Exp seg

G

7

H

I

5 R

J

R

4

6 ii

i

i

i reaches ii

ii

i does not reach ii

Figure 130:  Calanus helgolandicus. A, ♀ lateral view; B, ♂ lateral view; E, ♀ P5; F, ♂ P5; I, ♀ P5 coxa, basis and 1st seg of Enp and Exp, anterior view. Calanus finmarchicus. C, ♀ dorsal view; D, ♂ dorsal view; G, ♀ urosome ventral view; H, ♀ urosome lateral view; J, ♀ P5 coxa, basis and 1st seg of Enp and Exp, anterior view. A–D Rose (1933) after Sars (1903); E, F Fleminger and Hulsemann (1977); G, H Williams (1972); I, J Frost (1974).

Table 5:   Calanus species.

Total length

Calanus finmarchicus

Calanus helgolandicus

Calanus glacialis

Calanus hyperboreus

(Gunnerus, 1770)

(Claus, 1863)

Jaschnov, 1955

Krøyer, 1838

♀ 2.4–4.2 mm, ♂ 2.6–3.9 mm

♀ 1.9–3.5 mm, ♂ 2.4–3.2 mm

♀ 3.3–5.6 mm, ♂ 3.1–5.6 mm

Head rounded

Head rounded

Head slightly pointed

Head rounded

♀ 5.3–10 mm, ♂ 5.6–7.0 mm

♀ lateral End metasome rounded

End metasome rounded

(Vidal, 1971)

R

♀ and ♂

(Rose, 1933)

♂

♀ L

R

Coxa straight small teeth

L

End metasome rounded

♀ R

L

Coxa concave large teeth

(Vidal, 1971)

(Vidal, 1971)

♀

♂ R

End metasome with small point

L

R

L

♀

♂

Coxa concave large teeth

R

L

R

L

♂

Coxa straight or concave R

L

Coxa concave

P5

Elongate

(Fleminger and Hulsemann, 1977)

L-Enp reaches ⅓ Oval L-Exp 2nd seg (Fleminger and Hulsemann, 1977)

Elongate (Fleminger and Hulsemann, 1977)

(Conover, 1965)

Enp reaches ⅓ Exp 2nd seg

Similar spp.

C. glacialis, C. helgolandicus

C. finmarchicus, C. glacialis, Neocalanus gracilis

C. finmarchicus, C. helgolandicus, C.hyperboreus

C. glacialis

Distribution

Epi-bathypelagic. Boreal. CNRY (rare), NASE, NWCS, NASW, GFST, NADR, NECS, SARC, ARCT

Epi-mesopelagic. Temperate. CNRY, NASE, NWCS, NASW, GFST, NADR, NECS, SARC

Epi-bathypelagic. Arctic, subarctic. NWCS, NECS, SARC, ARCT

Epi-bathypelagic. Arctic, subarctic. CNRY (rare), NASE, NWCS, NASW (bathy), NECS, SARC, ARCT

crustacea: copepoda  289

L-Enp reaches ⅔ L-Exp 2nd seg

L-and R-Exp same length

L-Enp reaches ½–⅓ L-Exp 2nd seg

290 ta xonom y

DISTRIBUTION: Calanus finmarchicus

DISTRIBUTION: Calanus glacialis

DISTRIBUTION: Calanus helgolandicus

DISTRIBUTION: Calanus hyperboreus

crustacea: copepoda  291 Order Calanoida Superfamily Calanoidea  Family Calanidae Genus: Mesocalanus Bradford and Jillett, 1974 Mesocalanus tenuicornis (Dana, 1849) Total length: ♀ 1.5–2.4 mm, ♂ 1.5–2.2 mm Cephalosome: ♂ A1 not geniculate, with segs 1–2, 3–5, 7–8 and 9–10 fused (1) (unlike other members of the Calanidae), and approx. 1½ times longer than body (2). ♀ A1 approx. twice as long as total body length, distal segments very elongate. Mxp elongated and often outstretched (3). Head rounded Metasome: long slim, somewhat delicate body. 5 segs. End of metasome rounded

Urosome: ♀ 4 segs. ♂ 5 segs. ♀ GS with distinct ventral swelling in lateral view (5). ♀ urosome with short 2nd and 3rd segs (6). Caudal rami much longer than AS (7) and relatively longer and slimmer than those of Calanus Notes: M. tenuicornis is the only Mesocalanus species found in the Atlantic. For a summary of features used to distinguish members of the Calanidae please see Table 6 Similar taxa: Calanus spp., Neocalanus gracilis, Calanoides carinatus Distribution and ecology: epi-bathypelagic; cosmopolitan in warm waters. NWCS, GFST, CNRY, NASE, NADR, NECS

Legs: P5 unmodified, coxa with smooth inner border (4). ♀ P5 symmetrical. ♂ P5 slightly asymmetrical with 7 setae on Enp (unlike ♂ Neocalanus, Calanus and Calanoides)

D A

1

B

L

C R

4

L

♂

♀

3 E 6 7

5

2

Figure 131:  Mesocalanus tenuicornis. A, ♀ lateral view; B, ♂ dorsal view; C, ♂ P5; D, ♀ P5; E, distribution map of M. tenuicornis. A–D Bradford-Grieve (1994).

292 ta xonom y Order Calanoida Superfamily Calanoidea  Family Calanidae Urosome: ♀ 4 segs. ♂ 5 segs. R- and L- seminal receptacles of ♀ GS fused (5) (unlike Calanus). 1st urosome seg slightly asymmetrical in ♂ (6)

Genus: Nannocalanus Sars, 1925 Nannocalanus minor (Claus, 1863) Total length: ♀1.5–2.4 mm, ♂ 1.2–2.0 mm Cephalosome: ♂ A1 not geniculate, with segs 1–2 and 3–5 fused (1) (unlike in Calanus and Neocalanus). Head rounded Metasome: 4 segs, cephalosome fused with 1st metasome seg (5 segs in Calanus). Orange colouration present, persisting in formalin preserved specimens. ♀ last metasome seg strongly curved and partly overlapping GS (2) Legs: P5 unmodified, inner margin of coxa bearing small teeth (3); L-Exp of ♂ P5 with very long external spines on segs 1 and 2 (4)

A

B

C

Notes: N. minor is the only species in the genus Nannocalanus present in the North Atlantic. However, there appear to be 2 genetically distinct subspecies, N. minor f. major and N. minor f. minor (Bucklin et  al. 1996). For a summary of features used to distinguish members of the Calanidae please see Table 6 Similar taxa: Calanus spp. Distribution and ecology: epi-mesopelagic. Common in all subtropical and tropical oceans. Found throughout the North Atlantic, preferring warmer waters. Omnivorous filter feeder. CNRY, NASE, NWCS, NASW, GFST, NADR, NECS, SARC

D

E 1

♀

♀

♂

♂

R

3

2

F

6

R L

5

G

4

Figure 132:  Nannocalanus minor. A, ♀ dorsal view; B, ♀ lateral view; C, ♂ dorsal view; D, ♂ lateral view; E, ♀ P5; F, ♂ P5; G, distribution

map of N. minor. A, C, Owre and Foyo (1967); B, D, E, F, Bradford-Grieve (1994).

crustacea: copepoda  293 Order Calanoida Superfamily Calanoidea  Family Calanidae Urosome: ♀ 4 segs. ♂ 5 segs. ♀ GS produced in lateral view: with angular slope in Neocalanus gracilis (6) and truncate in N. robustior (7)

Genus: Neocalanus Sars, 1925 Total length: ♀1.8–4.7 mm; ♂ 1.6–3.6 mm Cephalosome: ♂ A1 not geniculate, approx. reaching to end of metasome; segs 1–2 and 24–25 fused. ♀ A1 long, 1 ½ times as long as total body length (1). Head rounded Metasome: ♀ 4 segs, cephalosome fused with first metasome seg (cf. unlike Calanus spp). ♂ 5 segs. End of metasome rounded Legs: P1 with hooked spine on basis (2). P2 distal portion of 1st Exp seg having recurved spiny process (3). P5 coxa without teeth (4). ♀ P5 unmodified, with 8 setae on Enp. ♂ P5 slightly modified; R-Enp with 8 setae; L-Enp rudimentary and without setae in Neocalanus robustior (5)

Notes: two species found in study area. Neocalanus tonsus has been reported off the coast of Woods Hole, however this may be a misidentification. For a summary of features used to distinguish members of the Calanidae please see Table 6 Similar taxa: Calanus spp., Mesocalanus tenuicornis, juvenile Undinula vulgaris. Distribution and ecology: epi-bathypelagic. CNRY, NASE, NWCS, GFST, NASW, NADR, NECS, SARC

Neocalanus gracilis A

(Giesbrecht, 1888) ♀ 1.8–4.4 mm ♂ 1.6–3.4 mm Epi-bathypelagic CNRY, NASE, NADR, NWCS, GFST, NASW, NECS, SARC

B ♀

♂

D

C

F 4

6 R

L

G

♀

2

Neocalanus robustior

3

(Dana, 1852) ♀ 3.2–4.7 mm ♂ 2.8–3.6 mm Epi-mesopelagic CNRY, NASE, NWCS, GFST, NASW, NADR, NECS

H

E

1

K

R

I

L

J

♀

7 5 Figure 133:  Neocalanus gracilis. A, ♀ lateral view; B, ♂ dorsal view; C, ♀ P1; D, ♀ P2; E, ♀ P5; F, ♂ P5; G, ♀ urosome lateral view; Neocalanus robustior. H, ♂ P5; I, ♀ urosome lateral view; J, distribution map of N. gracilis; K, distribution map of N. robustior. A–F, Bradford-Grieve (1994); H, Zheng et al. (1982); G, I, Owre and Foyo (1967).

294 ta xonom y Order Calanoida Superfamily Calanoidea  Family Calanidae Genus: Undinula A. Scott, 1909

Urosome: ♀ 4 segs. ♂ 5 segs

Undinula vulgaris (Dana, 1849)

Notes: this genus comprises only one species. For a summary of features used to distinguish members of the Calanidae please see Table 6

Total length: ♀ 1.8–3.3 mm, ♂ 2.0–3.2 mm Cephalosome: ♂ A1 segs 1–2 fused (1) and not geniculate. Head rounded Metasome: 4 segs, cephalosome fused with 1st metasome seg (cf. 5 segs in Calanus). End of metasome elongated into a strong lateral hook-like process in ♀ only (2) Legs: P2 Exp seg 2 with notch on outer border (3). P5 coxa lacking teeth along inner border. ♀ P5 unmodified with 7 setae on Enp (4). ♂ L-P5 large and highly modified; Exp segs 1 and 2 with very long external spines (5), seg 3 complex. ♂ R-P5 reduced

A

B

Similar taxa: stage CV lacks the metasome process present in the adult ♀ and therefore can be confused with other members of the Calanidae family; however, the CV will possess the P2 Exp notch Distribution and ecology: epi-mesopelagic with neritic tendencies. Subtropical to tropical distribution CNRY, NASE, NWCS, GFST, NASW

1

E

D ♂

♀

C

L

L

3

4

5

2

F

Figure 134:  Undinula vulgaris. A, ♀ lateral view; B, ♂ lateral view; C, ♀ P2; D, ♀ P5; E, ♂ P5; F, distribution map of U. vulgaris. A–D Bradford-Grieve (1994); E, Dakin & Colefax (1940).

Table 6:   Summary of key characters for the identification of the Calanidae species found in the North Atlantic. A1, antennule; C, concave; Exp, exopod; Enp, endopod; GS, genital segment; High. mod, entire P5 highly modified; L-, left; Met, metasome; R-, right; S, straight; segs, segments; TL, total body length; Uros, urosome. Table modified from Bradford (1988).

Total length (mm)

Prosome segs

P5 coxa teeth

Inner shape of P5 coxa

P2 Exp

P5 R P5 Enp setae Exp 3rd seg setae

Species

♀

♂

♀ and ♂

♀ and ♂

♀

♂

♀ and ♂

♂

♀

R-♂ L-♂

Calanus hyperboreus

5.3–10.0

5.6–7.0

5

Yes

S or C

S or C

–

0

7

8

8

Calanus glacialis

3.3–5.6

31.–5.6

5

Yes

C

C

–

0

8

8

Calanus finmarchicus

2.4–4.2

2.6–4.0

5

Yes

S

C

–

0

7

Calanus 1.9–3.5 helgolandicus

2.4–3.2

5

Yes

C

C

–

0

Nannocalanus 1.5–2.4 minor

1.2–2.0

4

Yes

S

S

–

Undinula vulgaris

1.8–3.3

2.0–3.2

4

No

S

Neocalanus gracilis

1.8–4.4

1.6–3.4

♀ 4, ♂ 5

No

Neocalanus robustior

3.2–4.7

2.8–3.6

♀ 4, ♂ 5

Mesocalanus tenuicornis

1.5–2.4

1.5–2.2

Calanoides carinatus

1.7–3.2

2.0–2.8

A1 segs fused

A1 length

Head shape End of lateral view ­metasome lateral view

♀ and ♂

♀ and ♂

♀ and ♂

1–2

Just past Uros

Rounded

Pointed

8

1–2

Just past Uros

Rounded

Rounded

8

8

1–2

Just past Uros

Rounded

Rounded

7

8

8

1–2

Just past Uros

Slightly pointed

Rounded

2

7

7

4

1–2, 3–5

~ End Uros Rounded

♀ Rounded, overlapping GS ♂ Rounded

High. mod

Seg 2 notch 0

7

3

0

1–2

~ End Uros Rounded

♀ Pointed ♂ Rounded

S

S

Seg 1 recurved spine

4

8

8

8

1–2, 24–25

♀ 1½ × TL Rounded ♂ ~end Met

Rounded

No

S

S

Seg 1 recurved spine

4

8

8

5

1–2, 24–25

♀ 1½ × TL Rounded ♂ ~end Met

Rounded

5

No

S

S

–

0

7

7

7

1–2, 3–5, ♀ 2 × TL 7–8, 9–10 ♂ 1½ × TL

5

No

S

S

–

0

6

4

0

1–2, 3–5

Rounded

Rounded

Rounded ♀ ~ end Met ♀ Keeled ♂ past Uros ♂ Rounded

296 ta xonom y Order Calanoida Superfamily Calanoidea  Family Paracalanidae Genus: Calocalanus Giesbrecht, 1888 Total length: ♀ 0.4–1.4 mm; ♂ 0.5–1.2 mm Cephalosome: ♀ A1 typically longer than body, up to twice as long as TL; may have several long setae and plumose setae (1) along its length; may possess short spinules on mid segs; last seg 2-6 times longer than penultimate seg (2). ♂ A1 not geniculate, reaching to end of urosome, proximal segs fused and swollen (3). Head rounded, rostrum with two long filaments. ♂ may have small dorsal cephalic hump Metasome: body may be elongate or stocky. 3 or 4 segs (5 segs in  Mecynocera clausi). Cephalosome typically fused with first metasome seg. End of metasome rounded Legs: P1 without inner seta on basis (4) (present in M. clausi). Posterior surfaces of P2-P4 with spinules (5) (without spinules in M. clausi). ♀ P5 uniramous, symmetrical, 3–4 segs: last seg with

a number of setae, spines or spinules (6). ♂ P5 asymmetrical, one leg longer than the other, uniramous; typically R-leg 3–4 segs, L-leg 5 segs; both legs terminating in a spine (7) Urosome: very short in ♀, 2–4 segs (2nd seg may be short and obscured by GS), GS bulbous, often with two clear spherical seminal receptacles in dorsal view (8). ♂ 5 segs, with well developed AS (longer than preceding seg) (9). Caudal rami articulated, may be divergent, with 4 feathered terminal setae, 2 of which typically thicker than the others (10); usually damaged during sampling Notes: 20 species reported for the study area Similar taxa: Mecynocera clausi Distribution and ecology: epi-bathypelagic. Temperate-tropical regions, absent from Polar seas. CNRY, NASE, NASW, NADR, NWCS, GFST, NECS

1

A

E

3

C

L

♂

2

D

♂

B

♀

R

8 9

7 6

10

5

G

F

4 Figure 135:  Calocalanus pavo. A, ♀ dorsal view; B, ♀ P5; C, ♂ dorsal view; D, ♂ urosome dorsal view; E, ♂ P5. Calocalanus contractus. F, ♀ P1; G, ♀ P3. A, C Rose (1933); B, D–G Corral Estrada (1970).

Table 7:   Calocalanus species.

Calocalanus pavo

Calocalanus contractus

(Dana, 1849)

Total length A1 length

Calocalanus styliremis

Farran, 1926

Giesbrecht, 1889

Calocalanus plumulosus (Claus, 1863)

Calocalanus tenuis Farran, 1929

♀ 0.8–1.4 mm, ♂ 0.6–1.2 mm

♀ 0.6–0.8 mm, ♂ 0.5–0.6 mm

♀ 0.5–1.0 mm, ♂ 0.5–0.6 mm

♀ 0.9–1.3 mm, ♂ 0.7–0.9 mm

♀ 0.9–1.3 mm, ♂ unknown

♀ A1 1.3–1.4 x longer than body

♀ A1 1.5–1.7 x longer than body

♀ A1 1.5–1.8 x longer than body

♀ A1 slightly longer than body

♀ A1 as long as body

♀ and ♂ Dorsal

♀

♂

♂

♂ ♀

♀

♀

GS wider than long

3 segs, mid seg may be obscured (Corral Estrada, 1972; Bradford-Grieve 1994)

3 segs, mid seg may be obscured (Corral Estrada, 1972)

GS longer than wide 3 segs without spinules

GS longer than wide

GS wider than long

GS wider than long

2 segs

(Corral Estrada, 1972)

♂ ♀

3 segs with spinules (Corral Estrada, 1972; asymmetrical Bernard, 1958) L- caudal ramus with one long plumose seta

♀ P5

symmetrical (Corral Estrada, 1972)

last seg very long short spine

long spine

1 seta

4 setae

2 setae (Corral Estrada, 1972)

♂ P5

R-leg with 3 segs

R-leg with 3 segs

L

(Corral Estrada, 1972)

(Corral Estrada, 1972)

(Corral Estrada, 1972)

2 setae

(Corral Estrada, 1972)

L unknown

R

L

L

2 setae R-leg with 4 segs

R

R

R

(Corral Estrada, 1972)

(Corral Estrada, 1972)

(Bernard, 1958)

Similar spp.

Mecynocera clausi

C. styliremis

C. contractus

C. tenuis

C. plumulosus

Distribution

Epi-bathypelagic. CNRY, NASE, NASW, NADR, NWCS, GFST, NECS.

Epi-mesopelagic. CNRY, NASE, NASW, NADR, NWCS, GFST, NECS.

Epipelagic. CNRY, NASE, NASW, NADR, NWCS, GFST, NECS.

Epipelagic. CNRY, NASE, NASW, NWCS, GFST.

Epipelagic. CNRY, NASE, NASW, NADR, NWCS, GFST, NECS.

crustacea: copepoda  297

R-leg with 4 segs

(Corral Estrada, 1972)

298 ta xonom y Order Calanoida Superfamily Calanoidea  Family Paracalanidae ♂ P5 asymmetrical and uniramous; both legs with 5 segs, L-leg slightly shorter than R-leg, seg 5 on both legs with a terminal spine and a spinule (7)

Genus: Mecynocera Thompson, 1888 Mecynocera clausi Thompson, 1888 Total length: ♀ 0.9–1.3 mm; ♂ 0.8–1.1 mm Cephalosome: ♀ A1 long, more than twice as long as TL, with several long setae along its length and with short spinules on mid segs (1). ♂ A1 not geniculate, reaching to end of urosome, proximal segs fused and swollen (2). Head rounded, rostrum with two long filaments. ♂ with small dorsal cephalic hump (3) Metasome: slender body, 5 segs (3-4 segs in Calocalanus). End of metasome rounded Legs: P1 with inner seta on basis (4) and distal spines on Exp seg 1 (5) (absent in Calocalanus). Posterior surfaces of P2-P4 without spinules (with spinules in Calocalanus). ♀ P5 uniramous, symmetrical, 5 segs: seg 4 with an inner seta (6), seg 5 with 5 setae.

Urosome: ♀ 3 segs. ♂ 5 segs. ♀ GS short, very rounded and swollen with 2 large seminal receptacles (8). Caudal rami articulated, may be divergent Notes: one species reported for the study area. Mecynocera is a monotypic genus Similar taxa: Calocalanus spp. Distribution and ecology: epipelagic. Cosmopolitan, associated with warm waters, absent from Polar seas; CNRY, NASE, NASW, NADR, NWCS, GFST, NECS

B

D

C 8

A

A

♀

1

E

F

2 3

5

4

♂

♂

L

R

G

6

H

I

J

7

Figure 136:  Mecynocera clausi. A, ♀ dorsal view; B, ♂ rostrum; C, ♀ urosome dorsal view; D, ♀ urosome lateral view; E, ♀ P1; F, ♀ P3; G, ♀ P5; H, ♂ dorsal view; I, ♂ lateral view; J, ♂ P5. A, Rose (1933); B–D, H, I Andronov (1970); E–G, J, Bradford-Grieve (1994).

crustacea: copepoda  299 Order Calanoida Superfamily Calanoidea  Family Paracalanidae ♀ P5 asymmetrical, uniramous: R-leg short with 2-3 segs (9); L-leg long with 5 segs; both legs terminating in two spines (10)

Genus: Paracalanus Boeck, 1864 Total length: ♀ 0.5–1.4 mm; ♂ 0.5–1.4 mm Cephalosome: ♀ A1 reaching from GS to just beyond caudal rami. ♂ A1 not geniculate, reaching to end of urosome, proximal segs fused and swollen (1). Head rounded, rostrum with two long filaments. ♂ may have small dorsal cephalic hump (2) Metasome: body oval, 3 or 4 segs. Cephalosome fused with first metasome seg. End of metasome rounded Legs: P1 with inner seta on basis (3). Exp seg 3 of P2-P4 with toothed outer border (4) (not toothed in Calocalanus and Mecynocera); two distal outer edge spines (5) and a long smooth distal spine (6), typical of Paracalanids (distal spine toothed in other similar families). Posterior surfaces of P2-P4 with spinules (7). ♀ P5 short, uniramous, symmetrical, 2-3 segs; last seg with 2 terminal spines of unequal length (8)

A

Urosome: ♀ 4 segs. ♂ 5 segs, with well developed AS (short in Clausocalanus and Pseudocalanus) (11). ♀ seminal receptacle thin and slit-like in lateral view (12) Notes: six species reported for the study area. Due to inconsistencies in the literature and morphological similarities, it may be difficult to distinguish species Similar taxa: Acrocalanus spp. (♀ P5 absent; not described here), Delibus nudus (not described here), Parvocalanus spp. (brackish; not described here), Clausocalanus spp. and Ctenocalanus vanus Distribution and ecology: epi-bathypelagic. Generally coastal, sometimes brackish, but also found in open ocean. CNRY, NASE, NASW, NADR, NWCS, GFST, NECS, SARC

C

B

D

2

1

♀

3

♂

F L

7

9

R

11

G

E

4 5 6

8 10

12

Figure 137:  Paracalanus parvus. A, ♀ lateral view; B, ♂ lateral view; D, ♀ P4; E, ♀ P5; F, ♂ P5. Paracalanus quasimodo. C, ♀ P1; G, ♀

pedigerous seg 5 and GS lateral view. A, B, D–F Rose (1933); C, G Bowman (1971).

Total length

Paracalanus parvus

Paracalanus aculeatus

Paracalanus nanus

Paracalanus denudatus

Paracalanus quasimodo

(Claus, 1863)

Giesbrecht, 1888

Sars, 1907

Sewell, 1929

Bowman, 1971

♀ 0.6–1.3 mm, ♂ 0.5–1.4 mm

♀ 0.8–1.4 mm, ♂ 0.7–1.4 mm

♀ 0.5–0.7 mm, ♂ 0.5–0.6 mm

♀ 0.6–1.0 mm, ♂ 0.8 mm

♀ 0.8–1.0 mm, ♂ 0.8 mm

body slender in L V

♂

♀ ♂

♀ and ♂ lateral

♀

♀

♂

♀

♂

♀

♂

dorsal

small lateral clusters of hairs on ♀ and ♂ GS ♀ A1 reaching to AS (Sars, 1903)

♀ A1 extends beyond caudal rami by 1–2 segs (Vervoort, 1963)

(Tanaka, 1960)

(Sars, 1903)

♀ A1 reaching AS (Bowman, 1971)

(Vervoort, 1963)

(Corral Estrada, 1970) no spines on Exp

spines on Exp seg 2

♀ A1 extends beyond caudal rami by 1–2 segs (Vervoort, 1963; Corral Estrada, 1970)

1st seg swollen and longer than 2nd seg

outer spine reaches almost half length of terminal seg (Vervoort, 1963)

♀ P5

no spines on Exp

♀ A1 reaching to GS

(Bowman, 1971) no spines on Exp

no spines on Exp

♀ P4 spines on Enp seg 2

(Sars, 1903)

spines on Enp segs 2 and 3 spines on Enp seg 3 spines on Enp segs 2 and 3 spines on Enp segs 2 and 3 (Vervoort, 1963) (Corral Estrada, 1970) (Corral Estrada, 1970) (Bowman, 1971) R-leg reaches beyond L-leg 2nd seg

R

♂ P5

Similar spp. Distribution

R

L (Vervoort, 1963)

(Sars, 1903)

P. quasimodo

P. denudatus

Epi-bathypelagic. CNRY, NASE, NASW, NADR, NWCS, GFST, NECS, SARC, ARCT.

Epipelagic. CNRY, NASE, NWCS.

L

L

L

R L

R

R (Corral Estrada, 1970)

Epi-mesopelagic. CNRY, NASE, NASW, NECS.

(Corral Estrada, 1970)

(Bowman, 1971)

P. aculeatus, P. quasimodo

P. parvus, P. denudatus, P. indicus

Epipelagic. CNRY, NASE.

Epi-mesopelagic. NWCS.

300 ta xonom y

Table 8:   Paracalanus species.

crustacea: copepoda  301 Order Calanoida Superfamily Clausocalanoidea  Family Aetideidae Genus: Aetideopsis Sars 1903 Total length: ♀ 2.2–4.6 mm; ♂ 2.4–4.2 mm Cephalosome: A1 reaching part way down urosome in ♀ and to end of metasome in ♂; not geniculate in ♂. Rostrum bifurcate: ♀ rostral rami may be parallel, divergent, widely (1) or narrowly (2) spaced; rostrum less developed in ♂. Mx2 typical of aetideids with a claw-like seta on 4th and 5th lobes, see Undeuchaeta Metasome: body robust, 4 segs. Cephalosome fused with 1st seg. 4th–5th segs usually separate. End of metasome expanded into spine-like projections (3), shorter in ♂ (4). Body surface sometimes with chitinous thickenings Legs: P1 Exp with an external spine on seg 1 (5), (absent in Aetideus). ♀ P4 coxa without small spinules near the inner seta (6). Outer coxa

A

B

C

of legs usually with small spinules (7). ♀ P5 absent. ♂ P5 biramous: exps long and slender; R-leg Exp 2 segs, last seg scythe-like (8); L-leg Exp 3 segs: Enps vestigial Urosome: ♀ 4 segs. ♂ 5 segs with very short AS (9). ♀ GS symmetrical. Notes: five species reported for the study area. The rostrum, head shape and total length are useful aids in species identification. Please refer to Markhaseva (1996) for a comprehensive key to species within the genus Similar taxa: Aetideus spp. Distribution and ecology: epi-abyssopelagic. Genus found throughout region: CNRY, NASE, NASW, NADR, NWCS, SARC, ARCT, NECS

D

E

F

♀

♀

G

R

L

7

♂

6

5

4 9

8

3

Aetideopsis armata (Boeck, 1872) ♀ 2.2–4.6 mm H ♂ 2.4–4.2 mm Epi-bathypelagic CNRY, NASE, NASW, NADR, NWCS, SARC, ARCT, NECS

♀

I ♀

L

M ♀

♂

K ♂

2

Aetideopsis rostrata Sars, 1903 ♀ 3.0–4.6 mm ♂ 3.1–4.2 mm Meso-abyssopelagic CNRY, NASE, NADR, NWCS, SARC, ARCT, NECS

J

♀

O

N 1

♂

♂

Figure 138:  Aetideopsis armata. A, ♀ dorsal view; B, ♀ lateral view; C, ♂ lateral view; D, ♀ P1; E, ♂ P5; F, P3; H, ♀ head lateral view; I, ♀ rostrum; J, ♂ head lateral view; K, ♂ rostrum. Aetideopsis carinata. G, ♀ P4. Aetideopsis rostrata. L, ♀ head lateral view; M, ♀ rostrum; N, ♂ head lateral view; O, ♂ rostrum. A Park (1975a); B, C, E Rose (1933); D, F–O Markhaseva (1996).

302 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Aetideidae Genus: Aetideus Brady, 1883 Total length: ♀ 1.3–2.3 mm; ♂ 1.1–2.1 mm Cephalosome: ♀ head may have crest (1). A1 reaching from end of metasome to end of urosome; not geniculate in ♂. ♀ rostrum bifurcate with strongly sclerotised, widely spaced rami (2), which may have thickenings between the rami (3), rostrum absent in ♂ (4); Mx2 typical of aetideids with a claw-like seta on 4th and 5th lobes, see Undeuchaeta Metasome: body robust, 3 segs. Cephalosome fused with 1st seg, 4th–5th segs fused. End of metasome expanded into spine-like (5) or wing-like projections (6), shorter in ♂ (7), rounded in Aetideus arcuatus (8). Body surface sometimes with chitinous thickenings

Urosome: ♀ 4 segs. ♂ 5 segs with very short AS (11). ♀ GS symmetrical, seminal receptacles with visible dorsal and ventral lobes linked by a curved duct (12). Caudal rami longer than wide: ♀ 2–3 times (13) and ♂ 1.6–3.7 times (shorter in Aetideopsis) Notes: five species reported for the study area. The rostrum, width of spermathecal duct and length of metasome spines are useful aids in species identification. Please refer to Markhaseva (1996) for a comprehensive key to species within the genus Similar taxa: Aetideopsis sp., Bradyidius spp. Distribution and ecology: epi-bathypelagic. Genus with a widespread distribution: CNRY, NASE, GFST, NASW, NADR, NWCS, ARCT, SARC, NECS

Legs: P1 Exp without an external spine on seg 1 (9) (present in Aetideopsis). P4 coxa may have some small spinules near the inner seta. ♀ P5 absent. ♂ P5 uniramous: L-leg long, 5 segs (10); R-leg absent

Aetideus armatus (Boeck, 1872) ♀ 1.3–2.3 mm ♂ 1.3–2.1 mm Epi-bathypelagic ARCT, NASE, GFST, NASW, NADR, NWCS, SARC, NECS

D

A B

♀

♀

♂

4

C

♂

♀

I

7

Aetideus giesbrechti

Aetideus acutus

(Cleve, 1904)

♀ 1.5–1.8 mm ♂ 1.2–2.0 mm Epi-mesoplagic CNRY, NASE

Farran, 1929

♀ 1.5–2.2 mm ♂ 1.1–1.7 mm Epi-mesoplagic CNRY, NASE, GFST, NASW, NADR 1

♀

L

1

♀

O

11

13 L

P

M

J

2

3

3

8 5

9

E

12 duct constricted

F

G

10

H

K

6

N

Q

duct not constricted

Figure 139:  Aetideus armatus. A, ♀ dorsal view; B, ♀ lateral view; C, ♂ head lateral view; D, ♂ dorsal view; G, ♂ P5; I, ♀ head lateral view; J, ♀ rostrum; K, ♀ urosome lateral view. Aetideus giesbrechti. E, ♂ P1; L, ♀ head lateral view; M, ♀ rostrum; N, ♀ urosome lateral view. Aetideus acutus. F, ♂ P4; O, ♀ head lateral view; P, ♀ rostrum; Q, ♀ urosome lateral view. Aetideus arcuatus. H, ♀ urosome lateral view. A, C, D, G Rose (1933); B, E, F, H–Q Markhaseva (1996).

crustacea: copepoda  303 Order Calanoida Superfamily Clausocalanoidea  Family Aetideidae Genus: Euchirella Giesbrecht, 1888 Total length: ♀ 2.0–8.7 mm; ♂ 2.5–7.4 mm Cephalosome: head with (1) or without (2) crest. A1 reaching from end of metasome to just beyond caudal rami; ♂ A1 not geniculate. Rostrum single and spiniform (3), rudimentary in Euchirella curticauda. Mx2 typical of aetideids with a claw-like seta on 4th and 5th lobes, see Undeuchaeta Metasome: body robust, 3-4 segs. Cephalosome may be fused with 1st seg, 4th–5th segs fused. End of metasome usually rounded, may end in a small point (4) or extend into a triangular lobe (Euchirella maxima) Legs: P2-P4 broad, Exp segs armed with strong outer spines. ♀ P4 coxa with an inner seta (5) and spines/teeth (6), number of spines

A

B

Urosome: ♀ 4 segs, very short (10); ♂ 5 segs. ♀ GS may (11) or may not be symmetrical, having projections (12). AS very short (13). Caudal rami short and divergent (14) Notes: twelve species reported for the study area. See identification Table 9 for the 9 of the most common species Similar taxa: Chirundina spp., Pseudochirella spp. Distribution and ecology: epi-hadopelagic, CNRY, NASE, GFST, NASW, NADR, NWCS, SARC, ARCT, NECS

C ♀

♀

varying with species. ♀ P5 absent. ♂ P5 long (7), asymmetrical and usually biramous; R-leg longer than L-leg, may form a chela (8); Exp 2 segs, Enp 1 seg. L-leg Exp 3 segs, last seg bifurcated and hairy (9); Enp with 1 seg, if present

D

E 6

♂

♂

R

F L 5

7

10 11

14 9

13 2

I

G

K

6

L ♀

♀

H

1

1

3

M ♀

12

L

R

♂

J ♂

O

N 6

13 14

8 4

Figure 140:  Euchirella rostrata. A, ♀ dorsal view; B, ♀ lateral view; C, ♂ dorsal view; D, ♂ lateral view; E, ♀ P4; F, ♂ P5. Euchirella messinensis.

G, ♀ head lateral view; H, ♂ head lateral view; I, ♀ P4 coxa; J, ♀ urosome dorsal view. Euchirella curticauda. K, ♀ dorsal view; L, ♀ lateral view; M, ♂ head lateral view; N, ♀ P4 coxa. Euchirella amoena. O, ♂ P5. A, C, Rose (1933); B, E, Park (1978); D, K-O, Bradford and Jillett (1980); G–J, Park (1976); N, Sars (1924).

E. amoena Giesbrecht, 1888

E. bella Giesbrecht, 1888

E. bitumida With, 1915

E. curticauda Giesbrecht, 1888

E. maxima Wolfenden, 1905

E. messinensis (Claus, 1863)

E. pulchra E. rostrata Lubbock, 1856 Claus, 1866

E. truncata Esterly, 1911

Total length (mm)

♀ 2.7–4.0 ♂ 2.9–3.9

♀ 3.3–4.9 ♂ 3.1–3.9

♀ 4.7–7.1 ♂ 4.8–6.1

♀ 2.5–4.8 ♂ 3.0–4.3

♀ 6.1–8.7 ♂ 6.1–7.4

♀ 3.8–6.2 ♂ 2.8–5.5

♀ 2.9–4.4 ♂ 3.0–4.2

♀ 2.0–4.1 ♂ 2.5–3.1

♀ 4.4–6.8 ♂ 4.5–5.6

Head crest

No

No

Yes, large

Yes

Yes, large

No

No

No

No

Rostrum

Small

Robust

Short robust

Rudimentary

Small

Large

Well developed Long

Small

♀ GS

Symmetrical

Smooth projection on right

Dorsal swellings

Symmetrical

Small projection on right

Large dorsal projection

Swelling on left

Symmetrical

Asymmetrical, small swellings on both sides

♀ P4 coxa

5 small teeth

3–4 robust spines

1 robust spine

6–13 spines

1 robust spine

2 spines

2 spines

7–9 teeth

1 long spine

Distribution

Epi-bathy Epi-hadal CNRY, NASE, CNRY, NASE NADR, SARC, NWCS, GFST, NASW

Epi-bathy CNRY, NASE, NASW, GFST, NWCS, NADR, SARC, NECS

Epi-bathy CNRY, NASE, NWCS, GFST, NASW, NADR, SARC, ARCT, NECS

Epi-meso CNRY, NASE, NWCS, NADR, NECS, SARC

Epi-bathy CNRY, NASE, GFST, NASW, NADR, SARC, NECS

Epi-bathy CNRY, NASE, GFST, NASW, NWCS, NECS

Epi-bathy CNRY, NASE, GFST, NASW, NADR, NWCS, SARC, ARCT, NECS

Epi-bathy CNRY, NASE, GFST, NASW, SARC, NECS

304 ta xonom y

Table 9:   Morphological and ecological characteristics used in the species identification of female Euchirella.

DISTRIBUTION: E. amoena

DISTRIBUTION: E. messinensis

DISTRIBUTION: E. curticauda

DISTRIBUTION: E. rostrata

306 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Aetideidae Genus: Gaetanus Giesbrecht, 1888 Total length: ♀ 1.7–9.7 mm; ♂ 1.7–7.0 mm Cephalosome: ♀ head with (1) or without (2) frontal spine. A1 reaching from end of metasome to twice as long as body; not geniculate in ♂. Rostrum single, ♀ may be bifid at tip (3). Mx2 typical of aetideids with a claw-like seta on 4th and 5th lobes, see Undeuchaeta. ♀ Mxp protopod with lateral plate (4), or absent, and with a distal ‘teat-like’ projection (5) Metasome: body robust, 3 segs. Cephalosome may be fused with 1st seg, 4th–5th segs fused. End of metasome expanded into large (6) or small (7) spine-like projections (smaller in ♂), rarely absent Legs: P2–P4 broad, Exp segs armed with strong spines. ♀ P4 coxa with an inner seta and a group of brush-like spines (8). ♀ P5 absent.

Gaetanus pileatus

1

Farran, 1903

C

B

♂ P5 biramous, exps long and slender; R-leg Exp 2 segs, last seg long and swollen medially (9); L-leg Exp 3 segs, last seg may be bilobed; Enps 1 seg Urosome: ♀ 4 segs and usually short. ♂ 5 segs with very short AS (10). ♀ GS symmetrical. ♀ caudal rami divergent (11) Notes: twenty species reported for the study area. The length and direction of the frontal spine and end of metasome spine are useful aids in species identification. Please refer to Markhaseva (1996) for a comprehensive key to the genus Similar taxa: Euchirella spp. Distribution and ecology: epi-hadopelagic, generally a mesobathypelagic genus. Most species with a widespread distribution: CNRY, NASE, GFST, NASW, NADR, NWCS, SARC, ARCT, NECS

D

L

A

♀ 4.1–6.7 mm

♂ 4.4–5.3 mm

♀

Meso-bathypelagic CNRY, NASE,

CNRY, NASE, NADR, NWCS,

NADR, NWCS,

E

NASW, GFST,

9

NECS

Gaetanus brevispinus

6

H

NASW,GFST, SARC, NECS

SARC, ARCT,

1

♀

♀ 1.7–2.4 mm ♂ 1.9–2.4 mm Epi-bathypelagic

F

♂

♀

Gaetanus minor Farran, 1905 G

R

♀

10 2

(Sars, 1900) ♀ 3.6–4.9 mm

J

♂ 2.1–4.0 mm Meso-hadopelagic CNRY, NASE,

K

♀

M

L

♀

O

P 8

♂

5

NADR, NWCS, NASW, GFST,

N

SARC, ARCT, NECS

4

I 3

7 11

Figure 141:  Gaetanus pileatus. A, ♀ dorsal view; B, ♀ lateral view; C, ♂ lateral view; D, ♀ urosome lateral view; E, ♂ urosome lateral view;

F, ♂ P5. Gaetanus minor. G, ♀ head dorsal view; H, ♀ head lateral view. Gaetanus brevispinus. I, ♀ rostrum; J, ♀ dorsal view; K, ♀ lateral view; L, ♂ dorsal view; M, ♀ urosome lateral view; N, ♂ urosome lateral view; O, ♀ P4; P, ♀ Mxp. A–C Rose (1933); D, Park (1978); E, F Park (1975b); G–P Markhaseva (1996).

crustacea: copepoda  307 Order Calanoida Superfamily Clausocalanoidea  Family Aetideidae Genus: Undeuchaeta Giesbrecht, 1888 Total length: ♀ 3.0–6.7 mm; ♂ 2.8–5.6 mm Cephalosome: head with (1), or without crest (2). A1 reaching from end of metasome to mid urosome; proximal segs bulbous (3);  ♂ A1 not geniculate. Rostrum single and spiniform (4). ♀ supralabrum not prominent, unlike in Euchaetidae. Mx2 typical of family, with claw-like setae on 4th and 5th lobes (5). ♀ and copepodites have large carnivorous Mxp (6); ♂ mouthparts reduced Metasome: body robust, 3–4 segs. ♀ end of metasome, asymmetrical, extended into triangular (7) or rounded lobes; rounded in ♂ (8)

than R-leg basis; R-leg Exp 2 segs, last seg either spiniform or blunt; L-leg Exp 3 segs, last seg styliform with hairs, 2nd seg with a toothlike projection (9); Enp 1 seg, longer on right Urosome: ♀ 4 segs. ♂ 5 segs. ♀ GS asymmetrical: with lateral spine on right in Undeuchaeta plumosa (10); with spine on genital field in Undeuchaeta major and Undeuchaeta incisa (11) AS very short. ♀ caudal rami short Notes: three species reported for the study area Similar taxa: Euchaetidae and ♂ Euchirella spp. Distribution and ecology: Epi-bathypelagic. CNRY, NASE, GFST, NWCS, SARC, NASW, NECS, NADR, ARCT

Legs: P4 coxa without large spines, may have small spinules. ♀ P5 absent. ♂ P5 large, asymmetrical and biramous: L-leg basis longer

Undeuchaeta plumosa B

♀

(Lubbock, 1856)

C ♂

♂

8

♀ 3.0–4.7 mm ♂ 2.8–3.9 mm Epi-mesopelagic CNRY, NASE, NADR, NWCS, NASW, GFST, SARC, NECS

4

Giesbrecht, 1888

Undeuchaeta incisa 5

E

Esterly, 1911 ♀ 5.5–6.7 mm ♂ 4.1–5.6 mm Meso-bathypelagic CNRY, SARC, NECS, ARCT

R

H 10 9 1

J

♀ 3.5–6.5 mm ♂ 3.0–5.5 mm Epi-bathypelagic CNRY, NASE, NADR, NWCS, NASW, GFST, SARC, NECS

6

G

F

Undeuchaeta major

D

L

2

I

3

A

K

L

L

M

R

7 11 1

N

O

P

Q

R

L

11

Figure 142:  Undeuchaeta plumosa. A, ♀ dorsal view; B, ♂ lateral view; C, ♂ dorsal view; D, ♀ Mxp; F, ♀ rostrum; G, ♀ urosome lateral view;

H, ♀ GS ventral view; I, ♂ P5. Undeuchaeta major. J, ♀ rostrum; K, ♀ urosome lateral view; L, ♀ GS ventral view; M, ♂ P5. Undeuchaeta incisa. E, ♀ Mx2; N, ♀ rostrum; O, ♀ urosome lateral view; P, ♀ GS ventral view; Q, ♂ P5. A, C, F, J, M, Sars (1924); B, D, G, H, I, K, L, Q, Markhaseva (1996); E, N, O, P, Park (1978); H, Corral Estrada (1970).

308 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Clausocalanidae Genus: Clausocalanus Giesbrecht, 1888

asymmetrical: L-leg long, thin with 5 segs, 5th seg very small and attached sub-apically to the last seg (7); R-leg short with 2-3 segs (R-leg longest in C. furcatus)

Total length: ♀ 0.7–2.0 mm; ♂ 0.5–1.5 mm Cephalosome: A1 reaching from end of metasome to past GS. ♂ A1 not geniculate. ♀ rostrum with 2 filaments (1), ♂ rostrum reduced to a single ventral knob, (not well developed in ♂ C. furcatus)

Urosome: ♀ 4 segs. ♂ 5 segs. GS in lateral view shows seminal receptacle with a dorsal (8) and ventral lobe (9), the shape and size of which is important in species identification. ♂ AS very short (10) Notes: eight species in the study area. In the review of the genus, Frost and Fleminger (1968) split Clausocalanus into 3 groups based on the characteristics of the ♀ seminal receptacle

Metasome: 3 segs, cephalosome fused with 1st metasome seg. Segs 4 and 5 fused. Body ovoid in shape, end of metasome rounded (2). The internal position of the spermatophore in the ♂ is used in species identification (3)

Similar taxa: Acrocalanus spp, Ctenocalanus vanus, Pseudocalanus spp., Microcalanus spp. and Paracalanus spp.

Legs: P2-P3 basis distally broad with spines, crown-like in appearance (4). P2-P4 with 3 outer-edge spines equally spaced on 3rd seg of Exp (5), unlike in Paracalanidae. ♀ P5 small, symmetrical and uniramous with 3 segs; last seg has bifurcate tip (6). ♂ P5

A

B

C

Distribution and ecology: epipelagic, widespread in warmtemperate waters (absent in Arctic waters). NECS, SARC, NADR, NASE, NASW, GFST, NWCS, CNRY

D

E

F L

♀

♂

♀

R

♂ 6

J 8 3

9

2 10

7

G

5 4

H 1

K I

Figure 143:  Clausocalanus arcuicornis. A, ♀ lateral view; B, ♀ dorsal view; C, ♂ lateral view; D, ♂ dorsal view; E, ♀ P5; F, ♂ P5; G, ♀ rostrum;

H, ♀ P3; I, ♀ P2 basis; J, ♀ GS lateral view; K, distribution map of Clausocalanus spp. A–G, I, J, Frost and Fleminger (1968); H, Rose (1933).

crustacea: copepoda  309 Table 10:   Clausocalanus species.

Species

Total Length

♀ rostrum and urosome ratios

♀ GS lateral

Group I Clausocalanus lividus Frost and Fleminger, 1968

Clausocalanus mastigophorus (Claus, 1863)

♀1.3–2.0 mm

R: straight

d.l: originating from anterior v.l large, directed dorsally

♂1.1–1.5 mm

Distribution CNRY, NASE, NWCS, GFST, NECS

P:U 2.7–2.2:1 U:GS 2.2–2.8:1 ventral profile non protruberant

♀ 1.2–1.9 mm d.l: thin, directed dorsally, originating from anterior v.l

R: long and rounded P:U 3.2–3.8:1 U:GS 2.0–2.6:1

♂ 1.1–1.5 mm

CNRY, NASE, NWCS, GFST, NECS

ventral profile non protruberant Group II

Clausocalanus jobei Frost and Fleminger, 1968

Clausocalanus arcuicornis (Dana, 1849)

♀ 1.0–1.6 mm

ventral profile protruberant

♂ 0.9–1.1 mm

♀1.1–1.6 mm

d.l: difficult to separate from v.l

♀ 0.8–1.8 mm

d.l: originating from posterior v.l and large

♂ 0.7–1.1 mm

R: slender, curved ventro-posteriad ventral profile undulate, with a step ventral profile undulate

v.l large

♀ 0.7–1.1 mm

Clausocalanus parapergens Frost and Fleminger, 1968

R: thick, short and slightly curved

CNRY, NASE, NWCS, NASW, GFST, NECS

CNRY, NASE, NWCS, NASW, GFST

CNRY, NASE, NWCS, GFST

P:U 2.0–2.7:1 U:GS 3.2–4.3:1

P:U 2.6–3.0:1 U:GS 2.4–3.0:1

v.l large ♂ 0.5–0.7 mm

P:U 2.6–3.0:1 U:GS 2.4–3.0:1

R: slender, curved ventro-posteriad

d.l: short and digitiform Clausocalanus pergens Farran, 1926

CNRY, NASE, NWCS, GFST, NECS, SARC

ventral profile may be concave

♂ 0.5–0.6 mm

Clausocalanus furcatus (Brady, 1883)

P:U 2.7–3.2:1 U:GS 2.2–3.0:1

♀ 0.7–0.9 mm

Group III

P:U 2.3–2.9:1 U:GS 2.3–2.9:1

R: short and straight

d.l: attached to anterior v.l digitform, directed dorso-posteriad

♂ 1.0–1.3 mm

Clausocalanus paululus Farran, 1926

R: slender, curved ventro-posteriad

d.l: small, directed dorso-posteriad difficult to separate from v.l

CNRY, NASE, NWCS, NASW GFST, NADR, NECS

ventral profile convex without step R: slender, curved ventro-posteriad

♀ 1.0–1.7 mm

CNRY, NASE, NWCS, NECS

d.l: short and bulb-like ♂ 1.0–1.2 mm

v.l large

ventral profile undulate

P:U 3.1–3.6:1 U:GS 2.2–2.9:1

All figures from Frost and Fleminger (1968). d.l, dorsal lobe; v.l, ventral lobe; R, rostrum; P:U, ratio of prosome length to urosome length; U:GS, ratio of urosome length to length of genital segment.

310 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Clausocalanidae Genus: Ctenocalanus Giesbrecht, 1888 Ctenocalus vanus Giesbrecht, 1888 Total length: ♀ 0.8–1.7 mm; ♂ 1.1–2.0 mm Cephalosome: head rounded, rostrum with filaments. ♀ and ♂ A1 reaching caudal rami. ♂ A1 not geniculate and thickened at base due to segmental fusion (1) Metasome: 3 segs. Body ovoid in shape, end of metasome rounded (2) Legs: P1 Enp with 5 setae, cf. 4 setae in Microcalanus. P2 and P3 basis with small spines, small in ♀ and larger in ♂, (3). P2–P4 with 3 outeredge spines equally spaced on 3rd seg of the Exp (4), unlike in

A

Paracalanidae. P3 and P4 with comb-like external spine on the 3rd Exp seg (5). ♀ P5 short, asymmetrical, only developed on left, consisting of 2–4 segs, somewhat variable in form (6). ♂ P5 asymmetrical, R-leg absent, L-leg with 5 segs, last seg with a tuft of hair (7) Urosome: ♀ 4 segs. ♂ 5 segs. ♀ seminal receptacle v-shaped in lateral view (8) Notes: one species recorded in study area Similar taxa: Clausocalanus spp., Acrocalanus spp., Pseudocalanus spp., Microcalanus spp. and Paracalanus spp. Distribution and ecology: epi-bathypelagic, warm oceanic and shelf waters. CNRY, NASE, NWCS, GFST, NADR, NASW, NECS

B

1

D

♀

♂

E

7

8

C

6 3

L

F L

2

H 4

G

5

Figure 144:  Ctenocalanus vanus. A, ♀ lateral view; B, ♀ GS lateral view; C, ♀ P5; D, ♂ lateral view; E, ♂ P5; F, ♂ P3 basis; G, P4 exopod segs 2–3; H, distribution map of C. vanus. A–G Bradford-Grieve (1994).

crustacea: copepoda  311 Order Calanoida, Superfamily Clausocalanoidea  Family Clausocalanidae Genus: Microcalanus Sars, 1901

Urosome: ♀ 4 segs. ♂ 5 segs. ♀ GS laterally swollen in dorsal view

Microcalanus pygmaeus (Sars, 1900)

Notes: this genus was originally described as consisting of two species, M. pygmaeus (Sars, 1900) (with ♀ A1 reaching to end of caudal rami) and M. pusillus Sars, 1903 (with ♀ A1 reaching to GS). However, Farran & Vervoort (1951) consider them variations of the same species, M. pygmaeus. Intermediate forms have also been recorded

Total length: ♀ 0.6–1.1 mm, ♂ 0.6–1.1 mm Cephalosome: head rounded, rostrum with filaments. ♀ and ♂ A1 length variable, reaching from GS to end of caudal rami. ♂ A1 not geniculate and thickened at base due to segmental fusion (1) Metasome: 3 segs. Body small and compact, end of metasome rounded (2) Legs: P1 Enp with 4 setae (3), cf. 5 setae in other members of the Clausocalanidae. P2-P4 with 3 outer-edge spines equally spaced on 3rd seg of the Exp, (4) unlike in Paracalanidae. ♀ P5 absent. ♂ P5 asymmetrical; L-leg long with 5-6 segs, R-leg shorter with 2-3 segs reaching end of 2nd seg of L-leg (5)

A

Similar taxa: Microcalanus pusillus (see note), Paracalanus spp., Pseudocalanus spp. and Ctenocalanus vanus Distribution and ecology: epi-bathypelagic. NWCS, ARCT, SARC, NECS, NADR

B

1

C

D L

R

2

5

F

3

4

G

E

Figure 145:  Microcalanus pygmaeus. A, ♀ lateral view; B, ♀ urosome lateral view; C, ♂ lateral view; D, ♂ P5; E, P1; F, P2; G, distribution

map of Microcalanus spp. A–D Vidal (1971); E, F Bradford-Grieve (1994).

312 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Clausocalanidae Genus: Pseudocalanus Boeck, 1873 Total length: ♀ 0.9–2.3 mm; ♂ 0.8–1.7 mm Cephalosome: head rounded, rostrum with 2 filaments. A1 reaching past GS, but not extending beyond caudal rami. ♂ A1 not geniculate and with segs 1-2 fused and thickened at base (1) Metasome: 3 segs, cephalosome fused with 1st metasome seg. Segs 4-5 fused. End of metasome rounded (2) Legs: P1 Enp with 5 setae (cf. 4 setae in Microcalanus). P2-P4 with 3 outer-edge spines (3), equally spaced on 3rd seg of the Exp (unlike in Paracalanidae). ♀ P5 absent. ♂ P5 asymmetrical, uniramous, both legs long and slender: L-leg longer than R-leg with 5 segs, last seg hairy and tipped with a spine (4); R-leg 4 segs, last seg styliform (5) Urosome: ♀ 4 segs. ♂ 5 segs. ♀ GS elongated (6), seminal receptacle shape irregularly rounded (7). ♂ AS very short (8)

A

1

Notes: according to Frost (1989), who revised the genus, 5 species are reported for the study area. Due to the complexity of morphometrics needed to separate the species, identification is usually limited to Pseudocalanus spp. Similar taxa: Paracalanus spp., Clausocalanus spp. and Microcalanus spp. Distribution and ecology: epi-bathypelagic, neritic, Arctictemperate waters P. elongatus (Boeck, 1865): NECS, SARC P. acuspes (Giesbrecht, 1881): NECS, SARC, ARCT, NWCS P. newmani Frost, 1989: NWCS P. moultoni Frost, 1989: NWCS, ARCT, SARC P. minutus (Krøyer, 1845): SARC, ARCT, NWCS

B

C

D L

♀

R

♂

5 2 3

4

E

egg sac 6

G

7

F 8

Figure 146:  Pseudocalanus elongatus. A, ♀ lateral view; B, ♂ lateral view; C, P2; D, ♂ P5; E, ♀ urosome lateral view; F, ♂ urosome dorsal view; G, distribution map of Microcalanus spp.. A–F, Sars (1903).

crustacea: copepoda  313 Order Calanoida Superfamily Clausocalanoidea  Family Diaixidae Genus: Diaixis Sars, 1902 Total length: ♀ 0.7–1.2 mm; ♂ 0.7–1.0 mm Cephalosome: A1 reaching from end of metasome to mid urosome; not geniculate in ♂. Head rounded: rostrum simple, may have two fine filaments. ♀ Mx2 Enp with 5 short brush-like and 3 long worm-like (1) sensory filaments Metasome: body oval and compact, 3-4 segs. Cephalosome fused with 1st seg, 4th–5th segs may be fused. End of metasome expanded into transparent irregular points (2), rounded and smaller in ♂ (3)

spinules on posterior surface. ♀ P5 absent. ♂ P5 aysmmetrical, uniramous, very irregular and elongate reaching beyond end of urosome (6). R-leg basis swollen with group of curved spines (7) Urosome: slender. ♀ 4 segs, GS with dorsal projection in Diaixis hibernica (8). ♂ 5 segs. ♂ AS very short Notes: three species reported for the study area. Members of the Diaixidae family are very similar to those in the Scolecitrichidae and further study is needed to determine if the family is valid Distribution and ecology: hyperbenthic, but also found in shallow coastal waters. CNRY, NASE, NADR, NWCS, SARC, NECS

Legs: P1 with setae on basis (4). Limbs somewhat spinulose in appearance (5). P2-P3 Enps and exps may be ornamented with

A Diaixis hibernica

B

D

♀ 0.7–1.2 mm ♂ 0.7–1.1 mm Hyperbenthic CNRY, NASE, NADR, SARC, NECS

F

G 7

♂

♀

(Scott, 1896)

E 4

1

8

3

C

R

L

5

2

H Diaixis pygmaea (Scott, 1896) ♀ 0.7–1.0 mm ♂ 0.7–1.0 mm Hyperbenthic CNRY, NASE, NADR, SARC, NECS

7

I

6

♀ L

R

J

Diaixis asymmetrica

K ♀

Grice & Hulsemann, 1970 ♀ 1.2 mm ♂ 1.2 mm Hyperbenthic NWCS

R L

Figure 147:  Diaixis hibernica. A, ♀ lateral view; B, ♀ Mx2 Enp; C, ♀ urosome dorsal view; D, ♂ lateral view; E, ♀ P1; F, ♀ P4; G, ♂ P5.

Diaixis pygmaea. H, ♀ lateral view; I, ♂ P5. Diaixis asymmetrica. J, ♀ lateral view; K, ♂ P5. A, C–G, Sars (1903); B, Ferrari and Markhaseva (1996); H-K, Andronov (2002).

314 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Euchaetidae Urosome: ♀ 4 segs. ♀ 5 segs. AS very short (8). Shape of ♀ GS species-specific. ♀ caudal rami with appendicular setae well developed, straight, thicker and much longer than other setae (9) (thinner and geniculated, or curved in Paraeuchaeta). ♂ appendicular caudal setae geniculated, thinner than, but nearly as long as 2nd or 3rd marginal caudal seta (10)

Genus: Euchaeta Philippi, 1843 Total length: ♀ 2.3–7.2 mm; ♂ 2.6–6.9 mm Cephalosome: head pointed (1). A1 with several very long setae (2). Rostrum single, strong, spiniform and anteriorly directed (3). ♀  has prominent supralabrum with long stiff hairs (4) (unlike in Undeuchaeta). Mouthparts atrophied in ♂, but large and carnivorous in ♀ and copepodites. ♀ Mx2 with 1 (5, a) or 2 (5, b) (1 or none in Paraeuchaeta) of the 6 apical setae with long spinules

Notes: five species reported for the study area. Species can be identified by the shape of the ♀ GS and the shape of the ♂ P5 L-Exp. See Park (1995) for a detailed review of the genus

Metasome: body long and robust. 3-4 segs. Cephalosome partially fused with 1st seg, 4th–5th segs fused. End of metasome rounded

Similar taxa: Paraeuchaeta spp., Undeuchaeta spp. and ♂ Euchirella spp. Distribution and ecology: carnivorous; epi-bathypelagic. Widely distributed, but absent from Arctic waters. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS, SARC

Legs: ♀ P5 absent. ♀ P5 large, asymmetrical and biramous; R-Exp 2 segs, last seg spiniform; Enp with 1 seg and long. L-Exp with segs 2 and 3 forming a chela, last seg ending in a long spine-like process (6) (short in E. spinosa and Paraeuchaeta); size, shape and position of processes, lamellae (7) and tuft of hairs on the chela are speciesspecific. L-Enp rudimentary

A

1

B

2

1

♂

♀

C

F E

3 4

R L

8

10

♀

Egg sac

D

a)

7 b)

5 9 6

Figure 148:  Euchaeta marina. A, ♀ dorsal view; B, ♂ lateral view; C, ♀ head lateral view; D b), ♀ Mx2 exploded image; E, ♂ P5; F, ♂ caudal

rami. Eucheata acuta. D a), ♀ Mx2 apical setae. A, B modified from Rose (1933); C–F Park (1995).

Table 11:   Euchaeta species.

Euchaeta acuta Total length

Euchaeta media

Euchaeta marina

Giesbrecht, 1893 [‘1892’] ♀ 3.4–4.7 mm, ♂ 3.2–4.8 mm

Euchaeta pubera

Giesbrecht, 1888

(Prestandrea, 1833)

♀ 2.3–3.9 mm, ♂ 2.6–3.6 mm

Euchaeta spinosa

Sars, 1907

♀ 3.3–4.8 mm, ♂ 3.1–4.2 mm

Giesbrecht, 1893 [‘1892’]

♀ 2.9–4.4 mm, ♂ 3.4– 3.7 mm

♀ 5.2–7.2 mm, ♂ 5.2–6.9 mm

Deep notch

Head lateral view

♂

♀

♀

(Park, 1995)

♀ Mx2

♀ GS

♀

♂

♂

(Park, 1995)

(Park, 1995)

(Park, 1995)

♂

(Park, 1995)

Mx2, two setae with long spinules

Mx2, one seta with long spinules

Mx2, one seta with long spinules

Mx2, one seta with long spinules

Asymmetrical, deep shoulder (1) on left

Asymmetrical, protuberance (1)

Asymmetrical, swelling on right (1)

Approx. symmetrical in DV, asymmetric

Almost symmetrical in DV,

side Conical GP placed high on GS (2)

placed low on right side

and notch on left side (2) Conical GP placed central on seg (3)

in VV. Right flange of GS hook shaped (1) Also body of ♀ and ♂ covered in fine hairs.

asymmetric in VV GF projecting past left side of GS (1)

LV

VV

1

LV

VV

LV

1

(Park, 1995)

SL (1) not reaching tuft of hair (2) SL round (3) at distal edge DP reaching end of SL (4) 3rd Exp seg long (5)

2

(Park, 1995)

SL (1) reaching beyond tuft of hair (2) DP placed distal to tuft of hairs (3) 3rd Exp seg long (4) 1

3 4

1 Hook

1

(Park, 1995)

3

SL (1) not reaching tuft of hair (2) SL bilobed (3) at distal edge 3rd Exp seg. long (4)

1 3

5

2

(Park, 1995)

SL (1) not reaching tuft of hair (2) SL pointed (3) at distal edge DP not reaching end of SL (4) 3rd Exp. seg. long (5)

1

4

LV

VV 1

3 1

1

DV

LV

2

2

2nd & 3rd segs of Left Exp

♀

Mx2, one seta with long spinules

VV

♂ P5

♂

♀

4

(Park, 1995) SL spoon-like (1) at distal edge SL teeth large at the distal edge (2) 3rd Exp. seg. short (3) 1

3 4

2

2 3

5

2

(Mauchline, 1999)

(Mauchline, 1999)

(Mauchline, 1999)

(Mauchline, 1999)

(Mauchline, 1999)

E. media

E. media

E. acuta

E. acuta

Paraeucheata barbata

Distribution

Epi-mesopelagic. CNRY, NASE, NWCS, NASW, NADR, GFST, NECS, SARC

Epi-mesopelagic. CNRY, NASE, NWCS, NASW, NADR, GFST, SARC, NECS

Epi-bathypelagic. CNRY, NASE, NWCS, NASW, NADR, GFST

Epipelagic. CNRY, NASE, NADR, NASW

Epi-bathypelagic. CNRY, NASE, NECS, NWCS, NADR, NASW, SARC

GP, genital prominence; GF, genital field; SL, serrated lamella; DP, digitiform process.

crustacea: copepoda  315

Similar spp.

DISTRIBUTION: Euchaeta acuta

DISTRIBUTION: Eucaheta marina

DISTRIBUTION: Euchaeta media

DISTRIBUTION: Euchaeta pubera

DISTRIBUTION: Euchaeta spinosa

crustacea: copepoda  317 Order Calanoida Superfamily Clausocalanoidea  Family Euchaetidae Genus: Paraeuchaeta A. Scott, 1909

Euchaeta); size, shape and position of processes (11), lamellae (12) and tuft of hairs (13) on chela is species-specific. L-Enp rudimentary

Total length: ♀ 2.6–12.0 mm; ♂ 3.2–10.0 mm

Urosome: ♀ 4 segs. ♀ 5 segs. AS very short (14). Shape of ♀ GS species-specific. Appendicular setae on caudal rami well developed in ♀ and may be geniculate or strongly curved (straight and thicker in Euchaeta), much thinner and longer than other setae (15); in ♂ much shorter and thinner than distal marginal setae (16)

Cephalosome: head pointed (1). A1 with several very long setae (2). Rostrum single, strong, spiniform and anteriorly directed (3). ♀  has supralabrum prominent with long stiff hairs (4) (unlike in Undeuchaeta). Mouthparts atrophied in ♂, but large and carnivorous in ♀ and copepodites. ♀ Mx2 with 1 or none (1 or 2 in Euchaeta) of the 6 apical setae (5) with long spinules

Notes: 20 species occur in the study area. Species can be identified by the shape of the ♀ GS and the shape of the ♂ P5 L-Exp. See Park (1995) for a detailed review of the genus

Metasome: body long and robust, 3-4 segs. Cephalosome partially fused with 1st seg, 4th-5th segs fused. End of metasome rounded, obtusely pointed or with toothlike process (6)

Similar taxa: Euchaeta spp., Undeuchaeta spp. and ♂ Euchirella

Legs: ♀ P5 absent. ♀ P5 large, asymmetrical and biramous; R-Exp with 2 segs, last seg either spiniform or blunt (7); Enp long with 1  seg. L-Exp, 2nd (8) and 3rd (9) segs forming a chela, last seg tapering into a blunt end with a minute spine (10) (long in

2

1

A

Distribution and ecology: carnivorous. Meso-hadopelagic. Widely distributed. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS, SARC, ARCT

F

E

B

3

♀

1

R

4

L

6

♀

♂

2

C

♀

G

♂ 12

D

11

13

14 15

5

16

7

8

9

10

Figure 149:  Paraeuchaeta norvegica. A, ♀ lateral view; B, ♂ dorsal view; C, ♀ head lateral view; E, ♀ end of metasome; F, ♂ P5 showing

detail of left Exp. Paraeuchaeta malayensis. D, Mx2 exploded image; G, ♂ caudal rami. N.B.: P. malayensis not found in Atlantic, used for illustrative purposes only. A, B, F, Rose (1933); C–E, G, Park (1995).

Total length

Paraeuchaeta barbata

Paraeuchaeta glacialis

Paraeuchaeta gracilis

Paraeuchaeta hebes

Paraeuchaeta norvegica

(Brady, 1883)

(Hansen, 1887)

(Sars, 1905)

(Giesbrecht, 1888)

(Boeck, 1872)

♀ 6.0–12.0 mm, ♂ 6.1–10.0 mm

Head lateral view

♀

♀ 9.8–11.0 mm, ♂ 6.2–8.0 mm

♀

♂

Mx2, no setae with long spinules

♀ GS

VV

LV

1

1 (Park, 1995)

♂ P5 2nd & 3rd segs of Left Exp

(Park, 1995)

1

3

GS with dorsal step in LV (1)

LV

2

VV

LV

(Park, 1995)

SL hollow and claw-shaped (1) Medial spines on 2nd seg (2) Proximal tooth on 2nd seg (3)

3

Mx2, no setae with long spinules ♀ end of metasome toothed (1) rounded in ♂ Ventral bumps (2) above GP. Large GP placed low on GS (3)

1 2

1 1

♂

(Park, 1995)

Mx2, one seta with long spinules

2

SL square in shape with rounded serrations (1) SL and DP short, not reaching tuft of hair on 3rd seg (2)

1

VV

LV

2

1

(Heptner, 1971)

SL square in shape (1) Proximal tooth on 2nd seg (2) DP reaching tuft of hair (3)

2

Squared extended GP placed centrally on GS (1) VV

LV

♀

(Rose, 1933)

Mx2, no setae with long spinules

Conical GP (1) in LV slightly swollen in VV (2)

♀ 7.0–11 mm, ♂ 5.5–7.5 mm

♀

(Park, 1995)

Mx2, no setae with long spinules

Tubercle on lower left side (1)

♀ 2.6–3.6 mm, ♂ 2.6–3.2 mm

♂

♀

♂

(Park, 1995)

(Park, 1995)

♀ Mx2

♀ 5.8–7.0 mm, ♂ 5.1–6.5 mm

2

1

2

3

(Mauchline, 1999)

(Park, 1995)

SL abruptly narrowing on distal end (1) Row of spines extending on most of 2nd seg (2)

SL hollow and claw-shaped (1) Medial spines on 2nd seg (2) also a small proximal (3) and distal (4) tooth

1 2

3

2

4 1

(Mauchline, 1999)

(Mauchline, 1999)

(Mauchline, 1999)

(Mauchline, 1999)

3rd seg of P2 Exp with middle outer spine reaching to end of seg

Note Similar spp.

P. glacialis

P. barbata

P. norvegica

Distribution

Meso-bathypelagic. ARCT, SARC, CNRY, NASE, NWCS, NASW, NECS

Epi-bathypelagic. ARCT, SARC, NWCS

Epi-bathypelagic. CNRY, SARC, Epipelagic, coastal. SARC, NECS, NADR, NASE NECS, CNRY, NASW, NASE

GP, genital prominence; SL, serrated lamella; DP, digiform process.

(Mauchline, 1999) 3rd seg of P2 Exp with middle outer spine not reaching end of seg

Euchaeta acuta

P. gracilis Epi-hadopelagic. ARCT, SARC, NECS, NWCS, NASW, NADR, GFST, NASE

318 ta xonom y

Table 12:   Paraeuchaeta species.

DISTRIBUTION: Paraeuchaeta glacialis

DISTRIBUTION: Paraeuchaeta hebes

DISTRIBUTION: Paraeuchaeta gracilis

DISTRIBUTION: Paraeuchaeta norvegica

320 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Scolecitrichidae Genus: Amallothrix Sars, 1925 and Pseudoamallothrix Vyshkvartzeva, 2000

Pseudoamallothrix ovata) and elongate; L-Enp as long as or longer than Exp in Pseudoamallothrix (8), approximately 2/3 length of Exp in Amallothrix; R-Enp rudimentary

Total length: 1.0–6.4 mm

Urosome: ♀ 4 segs. ♂ 5 segs. AS, very short, not longer than preceding segment (longer in Racovitzanus) (9)

Cephalosome: A1 reaching past GS but not extending beyond caudal rami. Head rounded. Rostrum with strong bifid branches (1): short in Pseudoamallothrix, long in Amallothrix; with 2 filaments (2): filaments longer than branches and thick in Pseudoamallothrix, shorter than branches in Amallothrix. Mx2 Enp with 3 worm-like and 5 brush-like sensory filaments (3)

Similar taxa: Archescolecithrix spp., Paracalanus spp., Racovitzanus spp., Scaphocalanus spp., Scolecithricella spp. Notes: fourteen species of Amallothrix and six species of Pseudoamallothrix reported from study area. Many species in the Scolecitrichidae are very similar and many have relatively recently changed genus name. Identification to genus level can rely on subtle features, making a correct identification difficult. See Vyshkvartzeva (2001) for a detailed review of the family

Metasome: 3 segs, cephalosome fused with 1st metasome seg. Segs 4–5 fused. End of metasome rounded Legs: P2-P3 Exp and Enp strongly spinulose on posterior surface and P4 with only a few spinules (4). P2-P4 coxa with projection on inner margin in Pseudoamallothrix (5). ♀ P5 with 2–3 segs (1 seg in Scolecithricella), last seg with 2 spines in Pseudoamallothrix and usually 3 spines in Amallothrix (6), inner spine strong and elongate (7). ♂ P5 asymmetrical, usually biramous (uniramous in

Distribution and ecology: meso-bathypelagic (P. ovata epipelagic). CNRY, NASE, NASW, GFST, NWCS, NADR, NECS (absent from North Sea), SARC, ARCT (Pseudoamallothrix only)

Pseudoamallothrix emarginata (Farran, 1905)

A

♀ 2.5–4.8 mm

♂ 3.8–4.1 mm

B

♀

Meso-bathypelagic CNRY, NASE,

E

D

Amallothrix valida

F

L

5

♂

R

(Farran, 1908)

1

NWCS, NASW,

2

SARC, NECS

C

4

1

♀ 2.1–4.5 mm ♂ 4.0–5.4 mm Meso-bathypelagic CNRY, NASE, NWCS, NASW, SARC, NECS

G

H

2

I

7

9 3

8

6

Figure 150:  Pseudoamallothrix emarginata. A, ♀ lateral view; B, ♂ lateral view; C, Mx2; D, P2; E, ♂ P5; F, rostrum; G, ♀ P5. Amallothrix valida. H, rostrum; I, ♀ P5. A, C, D, F, and G, Park (1980); B, E, Bradford et al. (1983); H, I, Tanaka (1962).

crustacea: copepoda  321 Order Calanoida Superfamily Clausocalanoidea  Family Scolecitrichidae Genus: Scaphocalanus Sars, 1900 Total length: ♀ 0.9–5.6 mm; ♂ 1.2–5.3 mm Cephalosome: A1 not reaching beyond posterior end of metasome. ♀ A1 with transparent flange along posterior edge of segments 5-22 (1), ♂ A1 non-geniculate. Head rounded (2) or with crest (♀ Scaphocalanus affinis and ♀ Scaphocalanus magnus) (3). Two long slender rostral filaments (4). Mx2 Enp with 5 short brushlike and 3 long worm-like sensory filaments (5). ♂ mouthparts greatly reduced Metasome: 3 segs, cephalosome fused with 1st metasome seg, segs 4–5 fused. End of metasome rounded (6), or extended into a broad point (7) Legs: P2-P4 Enp and Exp ornamented with spinules on posterior surface. ♀ P5 uniramous with 2-3 segs, with a strong inner and terminal  spine (8) (P5 absent in Scaphocalanus curtus). ♂ P5

aysmmetrical, biramous and elongate: Exp 3 segs, Enp 1-2 segs. L-leg with elongate basis (9), last Exp seg with hairs, Enp longer than Exp (10). R-leg with short basis, Enp shorter than Exp, last Exp seg long and blade-like (11) Urosome: posterior border of segs fringed with fine teeth or spinules (12). ♀ 4 segs, GS only slightly swollen ventrally (13). ♂ 5 segs, 2nd seg swollen and twice as long as seg 3 (14). AS very short Notes: 12 species reported for study area. Specimens are often incomplete as appendages are fragile Similar taxa: Archescolecithrix auropecten Distribution and ecology: majority of species are mesobathypelagic, only S. curtus, S. echinatus and S. magnus are regularly found in the epipelagic. CNRY, NASE, NWCS, NASW, NADR, NECS, SARC, ARCT

Scaphocalanus echinatus

Scaphocalanus magnus

(Farran, 1905)

(T. Scott, 1894)

A

B

♀ 1.6–2.6mm ♂ 1.3–2.4 mm Epi-bathypelagic CNRY, NASE, NWCS, NASW, NADR, NECS

C

2

♂

♀

G

3

♀ 3.6–5.6mm ♂ 4.0–5.3 mm Epi-hadopelagic NWCS, NASW, SARC, ARCT, NADR, NASE

H D

1

6

F R

4

E

L

14 5

9

I 13

10

7

12

11 8 Figure 151:  Scaphocalanus echinatus. A, ♀ A1; B, ♀ lateral view; C, ♂ lateral view; D, rostral filaments; E, ♀ P5; F, ♂ P5. Scaphocalanus magnus. G, ♀ head crest lateral view; H, ♀ Mx2; I, ♀ last metasome seg and urosome lateral view. A, B, Farran (1905) C, E–I, Park (1982); D, Tanaka (1961).

322 ta xonom y Order Calanoida Superfamily Clausocalanoidea  Family Scolecitrichidae Genus: Scolecithricella Sars, 1902

biramous, elongate and longer than urosome: R-Enp reduced; L-leg longer than R-leg, L-Enp half as long as Exp

Total length: ♀ 1.1–2.2 mm; ♂ 1.0–2.3 mm Cephalosome: A1 reaching between posterior end of metasome and caudal rami. ♂ A1 non-geniculate. Head rounded, rostrum biramous and strong, tapering into sensory filaments (1). Mx2 Enp with 3 long worm-like (2) and 5 short brush-like sensory filaments Metasome: body oval, 3 segs, cephalosome fused with 1st metasome seg, segs 4-5 fused. End of metasome rounded; in lateral view square-like in Scolecithricella minor (3), notched in Scolecithricella dentata (4) Legs: P2-P3 Enp and Exp with spinules on posterior surface, P2 basis may have row of inner spinules (5). ♀ P5 uniramous with 1 seg, flat oval-shaped with an inner and terminal spine, usually with a minute spine on external margin (6). ♂ P5 asymmetrical,

Notes: seven species reported for study area. Many species in the Scolecitrichidae are very similar and have relatively recently changed genus name. Identification to genus level can rely on subtle features, making a correct identification difficult. See Vyshkvartzeva (2001) for a detailed review of the family Similar taxa: Archescolecithrix spp., Racovitzanus Pseudoamallothrix spp., and Amallothrix spp.

Distribution and ecology: epi-bathypelagic. CNRY, NASE, NWCS, NASW, NADR, GFST, NECS, SARC, ARCT (S. minor only)

B ♀

D

C

♂

♂

2

E

5

8

7

R L

4

I

J 1

K

6

H

(Giesbrecht, 1893 [‘1892’]) ♀ 1.2–2.1 mm ♂ 1.3–1.9 mm Epi-mesopelagic CNRY, NASE, NWCS, NASW, NADR, GFST, NECS, SARC

F 3

spp.,

Scolecithricella dentata

Scolecithricella minor (Brady, 1883) A ♀ 1.6–2.6 mm ♂ 1.3–2.4 mm Epi-bathypelagic CNRY, NASE, NWCS, NASW, NADR, GFST, NECS, SARC, ARCT

Urosome: very slender. ♀ 4 segs, spermathecal sac thin and slitlike, GS characteristically flat (7), without ventral swelling. ♂ 5 segs. AS very short (8)

G

Figure 152:  Scolecithricella minor. A, ♀ lateral view; B, ♂ lateral view; C, ♂ dorsal view; D, P2; E, Mx2; F, rostrum; G, ♀ P5; H, ♂ P5. Scolecithricella dentata. I, ♀ last metasome seg and urosome lateral view; J, rostrum; K, ♀ P5. A-C, Brodsky (1967); D-I, Park (1980).

crustacea: copepoda  323 Order Calanoida Superfamily Clausocalanoidea  Family Scolecitrichidae Genus: Scolecithrix Brady, 1883

Enp with 1 seg and club-shaped (5); Exp with 3 segs. R-leg, uniramous, Exp with 2 segs, last seg very short in Scolecithrix danae (6) and divided into two branches in S. bradyi (7)

Total length: ♀ 1.1–2.5 mm; ♂ 1.0–2.4 mm Cephalosome: A1 may not reach end of metasome. ♂ A1 nongeniculate. Head rounded, with biramous rostrum tapering into sensory filaments (1). Mx2 Enp with 5 short brush-like and 3 long worm-like sensory filaments

Urosome: ♀ 4 segs and short: S. bradyi GS swollen dorsally on left side (8), caudal rami appearing somewhat elongate; S. danae GS swollen with large genital operculum (9). ♂ 5 segs. AS short

Metasome: body robust and ovoid in dorsal view, 4 segs, cephalosome fused with 1st metasome seg. ♀ end of metasome elongated (2), asymmetrical and reaching past GS in Scolecithrix bradyi (3)

Notes: two species found in the study area. According to Vyshkvartzeva (2001) there are only two species in this genus. S. danae may have red/brown colouration to body

Legs: P2-P3 Enps and exps ornamented with spinules on posterior surface. ♀ P5 absent (rudimentary in S. bradyi). ♂ P5 aysmmetrical longer than urosome. L-leg with elongate basis (4) and biramous:

Distribution and ecology: warm waters; epi-mesopelagic. CNRY, NASE, NWCS, NASW, NADR, GFST

Scolecithrix danae A

B

(Lubbock, 1856) ♀ 1.8-2.5 mm ♂ 1.7-2.4 mm Epi-mesopelagic CNRY, NASE, NWCS, NASW, NADR, GFST

D

C ♀

♀

E

R L

♂

2 4 9

G

Scolecithrix bradyi

6

5

I

L

H

Giesbrecht, 1888 ♀ 1.1–1.6 mm ♂ 1.0–1.6 mm Epipelagic CNRY, NASE, NWCS, NASW, NADR, GFST

Similar species: ♀ Euchirella rostrata, other ♂ scolecitrichids

♂

♀

F

R

4 1

J

8

3

5

7

K

Figure 153:  Scolecithrix danae. A, ♀ lateral view; B, ♀ dorsal view; C, ♂ lateral view; D, ♂ P5; E, ♀ last metasome seg and urosome lateral view. Scolecithrix bradyi. F, rostral filaments; G, ♀ dorsal view; H, ♂ lateral view; I, ♂ P5. J, distribution map of S. danae; K, distribution map of S. bradyi. F, Tanaka (1962); D, G-I, Bradford et al. (1983); A-C, Rose (1942); E, Ramirez (1971).

324 ta xonom y Order Calanoida Superfamily Diaptomoidea  Family Acartiidae Genus: Acartia Dana, 1846 Total length: ♀ 0.6–2.1 mm; ♂ 0.7–1.5 mm Cephalosome: somewhat angular apex to head (1). A1 ’hairy’ in appearance (2), reaching from end of metasome to end of caudal rami. ♂ R-A1 geniculate (3). Rostrum as paired filaments (4), or absent. Mouthparts somewhat plumose Metasome: elongated. 4 segs. Posterior end of metasome usually rounded (extended into short spine-like projections in A. danae) and armed with small spines/hairs (5) Legs: thin and slender. P1-P4 with long plumose setae; P2-P4 Exp with spine-like projections (6) (i.e. unarticulated unlike other copepods). ♀ P5 small, uniramous, symmetrical, 3 segs; 2nd seg with outer plumose seta (7), last seg forming a spine (8). ♂ P5 uniramous, asymmetrical; basis with outer seta (9); R-leg with inner processes, last seg claw-like (10) 2

A

♀

Urosome: ♀ 3 segs. ♂ 5 segs, 4th seg very short (11). Posterior border of segs often spinulose. In lateral view penultimate seg overlapping dorsally with AS (11). ♀ GS has looped spermathecal tubular-sac (12). Caudal rami slightly asymmetrical with 6 setae arranged in a characteristic fan-shape. One backward pointing seta on each caudal ramus (13) Notes: twenty species of Acartia found in the study area. The Acartiidae would benefit from taxonomic re-examination, due to the apparent extent of intra-specific and inter-specific variability. Please see Bradford-Grieve (1999a) for a review of the family Similar taxa: Paracartia spp. ♂ Distribution and ecology: epi-mesopelagic, littoral/brackishoceanic. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS, SARC, ARCT

1

F

D

C

B

♂

6

13

H

4

I

G

J

7

12 5

3

E

R

L 9

11 8

10

Figure 154:  Acartia clausi. A, ♀ dorsal view; B, ♀ lateral view; C, ♂ dorsal view; D, ♂ A1; F, P3; G, ♀ GS lateral view; H, ♂ urosome lateral view; I, ♀ P5; J, ♂ P5. A. tonsa. E, rostral filaments. A, B, G–J, Bradford-Grieve (1999a); C, D, Avancini et al. (2006); E, Rémy (1927); F, Sars (1903).

Table 13:   Acartia species.

Total length

Acartia clausi

Acartia longiremis

Acartia negligens

Acartia danae

Acartia tonsa

Giesbrecht, 1889

(Lilljeborg, 1853)

Dana, 1849

Giesbrecht, 1889

Dana, 1849

♀ 0.6–1.5 mm, ♂ 0.7–1.3 mm ♂ LV

♀ LV

♀ 0.8–1.3 mm, ♂ 0.8–1.2 mm ♂ LV

♀ LV

♀ 0.9–2.1 mm, ♂ 0.8–1.5 mm ♂ LV

♀ LV

Last metasome seg and urosome 1 dorsal long needle-like spine

row of dorsal spines

(Bradford, 1976)

(Avancini et al., 2006)

Absent

Absent

Rostral filaments

♀ 0.9–1.3 mm, ♂ 0.7–1.1 mm ♀ LV

row of dorsal spines

(Tanaka, 1965)

(Bradford-Grievea, 1999; Steuer, 1923)

Present

(Avancini et al., 2006; Gerber, 2000)

Present

NB: only one leg shown

Present NB: only one leg shown

NB: only one leg shown

seg length > width seg length > width

♀ P5 seta length ~ spine length (Sars, 1903)

(Sars, 1903)

spine long, thin and strongly angled inwards

seta length >3 x spine length (Giesbrecht, 1893 [‘1892’])

L

L

R

R

L

seg length > width inner projection

seta approx. 3 x as long as spine

(Rémy, 1927)

(Giesbrecht, 1893 [‘1892’])

small and large inner spine R

♂ DV

♀ LV

♂ DV

1 strong spine-like projection

some small spines and fine dorsal hairs

seg length = width

♀ 0.8–1.5 mm, ♂ 0.8–1.3 mm

L

R

RL L

R

♂ P5

(Bradford, 1976)

Similar spp. Distribution

A. margalefi, A. tonsa and A. hudsonica (North American waters) Epi-mesopelagic. Neritic. CNRY, NASE, NECS, NWCS, SARC, NADR.

(Bradford, 1976)

(Steuer, 1923)

A. danae

Epi-mesopelagic. Brackish and neritic. NASE, NASW, NWCS, ARCT, SARC, NECS.

Epi-mesopelagic. Oceanic and neritic in warm waters. CNRY, NASE, NASW.

short spine large pointed inner projections

wide inner projections

(Steuer, 1923)

A. negligens

Epi-mesopelagic. Oceanic and neritic in warm waters. CNRY, NASE, NWCS, NASW, GFST, NADR.

(Gerber, 2000)

A. clausi

Brackish and neritic. CNRY, NASE, NWCS, NASW, NECS.

crustacea: copepoda  325

long spine small inner projections

DISTRIBUTION: Acartia longiremis

DISTRIBUTION: Acartia danae

DISTRIBUTION: Acartia negligens

DISTRIBUTION: Acartia tonsa

crustacea: copepoda  327 Order Calanoida Superfamily Diaptomoidea  Family Acartiidae Urosome: ♀ 3 segs and short. ♂ 5 segs, 4th seg very short. In lateral view penultimate seg overlapping dorsally with AS (10). ♀ GS expanded laterally (11), spermatophore attached to GS with thin plate-like structure (12). Caudal rami broad in ♀ with one of the terminal setae often thickened (13)

Genus: Paracartia T. Scott, 1894 Total length: ♀ 0.8–1.3 mm; ♂ 0.8–1.2 mm Cephalosome: somewhat angular apex to head (1). A1 ‘hairy’ in appearance, almost reaching to end of metasome. ♂ right A1 geniculate, may be swollen and may have large spine (2). Rostral filaments present

Notes: strong sexual dimorphism. Two species found in the study area. For a review of the family please see Bradford-Grieve (1999a) and Boxshall & Halsey (2004)

Metasome: elongated. 4 segs. ♀ last seg with wing-like extensions (3), rounded in ♂ (4)

Similar taxa: ♂ Acartia

Legs: thin and slender. P1-P4 with long plumose setae; P2-P4 Exp with spine-like projections (i.e. unarticulated as in other copepods), see Acartia for figures. ♀ P5 large, uniramous, symmetrical: 2 segs; 1st seg with short outer setae (5), 2nd seg forming a strong curved spine with spinules (6). ♂ P5 uniramous, asymmetrical; basis with inner projection (7) (large in P. grani); R-leg long, last seg slender and claw-like (8); L-leg short with 2 terminal spines resembling a chela (9) 1

Paracartia grani

C

Sars, 1904 ♀ 0.9–1.3 mm ♂ 1.0–1.1 mm Coastal, littoral NECS, SARC, CNRY, NASE, NADR

Distribution and ecology: littoral, brackish. NECS, SARC, CNRY, NASE, NADR

D

E

♂

♀ B

L

7

9

5

R

2

A 3

8

6

11

K

12

Paracartia latisetosa

♀ G

F

R

J

(Kriczagin, 1873) ♀ 0.8–1.2 mm ♂ 0.8–1.2 mm Coastal, littoral CNRY

L

I

H

12 13

10

4

Figure 155:  Paracartia grani. A, ♀ dorsal view; B, ♀ P5; C, ♂ dorsal view; D, ♂ R-A1; E, ♂ P5. Paracartia latisetosa. F, ♀ dorsal view; G, ♀ P5;

H, ♀ urosome dorsal view; I, ♀ urosome lateral view; J, ♂ dorsal view; K, ♂ P5. A–E, Sars (1919); F-H, K, Crisafi and Crescenti (1972); I, J, Giesbrecht (1893 [‘1892’]).

328 ta xonom y Order Calanoida Superfamily Diaptomoidea  Family Candaciidae Genus: Candacia Dana, 1846 Total length: ♀ 1.6–3.9 mm; ♂ 1.4–3.8 mm Cephalosome: front of head squared (1) with pronounced ‘shoulders’ (2). Proximal segs of A1 strong and toothed (3). ♂ R-A1 geniculate. Geniculation in the form of a dark-coloured, serrated comb (4). Large and powerful carnivorous mouthparts (5) Metasome: robust body, 4 segs. Last seg of the metasome typically with prominent posterolateral asymmetrical projections (6), reduced in some species (7) Legs: P1-P4 exps are finely serrated on outer edge (8). ♀ P5 small, uniramous, symmetrical, 3 segs; last seg longest and armed with spines (9), or finger-like processes (10). ♂ P5 L-leg 4 segs; R-leg 3 segs, last seg claw-like (11), or feather-like (12)

Notes: 15 species reported for the study area. Many species, even after fixation in formaldehyde, possess characteristic black/brown pigmentation to various parts of the body (♂ A1 geniculation, tips of limbs and metasome). The family Candaciidae was previously comprised of two genera, Candacia (Dana, 1846) and Paracandacia (Grice, 1963), which were separated by the morphology of the ♂ and ♀ P5. Boxshall and Halsey (2004) however, describe the family as monogeneric and place the species previously belonging to Paracandacia in with Candacia Similar taxa: some members of the Pontellidae Distribution and ecology: epi-bathypelagic. Genus with widespread distribution. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS, SARC, ARCT

Urosome: ♀ 3 segs. ♂ 5 segs. GS of both sexes usually asymmetrical bearing processes or swellings (13) 3 1

A

4

B

2

C

13

♂

♀

15

E

6

13

13

19

17

F

20

G

L

H

11

8 9

K 5

R

Candacia bispinosa

I

L R

J

(Claus, 1863)

D

♀ 1.6–2.2 mm ♂ 1.4–2.2 mm Epi-mesopelagic CNRY, NASE, NASW, NWCS, GFST

7 13

10

12

Figure 156:  Candacia armata. A, ♀ dorsal view; B, ♂ dorsal view; C, ♂ A1; D, Mx2; E, P3; F, ♀ urosome ventral view; G, ♀ P5; H, ♂ P5. Candacia bispinosa. I, ♀ urosome lateral view; J, ♀ P5; K, ♂ P5. A, B, F, G, Rose (1933); C, Wilson (1932); D, E, Sars (1903); H, Farran (1948a); I–K, Grice (1962).

Table 14:   Candacia species.

Total length

Candacia armata

Candacia bipinnata

Candacia simplex

Candacia ethiopica

Candacia pachydactyla

(Boeck, 1872)

Giesbrecht, 1889

(Giesbrecht, 1889)

Dana, 1849

Dana, 1849

♀ 2.0–2.9 mm, ♂ 1.7–2.7 mm

♀ 2.2–3.2 mm, ♂ 1.9–3.0 mm

♀ 1.7–2.2 mm, ♂ 1.6–2.3 mm

♀ 2.0–3.0 mm, ♂ 2.0–2.9 mm

♀ 2.2–2.9 mm, ♂ 2.1–2.7 mm

Body strongly pigmented VV

♀ urosome dorsal

lateral swellings 2nd seg ventral twisted swelling (Farran, 1948a)

(Bradford-Grieve, 1999b)

right metasome projection not reaching past GS

♂ urosome dorsal

♀ P5

(Bradford-Grieve, 1999b)

irregular subchela

L R

♂ P5

Distribution Epi-mesopelagic.

CNRY,NASE, NWCS, SARC, NECS, NADR

(Grice, 1962)

(Owre and Foyo, 1967)

C. armata, ♂ C. bispinosa Epi-mesopelagic. CNRY, NASE, NWCS, NASW

broad rounded process

2 small processes

no GS projection

(Bradford-Grieve, 1999b)

(Owre & Foyo, 1967) 7 spines, terminal spines heavily chitinised

7 spines, 3rd inner spine longest

6 spines, 2nd inner spine longest (Grice, 1962)

(Grice, 1962)

(Bradford-Grieve, 1999b) L R

L R

L R

chela

(Grice, 1962)

broad chela

long terminal spines

feather-like seta (Grice, 1962)

C. bispinosa, ♀ C. ethiopica Epi-mesopelagic. CNRY, NASE, NWCS, NASW, NADR

(Grice, 1962)

(Bradford-Grieve, 1999b)

♀ C. simplex, ♀ C. bispinosa

♂ C. armata, ♀ C.bispinosa ♀ C. ethiopica

Epi-mesopelagic. CNRY, NASE,NWCS, NASW

Epi-mesopelagic. CNRY, NASE, NWCS, NASW

crustacea: copepoda  329

(Farran, 1948a)

♂ C. longimana C. bipinnata, C. giesbrechti

L R

(Bradford-Grieve, 1999b)

small metasome projections

3 short outer spines (Bradford-Grieve, 1999b)

2 long thin lateral process

small lateral spines

fine hairs

(Bradford-Grieve, 1999b)

GS with tapering hook-like projection

3 short outer spines (Rose, 1933)

small metasome projections

swollen laterally and ventrally

right metasome projection thin, reaching past GS

GS with tapering projection (Rose, 1933)

Similar spp.

triangular lateral projections

2nd seg with ventral swelling

DISTRIBUTION: Candacia armata

DISTRIBUTION: Candacia bipinnata

DISTRIBUTION: Candacia simplex

DISTRIBUTION: Candacia ethiopica

DISTRIBUTION: Candacia pachydactyla

crustacea: copepoda  331 Order Calanoida Superfamily Diaptomoidea  Family Centropagidae Genus: Centropages Krøyer, 1849 Total length: ♀ 0.9–2.5 mm; ♂ 0.9–1.8 mm Cephalosome: head may be square-like (1), sometimes with pronounced ‘shoulders’ (2). A1 usually reaching to or beyond caudal rami; segs 1, 2 and 5 may be tooth-like (3). ♂ R-A1 geniculate (4), region of geniculation serrated (5), sometimes with proximal spines (6). Rostrum bifurcated. Eye spot well developed (7). Mx2 well developed and characteristically plumose (8) Metasome: 5 segs. End of metasome rounded or expanded into spine-like projections (9). May be slightly asymmetrical Legs: ♀ P5 biramous and symmetrical, resembling P2-P4, except Exp seg 2 with a large inner spine (10). ♂ P5 aysmmetrical and biramous: Enp 3 segs; L-Exp 2 segs; R-Exp 3 segs, forming a chela (11)

1

Urosome: ♀ usually 3 segs. ♂ typically 4 segs (5 in Centropages violaceus). ♀ GS swollen and without seminal receptacles, often asymmetrical possessing hairs or spines (12); 2nd seg may have ventral process. Caudal rami somewhat elongate and distally broad (13) Notes: nine species reported for study area. Some species may have diffuse/speckled colouration to metasome Similar taxa: Isias clavipes, Pseudodiaptomus spp. Distribution and ecology: epi-bathypelagic, mostly epipelagic. Widespread. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS, SARC, ARCT

D

3

A

B

2

7

3

6

3

8

4

C

6 5

E

4

F

L

R

10

12

11 13

9

Figure 157:  Centropages typicus. A, ♀ dorsal view; B, ♂ dorsal view; C, ♂ R-A1; D, Mx2; E, ♀ P5; F, ♂ P5. Boxshall and Halsey (2004); D, E, modified.

Centropages typicus

Centropages hamatus

Centropages chierchiae

Centropages bradyi

Centropages violaceus

Krøyer, 1849

(Lilljeborg, 1853)

Giesbrecht, 1889

Wheeler, 1901

(Claus, 1863)

♀ 1.3–2.5 mm, ♂ 1.5–2.4 mm

♀ 1.8–2.2 mm, ♂ 1.8–2.2 mm

♀ 1.5–2.0 mm, ♂ 1.0–1.9 mm

Total length

♀ 0.9–1.4 mm, ♂ 0.9–1.3 mm

1

1 1

♀

2

1

2

small projections

(Gerber, 2000)

♀ urosome

lateral swelling high on right side

curved and thickened

♀

♂  

DV

GS ventral hook

(Farran, 1948b; Lee, 1972) DV

LV

LV

lateral swelling and tufts of hair

ventral process lateral swelling low on right side

(Lee, 1972)

(Farran, 1948b; Bradford-Grieve, 1999b) LV

VV

ventral swellings caudal rami with a ‘peg’

(Brodsky, 1962) slender and sigmoid

slender and slightly angular

Chela of ♂ R- P5

5 urosome segs in ♂

Round Peg-like

large projections

(Gerber, 2000)

(Lee, 1972; Sars, 1903)

(Sars, 1903)

♀

A1 without teeth. No spine before geniculation in ♂

♂  

A1 without teeth (1); No spine before geniculation in ♂ (2)

large projections

LV

♀

A1 without teeth. No spine before geniculation in ♂

♂  

A1 with teeth (1); 1 spine before geniculation in ♂ (2)

DV

A1 with teeth (1); 2 spines before geniculation in ♂ (2) 2

1

♀

♂  

♀ and ♂ dorsal

♀ 1.5–2.1 mm, ♂ 1.5–1.9 mm

(Farran, 1948b; Mori, 1937) LV

spinules Right-side

VV

ventral swellings

(Bradford-Grieve, 1999b) slender, sigmoid and very elongated

angular, slender and denticulate at tip

(Farran, 1948b)

(Farran, 1948b)

(Giesbrecht 1893 [‘1892’])

Similar spp.

C. furcatus (NWCS, CNRY, NASE), C. chierchiae

C. kroyeri (CNRY, NASE), C. typicus, Pseudodiaptomus.

Distribution

Epipelagic. CNRY, NASE, NWCS, NADR, NECS, SARC.

Epipelagic. CNRY, NASE, NWCS, Epipelagic. CNRY, NASE, NASW, NADR, NECS. NADR, NECS, SARC, ARCT

C. brachiatus (CNRY, NASE), C. typicus

(Farran, 1948b)

(Farran, 1948b)

C. violaceus

C. gracilis (NASE, NASW), C. bradyi

Epi-bathypelagic. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS.

Epipelagic. CNRY, NASE, NASW, GFST, NWCS, NADR, NECS.

332 ta xonom y

Table 15:   Centropages species.

DISTRIBUTION: Centropages typicus

DISTRIBUTION: Centropages hamatus

DISTRIBUTION: Centropages chierchiae

DISTRIBUTION: Centropages bradyi

DISTRIBUTION: Centropages violaceus

334 ta xonom y Order Calanoida Superfamily Diaptomoidea  Family Centropagidae Genus: Isias Boeck, 1864

R-Enp absent; Exp 2 segs; last seg broad with 4 spines, right distal spine large (8)

Isias clavipes (Boeck, 1864) Total length: ♀ 1.3–1.7 mm; ♂ 1.3–1.4 mm Cephalosome: A1 reaching to end of metasome. ♂ R-A1 geniculate (1), region of geniculation serrated (2), with a proximal spine (3). Rostrum bifurcated. Mx2 with plumose setae, not as strongly developed as in Centropages

Urosome: ♀ 3 segs. ♂ 5 segs. ♀ GS swollen and without seminal receptacles, asymmetrical with 2 ventral hooks (9) and dorsal swelling on left (10). ♂ 3rd seg with projection on right side (11). Caudal rami somewhat elongated, with oblique outer edge (12) Notes: one species reported for the study area

Metasome: body oval, 4 segs, 4th and 5th segs fused. End of metasome rounded

Similar taxa: Centropages hamatus, Parapontella brevicornis, Temora longicornis

Legs: ♀ P5 biramous and symmetrical, resembling P2-P4; basis with large outer distal spines (4), Exp seg 2 with a large inner spine (5). ♂ P5 aysmmetrical: basis with inner spine (6); L-Enp reduced (7),

Distribution and ecology: coastal. CNRY, NASE, NADR, NECS, NWCS

A

D

B

E

L

1

♀

3

♂

R

4

2

C 6

1

F

G 8

9

10

7

5

H

11

10

12

Figure 158:  Isias clavipes. A, ♀ dorsal view; B, ♂ dorsal view; C, mid-section of ♂ A1; D ♀ P5; E ♂ P5; F ♀ GS lateral view; G, ♀ urosome dorsal view; H, distribution map of I. clavipes. A, B, D-F, Rose (1933); C and G, Sars (1903).

crustacea: copepoda  335 Order Calanoida Superfamily Diaptomoidea  Family Parapontellidae Genus: Parapontella Brady, 1878 Parapontella brevicornis (Lubbock, 1857) Total length: ♀ 1.3–1.6 mm; ♂ 1.3–1.6 mm Cephalosome: head rounded, with a small ventral eye and a large dorsal eye (1), but without cuticular lenses. A1 reaching mid-point of metasome. R-A1 geniculate in ♂, region of geniculation serrated and with spines (2), first 3 segs and mid segs swollen, last seg ending in a strong spine (3). Rostrum with two fine filaments (4). Mx2 short, strong and curved with short crossed setae (5) Metasome: body robust; 4 segs, 4th and 5th segs fused. End of metasome rounded in ♀ (6) and asymmetrical and pointed in ♂ (7)

A

B

Legs: ♀ P5 large, biramous: Exp with 1 seg, longer than Enp and terminating is a spine (8); Enp with 1 seg, bifurcated at tip (9). ♂ P5 aysmmetrical, uniramous, broad and with 3 segs: R-leg with inner process on 1st seg (10), last seg tapering into a spine (11) Urosome: ♀ 3 segs. ♂ 5 segs. ♀ GS ventrally swollen (12), 2nd seg with 2 dorsal spines on rear margin (13); ♂ with small blunt lateral projections on 3rd and 4th segs (14). Caudal rami somewhat elongated with oblique outer edge (15) Notes: Parapontella is a monospecific genus Similar taxa: Isias clavipes, ♂ Centropages spp. Distribution and ecology: neritic; hyperbenthic. NECS, NASE

D

1

♀

♀

E

C

♂

4

3

F

7

12 13

14 15

G

H

2 L

5

R

I 10 9 11 8

Figure 159:  Parapontella brevicornis. Figures: A, ♀ lateral B, ♀ dorsal view; C, ♂ dorsal view; D, ♂ A1; E, rostrum; F, ♀ Mx2; G, ♀ P5; H, ♂ P5; I, distribution map of Parapontella brevicornis. A–C, E–H, Sars (1903); D, Giesbrecht (1893 [‘1892’]).

336 ta xonom y Order Calanoida, Superfamily Diaptomoidea  Family Pontellidae Genus: Anomalocera Templeton, 1837

L-leg 4 segs, last segment short with up to 4 spines (10); R-leg 4 segs with a chela (11)

Total length: 2.1–4.8 mm; ♂ 2.5–4.3 mm Cephalosome: triangular head with lateral hooks (1). Two pairs of dorsal eye-lenses (2) and a ventral eye, large in ♂ (3). A1 approx. reaching 3rd metasome seg in ♀ and almost end of metasome in ♂. R-A1 geniculate in ♂, region of geniculation serrated; mid segs robust and swollen (4). Paired rostral processes. Mx2 well developed and plumose (5) (similar to Centropages)

Urosome: asymmetrical: ♀ 3 segs. ♂ 5 segs. GS with species specific asymmetric processes (12). Caudal rami asymmetrical in ♀ (13), elongated in male (14)

Metasome: body robust; 5 segs. End of metasome expanded into strong points (6), strongly asymmetrical in ♂ (7)

Similar taxa: Labidocera spp., Pontella spp., Pontellopsis spp., Pontellina spp., ♂ Candacia spp.

Legs: ♀ P5 large, biramous: Exp with 2 segs, 1st seg elongate (8); Enp with 1 seg, short and bifurcate (9). ♂ P5 aysmmetrical, uniramous,

Distribution and ecology: neustonic. Common in coastal waters. CNRY, NECS, NASE, NWCS, ARCT, SARC

Anomalocera patersoni

A

Templeton, 1837 ♀ 3.0-4.1 mm ♂ 2.5-4.0 mm Epipelagic CNRY, NECS, NASE, NWCS, ARCT, SARC

C

♀

1

E

D

B

Notes: three species in the genus, all found in the study area (A. patersoni, A. opalus and A. ornata). May be colourful with blue and red pigments, even in preserved specimens

F

G L

H

R

2

♂

4 4

–1

13

9 17

3

8

13

11

6

M

10

7

2

14

♂

Anomalocera ornata Sutcliffe, 1949

Anomalocera opalus Pennel, 1976 ♀ 2.1–4.0 mm (prosome) ♂ 2.7–3.4 mm (prosome) NWCS 12

I

J

♀ 4.0–4.8 mm ♂ 4.0–4.3 mm Neritic NWCS

K

5

L

12

12

Figure 160:  Anomalocera patersoni. A, ♀ dorsal view; B, ♂ dorsal view; C, ♂ head lateral view; D, ♂ A1; E, ♀ P5; F, ♂ P5; G, ♀ urosome ventral view; H, ♂ urosome dorsal view. Anomalocera opalus. I, ♀ urosome ventral view; J, ♂ urosome dorsal view. Anomalocera ornata. K, Mx2; L, ♀ urosome dorsal view; M, ♂ dorsal view. A–C, E, and F, Sars (1903); G-J, Pennell (1976); D, Wilson (1932); K-M, Wilson (1950).

crustacea: copepoda  337 Order Calanoida Superfamily Diaptomoidea  Family Pontellidae ♂ P5 aysmmetrical, uniramous; 4 segs, last 2 segs forming a claw in  R-leg (6). Claw of C. elliptica with 3 strong blunt and 3 short spines (7). L-leg terminating in a spine (8) and with an inner projection on 2nd seg in C. americana (9)

Genus: Calanopia Dana, 1853 Total length: ♀ 1.3–2.1 mm; ♂ 1.2–1.9 mm Cephalosome: head rounded without lateral hooks in North Atlantic species (may have lateral hooks in other regions) and without eye lenses. A1 reaching to end of metasome. ♂ R-A1 geniculate (1). Rostrum bifurcate

Urosome: ♀ 2 segs. ♂ 5 segs. ♂ C. elliptica with tooth on right 2nd seg (10). ♀ GS and AS elongated. ♀ GS not swollen ventrally in C.  americana (11). Caudal rami somewhat elongate with long plumose setae

Metasome: body oval, 4 segs, 4th and 5th segs fused. End of metasome produced into pointed projections (2), long in Calanopia elliptica (3)

Notes: two species found in the study area Distribution and ecology: neritic; epi-mesopelagic. West Atlantic. NWCS, NASW

Legs: ♀ P5 uniramous, asymmetrical in C. elliptica (4); 3 segs in C. americana, 4 segs in C. elliptica; last seg with terminal spines (5).

Calanopia elliptica (Dana, 1849) ♀ 1.6–2.1 mm

E

A

♀

Epi-mesopelagic, neritic NASW (Bermuda)

L

♂

C

♂ 1.4–1.9 mm

F

R

1

B

D 4

7 5 10

3

G

Calanopia americana

6 8

I K

♀ J

L

L

R 9

Dahl, 1894 ♀ 1.3–2.1 mm ♂ 1.2–1.4 mm Epipelagic, neritic NWCS, NASW

H 6 2 5 11

8

Figure 161:  Calanopia elliptica. A, ♀ dorsal view; B, ♀ GS lateral view; C, ♂ dorsal view; D, ♂ urosome dorsal view; E, ♀ P5; F, ♂ P5.

Calanopia americana. G, ♀ dorsal view; H, ♀ GS lateral view; I, ♂ R-A1 grasping region; J, ♂ urosome dorsal view; K, ♀ P5; L, ♂ P5. A, C Mori (1937); B, D–F, Bradford-Grieve (1999b); G, I, Owre and Foyo (1967); H, J–L Scott (1909).

338 ta xonom y Order Calanoida, Superfamily Diaptomoidea  Family Pontellidae Genus: Labidocera Lubbock, 1853

with 4 segs, last seg terminating in spines (12), may have rudimentary Enp (13)

Total length: ♀ 1.8–4.3 mm; ♂ 1.8–4.2 mm Cephalosome: head rounded or triangular, with (1) or without lateral hooks. Forehead with crest in L. acutifrons (2). One pair of dorsal eye-lenses (3) and a ventral eye (4). A1 reaching from end of metasome to mid urosome. ♂ R-A1 geniculate, usually with various processes (5), mid segs swollen, region of geniculation comb-like (6) . Rostrum deeply bifurcated with two fine filaments. Mx2 well developed and plumose (7) Metasome: body robust, 4-5 segs, 4th and 5th segs may be fused. End of metasome expanded into strong (8) or small points Legs: ♀ P5 large, biramous, may be asymmetrical: Exp with 1 seg, longer than Enp, and may have terminal processes (9); Enp with 1  seg, usually short (10). ♂ P5 asymmetrical: R-leg larger than L-leg, uniramous, with 4 segs ending in a large chela (11); L-leg Exp

A

Notes: eight species found in the study area. Several of these species have been assigned to separate species groups by Fleminger et al. (1982). May be colourful with blue, greenish and red pigments even in preserved specimens. See Fleminger (1979) for a key to west Atlantic species Similar taxa: Anomalocera spp., Pontella spp., Pontellopsis spp., Pontellina spp. Distribution and ecology: neustonic, epipelagic. Common in coastal waters. CNRY, NASE, NECS, NWCS

5

C

B 1

Urosome: ♀ 2-3 segs. ♂ 4-5 segs and symmetrical. ♀ GS typically asymmetrical with projections (14). ♀ caudal rami may be asymmetrical

6

♂

♀

2

3

D

♀

F

3

E

7

4 8

14

I H

L

R

G 11

12

10

13

9

Figure 162:  Labidocera wollastoni. A, ♀ dorsal view; B, ♂ lateral view; C, ♂ A1; D, ♀ head lateral view; G ♀ P5; H, ♂ P5. Labidocera acutifrons.

E, ♀ head dorsal view; F, ♀ Mx2; I, ♀ P5. A–D, G, H, Sars (1903); E, I, Wilson (1932); F, Jeong et al. (2009).

Table 16:   Labidcoera species.

Total length

Labidocera wollastoni

Labidocera acutifrons

Labidocera aestiva

Labidocera nerii

Labidocera detruncata

(Lubbock, 1857)

(Dana, 1849)

Wheeler, 1901

(Krøyer, 1849)

(Dana, 1849)

♀ 2.2–2.4 mm, ♂ 2.0–2.6 mm

♀ 3.2-4.3 mm, ♂ 3.3-4.2 mm

♀ 1.8-3.0 mm, ♂ 1.8-2.5 mm

rounded crest

♂

♀  ♀ and   dorsal view

LV

♀

♂

♀ 2.3–2.8 mm, ♂ 2.2–2.7 mm

♀

♀

♂

lateral spine lateral projection

large lateral projection

large lateral projection

(Sars, 1903) strongly hooked process

Region of geniculation on ♂ right A1 denotes geniculation point

hooked process

(Sars, 1903) large conical Enp

♀ P5

(Sars, 1903) LV

L

asymmetrical, R-leg longer

(Bradford-Grieve, 1999b)

R

pointed inner process

L

R

Enp with corrugated knob

L

(Sars, 1903)

Similar spp.

Anomalocera patersoni, Pontella spp.

Distribution

Epipelagic, coastal. CNRY, NASE, NECS, NWCS (rare)

(Jeong et al, 2009)

♂ L. aestiva Epipelagic, coastal. CNRY, NASE, NWCS

(Gerber, 2000)

asymmetric, L-leg longest outer and apical processes (Wilson, 1932)

symmetrical without hairs or projections

(Wilson, 1932)

rounded inner process bifid Enp

Enp small

GS symmetrical with lateral hairs

(Bradford-Grieve, 1999b) twisted Enp filament

L

apical processes

finger-like process

(Giesbrecht, 1893[‘1892’]) L R

R

rounded inner process

♂ L. acutifrons Epipelagic, coastal and brackish. CNRY, NWCS

crest

(Giesbrecht, 1893[‘1892’]) R

(Gerber, 2000)

(Jeong et al, 2009)

finger-like inner process

no Enp

(Jeong et al, 2009) conical Enp outer and apical processes

( Jeong et al, 2009)

uneven surface on GS extended operculum (Bradford-Grieve, 1999b) L R 4 processes no Enp

(Giesbrecht, 1893[‘1892’])

♂ L. detruncata Epipelagic, oceanic. CNRY, NWCS

no inner process (Jeong et al, 2009)

♂ L. nerii Epipelagic. NWCS

crustacea: copepoda  339

L

♂ P5

without process

large conical Enp

apical processes

extended operculum

(Giesbrecht, 1893[‘1892’])

(Wilson, 1932)

Enp slender

large dorsal swelling

(Sars, 1903)

asymmetrical

long denticulate process

(Jeong et al, 2009) R

CR oval and asymmetrical

CR symmetrical

(Wilson, 1932)

(Wilson, 1932)

small process

without large lateral projection

CR symmetrical and elongate

CR asymmetrical

♀ urosome dorsal view

♂

♀

♀ 2.8–3.7 mm, ♂ 2.5–3.6 mm

340 ta xonom y Order Calanoida Superfamily Diaptomoidea  Family Pontellidae Genus: Pontella Dana, 1846 Total length: ♀ 2.4–7.0; ♂ 2.4–6.0 mm Cephalosome: head triangular with lateral hooks (1). One pair of dorsal eye-lenses (2), a ventral eye (3) and usually a rostral lens (may be large in ♂) (4). A1 reaching just below the middle of the metasome. ♂ R-A1 geniculate, region of geniculation serrated, with only two segs posterior to geniculation (5); mid segs swollen (6). Mx2 well developed and plumose Metasome: body robust; 5 segs. End of metasome expanded into strong (7) or small points (8), often asymmetrical and sexually dimorphic Legs: ♀ P5 large, biramous, may have slight asymmetry: Exp with 1 seg, longer than Enp, usually with external and terminal spines (9);

A

2

Enp with 1 seg and bifurcate (10). ♂ P5 asymmetrical, uniramous: R-leg 4 segs ending in a large chela (11); L-leg 4 segs, last seg irregular terminating in spines (12) or a lamella Urosome: ♀ 2-3 segs. ♂ 4-5 segs. GS and caudal rami asymmetrical in ♀ (13), GS may be asymmetrical in ♂ Notes: seven species found in the study area. May be colourful: may have red, brown, blue, green, black pigment in rostrum, eyes, and along mid-line of body. Different authors report slight variations in morphology within a species for the shape of the ♀ GS and the number of spines on the ♀ P5 Similar taxa: Anomalocera spp., Labidocera spp. and Pontellopsis spp. Distribution and ecology: epipelagic, neustonic. CNRY, NASE, NECS, NWCS

C

4

D

1

B

♀

♂

3

5 6

F 8

R

L

E 7

13

10

12

11

9 Figure 163:  Pontella securifer. A, ♀ dorsal view; B, ♀ head lateral view; C, ♂ dorsal view; D, ♂ R-A1; E, ♀ P5; F, ♂ P5. A, B, and E, Giesbrecht (1893 [‘1892’]); C, D, Wilson (1950); F, Wilson (1932).

Table 17:   Pontella species.

Total length

Pontella lobiancoi

Pontella securifer

Pontella atlantica

Pontella spinipes

Pontella mediterranea

(Canu, 1888)

Brady, 1883

(Milne Edwards, 1840)

Giesbrecht, 1889

(Claus, 1863)

♀ 4.0–7.0 mm, ♂ 3.3–6.0 mm

♀ 3.6–4.6 mm, ♂ 3.2–4.6 mm

♀ 3.2–4.6 mm, ♂ 4.3–5.7 mm

♀ 4.5–4.8 mm, ♂ 3.1–4.4 mm

♀ 2.8–3.3 mm, ♂ 2.4–2.9 mm

LV

♀ 

  ♀ and ♂ dorsal view

♀

♂

♀

♂

♀

♂

strongly asymmetrical

strongly asymmetrical

strongly asymmetrical

(Giesbrecht, 1893['1892'])

(Giesbrecht, 1893['1892']; Wilson, 1950)) (Giesbrecht, 1893['1892']; Wilson, 1950)) (Wilson, 1932; Sewell 1912) Please note: only one leg shown

♀ P5

outer and terminal spines

strongly curved and acuminate

(Giesbrecht, 1893 ['1892'])

  ♀ urosome

VV

(Giesbrecht, 1893 ['1892'])

3 segs

L

L

opposing finger

(Giesbrecht, 1893 ['1892'])

(Wilson, 1932)

VV

1 angular process opposing finger

2 processes

long caudal rami

(Giesbrecht, 1893 ['1892'])

R

(Giesbrecht, 1893 ['1892'])

opposing finger

inner processes on thumb

R

2 segs symmetrical

VV

lumpy dorsal appearance

(Giesbrecht, 1893['1892']) L

(Giesbrecht, 1893 ['1892']) LV

2 segs

dorsal and lateral processes

no opposing finger, but corrugated process

R

DV

L

(Seymour-Sewell, 1912)

(Giesbrecht, 1893 ['1892']) no opposing finger

blunt ended thumb

R

L

(Giesbrecht 1893 ['1892'])

Similar spp.

P. mediterranea

P. spinipes, P. atlantica

P. securifer

P. securifer

P. lobiancoi

Distribution

Epipelagic. NECS

Epipelagic. CNRY, NASE, NWCS

Epipelagic. NASE, NWCS

Epipelagic, neritic. CNRY, NASE, NWCS

Epipelagic. CNRY, NASE

crustacea: copepoda  341

R

inner, outer and terminal spines

(Wilson, 1932)

3 segs

(Giesbrecht, 1893 ['1892'])

Please note: only one leg shown

strongly curved and acuminate

(Giesbrecht, 1893 ['1892']) VV

(Giesbrecht, 1893['1892'])

Please note: only one leg shown

endopod ½ length of exopod

strongly asymmetrical

(Giesbrecht, 1893 ['1892']) 1 rounded process

DV

2 segs dorsal processes

VV

lumpy appearance

♂ P5

♂

♂

342 ta xonom y Order Calanoida Superfamily Diaptomoidea  Family Pontellidae Genus: Pontellina Dana, 1852 Pontellina plumata (Dana, 1849) Total length: ♀ 1.0–1.9 mm; ♂ 1.3–1.9 mm Cephalosome: head rounded with strong rostrum visible in dorsal view (1), without lateral hooks (unlike in Pontella). ♀ without eye lenses, ♂ with one pair of dorsal eye-lenses (2) and a small ventral eye. A1 reaching to caudal rami. ♂ R-A1 geniculate (3), region of geniculation serrated, with only 2 segs posterior to geniculation; mid segs swollen (4). Rostrum with filaments (5). Cephalic appendages with long plumose setae (6) Metasome: broad and oval; 4 segs, 4th and 5th segs fused. End of metasome expanded into strong points in ♀ (7), rounded or with a small point in ♂ (8)

Legs: ♀ P5 usually symmetrical, biramous with 1 seg: Exp 3 x length of Enp, with 3 long terminal setae (9); Enp usually bifurcate (10). ♂ P5 asymmetrical, uniramous, basis with long plumose seta (11): R-leg 4 segs, last two segs forming a large chela (12); L-leg 4 segs, last seg with 4 spines (13) Urosome: ♀ 2 segs. ♂ 5 segs. ♀ GS with 4 groups of lateral spinules (14); AS fused with right caudal ramus (15) Notes: one species found in the study area. Body with dark violet colouration (Wilson, 1932) Distribution and ecology: neritic and oceanic. Epi-mesopelagic. CNRY, NASE, NWCS, NASW, GFST, NECS

1

D

A

♂ ♀

4

5

6

B

3

2

C

♂ 7

14

6

15

E

F

L

8 R

G 11

10 11 9

12 13

Figure 164:  Pontellina plumata. A, ♀ dorsal view; B, ♀ head lateral view; C, ♂ dorsal view; D, ♂ lateral view; E, ♀ P5; F, ♂ P5; G, distribution map for Pontellina plumata. A, D, Bradford-Grieve (1999b); C, E, F, Owre and Foyo (1967); B, Rose (1933).

crustacea: copepoda  343 Order Calanoida Superfamily Diaptomoidea  Family Pontellidae Enp with 1 seg and bifurcate (9). ♂ P5 asymmetrical; uniramous, R-leg 4 segs ending in a large chela (10); L-leg 4 segs, last seg terminating in spines and hairs (11)

Genus: Pontellopsis Brady, 1883 Total length: ♀ 2.0–4.5 mm; ♂ 1.6–3.6 mm Cephalosome: head rounded without lateral hooks and without dorsal and rostral lenses. Ventral eye present (1). A1 not reaching end of metasome. ♂ R-A1 geniculate, region of geniculation serrated (2), with only two segs posterior to geniculation (3); mid segs swollen (4). Rostrum bifurcate with filaments (5). Mouthparts with reduced plumose setation (6), unlike Labidocera

Urosome: ♀ 1–2 segs. ♂ 5 segs. Asymmetrical, usually with lateral projections (12) Notes: five species found in the study area. May be colourful, even in preserved specimens. Within a species, some authors report variability in shape of the ♀ GS Similar taxa: Labidocera spp.

Metasome: body robust, 4 segs; 4th and 5th segs fused. End of metasome expanded into pointed projections, usually strongly asymmetrical in ♂ (7)

Distribution and ecology: epi-mesopelagic, neritic to oceanic. CNRY, NASE, NWCS, NASW

Legs: ♀ P5 large, biramous, may have slight asymmetry: Exp with 1 seg, longer than Enp, usually with external and terminal spines (8);

A

B

C

♀

D

E

♂ 5

1

4 7

12

H G

F

6

12

L

2 R

3

9 11

8

10

Figure 165:  Pontellopsis regalis. A, ♀ dorsal view; B, ♂ lateral view; C, ♂ dorsal view; D, ♂ R-A1; E, ♀ rostrum; F, ♀ Mx2; G, ♀ P5; H, ♂ P5. A, C, E, G, and H, Zheng et al. (1982); B, Giesbrecht (1893 [‘1892’]); D, Wilson (1932); F, Park (1968).

Total length

Pontellopsis brevis

Pontellopsis villosa

Pontellopsis regalis

Pontellopsis perspicax

(Giesbrecht, 1889)

Brady, 1883

(Dana, 1849)

(Dana, 1849)

♀ 2.0–2.5 mm, ♂ 1.8–2.2 mm

♂ urosome

♀ 2.0–3.0 mm, ♂ 2.1–2.6 mm

♀

♀ 2.7–4.5 mm, ♂ 1.6–3.6 mm

♂

♀

♂

Pontellopsis strenua (Dana, 1849)

♀ 2.6–4.3 mm, ♂ 1.8–2.6 mm

♀

♀ 2.7-2.8 mm, ♂ 2.4–2.9 mm

♂

♂ ♀

♀ and ♂ dorsal view

asymmetrical

(Giesbrecht, 1893 ['1892'])

(Giesbrecht, 1893 [‘1892’])

2 processes on right

(Giesbrecht, 1893 ['1892'])

♂ P5

endopod very short, < half length of exopod

(Giesbrecht, 1893 ['1892'])

L

R

opposing finger-like process absent

R

L

L

process on left

extended anal operculum

(Giesbrecht, 1893['1892'])

(Giesbrecht, 1893['1892'])

(Giesbrecht, 1893 ['1892']) opposing finger short

almost symmetrical rounded lateral processes

variable lateral swellings

lobe on left

2 processes on right

(Giesbrecht, 1893['1892'])

(Mori, 1937)

(Zheng et al., 1982)

ventral view

♀ urosome dorsal view

(Giesbrecht, 1893 ['1892']; Chen and Zhang, 1965) Please note: only one leg shown

inner spine approx. mid-length along exopod

inner spine approx. mid-length along exopod

(Giesbrecht, 1893 ['1892'])

strongly asymmetric

(Owre and Foyo, 1967)

(Zheng et al., 1982)

Please note: only one leg shown

Please note: only one leg shown

♀ P5

strongly asymmetrical

strongly asymmetrical

almost symmetrical

opposing finger

R

long, thin, opposing finger

R

L

(Giesbrecht, 1893['1892']) opposing finger

R

L

medial process

(Giesbrecht, 1893 ['1892'])

(Giesbrecht, 1893 ['1892'])

(Giesbrecht, 1893['1892'])

(Zheng et al., 1982)

(Giesbrecht, 1893['1892'])

Similar spp.

P. villosa

P. brevis

P. strenua

P. regalis

P. regalis

Distribution

Epipelagic. CNRY.

Epipelagic. Oceanic and neritic. CNRY, NASE, NWCS.

Epipelagic. Oceanic and neritic. CNRY, NASE, NWCS.

Epipelagic. Oceanic and neritic. CNRY, NWCS.

Epi-mesopelagic. NASW.

344 ta xonom y

Table 18:   Pontellopsis species.

crustacea: copepoda  345 Order Calanoida Superfamily Diaptomoidea  Family Pseudodiaptomidae Genus: Pseudodiaptomus Herrick, 1884 Total length: ♀ 1.1–1.5 mm; ♂ 0.9–1.2 mm Cephalosome: A1 reaching past end of metasome. ♂ R-A1 geniculate (1). Rostrum with paired filaments. Eye spot may be well developed Metasome: slender, 3–5 segs. Posterior borders of segs may be ornamented with spinules (e.g. Pseudodiaptomus pelagicus (2)). End of metasome rounded or expanded into spine-like projections (e.g. Pseudodiaptomus marinus (3)) Legs: P2-P4 Exp seg 3 with 2 outer spines (4) (cf. 3 outer spines in Centropagidae). ♀ P5 long and uniramous: 5 segs, last seg plumose and spine-like (5) with a small inner process (6). ♂ P5 aysmmetrical, usually biramous and may be spinulose: R-leg with projection on

A

♀ 1.1–1.5 mm ♂ 0.9–1.2 mm Brackish-neritic NECS

Urosome: ♀ usually 3-4 segs, ♂ 5 segs. ♀ GS often swollen with asymmetrical processes and hairs (10); posterior borders of segs denticulated (11). Caudal rami somewhat elongate (12) Notes: two species found in the study area. P. marinus, native to the Pacific, has recently been introduced to the North Sea and is believed to have been introduced to Atlantic waters via ballast water (Brylinski et al., 2012) Similar taxa: Centropages spp. and Isias clavipes Distribution and ecology: shallow coastal and brackish waters, demersal. NECS, NWCS Pseudodiaptomus pelagicus

Pseudodiaptomus marinus Sato, 1913

coxa (7); Exp with 3 segs, last seg usually curved (8), Enp with 1 seg; L-leg Exp with 2 segs, last Exp seg with apical spines (9); Enp with 1 seg

B ♀

Herrick, 1884

C

I

♀ 1.2–1.4 mm ♂ 0.9–1.0 mm

♀ 10 11

K

J

♀

Brackish-neritic

2

NWCS

G 6 10

3

D

12

L

E

♂

F

♂

5

M

R

L

H

7

1

4

9

8

Figure 166:  Pseudodiaptomus marinus. A, ♀ dorsal view; B, ♀ lateral view; C, ♀ GS lateral view; D, ♂ R-A1; E, ♂ dorsal view; F, ♀ P3; G, ♀ P5; H, ♂ P5. Pseudodiaptomus pelagicus. I, ♀ dorsal view; J, ♀ metasome and urosome lateral view; K, ♀ P5; L, ♂ dorsal view; M, ♂ P5. A–D, F, Grindley and Grice (1969); E, G, Brodsky (1967); H–M, Walter (1989).

346 ta xonom y Order Calanoida Superfamily Diaptomoidea  Family Temoridae Genus: Temora Baird, 1850 and Eurytemora Giesbrecht, 1881 Total length: Temora spp. ♀ 0.8–2.1 mm; ♂ 0.8–1.9 mm; Eurytemora spp. ♀ 0.7–2.2 mm; ♂ 0.8–1.9 mm Cephalosome: widest part of body in Temora (1). A1 reaching past end of metasome in Temora (2) and may not reach end of metasome in Eurytemora (3). ♂ R-A1 geniculate (4). Rostrum with two filaments (5) Metasome: body somewhat coffin-shaped in Temora with 4 segs (4th and 5th segs fused) and 5 segs in Eurytemora. End of metasome rounded (6) or expanded into wing or spine-like projections (7)

uniramous; L-leg with 4 segs forming a chela in Temora (10) and somewhat claw-like in Eurytemora. R-leg with 3 segs Urosome: ♀ 3 segs. ♂ 5 segs. Caudal rami narrow and long (11) Notes: three species of Temora and five species of Eurytemora reported for the study area Similar taxa: Lucicutia spp. Distribution and ecology: in general Temora are epipelagic, neritic and Eurytemora freshwater, brackish. CNRY, NASE, NECS, NWCS, SARC, ARCT

Legs: ♀ P5 uniramous 3 segs; last seg with terminal spines (8), 2nd seg with inner projection in Eurytemora (9). ♂ P5 aysmmetrical,

B

A

C

♀

D 4

♂

♀

5

1

H 6 11

F

E

8

♀

3

2

10

G

7

9

Figure 167:  Temora longicornis. A, ♀ dorsal view; B, ♀ lateral view; C ♂ dorsal view; D, ♀ rostrum; E, ♀ P5; F, ♂ P5. Eurytemora affinis. G,

♀ P5; H, ♀ dorsal view. A–C, E–H, Gerber (2000); D, Sars (1903).

Table 19:   Temoridae species.

Total length

♀ and ♂ dorsal view

Temora longicornis

Temora stylifera

Temora turbinata

Eurytemora affinis

Eurytemora americana

(Müller, 1792)

Dana, 1849

(Dana, 1849)

(Poppe, 1880)

Williams, 1906

♀ 0.8–1.7 mm, ♂ 0.8–1.7 mm

♂

♀

♀

♀ and ♂ with projections

♀ AS longer than preceding seg

♀ 0.9–.7 mm, ♂ 0.9–1.6 mm

1

♀ 1.2–1.73 mm, ♂ 1.1–1.7 mm

long projections

1

( Gerber, 2000)

( Gerber, 2000) Large inner process (1) 2 long terminal spines (2)

2 1

(Mod. Rose, 1933)

(Gerber, 2000) Apical spine (1) and seta (2) of unequal length

♂ P5

Two apical spines of same length (1)

Last seg very broad (1)

R

R

L 1

1

2

(Rose, 1933)

T. turbinata, Lucicutia spp.

Distribution Epipelagic. CNRY, NASE, NECS, NWCS, SARC, ARCT

1

(Gerber, 2000)

(Gerber, 2000)

Left leg last seg with 3 processes (1)

Left leg last seg with two rounded spiny lobes (1)

1

L

L

R

2

R

L

1

1

1

1

(Rose, 1933)

(Bradford-Grieve, 1999b)

(Gerber, 2000)

(Rose, 1933)

Eurytemora spp.

T. longicornis

Eurytemora herdmani, Eurytemora velox, E. americana

E. affinis

Epipelagic. CNRY, NECS, NASE, NWCS, NASW

Epipelagic. CNRY, NASE, NWCS, NASW

Freshwater, brackish. NWCS, NECS, SARC, ARCT

Littoral, brackish. NWCS, NECS, SARC

crustacea: copepoda  347

Similar spp.

(Mod. Bradford-Grieve, 1999b)

no projections

hairy AS and caudal rami

Large inner process (1) 1 long and 1 short terminal spine (2)

1

1

♂

short projections

no projections

hairy AS and caudal rami

(Bradford-Grieve, 1999b) Inner spine shorter than terminal spines (1)

♀ 1.1–1.9 mm, ♂ 0.8–1.7 mm ♀

♂

♀

♀ AS shorter than preceding seg

Inner spine longer than terminal spines (1)

Inner spine shorter than terminal spines (1)

♂

♀

♂

(Rose 1933, Ramirez, 1966 )

(Gerber, 2000)

♀ P5

♀ 1.2–2.1 mm, ♂ 1.0–1.9 mm

DISTRIBUTION: Temora stylifera

DISTRIBUTION: Temora longicornis

DISTRIBUTION: Temora turbinata

crustacea: copepoda  349 Order Calanoida Superfamily Diaptomoidea  Family Tortanidae Urosome: usually asymmetrical with projections (8); ♀ 3 segs. ♂ 5 segs. ♀ GS without seminal receptacles. Caudal rami narrow and long, may be fused to AS; strongly asymmetrical in ♀ T. discaudatus, with spine-like projection on right ramus (9)

Genus: Tortanus Giesbrecht, 1898; emend. Ohtsuka and Reid, 1998 Total length: ♀ 1.3–3.1 mm; ♂ 0.8–2.6 mm Cephalosome: A1 reaching end of caudal rami. ♂ R-A1 geniculate (1). Rostrum absent. Large median eye, blue in colour in Tortanus seticaudatus. Mx2 with claw-like setae (2)

Notes: two species in study area. In 1992 Ohtsuka reviewed the genus and divided it into four subgenera Similar taxa: Centropages spp., Temora spp.

Metasome: body slender. 4–5 segs (4th and 5th segs may be fused). End of metasome may be pointed in ♀ (3), usually rounded in ♂ (4)

Distribution and ecology: typically warm neritic and brackish waters. NWCS

Legs: ♀ P5 uniramous 3 segs, last seg elongate in Tortanus discaudatus (5), with short terminal spines in T. setacaudatus (6). ♂  P5 aysmmetric, uniramous: L-leg 4 segs, longer than R-leg in T. setacaudatus; R-leg 3 segs, last 2 segs forming a claw (7)

Tortanus discaudatus

A

(Thompson and Scott, 1897)

B

♀

♀

♀ 1.4–3.1 mm

C

H

G

L

R

♂

♂ 1.4–2.6 mm Neritic, brackish NWCS

1

2

7

D

4

3 8

9

F Tortanus setacaudatus

♀

E I

Williams, 1906 ♀ 1.3–1.4 mm ♂ 0.8–1.0 mm Neritic NWCS

5

J

R

L

7

8

6

Figure 168:  Tortanus discaudatus. A, ♀ lateral view; B, ♀ dorsal view; C, ♂ dorsal view; D, Mx2; G, ♀ P5; H, ♂ P5. Tortanus setacaudatus. E,

♂ urosome dorsal view; F, ♀ dorsal view; I, ♀ P5; J, ♂ P5. A–D, G, and H, Mori (1937); E, Ohtsuka (1992); F, I, and J, Wilson (1932).

350 ta xonom y Order Calanoida Superfamily Eucalanoidea  Family Eucalanidae Genus: Eucalanus Dana, 1852 Eucalanus hyalinus (Claus, 1866) Total length: ♀ 4.4–8.3 mm; ♂ 3.2–5.0 mm Cephalosome: head pointed and triangular in dorsal view in ♀ (1), rounded in ♂ (2). Long paired rostral filaments joined at base (separate in Rhincalanus) (3). A1 extending beyond caudal rami (4); length asymmetrical in ♀; non geniculate in ♂ Metasome: long slim body; 3 segs, cephalosome and 1st metasome seg fused, 4th and 5th segs partially fused. Last seg with small lateral projections, more pronounced in ♀ (5) Legs: ♀ P5 absent; ♂ P5 uniramous, 4 segs, asymmetrical, with an outer seta on 3rd seg (6) and terminating in a spine (7); L-leg longer with fine setules on 3rd and 4th segs (8)

AS fused with caudal rami (10); ♀ AS with small dorsal tooth (11). Caudal rami asymmetrical (12), longest ramus on opposite side to longest A1 in ♀; caudal rami with one very elongate 2nd inner seta (13) Notes: two species reported for the study area: E. hyalinus and Eucalanus spinifer, which can be distinguished mainly by the head shape and the asymmetry of the A1 (Goetze and Bradford-Grieve, 2005). The identifications given by the majority of studies cannot be reliably assigned to either species Similar taxa: Eucalanus spinifer, Pareucalanus spp., Subeucalanus spp. Distribution and ecology: epi-bathypelagic. CNRY, NASE, NWCS, NASW, NADR, NECS, SARC

Urosome: ♀ 4 segs and short. ♂ 5 segs. ♀ GS swollen ventrally, with horizontal thin slit-like seminal receptacles (9) (unlike Subeucalanus). 4

2 2

C A B

♀

4

1

3

♂ 6 7

D

8

R L

F

E

G

5 5 9

13 11

13 10 12 Figure 169:  Eucalanus hyalinus. A, ♀ dorsal view; B, ♀ head ventral with rostrum; C, ♂ dorsal view; D, ♂ P5; E, ♀ urosome lateral view; F, ♂ urosome dorsal view; G, distribution map for E. hyalinus. A–D, Giesbrecht (1893 [‘1892’]); E, F, Bradford-Grieve (1994).

crustacea: copepoda  351 Order Calanoida Superfamily Eucalanoidea  Family Eucalanidae Genus: Pareucalanus Geletin, 1976 Pareucalanus sewelli (Fleminger, 1973) Total length: ♀ 3.7–6.1 mm; ♂ 2.9–4.6 mm Cephalosome: head triangular and sharply pointed in dorsal view (1). Long rostral filaments joined at base (2) (separated in Rhincalanus). A1 extending beyond caudal rami (3); non geniculate in ♂ Metasome: long slim body; 3 segs, cephalosome and 1st metasome seg fused, 4th and 5th segs partially fused. Last seg rounded (4) (small projections in Eucalanus, rounded in Subeucalanus) Legs: ♀ P5 absent. ♂ P5 uniramous, 4 segs, asymmetrical and terminating in a spine; L-leg much longer with fine setules on 3rd and 4th segs (5); R-leg only reaching 2nd seg of L-leg 1

A

Urosome: ♀ 3 segs and short. ♂ 5 segs. ♀ GS swollen ventrally with thin slit-like, angled, seminal receptacles when viewed laterally (6) (horizontal in Eucalanus). AS fused with caudal rami (7); caudal rami slightly asymmetrical, L-ramus longest (8); caudal rami with one very elongate 2nd inner seta (9) Notes: this species has been confused with Pareucalanus attenuatus (not found in the Atlantic according to Fleminger 1973) which differs slightly from P. sewelli by the shape of the head and the pattern of integumental pores (Fleminger 1973) Similar taxa: Pareucalanus attenuatus, Eucalanus spp., Subeucalanus spp., Rhincalanus spp. Distribution and ecology: epipelagic. Circumglobal in warm waters. CNRY, NASE, NWCS, NASW 3

B

G F

♀

♂

4

D

C

2

♂

6

♀

7 8

5

E 9

R L

Figure 170:  Pareucalanus sewelli. A, ♀ dorsal view; B, ♂ dorsal view; C, ♀ head ventral with rostrum; D, ♂ head dorsal view; E, ♂ P5; F, ♀

GS lateral view; G, ♂ urosome dorsal view. A, F, and G, Bradford-Grieve (1994); B, C, Giesbrecht (1893 [‘1892’]); D, Corral Estrada (1970); E, Rose (1933).

352 ta xonom y Order Calanoida Superfamily Eucalanoidea  Family Eucalanidae ID species (7). AS fused with caudal rami (8); ♀ caudal rami may have slight asymmetry (9). Caudal rami with one elongate 2nd inner seta (10)

Genus: Subeucalanus Geletin, 1976 Total length: ♀ 1.8–4.6 mm; ♂ 1.6–3.8 mm Cephalosome: head rounded (1), or pointed and triangular in dorsal view (2). Paired rostral filaments joined at base (3). A1 extending to or beyond caudal rami; non geniculate in ♂

Notes: six species reported in the study area. Fleminger (1973) uses integumental pore patterns to distinguish between species and genera within the Eucalanidae; he also states that pore patterns concur with differences in shape of seminal receptacles. Males are all very similar and poorly described

Metasome: elongate or broad; 3 segs, cephalosome and 1st metasome seg fused, 4th and 5th segs partially fused. Last seg rounded (small projections in Eucalanus, rounded in Pareucalanus). ♂ 2nd and 3rd seg may bear two lateral stout setae (4).

Similar taxa: Pareucalanus spp., Eucalanus spp. and Rhincalanus spp.

Legs: ♀ P5 absent. ♂ P5 asymmetrical, R-leg absent, L-leg uniramous, 4 segs, last seg with fine hairs and terminating in a spine (5)

Distribution and ecology: epi-bathypelagic. Circumglobal, widespread from temperate to warm waters. CNRY, NASE, NECS, NADR, NWCS, GFST, NASW, SARC

Urosome: ♀ 3 segs and short. ♂ 5 segs. ♀ GS swollen ventrally (6). Shape and orientation of seminal receptacles in ♀ can be used to 1

A

B

F

♂

♀

2

♀

G

♂

4

9

C

3

D

E 6 L

7

10

10

8 5 Figure 171:  Subeucalanus crassus. A, ♀ dorsal view; B, ♂ dorsal view; C, rostrum; D, ♂ P5; E, ♀ GS lateral view. Subeucalanus mucronatus. F, ♀ dorsal view. Subeucalanus subcrassus. G, ♂ lateral view. F, G, Mori (1937); A, E, Bradford-Grieve (1994); B–D, Giesbrecht (1893 [‘1892’]).

Table 20:   Subeucalanus species.

Subeucalanus crassus

Subeucalanus monachus

Subeucalanus mucronatus Subeucalanus pileatus

Subeucalanus subtenuis

(Giesbrecht, 1888)

(Giesbrecht, 1888)

(Giesbrecht, 1888)

(Giesbrecht, 1888)

♀ 2.1–4.6 mm, ♂ 2.4–3.5 mm

Total length

1

♀

♀ and ♂ dorsal ♀ head lateral

♀ 1.8–2.8 mm, ♂ 1.9–2.6 mm 1

1

♂

♀

1

♂

♀

♀ ♀ and ♂

(Owre and Foyo, 1967; Bradford-Grieve, 1994) GS wider than long, widest in middle (1) SR broad and enlarged distally (2)

♀

1

♂

♀

2

♀

♂

♀ ♀ head triangular and

♀ head pointed,

pointed (1);

may vary (1);

♂ rounded (2)

♂ rounded (2)

(Owre & Foyo, 1967)

(Giesbrecht, 1893 ['1892'])

GS longer than wide (1) SR thin and enlarged distally (2)

GS as broad as long (1) SR bilobed (2)

(Giesbrecht, 1893 ['1892']) GS longer than wide (1) SR directed upward (2)

1

1

1

2 1

♂

♀

♀ 1.8–3.53 mm, ♂ 2.6–3.8 mm

2

1

♂ rounded (2)

(Owre and Foyo, 1967; Corral Estrada, 1970)

DV

2

pointed (1);

head rounded (1)

GS rounded, as wide as long (1) SR broad and curling upward (2)

♀ 1.8–2.5 mm, ♂ 1.7–2.3 mm

♀ head acutely

♀ and ♂

head rounded (1)

♀ urosome ventral and ♀ GS lateral

♀ 2.8–3.5 mm, ♂ 2.5–3.3 mm

1

♀

(Giesbrecht, 1888)

2

1

2

2

2

(Bradford-Grieve, 1994; Fleminger, 1973)

(Corral Estrada 1970; Fleminger, 1973)

(Giesbrecht, 1893 ['1892'] Fleminger, 1973) (Giesbrecht, 1893 ['1892']; Fleminger, 1973) (Giesbrecht, 1893 ['1892']; Fleminger, 1973)

terminal spine shorter than last seg (1)

terminal spine shorter than last seg (1)

terminal spine shorter than last seg (1)

terminal spine shorter than last seg (1)

NB: longer in S. subcrassus (not illustrated).

L L

♂ P5

1

(Bradford-Grieve, 1994)

Distribution

SR = seminal receptacle

1

1

(Giesbrecht, 1893 ['1892'])

like in S. subcrassus (not illustrated)

L

1

1 L

(Owre and Foyo, 1967)

(Giesbrecht, 1893 ['1892'])

(Giesbrecht, 1893 ['1892'])

S. subcrassus, S. monachus, other ♂ Subeucalanus

S. pileatus, S. subcrassus, other ♂ Subeucalanus

S. crassus, other ♂ Subeucalanus

S. monachus, S. subcrassus, other ♂ Subeucalanus

S. subcrassus, other ♂ Subeucalanus

Epi-mesopelagic. CNRY, NASE, NECS, NADR, NWCS, GFST, NASW, SARC

Meso-bathypelagic. CNRY, NASE, NADR, NWCS, GFST, NASW

Epi-mesopelagic. CNRY, NASE, NADR, NWCS

Coastal, neritic. Epipelagic. CNRY, NASE, NWCS, GFST

Epipelagic. CNRY, NASE, NWCS, NASW, GFST

crustacea: copepoda  353

Similar spp.

L

terminal spine longer than last seg (1)

DISTRIBUTION: Subeucalanus crassus

DISTRIBUTION: Subeucalanus mucronatus

DISTRIBUTION: Subeucalanus monachus

DISTRIBUTION: Subeucalanus pileatus

crustacea: copepoda  355 Order Calanoida Superfamily Eucalanoidea  Family Rhincalanidae Genus: Rhincalanus Dana, 1852

R.  cornutus (6), L-leg biramous, Enp 2 segs and Exp 1 seg; R-leg 3 segs, distal seg terminating in a strongly curved spine (7)

Total length: ♀ 2.8–6.1 mm; ♂ 2.4–4.5 mm Cephalosome: head very pointed (triangular in dorsal view in Rhincalanus nasutus) (1), with long rostral filaments (visible dorsally in Rhincalanus cornutus (2)), separated at base (3) (cf. joined at base in Eucalanus). A1 extending beyond caudal rami in ♀; ♂ non geniculate Metasome: long slim body; 3 segs, cephalosome and 1st metasome seg fused, 4th and 5th segs partially fused. Spines on dorsal and postero-lateral borders of the 3 segs (4) Legs: ♀ P5 small and uniramous, 3 segs terminating in 1 or more setae (5); ♂ P5 small, asymmetrical, coxa and basis elongate in

Rhincalanus nasutus

Urosome: ♀ 3 segs and short, ♂ 5 segs. GS with 2 dorsal spines (8). AS fused with caudal rami (9); L-caudal ramus with a very elongate 2nd seta (10) Notes: two species in study region. There appear to be two forms of R. cornutus in the study area, based on differences in the ♀ P5, a forma typica and a forma atlantica (Schmaus, 1917). Both species are quite transparent Similar taxa: Eucalanus spp. Distribution and ecology: epi-bathypelagic

C 1

A

Giesbrecht, 1888 Epi-bathypelagic

B

3

♂

♀

♀ 2.8–6.1 mm

♂ 2.7–4.5 mm

J

CNRY, NASE, NWCS, NASW,NADR, GFST, NECS, SARC, ARCT

4

2 8

F

♀

Rhincalanus cornutus

K

D outer spine longer than 3rd seg

10

(Dana, 1849) Epi-bathypelagic

5

♀ 2.8–4.2 mm ♂ 2.4–3.7 mm CNRY, NASE, NWCS, NASW, NADR, GFST, NECS

E 4 8

7

9

G

outer spine shorter than 3rd seg

H

I 6 7

5 one seta on 3rd seg

2 setae on 2nd and 3rd segs

Figure 172:  Rhincalanus nasutus. A, ♀ lateral view; B, ♀ head ventral view; C, ♂ dorsal view; D, ♀ P5; E, ♂ P5. Rhincalanus cornutus. F, ♀ dorsal view; G, ♀ P5 forma typica; H, ♀ P5 forma atlantica; I, ♂ P5. J, Distribution map of R. nasutus; K, distribution map of R. cornutus. A, B, Rose (1933); E-G, and I, Giesbrecht (1893 [‘1892’]); C, Bradford-Grieve (1994); D, Owre and Foyo (1967); H, Bowman (1971).

356 ta xonom y Order Cyclopoida  Family Oithonidae Genus: Oithona Baird, 1843 Total length: ♀ 0.3–2.0 mm; ♂ 0.3–1.8 mm Cephalosome: ♀ A1 reaching from mid to end of metasome (1) with numerous long setae (2); geniculate on both sides in ♂ (3). Rostrum directed antero-ventrally (4) or ventrally (5); absent in some species (e.g. O. nana (6)) and in ♂ (7). Mouthparts with strong stiff setae (8) Metasome: oval and slender (9), 4 segs, last seg rounded (10) Legs: number of spines on P1-P4 Exp segs (11) used to ID ♀ species; P5 and P6 reduced into lateral setae; P5 biramous and located on 1st urosome seg. (12). P6 uniramous (13) or biramous (14) located on GS

♀

A

Distribution and ecology: epi-bathypelagic. Neritic and oceanic; fresh-water and marine habitats. Sac spawner. This genus is considered the most abundant copepod taxon with worldwide distribution. ARCT, SARC, NECS, NASE, NWCS, NASW, GFST, NADR, CNRY

3

G

4

3

D

I

10 16

5

♂

1 9

Remarks: 21 species in study area. The ♂ are considered very difficult to ID to species. For a review of the family see Nishida (1985) and Abiahy (2000)

C

B

2

Urosome: ♀ 5 segs, long and slender. ♂ 6 segs. ♀ GS elongated (15), proximally swollen (16); ♀ caudal rami often divergent with elongated setae (17)

15

6

H

7

8

17

E F

P1: 1,1,2

K

J

12 12

2 spines 1 spine 1 spine

13 3 2 1

11

Spermatophores

12

12

14 15

15

Figure 173:  Oithona similis. A, ♀ adult dorsal view; B, ♂ adult dorsal view; C, ♀ head lateral view; D, Mx2; E, P1 with exopodite spine

formula; F, ♀ P5, P6 and GS lateral view. Oithona atlantica. G, ♀ head lateral view; H, ♂ head lateral view. Oithona nana. I, ♀ head lateral view; J, ♀ P5, P6 and GS lateral view; K, distribution map of Oithona spp. A, B, H, Gerber (2000); C–G, I, J, Nishida, 1985.

Table 21:   Oithonidae species.

Oithona atlantica

Oithona plumifera

Farran, 1908

Total length

♀ 0.6–1.4 mm, ♂ 0.6–1.0 mm

♀ 0.7–1.5 mm, ♂ 0.6–1.0 mm

rostrum directed antero-ventrally

♀

♀

♀ dorsal view and ♀ Head lateral view

Oithona similis

Oithona setigera

Baird, 1843

rostrum directed antero-ventrally

(Dana, 1849)

Claus, 1866

♀ 1.1–2.0 mm, ♂ 0.5–1.2 mm

♀ 0.4–1.2 mm, ♂ 0.4–1.8 mm

♀

rostrum directed antero-ventrally

♀ rostrum directed ventrally

♀

♀

♀

(Nishida, 1985)

(Nishida, 1985)

♀

(Nishida, 1985)

(Nishida, 1985) dorsal

♀ GS lateral view P5 and P6

P6 uniramous P6 uniramous (Nishida, 1985)

P1

P1: 1,1,2 P2: 1,0,2 P3: 1,0,1 P4: 0,0,1 P2

P3

Distribution

P4 P1

(Nishida, 1985)

Similar species

(Nishida, 1985)

P2

Broad GS (cf. O. decipiens)

(Nishida, 1985) P1: 1,1,3 P2: 1,0,2 P3: 1,0,1 P4: 0,0,1

P3 P4

(Nishida, 1985)

P1

P2

(Nishida, 1985) P1: 1,1,2 P2: 1,0,1 P3: 1,0,1 P4: 0,0,1

P3

P4

P1

P2

P3

(Nishida, 1985)

(Nishida, 1985)

O. plumifera, O. setigera,

O. atlantica, O. setigera

O. plumifera, O. atlantica

O. decipiens

Epi-mesopelagic

Epi-mesopelagic, oceanic and neritic, also brackish, warm water NECS, NASE, CNRY, NWCS, NASW, GFST, NADR

Epi-mesopelagic, oceanic

Epi-bathypelagic, cosmopolitan

NECS, NASE, CNRY, NWCS, NASW, GFST, NADR

ARCT, SARC, NECS, NASE, CNRY, NWCS, NASW, GFST, NADR

ARCT, SARC, NECS, NASE, NWCS, NASW, GFST, NADR

P4

crustacea: copepoda  357

♀ P1- P4 and Exp spines formula

P1: 1,1,2 P2: 1,0,2 P3: 1,0,1 P4: 0,0,1

P6 uniramous

Tuft of hairs

P6 uniramous

Oithona decipiens

Oithona davisae

Oithona nana

Oithona parvula

Farran, 1913

Ferrari & Orsi, 1984

Giesbrecht, 1892

(Farran, 1908)

♀ 0.5–0.6 mm, ♂ 0.5 mm

♀ 0.3–1.0 mm, ♂ 0.3–0.6 mm

♀ 0.6–1.4 mm, ♂ unknown

Total length

rostrum directed ventrally

rostrum directed ventrally

♀

♀ dorsal view and ♀ Head lateral view

rostrum absent

♀

♀

♀

♀

♀

rostrum absent

♀

Egg sac

(Nishida, 1985)

♀ 0.5 mm ♂ unknown

DV LV

358 ta xonom y

Table 21:  Continued

♀

(Ferrari and Orsi, 1984)

(Rose, 1933; Nishida, 1985)

(Farran, 1908)

dorsal

♀ GS lateral view P5 & P6

Narrow GS (c.f. O. similis)

P6 uniramous

P6 biramous

(Nishida, 1985)

♀ P1-P4 and Exp spines formula

P1

P1: 1,0,2 P2: 1,1,2 P3: 1,0,1 P4: 0,0,0

P2

P3

(Nishida, 1985)

Similar species Distribution

P4

P6 biramous

(Ferrari and Orsi, 1984) P1: 1,1,3 P2: 1,1,3 P3: 1,1,3 P4: 1,1,2 P2

P1

P3

(Nishida, 1985)

P4

(Ferrari and Orsi, 1984)

P1

P1: 1,1,3 P2: 1,1,3 P3: 1,1,3 P4: 1,1,2

P2

P3

P4

P1: 1,1,2 P2: 1,1,2 P3: 1,1,2 P4: 1,1,1 P1

P3

P2

(Nishida, 1985)

(Nishida, 1985)

O. similis

O. nana

O. davisae

O. nana

Epi-mesopelagic, coastal, warm waters NASE

Epipelagic, neritic, estuarine, harbors NASE

Epipelagic, neritic, estuarine, harbors NECS, SARC, NASE, CNRY, NWCS, NASW, GFST?, NADR?

Epi- bathypelagic NECS, NASE, NWCS, GFST

P4

crustacea: copepoda  359 Order Poecilostomatoida  Family Corycaeidae Corycaeidae Dana, 1852 Total length: 0.5–3.1 mm Cephalosome: A1 very short, not reaching metasome, typically 6 segs, ♂ A1 not geniculate. A2 with 4 segs, terminal seg claw-like and stronger in ♂ (1), coxobasis setae with long spinules in Farranula (2). Mxp terminating in a claw, stronger in ♂ (3). Rostrum weakly developed. Paired frontal eyes with large cuticular lenses (4) Metasome: usually robust with 2-3 segs in Corycaeus and 1 seg in Farranula, 3rd and 4th segs often fused dorsally. 2nd and 3rd seg with strong lateral spinal projections (5), inner spines absent in Farranula. Beak-like ventral process in Farranula ♀ (6), absent in ♂ Farranula and other corycaeids Legs: P4 Exp with 3 segs, Enp reduced to a small knob with 1 or 2 setae (7), Enp absent in Farranula. P5 reduced to 2 setae (1 in Farranula) located ventro-laterally. P6 located on GS, in the form of

A

4

B

C

♀

♀

paired operculae: well developed, bearing a long seta and located ventrally in ♂ (8); less developed in ♀ and located dorso-laterally (9), often with a single plumose seta in smaller species Urosome: ♀ 2–3 segs (2 segs in Agetus and Farranula). ♂ 3 segs (2  segs in Farranula). First seg short and narrow, GS large and swollen, caudal rami narrow and elongate (10) Notes: 17 apparent species belonging to six genera are found in the study area, please see key to genera. For a species level key to the family Corycaeidae, please see Boxshall and Halsey (2004). Body may have blue-green or red pigmentation, eggs may also be colourful Similar taxa: Oncaea spp., Vettoria spp. Distribution and ecology: Epi-mesopelagic. Widely distributed in world oceans from coastal to open waters. Carnivorous. SARC, NWCS, NECS, NADR, NASE, NASW, GFST

D

E

2

1

1

10

H I

♂

J

♀

K 9

5

G

♂

♂ 6

♀

F

8

L 7

5 3

Figure 174:  Farranula gracilis. A, ♀ dorsal view; B, ♀ lateral view; C, ♀ A2; D, ♂ dorsal view; E, ♂ lateral view; F, ♂ A2; G, ♀ P4. Corycaeus

speciosus. H, ♀ dorsal view; I, ♀ lateral view; J, ♂ lateral view; K, ♂ Mxp; L, ♀ P4. A–L, (Dahl 1912).

360 ta xonom y Table 22:   Key to North Atlantic adult Corycaeidae genera. Adapted from Boxshall and Halsey (2004).

1

2 3 4

5

Seta on coxobasis of A2 with long spinules; ♀ with posteriorly directed ventral cephalothoracic process; leg 4 uniramous, (lacking endopod)

Farranula

Seta on coxobasis of A2 without spinules; ♀ without ventral cephalothopracic process; leg 4 with reduced endopod

2

♂ caudal rami longer than urosome, ♀ caudal rami almost as long as urosome, but not divergent

Urocorycaeus

♂ caudal rami shorter than urosome, ♀ caudal rami as long as urosome and strongly divergent

3

Endopod of P4 with 2 setae

Ditrichocorycaeus

Endopod of P4 with 1 setae

4

Small forms (up to 1.18mm): ♀ with large prosome, prosome usually two thirds as wide as long; ♂ A2 claw longer than first two segments.

Onychocorycaeus

Large forms: ♀ with slender prosome, prosome less than two thirds as wide as long; ♂A2 claw shorter than second segment.

5

Proximal setae (basal seta and first endopodal seta) on A2 about equal in length. Genital double somite and AS fused in ♀

Agetus

These seta unequal; basal seta about twice as long as first endopodal seta

Corycaeus

DISTRIBUTION: Corycaeus speciosus

DISTRIBUTION: Farranula spp.

DISTRIBUTION: Urocorycaeus spp.

Total length

Corycaeus speciosus

Ditrichocorycaeus anglicus

Agetus typicus

Dana, 1849

(Lubbock, 1857)

(Krøyer, 1849)

♀ 1.4–2.6 mm, ♂ 0.8–2.0 mm

♀ 0.6–1.2 mm, ♂ 0.5–1.0 mm

♀

♀

♂

♀ and ♂

♂

♀ 1.3–1.8 mm, ♂ 1.3–1.6 mm

♀

♂

Onychocorycaeus giesbrechti (Dahl, 1894)

Urocorycaeus lautus

♀ 0.8–1.3 mm, ♂ 0.8–1.1 mm

♀ 2.3–3.1 mm, ♂ 1.5–2.6 mm

♂

♀

(Dana, 1849)

♂

♀

Dorsal

spine reaching past mid GS

(Dahl, 1912)

caudal rami almost same length as urosome; strongly divergent in ♀

egg sacs

GS and AS fused

caudal rami almost same length as urosome; divergent in ♀ (Sars, 1918)

♂ A2

(Dahl, 1912)

inner distal edge finely serrated

strong tooth (Dahl, 1912)

caudal rami almost same length as urosome

(Sars, 1918)

(Dahl, 1912)

(Dahl, 1912)

(Dahl, 1912)

(Dahl, 1912)

spines almost equal in length spines unequal in length

spines unequal in length (Dahl, 1912) Enp present with 1 seta

caudal rami longer than urosome; not divergent

claw longer than segs 1 and 2

(Dahl, 1912)

spines unequal in length

♀ A2

caudal rami shorter than urosome; divergent

spine not reaching past mid GS

(Sars, 1918) Enp present with 2 plumose setae

P4

(Dahl, 1912) Enp present with 1 seta

(Dahl, 1912)

(Dahl, 1912)

Enp present with 1 seta

♀

♀

spines unequal in length

♀

♀

Enp present with 2 setae

(Dahl, 1912)

(Sars, 1918)

(Dahl, 1912)

(Dahl, 1912)

(Dahl, 1912)

Similar spp.

U. lautus, ♂ Agetus

Other Ditrichocorycaeus, ♂ Agetus

Other Agetus, ♂ D. anglicus, ♂ C. speciosus

Other Onychocorycaeus

Urocorycaeus furcifer

Distribution

Epi-mesopelagic. CNRY, NASE, NASW, GFST, NWCS, NADR

Epipelagic. CNRY, NASE, NWCS, NECS

Epi-mesopelagic. CNRY, NASE, NASW, GFST, NWCS

Epipelagic. CNRY, NASE, NASW, GFST, NWCS

Epi-mesopelagic. CNRY, NASE, NASW, GFST, NWCS, NADR

362 ta xonom y

Table 23:   Corycaeidae species.

crustacea: copepoda  363 Order Poecilostomatoida  Family Lubbockiidae Genus: Lubbockia Claus, 1862 Total length: ♀ 1.0–2.8 mm; ♂ 1.4–2.7 mm Cephalosome: A1 short, not reaching metasome, ♀ with up to 7 segs, ♂ with 5 segs and long terminal aesthetascs (1). ♂ A1 not geniculate. Mxp claw-like with robust basis; denticle on syncoxa of ♀ Lubbockia aculeata (2), denticle absent in ♀ Lubbockia squillimana. Rostrum directed ventrally Metasome: long and slender; 4 segs. End of metasome rounded in L. squillimana (3), with lateral projections in L. aculeata (4) Legs: P5 with 1 seg and 2 setae with serrate leaf-like hyaline flanges; located laterally on first urosome seg. ♀ P5 reaching beyond GS in L. squillimana (5), not reaching beyond GS in L. aculeata (6)

Claus, 1863 ♀ 1.0–2.0 mm ♂ 1.5–2.4 mm Mesopelagic CNRY, NASE, NASW, GFST, NWCS, NADR

C

B

Lubbockia A squillimana

Urosome: long and slender. ♀ 5 segs. ♂ 6 segs. ♀ GS elongate and proximally swollen. May have denticles on posterior margin of segs. ♂ AS constricted in L. squillimana (7). Caudal rami narrow and elongate (8) Notes: two species found in the study area. For a species level key to the family Lubbockiidae, please see Boxshall and Halsey (2004) Similar taxa: Oithona spp., Mormonilla phasma Distribution and ecology: epi-mesopelagic. Widely distributed in world oceans from coastal to open waters. CNRY, NASE, NASW, GFST, NWCS, NADR

E

D

♀

♂

♀

F

♂

♀

5

1

3

G

J ♀ eggs

I

♂

7 8

H

Lubbockia aculeata

K

Giesbrecht, 1891 ♀ 1.3–2.8 mm ♂ 1.4–2.7mm Epi-mesopelagic CNRY, NASE, NASW, GFST, NWCS, NADR

4 6 2

Figure 175:  Lubbockia squillimana. A, ♀ dorsal view; B, ♀ lateral view; C, ♀ urosome ventral view; D, ♀ Mxp; E, ♂ dorsal view; F, ♂ urosome ventral view. Lubbockia aculeata. G, ♀ dorsal view. H, ♀ cephalosome ventral view; I, ♂ dorsal view; J, ♂ Mxp. K, distribution map of Lubbockia spp. A, B, and E, Rose (1933); C, D, F, G, I, and J, Boxshall (1977); H, Heron and Damkaer (1978).

364 ta xonom y Order Poecilostomatoida  Family Oncaeidae

Total length: 0.2–1.5 mm

Urosome: ♀ 5 segs. ♂ 6 segs. GS large and elongate, more so in ♂ (9). Caudal rami with dorsal expansion in Epicalymma and Conaea

Cephalosome: A1 short, not reaching metasome; 3rd seg longest (1): ♀ 6 segs, ♂ 4 segs. ♂ A1 not geniculate. A2 with 3 segs, length of terminal seg used to ID species (2). Mxp short and claw-like, ♂ with 3 segs, ♀ with 4 segs; length of basis used in species ID (3). Rostrum small and ventrally directed

Notes: 38 species, from six of the seven genera of oncaeids are found in the study area (Conaea, Monothula, Triconia, Epicalymma, Oncaea and Spinoncaea). The species page illustrates one example from each genus. Please see Boxshall and Halsey (2004) for a full key to the family. Body may have red, purple or orange pigmentation

Metasome: compact, 4 segs. ♀ may have dorsal hump in some Triconia (4). End of metasome rounded or tapering into a point

Similar taxa: Corycaeus spp., Vettoria spp.

Oncaeidae Giesbrecht, 1893 [‘1892’]

Distribution and ecology: epi-bathypelagic. Widely distributed in world oceans from coastal to open waters. Scavengers, also carnivorous. Associated with marine snow and gelatinous plankton. Sac-spawner, typically producing dorsally held paired sacs (10), single egg sac in Monothula and Epicalymma

Legs: P2 to P3 often with distal conical process on Enps (5), also present on P4 of Triconia. P1-P4 with species specific differences in seg and spine length. P5 reduced to 1 seg, with 1 or 2 setae located laterally on 1st urosome seg (6), often fused to seg; length of P5 Exp and number of setae used in species ID. P6 located on GS in the form of paired operculae, located dorsally in ♀ with 1 seta (7) and ventrally in ♂ (8)

L

Triconia borealis

C

♀

A

♂

♀

(Sars, 1918) ♀ 0.6–0.8 mm ♂ 0.4–0.6 mm Epi-mesopelagic NWCS, ARCT, SARC,NECS

6

B 4 9

♀

H

10 1

D Oncaea venusta f. typica Philippi, 1843 ♀ 1.0–1.2 mm ♂ 0.8–0.9 mm Epi-mesopelagic NWCS, NASW, NECS

E

F

G

2

K

I

7

J 8 3 5

Figure 176:  Triconia borealis. A, ♀ dorsal view; B, ♀ lateral view; C, ♂ dorsal view. Oncaea venusta f. typica. D, ♀ dorsal view; E, ♀ urosome lateral view; F, ♂ urosome lateral view; G, ♂ urosome ventral view; H, ♀ A1; I, ♂ A2; J, ♂ Mxp; K, ♀ P3. L, distribution map of Oncaeidae. A, Boxshall and Halsey (2004), modified; B, C, Rose (1933); D–K, Böttger-Schnack (2001).

Table 24:   Oncaeid species.

Total length

Conaea rapax

Monothula subtilis

Triconia minuta

Epicalymma exigua

Oncaea media

Spinoncaea ivlevi

Giesbrecht, 1891

Giesbrecht, 1893 [‘1892’]

Giesbrecht, 1893 [‘1892’]

(Farran, 1908)

Giesbrecht, 1891

(Shmeleva, 1966)

♀ 0.4–0.7 mm, ♂ 0.3–0.4 mm

♀ 0.5–0.8 mm, ♂ 0.4–0.6 mm ♀ 0.4–0.6 mm, ♂ 0.3–0.6 mm ♀ 0.7–1.0 mm, ♂ 0.4–0.9 mm

♀ 0.9–1.2 mm, ♂ 0.8–1.0 mm ♀

♂

♂

♀

♀

♀

♂

♂

unpaired egg-sac

basis elongate length:width >2.7

(Böttger-Schnack, 1999)

(Böttger-Schnack and Huys, 2001)

♀ Mxp

basis robust and elongate length:width